Running head:
The role of HMGRs in synthesizing ginseng saponins
Correspondence:
Deok-Chun Yang
Dept. OMMP, College of Life Science, Kyung Hee University
Suwon 449-701, Korea
E-mail: [email protected]
Tel: +82 (0) 31 201 2100
Fax: +82 (0) 31 202 2687
Title of article:
Functional Analysis of HMGRs in Biosynthesizing Triterpene Saponin in Panax ginseng
Authors’ names:
Yu-Jin Kim, Ok Ran Lee, Ji Yeon Oh, Moon-Gi Jang, Deok-Chun Yang
Addresses:
Department of Oriental Medicinal Materials and Processing, College of Life Science, Kyung Hee
University, Suwon 449-701, Korea
One-sentence Summary
PgHMGR, functional ortholog of AtHMGR1, contributes to the production of triterpene saponin
in Panax ginseng.
Footnotes:
The author responsible for the distribution of materials integral to the findings presented in this
article in accordance with the policy described in the Instructions for Authors
(www.plantphysiol.org) is: Deok-Chun Yang ([email protected]).
ABSTRACT
Ginsenosides are glycosylated triterpenes, which are considered to be the most pharmaceutically
active ingredients in ginseng (Panax ginseng), known as an adaptogenic herb. However, the
mechanism underlying the biosynthesis of triterpene saponin in ginseng remains unclear. In this
study, we characterized the role of 3-hydroxy-3-methylglutaryl coenzyme A reductase (HMGR)
in ginseng saponin synthesis. Firstly we showed that P. ginseng has two HMGRs, PgHMGR1
and PgHMGR2 with high sequence identity by analyzing the full-length cDNAs. The expression
of PgHMGR1 and PgHMGR2 is strong in the root vesiculture, and increase in the older ginseng,
supporting its role in accumulation of saponin in roots. In addition, treatment with mevinolin, a
competitive inhibitor of HMGR, on ginseng adventitous roots caused significant reduction of
total ginsenoside. Moreover, we showed that continuous dark exposure for 2 days to 3-year-old
ginseng increased total ginsenoside contents, and PgHMGR1 showed dark-induced expression,
suggesting that PgHMGR1 may be associated with dark-dependent promotion of secondary
metabolite biosynthesis in the ginseng plant. Furthermore, overexpression of PgHMGR1
enhanced the production of sterol and triterpene in Arabidopsis and ginseng. The conserved
biochemical role of PgHMGR1 was shown by the genetic complementation of hmgr1-1 in
Arabidopsis. Taken together, our data suggest that ginseng HMGRs play a key role in the
triterpene pathway for ginseng.
INTRODUCTION
Ginseng (Panax ginseng Meyer), a member of the Araliaceae family, is a perennial
herbaceous plant and has been cultivated for its highly valued roots used for medicinal purposes.
The root of ginseng has been used as a drug by the people in the Eastern Asia for thousands of
years because it contains polyacetylenes, polysaccharides, peptidoglycans, phenolic compounds,
and saponin (Park, 1996; Kitagawa et al., 1987; Radad et al., 2006). The triterpene saponin in
ginseng, referred to as ginsenoside, has especially been noted as the active compound
contributing to the efficacy of ginseng. Triterpene saponins are secondary metabolites of
isoprenoid compounds and are common in a large number of plant species, particularly in dicot
plants. They exhibit great structural diversity and notable biological activity (Hostettmann and
Marston, 1995; Augustin et al., 2011). Ginsenosides are found exclusively in the plant genus
Panax, with a content of about 6 to 10%. More than 150 naturally occurring ginsenosides have
been isolated from Panax species (Shi et al., 2010). To date, more than 40 ginsenosides have
been isolated and identified from white and red ginseng, showing different biological activities
based on their structural differences (Gillis, 1997; Fuzzati, 2004; Lu et al., 2009; Xie et al., 2005;
Tung et al., 2009).
The main ginsenosides are glycosides that contain an aglycone with a dammarane skeleton
(Fig. 1A). They include protopanaxadiol-type (PPD) saponins (where sugar moieties are attached
to the β-OH at C-3 and/or C-20) such as ginsenosides Rb1, Rb2, Rc, and Rd and
protopanaxatriol-type (PPT) saponins (where sugar moieties are attached to the α-OH at C-6
and/or the β-OH at C-20) such as ginsenosides Re, Rg1, Rg2, and Rf, together constituting more
than 80% of the total ginsenosides (Kim, 1987). The oleanane group has a pentacyclic structure,
and only one ginsenoside, Ro, was identified, which is found in minor amounts in P. ginseng.
These ginsenoside compounds contribute to the various pharmacological effects of ginseng, such
as anti-aging (Cheng et al., 2005), anti-diabetes (Attele et al., 2002), anti-inflammatory (Wu et al.,
1992), and anti-cancer activities, such as the inhibition of tumor-induced angiogenesis (Liu et al.,
2000; Nakajima et al., 1998; Yue et al., 2007), anti-tumor activity, and the prevention of tumor
invasion and metastasis (Mochizuki et al., 1995; Sato et al., 1994).
Ginsenosides are synthesized from the 30-carbon intermediate 2,3-oxidosqualene, a
common precursor of sterols and which undergoes further cyclization, hydroxylation, and
glycosylation (Fig. 1B). Triterpene saponins, including ginsenosides, are derived from a
universal precursor, isopentenyl diphosphate (IPP), which can be synthesized via the cytoplasmic
mevalonate (MVA) pathway, conserved in some prokaryotes and in all eukaryotes. The MVA
pathway is catalyzed by the key regulatory enzyme 3-hydroxy-3-methylglutaryl coenzyme A
reductase (HMGR, EC1.1.1.34) (Stermer et al., 1994). HMGR is known as a rate-limiting
enzyme in the isoprenoid pathway in plants (Bach and Lichtenthaler, 1982) and in mammals
(Goldstein and Brown, 1990). Since HMGR is a rate-controlling enzyme for cholesterol
biosynthesis, its role has been studied in human health, such as how the inhibition of HMGR is a
major intervention strategy for the treatment of cardiovascular disease and blood pressure
reduction (Liao and Laufs, 2005). In contrast to the single HMGR gene in animals, plant HMGR
is encoded by a multigene family, where the different isoforms exhibit different tissue and
developmental patterns of expression. Two HMG1 and HMG2 proteins of Arabidopsis have the
same structural organization and intracellular localization, but their expression profiles are
different. The broad expression of HMG1 suggests that it may encode a housekeeping form of
HMGR; correspondingly, the loss of HMG1 caused senescence and sterility as well as a dwarf
phenotype, which are likely related to the reduced sterol content (Suzuki et al., 2004). In contrast,
the hmg2 mutant did not display a distinct phenotype. hmg1 hmg2 double mutants are not viable
because of the requirement of two genes for gametophyte development (Suzuki et al., 2009).
Both HMG1 and HMG2 genes were also shown to play a major role in the biosynthesis of
triterpene metabolites (Ohyama et al., 2007).
So far, even though much advance revealing the pharmacological effects of ginsenosides
and the mass production of ginsenosides by genetic engineering and biotechnology, little is
known about the regulatory function of ginsenoside biosynthesis at the molecular level in
ginseng. Because of the lack of genome sequence information, some attempts have been tried to
clone complete cDNA clones encoding several enzymes from ginseng involved in the post-
squalene step (indicated according to the accession number in Fig. 1B) (Kushiro et al., 1998; Lee
et al., 2004; Kim et al., 2010; Han et al., 2006; 2010; 2011; 2012). In this present study, the full-
length cDNA sequences, together with promoter sequences of two HMGR genes, were isolated
and characterized from P. ginseng for the first time. The phylogenetic analysis, the expression
profiles of two HMGR genes, and functional analysis by heterologous overexpression of HMGR1
in Arabidopsis and ginseng were also investigated. Our finding suggests that HMGR is essential
for the formation of triterpene ginsenoside.
RESULTS
Isolation and Sequence Analysis of PgHMGR1 and PgHMGR2
In order to identify the first committed enzyme of ginseng in the mevalonic acid (MVA)
pathway for the biosynthesis of isoprenoids, two expressed sequence tag (EST) clones coding 3-
hydroxy-3-methylglutaryl coenzyme A reductases, (HMGR from Panax ginseng, PgHMGR, EC
1.1.1.34) were selected from previously constructed EST libraries of 14-year-old ginseng and
hairy root, respectively (Kim et al., 2006b). By RACE PCR, full-length complementary DNA
(cDNA) sequences of PgHMGR1 and PgHMGR2 were obtained. PgHMGR1 has a length of
1,722 bp encoding 573 amino acids, whereas PgHMGR2 has 1,785 bp encoding 594 amino acids
(Supplementary Fig. S1A, B). Both genes contain four exons and three introns (Fig. 2A), a
typical feature of the HMGR genes from other plant species. PgHMGR2 is 63 bp longer than
PgHMGR1 in the first exon region, although the other three exons are the same length. Moreover,
a full genomic DNA sequence of each PgHMGR with the promoter sequence was obtained by
genomic DNA walking. The promoter region of PgHMGR1 contains G-box (CACGTG),
although PgHMGR2 does not (Supplementary Fig. S1 and S2).
HMGR proteins are comprised of three domains (Campos and Boronat, 1995); the membrane-
anchor domain, a flexible linker domain, and the catalytic domain. PgHMGR1 and PgHMGR2
also contain conserved substrate and cofactor binding sites that are proposed to be characteristic
of the HMGR protein (Fig. 2C and Supplementary Fig. S7): two putative HMG-CoA binding
sites (EMPI/VGYVQ, TTEGCLVA) and two NADP(H)-binding sites (DAMGMNM,
GTVGGGT) in the catalytic domain. Transmembrane prediction and the hydrophobicity profile
suggest that both the PgHMGR1 and PgHMGR2 proteins have two membrane-spanning domains,
indicated as H1 and H2 in Fig. 2C. Both proteins contained a motif rich in arginines (RRR)
(Supplementary Fig. S1 and S2), which is predicted to be specific for endoplasmic reticulum (ER)
retention (Schutze et al., 1994).
The three-dimensional (3D) structures of PgHMGR1 and PgHMGR2 were predicted by the
Swiss-Model using sequence homology-based structural modeling. The molecular modeling
results (Fig. 2B) showed an intricate spatial architecture, which is very similar to the human
HMGR structure (Istvan et al., 2000). The catalytic domain consists of three domains: the small
and helical N-terminal N-domain; the large, central L-domain harboring two HMG-CoA binding
sites; and the small helical S-domain which is inserted into the L-domain. The PgHMGR1 and
PgHMGR2 catalytic domains possess three catalytic active residues EGC, DKK, and GQD in the
three conserved motif sequences predicted by Multiple EM for Motif Elicitation (MEME) (Fig.
2B, 2C and Supplemental Fig. S8). These are similar to Cantharanthus roseus HMGR, which
Abdin et al. (2012) recently showed as responsible for the active site that can bind to HMG-COA
and NADPH2.
PgHMGR1 and PgHMGR2 Are Mainly Expressed in Roots
Ginseng is a perennial medicinal plant; an age-dependent increase of ginsenosides was
reported in ginseng roots (Shi et al., 2007). To investigate the expression patterns of PgHMGR1
and PgHMGR2, quantitative reverse transcription (qRT)-PCR was performed on RNA extracted
from ginseng seed, an early seedling and leaf, petiole and root of a 2-week-old seedling, as well
as on RNA extracted from flower, leaf, stem, main root, and lateral root of 3-year-old and 6-year-
old plants (Fig. 3A). A distinct anatomical feature of ginseng is the long petiole structure having
a relatively shortened stem in a 2-week-old seedling (Fig. 3A). The number of leaf petioles
corresponds to the number of years cultivated, and 2-year-old ginseng possesses five leaves.
PgHMGR1 was most highly expressed in the petiole in the 2-week-old seedling (Fig. 3B). In the
3-year-old ginseng, when most of the known ginseng organs are formed, and the 6-year-old
ginseng, transcripts of PgHMGR1 were highly observed in the main roots and lateral roots (Fig.
3C, D). Interestingly, its expression was similarly expressed to high level in 3-year and 6-year
old ginseng (Fig. 3C, D). PgHMGR2 was also comparatively highly expressed in root, but the
level of expression of PgHMGR2 in the root was at least 4-fold lower than that of PgHMGR1
(Fig. 3E, F, G). These results suggest that PgHMGR1 and PgHMGR2 are closely linked with the
age-dependent production of ginsenosides in roots.
When a promoter sequence of PgHMGR1 (-1317~1 bp)-fused GUS was transformed into a
ginseng adventitious root, it was expressed highly in the whole root, with more restriction in the
vasculature and root tip (Fig. 3J). When PgHMGR1::GUS and PgHMGR2::GUS were expressed
in heterologous Arabidopsis, expressions were also observed in the whole root vasculature (Fig.
3Hb, Ib) and in an 8-day-old main root tip (Fig. 3Hc, Ib), similar to the ginseng plant, which
suggests that Arabidopsis is an ideal plant for functional characterization of PgHMGR. Precise
expression of PgHMGR1::GUS and PgHMGR2::GUS in all tissue of a germinating embryo (Fig.
3Ha, Ia), cotyledon, true leaf, and root were also observed in Arabidopsis (Fig. 3Hd, Id).
PgHMGR1 and PgHMGR2 GUS expression was high in the filament, flower sepal and pistil
style (Fig. 3He, Ie) and in the junction between the silique and pedicel of a 50-day-old plant (Fig.
3Hf, If).
HMGR Activity Positively Correlates with Ginsenoside Production
To understand whether HMGR activity is involved in ginsenoside biosynthesis, inhibition of
HMGR activity by mevinolin was conducted. Mevinolin (6α-methylcompactin), also referred to
as lovastatin, competitively inhibits the binding of substrate HMG-CoA to the active site of the
enzyme HMGR and consequently blocks the synthesis of cytosolic IPP (Bach and Lichtenthaler,
1982) and phytosterol biosynthesis (Bach and Lichtenthaler, 1987; Bach et al., 1990). Mevinolin
treatment for 1 day in 4-week-old adventitious roots significantly decreased the total ginsenoside
content to about 34% lower than the control (Fig. 4A), with a decrease in the expression of
PgHMGR1 and PgHMGR2 (Fig. 4B). On the other hand, methyl jasmonate (MJ), known as an
elicitor for triterpene biosynthesis, increased the ginsenoside contents (Fig. 4D) and an increase
in PgHMGR1 expression were observed in ginseng adventitious root and pHMGR1::GUS (Fig.
4E and 4F), confirming the essential role of HMGRs in ginsenoside biosynthesis (Supplementary
Fig. S3).
Expression of PgHMGR1 Is Light-Inhibited and Possibly Associated with Ginseng
Shadowing Growth
The evidence for a dark-dependent hypocotyl expression pattern of PgHMGR1::GUS (Fig. 5G)
led us to hypothesize that ginsenoside synthesis may be inhibited by light. As shown in Fig. 5,
placing the ginseng plants under completely dark conditions for 48 h caused a dramatic increase
in the ginsenoside contents compared to the control grown under a 16 h light and 8 h dark
condition. Dark conditions for 72 h also resulted in an increase in ginsenosides. The total
ginsenoside contents were decreased in both leaves and roots, in spite of different patterns
demonstrated by the individual ginsenosides (Supplementary Fig. S4). Transcriptions of
PgHMGR1 and PgHMGR2 were significantly decreased in the ginseng leaves and roots exposed
to darkness for three days (Fig. 5G, E). However, the HMGR activity was increased in both
leaves and roots (Fig. 5C).
PgHMGR1::GUS promoter activity was differently modulated by light exposure in the leaf
and hypocotyl. In a young seedling that had not yet undergone true leaf emergence, dark-induced
hypocotyl expression of PgHMGR1::GUS was decreased by light exposure within 2 h (Fig. 5G).
However, GUS expression in a 7-day-old seedling was decreased by dark treatment in the true
leaves (Fig. 5H).
PgHMGR1 Localizes to the ER and Peroxisome in Arabidopsis
Fractionation analysis detected plant HMGR in three subcellular sites – the plastids,
mitochondria, and ER (Bach et al., 1999). Despite enzyme activity detection in several organelles,
immunofluorescence confocal microscopy and immunogold electron microscopy analyses
showed that AtHMGR1 is localized in the ER and unidentified spherical vesicles and is not co-
localized with peroxisomal catalase (Leivar et al., 2005). Cyan fluorescent protein (CFP) or
monomeric red fluorescent protein (mRFP) was tagged to the C-terminal ends of PgHMGR1 and
PgHMGR2 to identify subcellular localization patterns by confocal laser scanning microscopy.
Both PgHMGR1 and PgHMGR2 were localized in ER-like vesicles (Fig. 6A, B, C). But the
spherical vesicles and other unidentified vesicles of PgHMGR1 were co-merged with
peroxisome and plastids when PgHMGR1-mRFP was crossed with endosomal markers (Fig. 6E).
Only the catalytic domain of PgHMGR1 (H1CD) was localized to the cytosol and nucleus (Fig.
6D), as in the case of Arabidopsis HMGR1 (Leivar et al., 2005).
Overexpression of PgHMGR1 Enhances Production of Triterpenes in Arabidopsis and
Ginseng
To investigate whether increased HMGR activity contributed to the metabolite profiles, sterol
contents in Arabidopsis HMGR1ox lines were analyzed using gas chromatography-mass
spectrometry (GC-MS) analysis. Higher plants synthesize a mixture of phytosterols, including
campesterol, stigmasterol, and β-sitosterol (Hartmann and Benveniste, 1987); thus, three major
phytosterols were analyzed. Pentacyclic β-amyrin, one of the most common triterpenes in plants,
and α-amyrin are present in Arabidopsis; therefore, these two triterpenes were also analyzed
together with squalene, a common precursor of triterpene and sterol. Total sterols were extracted
from the rosette leaves and inflorescence of 5-week-old plants. Compared to wild type,
HMGR1ox No. 15-8 rosette leaves were 2-times higher in β-sitosterol, 1.8-times higher in
campesterol and cycloartenol, 2.5-times higher in β-amyrin, and 2-times higher in α-amyrin (Fig.
7A). The inflorescence was 1.6-times higher in campesterol, β-sitosterol, and β-amyrin, 2.6-
times higher in cycloarternol, and 3.7-times higher in α-amyrin (Fig. 7B) than the WT. The
squalene and stigmasterol contents were not significantly changed compared with the control.
Volatile (-)-E-b-caryophyllene, which is a sesquiterpene, was also increased in the HMGR1ox
line analyzed by SPME GC-MS (data not shown).
To test whether PgHMGR1 can constitute a functional triterpene pathway in planta, PgHMGR1
was constitutively expressed in Arabidopsis as well as ginseng under the control of the 35S
promoter.
To understand the contribution of HMGR to ginsenoside biosynthesis, adventitious roots were
induced from transgenic ginseng callus after transformation of the PgHMGR1 gene to ginseng.
The ginsenoside contents of different transgenic lines were 1.5-2 times higher than the control,
with no alteration of the ratio of individual major ginsenosides (Fig. 7C).
Conserved Role of PgHMGR1
To understand the evolutionary role of PgHMGRs, we performed amino-acid sequence
alignments of HMGRs from various plants searched by GenBank BlastX from NCBI database
(Supplementary Fig. S5). Alignment analysis revealed that two PgHMGRs had high similarity
with other plant HMGRs. PgHMGR1 shares 93.7% identity with HMGR from Eleutherococcus
senticosus (EsHMGR, AFM77981), which belongs to the same Araliaceae family. PgHMGR2
shared 88.4% identity with HMGR from Panax quinquefolius (PqHMGR, ACV65036). Both
PgHMGR1 and PgHMGR2 showed 74.8% and 67.8% amino acid sequence identity to
AtHMGR1 (At1g76490) and AtHMGR2 (At2g17370) (Enjuto et al., 1994), respectively, and
shared a highly conserved domain with other plant homologs containing a membrane domain
(Campos et al., 1995) and a catalytic domain harboring two HMG-CoA binding motifs and two
NADP(H)-binding motifs, which is comparable to other HMGRs reported previously (Shen et al.,
2006), except for the highly divergent N-terminal region and the linker domain (Supplementary
Fig. S5).
PgHMGR was clustered into plant HMGRs and was structurally distinct from fungus and
animal enzymes (Fig. 8A). PgHMGR1 has less than 30% identity with HMGR from animals,
such as Homo sapiens (NP_000850), Drosophila melanogaster (P14773), and Mus musculus
(NP_032281) and with the fungi, Pichia jadinii (O74164). Three conserved motif sequences
were identified in plant as well as animal HMGRs (Fig. 8B, Supplementary Fig. S5).
The conserved role of ginseng HMGRs was revealed by the genetic complementation of
Arabidopsis null mutant hmgr1-1 (Fig. 8C and 8D). Overexpression of the full length of the
cDNA sequence of PgHMGR1 and the catalytic domain of PgHMGR1 under the 35S promoter
complemented the phenotypic defect of hmgr1-1, whereas PgHMGR2 did not, indicating that
PgHMGR1 has a conserved role with AtHMGR1.
DISCUSSION
Ginseng root, one of the most famous well-known medicinal plants, particularly in traditional
oriental medicine, contains more than 4% ginsenoside by dry weight (Shibata, 2001), which
contribute to its many pharmaceutical activities. Ginsenoside, dammarane-type tetracyclic
triterpene, is unique to ginseng (Fig. 1), although other oleanane-type triterpenes are observed in
other plants. Manipulation of ginsenoside biosynthesis is of particular importance to the increase
of ginsenoside in ginseng or other species. Currently, much of the research about ginsenoside has
been conducted for pharmaceutical efficacy, and there is less information about ginsenoside
biosynthesis. In common, all triterpenes and sterols are biosynthesized via the MVA pathway.
The enzyme HMGR catalyzes the formation of MVA, a rate-limiting step in isoprenoid
metabolism. Here, we report the characterization of the HMGR gene, an essential gene for
ginsenoside biosynthesis in ginseng. Although one HMGR sequence of Panax quinquefolius
(ACV65036) was reported in NCBI, having 76% identity to the amino acid sequence of
PgHMGR1, a functional study of HMGR has not yet been studied in ginseng.
The roots of P. ginseng plants are usually used as important components of traditional oriental
medicine, as the ginsenoside content increases with plant age (Shi et al., 2007). Specifically, the
transcript of PgHMGR1 was enhanced in the roots of 3-year-old and 6-year-old ginseng,
corresponding to the identification of EST homologous from the ginseng root EST library. This
suggests that PgHMGR1 has a major role in ginseng roots with regard to providing sterol or
triterpene. Sterol biosynthesis-related genes were supposed to be expressed constitutively in all
plant tissues in order to synthesize sterol, which is necessary for plant development (Shen et al.,
2006, He et al., 2003). A detailed functional understanding of sterol and triterpene in plant
development needs to be clarified further. Corresponding with the expression of PgHMGR1 and
PgHMGR2 in the vasculature tissue of the roots (Fig. 3Hb, 3Ib), ginsenoside biosynthesis-related
genes PgSSs and PgSEs were also characterized as being expressed in vascular bundle tissues
including phloem cells, parenchymal cells near xylem, and resin ducts in the petioles of ginseng
(Han et al., 2010; Kim et al., 2011). It can be postulated that the phloem and resin ducts might
serve as the metabolically active sites for sterol and saponin biosynthesis and play a role in
conducting squalene oil. In fact, ginsenoside is preferably accumulated more in the phloem than
in the xylem (Fukuda et al., 2006) and is observed in parenchymal and phloem cells (Yokota et
al., 2011).
PgHMGR1 is expressed in a range of ginseng tissues in addition to the roots. Our qRT-PCR
results showed that PgHMGR1 was expressed not only in ginseng roots, but also in the leaves
and flowers, followed by weaker expression in the stem, and PgHMGR2 was expressed at a very
low level (Fig. 3). The results indicate that the expression of PgHMGR1 is in agreement with the
ginsenoside biosynthesis in different tissues, where there is lower ginsenoside in the stems
compared to the roots, leaves, and fruit (Shi et al., 2007). PgHMGR1::GUS and
PgHMGR2::GUS were detected in ginseng seedlings at early post-germination stage, and GUS
expression was also triggered during the germination stage in Arabidopsis. This suggests that
triterpene is required in the seedling stage during germination, corresponding to active β-amyrin
production in pea seedlings (Baisted, 1971). Interestingly, PgHMGR1 was highly expressed in
petiole tissue (Fig. 3B), which is similar to the high expression of other ginsenoside biosynthesis-
related SS (Kim et al., 2011) and SE genes (Han et al., 2010). A long petiole is a distinct anatomic
structure that can be found in the ginseng plant. Triterpene production by petiole-specific gene
expression might provide metabolites required for special petiole development in ginseng,
although there has been no study on the relationship between petioles and triterpene. The
promoter activity of PgHMGR1 and PgHMGR2 was high in the early flowering stage, with no
detection in matured siliques or seed. High HMGR expression in the early developmental stage
has also been shown in other plants: potato HMGR1 is expressed in early flower development
(Korth et al., 1997), and tomato HMGR1 is expressed in young tomato fruit (Jelesko et al., 1999).
The flower phenotype in tobacco is altered by overexpression of AtHMGR1 (Hey et al., 2006),
which suggests that HMGR-derived phytosterols and metabolites play roles in flower
development. Interestingly, overexpression of PgHMGR1 resulted in an increase of volatile oil,
caryophyllene (a sesquiterpene), triterpene, and sterol in the inflorescence of Arabidopsis (Fig.
7B), which supports the idea that higher activity of PgHMGR1 provides more isoprenoid
compounds required for flower development. It is noticeable that the constitutive inhibition of
HMGR by mevinolin ultimately blocked fruit development (Narita and Gruissem, 1989).
All of these results imply that PgHMGR1 plays a key role in ginsenoside biosynthesis. Hence,
it is interesting to determine whether or not PgHMGR1 is positively correlated with the
ginsenoside content in the P. ginseng plant. Unfortunately, ginseng is not practically amenable to
loss- or gain-of-gene-function studies. To gain insight into the effects of the loss of HMGR
activity on ginsenoside biosynthesis, an inhibitor was used to deplete the metabolic flux through
the MVA pathway. Treatment with mevinolin, a highly potent competitive inhibitor of HMGR,
reduced the total ginsenoside content in adventitious roots compared to the control after 24 h due
to reduced gene expression, in contrast to the effect of the elicitor MJ, which increased the
ginsenoside level expression of PgHMGR1 (Fig. 4). This is the first report that the activity of
HMGR is possibly playing an important role in ginsenoside biosynthesis. It was confirmed again
with increased ginsenoside levels in transgenic ginseng adventitious roots overexpressing
PgHMGR1 (Fig. 7C).
Light is known to be one factor that has an effect on plant HMGR. Light-regulated
suppression of HMGR1 promoter activity in Arabidopsis immature leaves and seedlings
(Learned and Connolly, 1997) as well as increased HMGR activity in etiolated pea seedlings
(Brooker and Russsell, 1979) are in direct contrast to the suppression of HMGR activity in dark-
treated potato plants (Korth et al., 2000). In the case of ginseng HMGR, in contrast to the
enhanced gene expression in the etiolated hypocotyl region (Fig. 5G), dark treatment caused a
significant decrease in GUS activity in true leaves of PgHMGR1::GUS in Arabidopsis (Fig. 5H)
and a decrease in the expression level of PgHMGR1 and PgHMGR2 in ginseng (Fig. 5D and E),
consistent with reduced HMGR2 in potato leaves due to dark treatment (Korth et al., 2000).
Differential regulation of PgHMGR1::GUS expression in leaf and hypocotyl tissue can be
explained by the idea that light-mediated control of HMGR1 is an organ-autonomous response
(Learned and Connolly, 1997). Downregulation of HMGR in Arabidopsis due to light was shown
to occur via photoreceptor phytochrome B (PHYB) and transcription factor HY5 (Rodríguez-
Concepción et al., 2004). Analysis of the promoter sequence implied that PgHMGR1 was
regulated by binding PIF3 to the G-box in the promoter of the PgHMGR1 gene, although further
experimental analysis is needed for confirmation. PIFs, basic helix-loop-helix (bHLH)
transcription factors, play a central role in the light-mediated response (Castrillon et al., 2007).
Shin et al. (2007) showed that PIF3 and HY5 regulate anthocyanin biosynthesis by binding to the
promoters of anthocyanin biosynthesis-related genes. PIF3 is also known as a negative
component in the PHYB-mediated inhibition of hypocotyl elongation (Kim et al., 2003). The
PgHMGR1 gene contains a PIF3 binding motif corresponding with dark-dependent hypocotyl
expression, suggesting that ginsenoside biosynthesis might be tightly regulated by light, possibly
through PIF3-mediated suppression of PgHMGR1. When the whole ginseng plant was in dark
conditions for up to 48 h, the ginsenoside content was increased. This could be affected by dark
stress via the light-to-dark switch (Souret et al., 2003). HMGR activity in ginseng was also
increased under dark treatment for 2-3 days, possibly at the post-transcriptional level (Korth et
al., 2000), and it was positively correlated with total ginsenoside content. Our research may
provide new insight into the relationship between light and ginsenosides.
Although HMGR activity was reportedly detected in the ER, plastid, and mitochondria in
plants (Bach et al., 1999), there has been no observation of subcellular localization of plant
HMGR in multiple organelles, except for the localization of AtHMGR1 in the ER and
unidentified spherical vesicles (Leivar et al., 2005). Both PgHMGR1 and PgHMGR2 contain an
RRR motif at their N terminus, which may act as an ER retention signal (Schutze et al., 1994).
When PgHMGR1 and PgHMGR2 were expressed individually in Arabidopsis as C-terminal CFP
fusions, both proteins were also targeted to the ER and spherical vesicles. We further confirmed
that the spherical vesicles and other unidentified vesicles of PgHMGR1 were co-merged with the
peroxisome and plastids (Fig. 6E). Discovery of the subcellular localization of ginseng HMGR
suggests that ginseng HMGR could be targeted to different subcellular organelles through post-
translational modifications. In addition, alternative mRNA processing could contribute to the
different localization. For example, the IPP isomerase long isoform is targeted to the chloroplasts
and mitochondria, and its short isoform is localized to the peroxisome (Sapir-Mir et al., 2008).
The catalytic domain of ginseng HMGR (H1CD) was localized in the nucleus and cytosol,
confirming that the N-terminal domain of PgHMGR is necessary for binding to the membrane,
which confirms the previous report (Leivar et al., 2005). In mammalian research, peroxisomal
localization of HMGR in rat (Keller et al., 1985) was reported in addition to ER localization
(Goldfarb, 1972); thus, dual localization of HMGR in the peroxisome and ER is now commonly
accepted (Kovacs et al., 2007). Localization of HMGR in the peroxisome could have related
roles in lipid metabolism and cholesterol degradation in animals (Keller et al., 1985). Recently,
in plants, acetoacetyl-CoA thiolase (Reumann et al., 2007) and IPP isomerase (Sapir-Mir et al.,
2008) were reported to be localized in peroxisomes (Reumann et al., 2007), and other MVA
pathway genes except HMGR, HMGS, MVK, PMK, DMD, and FPS were predicted to be
partially localized in the peroxisome (Sapir-Mir et al., 2008). The presence of PgHMGR1 in
multiple subcellular localizations supports that MVA biosynthesis can occur in different cell
compartments, in parallel with the localization of ginsenoside Rb1 in the cytosol, plastid, and
peroxisome in the ginseng plant (Yokota et al., 2011). Peroxisome-localized HMGR in animals is
functionally and structurally different from ER-localized HMGR (Aboushadi et al., 2000).
Different compartments could contain separate biosynthetic pathway with separate enzymes for
producing organelle-specific isoprenoids, such as sterol in the ER, pigments in plastids,
ubiquinone in mitochondria, or rubber particles in specialized vacuoles (McGarvey and Croteau,
1995).
Furthermore, to test whether PgHMGR1 can contribute to the triterpene pathway in other
plants, PgHMGR1 was expressed in Arabidopsis. Although HMGR is known as the key
regulatory enzyme in sterol biosynthesis in plants (Babiychuk et al., 2008), overexpression of
HMGR in Arabidopsis did not show any altered morphology or isoprenoid content (Re et al.,
1995). However, overexpression of the HMGR gene in tobacco plants increased the amount of
sterol and intermediates of the sterol pathway in the form of fatty acyl esters in accordance with
the increased activity of HMGR (Chappell et al., 1995; Schaller et al., 1995). In the case of
overexpression of HMGR in tomato, phytosterols were elevated without alteration of the HMGR
activity (Enfissi et al., 2005). The constitutive overexpression of PgHMGR1 driven by the 35S
promoter resulted in the accumulation of not only sterol, but also sesquiterpene, volatile oil, and
triterpenes in Arabidopsis (Fig. 5A, B), with modest increases in the levels of HMGR activity.
This result is comparable with the previous study where overexpression of AtHMGR1 resulted in
co-activation of the HMGR1 gene without altering the accumulation of the isoprenoids end-
products (Re et al., 1995).
Although PgHMGR1 was expressed in different levels among tested organs, the PgHMGR1
gene appears to be expressed constitutively throughout ginseng development, which was similar
to characterized homologs from Salvia miltiorrhiza (Liao et al., 2009) and Eucommia ulmoides
(Jiang et al., 2006). In Arabidopsis, AtHMGR1 can be detected in all tissues (Enjuto et al., 1994),
but AtHMGR2 is expressed exclusively in meristematic and floral tissues (Enjuto et al., 1994).
As reported for the AtHMGR1 gene, PgHMGR1 may play a housekeeping role in ginseng,
whereas PgHMGR2 may play a more specialized role. Indeed, heterologous expression of two
full-length ginseng HMGRs in an Arabidopsis hmgr1-1 mutant background exhibited that only
PgHMGR1 could complement the dwarf and sterile phenotype of hmgr1-1 (in this study; Suzuki
et al., 2004), which suggests that PgHMGR1 is a functional ortholog of AtHMGR1. Phylogenetic
analysis also showed that both PgHMGR1 and PgHMGR2 have three conserved motifs with
conserved amino acid sequences in the homologues (Fig. 9A, B and Supplemental Fig. S6).
Among all of the identified plant HMGR genes, corresponding with the topology of HMGR as a
membrane-binding protein, the N-terminal region exposing to the cytosol and linker domain
differs greatly both in length and amino acid sequence, while the membrane-spanning domain
that serves as a membrane anchor (Campos & Boronat, 1995) and C-terminal catalytic domain
are highly conserved. These two domains contain the binding motifs of two HMG-CoA substrate
and NADP (H) cofactor (Ruiz-albert et al., 2001; Shen et al., 2006). A defect of AtHMGR1 could
be complemented by only the catalytic domain of PgHMGR1, suggesting that the product of
PgHMGR1 is enough to complement without proper targeting to an organelle. However,
overexpression of PgHMGR2 driven by the 35S promoter and not the native promoter could not
complement. This implies that PgHMGR1 and PgHMGR2 could possibly be involved in
different metabolite pathways, ruling out the difference of the promoter or membrane domain
region. Although PgHMGR1 shares more identity with AtHMGR1 than AtHMGR2, it has a
closet relationship with EsHMGR in Acanthopanax, which is a small and woody shrub in the
same family, that is known to have similar herbal properties to those of P. ginseng (Huang et al.,
2011), and that clusters separately from the AtHMGR1 in Arabidopsis (Fig. 9A). PgHMGR2
shares high identity with HMGR2 (CaHMGR, AAB69727) from Camptotheca acuminata
(Maldonado-Mendoza et al., 1997), which also showed lower expression than HMGR1. This
suggests that PgHMGR1 and PgHMGR2 originated from a gene evolution event after the split
from Arabidopsis and before the separation of PgHMGR1 and its homologues. Considering these
findings, it is possible that each of the isozymes of HMGR could have evolved separately
according to the production of specific products rather than duplication.
In conclusion, our data suggest that PgHMGR1 is very likely to contribute to the ginsenoside
synthesis pathway, playing a housekeeping role as a functional ortholog of AtHMGR1. However,
despite the PgHMGR2 expression in ginseng roots, the specific role of PgHMGR2 is unclear. To
understand the contribution of PgHMGR2 to the ginsenoside pathway, we are now generating
PgHMGR2-overexpressing ginseng. Examining this transgenic ginseng should further clarify the
function of PgHMGR2. Further work will be required to clarify the specific role of the HMGR
isozymes in plants.
MATERIAS AND METHODS
Plant Materials and Growth Conditions
The Columbia ecotype of Arabidopsis thaliana was used as a model plant in this study. ER -
YFP (ER-yk, CS16251; Nelson et al., 2007)-, PX-YFP (PX-yk, CS16261; Nelson et al., 2007)-,
MT-YFP (MT-yk, CS16264; Nelson et al., 2007)-, and PT-YFP (PT-yk, CS16267; Nelson et al.,
2007)-expressing lines and hmgr1-1 (SALK_125435) were purchased from the Arabidopsis
stock center (http://www.Arabidopsis.org/). Seeds were surface sterilized and then sown on 1/2
MS medium (Duchefa Biocheme, The Netherlands) containing 1% sucrose, 0.5 g/L MES (2-[N-
morpholino]ethanesulphonic acid), pH 5.7 with KOH, and 0.8% agar. Three-day cold-treated
seeds were germinated under a long-day condition of 16 h light/8 h dark at 23°C. Transformants
were selected on hygromycin-containing plates (50 µg/mL). Ten-day-old seedlings were
transplanted into soil and allowed to grow for up to 5 weeks under the same light/dark conditions.
For RNA extraction, seedlings were grown on plates for 15 days. For metabolite analysis, leaves
and the inflorescence from 5-week-old plants were collected.
Ginseng Materials and Treatment
Mevinolin (Mev) and methyl jasmonate (MJ) were purchased from Sigma (St. Louis, MO,
USA). Stock solutions of Mev (10 mM) and MJ (100 mM) in ethanol were stored at -20°C. For
inhibitor treatment of ginseng, after 4 weeks of pre-cultivation of ginseng adventitious roots, 10
µM Mev or 10 µM MJ was added. After 1, 3, and 7 days, harvested adventitious roots were
frozen and used for RNA extraction, an HMGR activity assay, and ginsenoside analysis.
Three-year-old ginseng plants hydroponically cultured in perlite and peatmoss grown at
23°C±2 under white fluorescent light (60-100 µmol m-2
s-1
) in a controlled greenhouse (kindly
provided by i-farm in Yeo-Ju, Korea) were used for dark treatment. Control plants were grown in
16 h light/8 h dark and sampled in light conditions, while dark treatments lasted for 2 and 3 days
using a black-cover with an aerial part. Leaf and root parts were separately used for RNA
isolation, an HMGR activity assay, and ginsenoside analysis.
Identification of PgHMGR Genes and Sequence Analysis
In order to obtain a full-length coding sequence of the PgHMGR gene, rapid amplification of
cDNA ends (RACE) PCR was performed with a CapfishingTM
full-length cDNA premix kit
(Seegene, Korea) according to the product manual, with HotStarTaq (Qiagen) as DNA
polymerase. The first-strand cDNA synthetic reaction from total RNA was catalyzed by
superscript III RNase H reverse transcriptase (Invitrogen Life Technologies), according to the
product instruction manual, and the cDNA was prepared using a commercial cDNA synthesis kit
according to the manufacture's instruction manual (Clontech, US). Specific primers were
designed according to the 3’-end and 5’-end sequences of partial PgHMGR ESTs, which were
derived from our ginseng EST library. The primer sequences used were as follows: (HMGR1;
HMG_F2: 5’- ATG GAG GCC ATT AAC GAT GGA AAA G-3’, HMG_R2: 5’- ACC AAC CTC
AAT TGA TGG CAT TGT G-3’, HMG_R3: 5’- CCT TTT TGC CGT AGA AAA CCT AAC CA-
3’; HMGR2: HMG2_F2: 5’- TAC TCA CTC GAG TCC AAA CTG GGA GAC -3’, HMG2_R1:
5’-GGC ATG CTA ATT GGG TCC CAC CTC CT-3’, HMG2_R3: 5’- ATT GCC TCA CAA ACA
ACC GAT TTA CC -3’). RACE PCR was performed by the hot start method with the following
conditions: 30 cycles of 94ºC for 40 s, 60 - 66ºC for 40 s, 72ºC for 1 min 20 s, and a final
extension of 72ºC for 5 min. The PCR product was purified and ligated into a pGEM-T vector
(Promega, USA) followed by sequencing. By assembling the sequence of the 3’-RACE and 5’-
RACE products, the full-length cDNA sequence of PgHMGRs was deduced. A 60 ng genomic
DNA of ‘Yunpoong’ and a pair of PCR primers (Start codon and Stop codon) were used for
amplifying the genomic sequence of PgHMGR.
Deduced amino acid sequences were searched for homologous proteins in the databases using
BLASTX network services at the NCBI. ClustalX with default gap penalties was used to perform
multiple alignment of HMGRs isolated in ginseng and previously registered in other species. A
phylogenetic tree was constructed by the neighbor-joining method, and the reliability of each
node was established by bootstrap methods using MEGA4 software. Identification of conserved
motifs of HMGR was accomplished with MEME. A three-dimensional model was prepared by
using PgHMGR as a template on a SWISS-MODEL WORKSPACE in automated mode. The
generated 3-D structure was visualized by the UCSF Chimera package.
Isolation of Promoter Sequences
The Universal Genome Walker™ Kit (Clontech Laboratories, Inc., Palo Alto, CA, USA) was
used to isolate fragments of the PgHMGR1 and PgHMGR2 promoter. Ten 6 bp-recognizing and
blunt end-forming restriction enzymes (DraI, EcoRV, PvuII, StuI, SspI, SmaI, MscI, ScaI,
Eco105I, and HpaI) were used to digest the isolated ‘Yungpoong’ genomic DNA. DNA
fragments containing the adaptors at both ends were used as templates for amplifying the
PgHMGR promoter regions. The first PCR, using AP1 and H1–GSP1 and H2-GSP1, respectively,
(H1-GSP1, 5’-ATA ACA GTC CCC GAG TTT TGA TTC CAG-3’; H2-GSP1, 5’-GCT TGG
GAG GAA GAG AAC AGT CGA TAG C-3’) and the second nested PCR, using AP2 and
H1_GSP2 and H2_GSP2, respectively, (H1_GSP2, 5’-ACG TGA GAC AAA TGA TTG GAC
GAA ATC-3’; H2_GSP2, 5’-ATG GCT CTC GAC GAC TAT CTT CCT CGT T-3’) were
performed using i-MAXII (Intron, Korea). The amplified fragment was cloned in the pGEM-T
vector and sequenced. The promoters were analyzed using PlantPan (Plant Promoter Analysis
Navigator).
Vector Construction and Arabidopsis Transformation
To visualize the subcellular localization patterns of ginseng HMGRs, cDNA sequences of
PgHMGR1 and PgHMGR2 were cloned into a pCAMBIA1390 vector containing the cauliflower
mosaic virus (CaMV) 35S promoter and eCFP. The PgHMGR1 cDNA and its catalytic domain
(PgHMGR1CD; amino acid residues 154 to 573) were amplified using primers with XhoI and
EcoRI sites (PgHMGR1: 5’-TA CTC GAG ATG GAC GTC CGC CGG AGA-3’ and 5’-TG GAA
TTC AGA TCC AAT TTT GGA CAT-3’, PgHMGR1CD: 5’-TA CTC GAG ATG CCC GTA GTG
ATG TCA-3’ and 5’-TG GAA TTC AGA TCC AAT TTT GGA CAT-3’). The PgHMGR2 cDNA
was amplified using primers with SalI and AvrII sites: 5’-CA GTC GAC ATG GAC GTT CGC
CGG CGA CCT-3’ and 5’- GA CCT AGG CTA GGA GGA GAG TTT TGT-3’. Enzyme-digested
PCR products were cloned into the PLA2a gene site of Pro35S:PLA2a-
eCFP/pCAMBIA1390 (provide by Dr. Lee) as eCFP fusion proteins (Pro35S:PgHMGR1-eCFP,
Pro35S:PgHMGR1CD-eCFP or Pro35S:PgHMGR2-eCFP). To express the PgHMGR1 fusion
protein with mRFP at the C terminus, the mRFP was amplified from 326-mRFP using primers
with EcoRI and SpeI sites (5’-TG GAA TTC ATG GCC TCC TCC GAG GAC-3’ and 5’-GG
ACT AGT TTA GGC GCC GGT GGA GTG-3’) and was replaced with eCFP of
Pro35S:PgHMGR1-eCFP.
The 1317 and 477 nucleotides of genomic PgHMGR1 and PgHMGR2 DNA, respectively,
were amplified with HindIII and BamHI sites (ProPgHMGR1; 5’-TG AAG CTT AGC TAA
AGA AAG TTA GGC-3’ and 5’-TA GGA TCC GGA AGA GTA TAT TCC GGC-3’,
ProPgHMGR2; 5’-CG AAG CTT GTA ATG TGT TCC CAC TTC CAT-3’ and 5’-TA GGA TCC
CTT TAT GGT GGG GGA ACT CCG-3’) and cloned to a pCambia 1390 vector containing the
GUS gene. All transgene constructs were confirmed by nucleotide sequencing before plant
transformation. The constructs were transformed into Arabidopsis using Agrobacterium
tumefaciens C58C1 (pMP90) (Bechtold and Pelletier, 1998). The insertion of transgenes into the
transformants was confirmed by confocal microscopy or PCR analysis of the genomic DNA
from the transformants. Homozygous plants with a 3:1 segregation ratio were selected on
antibiotic plates for further analyses. For each construct, 20-50 T1-independent lines were
obtained, and the phenotypic significance of the transgenes was analyzed in the chosen lines.
Transformation of Ginseng
Zygotic embryos of P. ginseng cv. “Yunpoong” seeds (provided by Ginseng Genetic Resource
Bank) were stratified in humidified sand to mature for three months at 5°C. After stratification,
the seeds, in which zygotic embryos were in a mature state (4 mm in length), were immersed in
70% ethanol for 1 min, surface sterilized in 2% NaOCl for 15 min, and rinsed three times with
sterilized distilled water. After carefully dissecting the zygotic embryos, they were placed on MS
basal medium containing 3% sucrose and 0.8% agar. Five-day-old zygotic embryos were excised
transversely, and cotyledon explants (4 mm) were used for transformation. The explants were
dipped in the bacterial solution for 15 min, subsequently blotted with sterile filter paper, and co-
cultivated with A. tumefaciens C58C1 on MS medium containing 3% sucrose and 1 µg/mL 2,4-
dichlorophenoxyacetic acid (2,4-D) for 2 days. Thereafter, the explants were cultured on
selection MS medium with 3% sucrose, 1 µg/mL 2,4-D, 0.5 µg/mL 6-benzylaminopurine (BA),
250 µg/mL cefotaxime, and 50 µg/mL hygromycin. After subculturing six times every two weeks
on selection medium, the surviving cotyledons producing calli were cultured on the same
medium without antibiotics, and adventitious roots were induced from calli on B5 medium with
3% sucrose and 3 mg/L indole-3-butyric acid (IBA). More than 10 transgenic lines were
generated, and adventitious roots were excised from the maternal explants and sub-cultured in
liquid B5 medium with 3% sucrose and 2 mg/L IBA every 5 weeks.
Expression Analysis
Total RNA was extracted from frozen samples with the RNeasy plant mini kit (Qiagen, USA),
including the DNase I digestion step. Next, 2 µg of the total RNA was reverse transcribed with
RevertAid™ H Minus M-MuLV Reverse Transcriptase (Fermentas, USA). RT-PCR was
performed in a 25 µl reaction volume consisting of 0.2 µl of the cDNA product and 5 pmol of
each primer using Super-Taq DNA polymerase (Super Bio, Korea) by a Bio-Rad PCR machine
(3 min at 95ºC, followed by 28 cycles at 94ºC for 20 s, 60ºC for 20 s, and 72ºC for 30, with a
final extension at 72ºC for 5 min). The products were analyzed on 1.2% agarose gels.
Real-time quantitative PCR was performed using 100 ng of cDNA in a 10-µl reaction volume
using SYBR®
Green Sensimix Plus Master Mix (Quantace, Watford, England). The following
thermal cycler conditions recommended by the manufacturer were used: 10 min at 95ºC,
followed by 40 cycles at 95ºC for 10 s, 60ºC for 10 s, and 72ºC for 20 s. The fluorescent product
was detected during the final step of each cycle. Amplification, detection, and data analysis were
carried out on a Rotor-Gene 6000 real-time rotary analyzer (Corbett Life Science, Sydney,
Australia). To determine the relative fold-differences in template abundance for each sample, the
Ct value for each of the gene-specific genes was normalized to the Ct value for ß-actin and
calculated relative to a calibrator using the formula 2ㅡΔCt
or 2ㅡΔΔCt
. Three independent
experiments were performed, and the primer efficiencies were determined according to the
method of Livak and Schmittgen (2001) to validate the ΔΔCt method used in our experiment.
The observed slopes were close to zero, indicating that the efficiencies of the gene and the
internal control ß-actin were equal.
Confocal Microscopy Analysis
The fluorescence from reporter proteins and organelle markers was observed by confocal laser
scanning microscopy (LSM 510 META, Carl Zeiss, Jena, Germany). GFP, CFP, YFP, and mRFP
were detected using 488/505-530, 458/475-525, 514/>530, and 543/560-615 nm
excitation/emission filter sets, respectively. Fluorescence images were digitized with the Zeiss
LSM image browser.
HMGR Activity Assay
Extraction of crude proteins was performed as described (Rodríguez-Concepción et al., 2004),
with minor modifications. Two-week-old seedlings of Arabidopsis, leaves or roots of 3-year-old
ginseng, or adventitious roots of ginseng (~200 mg) were homogenized in liquid nitrogen and
mixed with 1 ml of prechilled extraction buffer containing 100 mM sucrose, 40 mM sodium
phosphate pH 7.5, 30 mM EDTA, 50 mM NaCl, 10 mM DTT, 1 mM AEBSF, 1 µM bestatin, 15
µM E64, 20 µM leupeptin, 15 µM pepstatin, 0.5 mM pheanthroline, 0.5 mM
phenylmethylsulfonyl fluoride, and 0.25% (w/v) Triton X-100. The slurry was centrifuged at 200
g and 4°C for 10 min to remove cell debris, and the supernatant was used for HMGR activity as
described (Dale et al., 1995) with the following modifications, and the protein concentration was
determined with the BCA Protein Assay kit (Intron, Korea). The supernatant was analyzed using
a spectrophotometric assay at 37°C (total volume 200 µl) containing 0.3 mM of HMG-CoA, 0.2
mM of NADPH, and 4 mM of dithiothreitol in 50 mM Tris-HCl, pH 7.0. The decrease in
absorbance at 340 nm was monitored for 20 min after the addition of HMG-CoA. One unit of
HMGR activity is defined as the amount of enzyme which oxidizes 1 µmol of NADPH per
minute at 37°C.
GUS Histochemical Analysis
Four-day-old seedlings were treated with the indicated chemical for 3 h before visualizing the
GUS activity. GUS staining was performed by incubating whole seedlings in the staining buffer
containing 1 mM 5-bromo-4-chloro-3-indoyl-β-D-glucuronic acid cyclohexylammonium salt (X-
Gluc, Duchefa Biocheme, The Netherlands), 0.1 M NaH2PO4, 0.1% Triton-X, and 0.5 mM
potassium ferri- and ferrocyanide at 37°C until a blue color appeared (1-3 h). Stained seedlings
were cleared in 70% ethanol for 2 h and 100% ethanol for 2 h. In the final step of dehydration,
samples were sequentially exposed to 10% (v/v) glycerol/50% (v/v) ethanol and 30% (v/v)
glycerol/30% (v/v) ethanol. Seedlings were photographed under a microscope (ZEISS Axio
Observer D1, Germany).
HPLC Analysis of Ginsenosides Extracted from Ginseng
For the analysis of ginsenosides from different ginseng samples, 0.3 - 1 g of milled powder of
freeze-dried adventitious roots, leaves, and roots were soaked in 80% MeOH at 70°C. After the
liquid was evaporated, the residue was dissolved in H2O, followed by extraction with H2O-
saturated n-butanol. The butanol layer was then evaporated to produce the saponin fraction. Each
sample was dissolved in MeOH (1 g/5 ml) and then filtered through a 0.45 µm filter and used for
HPLC analysis. The HPLC separation was carried out on an Agilent 1260 series HPLC system
(Palo Alto, CA, USA). This experiment employed a C18 (250 x4.6 mm, ID 5 µm) column using
acetonitrile (solvent A) and distilled water (solvent B) mobile phases, with a flow rate of 1.6
ml/min and the following gradient: A/B ratios of 80.5:19.5 for 0-29 min, 70:30 for 29-36 min,
68:32 for 36-45 min, 66:34 for 45-47 min, 64.5:35.5 for 47-49 min, 0:100 for 49-61 min, and
80.5:19.5 for 61-66 min. The sample was detected by UV 203 nm. Quantitative analysis was
performed with a one-point curve method using external standards of authentic ginsenosides.
GC-MS Analysis of Sterols and Triterpenes Extracted from Arabidopsis
Using the method modified from Suzuki et al. (2004), freeze-dried plant materials (rosette
leaves, 200 mg; inflorescence, 25 mg) were powdered and then extracted twice with 1 ml CHCl3-
MeOH (7:3) at room temperature. 5-α-Cholestane (20 µg) was used as an internal standard. The
extract was dried in a rotary evaporator and saponified with 1.5 ml each of MeOH and 20%
KOH aq. for 1 h at 80°C to hydrolyze the sterol esters. After saponification, 1.5 ml each of
MeOH and 4 N HCL were added for 1 h at 80°C, and these reaction mixtures were extracted
three times with 4 ml of hexane. When the phases separated, the sterols partitioned to the hexane
layer, and the combined hexane layer was evaporated to dryness. The residue was
trimethylsilylated with pyridine and BSTFA+1% TMCS (1:1) at 37°C for 90 min and analyzed
by GC-MS.
Capillary gas chromatography-mass spectrometry (GC-MS) analysis was performed using a
mass spectrometer (HP 5973 MSD) connected to a gas chromatograph (6890A, Agilent
Technologies) with a DB-5 (MS) capillary column (30 m×0.25 mm, 0.25 μm film thickness). The
analytical conditions were as follows: EI (70 eV), source temperature 250°C, injection
temperature 250°C, column temperature program: 80°C for 1 min, then increased to 280°C at a
rate of 10°C/min and held at this temperature for 17 min; post temperature of 300°C, carrier gas
He, flow rate 1 ml/min, run time of 38 min; splitless injection. The endogenous sterol levels were
determined as the peak area ratios of molecular ions of the endogenous sterol and internal
standard. Standards including squalene, phytosterols (campesterol, β-sitosterol, stigmasterol),
and triterpene (β-amyrin, α-amyrin) were purchased from Sigma (USA).
For sesquiterpene analysis, the solid-phase microextraction (SPME)-trapped volatiles
(Aharoni et al., 2008) were analyzed by GC-MS as described by Verhoeven et al. (1997). Exactly
0.3 g of inflorescence of 5-week-old Arabidopsis was introduced into a 20 ml vial (Gerstel,
Mu l̈heim, Germany). The fused silica fiber of the SPME device coated with 100 µm of
polydimethylsiloxane was inserted into the vial through an aluminum cap, and volatiles were
trapped by exposing the fiber to the headspace for 30 min. A 65 µm PDMS-DVB
(polydimethylsiloxane – divinylbenzene)-coated fiber was used. The SPME fiber was exposed
for 20 min in the head-space at 45°C, and then fiber was withdrawn into the needle and
transferred to the injector of the GC-MS. An HP-5 column (50 m x 0.32 mm, film thickness 1.05
pro) was used with He (37 kPa) as the carrier gas. The GC oven temperature was programmed as
follows: 80°C for 2 min, increased to 250°C at a rate of 8°C /min, and held at 250°C for 5 min.
Mass spectra in the electron impact mode were generated at 70 eV. The compounds were
identified by comparison of GC retention indices and mass spectra with those of authentic
reference compounds.
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FIGURE LEGENDS
Figure 1. Biochemical pathway for biosynthesis of Panax ginseng saponins. Classification of
main ginsenosides based on attached glycosides and the dammarendiol-type structure (A). Glc,
β-D-glucopyranosyl; Ara (pyr), α-L-glucopyranosyl; Ara (fur), α-L-arabinofuranosyl; Rha, α-L-
rhamncpyranosyl. Ginsenoside biosynthesis pathway (B). FPS, farnesyl diphosphate synthase;
SS, squalene synthase; SE, squalene epoxidase; DDS, dammarenediol synthase; β-AS, β-amyrin
synthase; CAS, cycloartenol synthase; P450, cytochrome P450; GT, glucosyltransferase;
Mevinolin, a competitive inhibitor of HMGR. Dotted line displays putative pathway. Reported
enzymes in ginseng are shown with NCBI accession number in bracelet.
Figure 2. Analysis of sequence and domain characteristics of PgHMGR1 and PgHMGR2. The
DNA structures of PgHMGR1 and PgHMGR2 genes. Exons are represented by dark-filled
square boxes. Lines between boxes correspond to introns. Numbers above exons and below
introns indicate their length in bp. Indicated triangle represents the starting site used for H1CD
cloning in this study (A). The predicted 3-D structures of PgHMGR1 and PgHMGR2, which
consist of three domains: the small and helical N-terminal N-domain; the large, central L-domain;
and the small helical S-domain which is inserted into the L-domain. The catalytically active
residues (Abdin et al., 2012) are highlighted in yellow, and the sequences are indicated in
Supplemental Figure S5. (B). Schematic representation of the domain structures of PgHMGR1
and PgHMGR2. HMGR proteins are comprised of three domains: the membrane-anchor domain,
a flexible linker domain, and the catalytic domain. Within the catalytic domain, three subdomains
have been defined. H1 and H2 indicate two probable trans-membrane hydrophobic regions. LS,
lumenal sequence. The catalytic domain contains HMG-CoA- and NADH(H)-binding regions
and three conserved motifs. The active sites are indicated as *. The numbers indicate the position
in the amino acid sequence of the start and end of each domain and motif (C).
Figure 3. Ubiquitous expression of PgHMGR1 and PgHMGR2 in vascular tissue. Different aged
ginseng materials used for RT-PCR (A). Expression patterns of PgHMGR1 in seed, 1-week-old
seedlings, and leaves, petioles, and roots of 2-week-old of seedlings (B). Expression patterns of
PgHMGR1 in flower buds, leaves, stem, main roots, and lateral roots of 3-year-old (C) and 6-
year-old ginseng (D). Expression patterns of PgHMGR2 (E, F, G) were analyzed using the same
tissue as with PgHMGR1. Vertical bars indicate the mean value ± SE from three independent
experiments. Scale bars indicate 1 cm in A. Histochemical analysis of GUS expression in
transgenic Arabidopsis plants harboring the pHMGR1::GUS (H) and pHMGR2::GUS (I)
constructs at different developmental stages. (a) GUS expression in 2-day-old germinated
seedlings. Main roots (b), lateral roots (c), true leaves, and cotyledons (d) of 8-day-old seedlings.
Mature flowers (e) and siliques (f) of 50-day-old plants. All scale bars indicate 100 µm in H, I,
and J. PgHMGR1::GUS expression is shown in the root vasculature and root tip of ginseng (J).
Figure 4. Expression of PgHMGR1 and PgHMGR2 is associated with the production of
ginsenoside. Mevinolin (10 µM), an inhibitor of HMGR, was administered to 4-week-old
adventitious roots for 1, 3, and 7 days. We analyzed total ginsenoside contents (mg/g dry weight)
(A), gene expression of PgHMGR1 and PgHMGR2 (B), and relative HMGR activity (C). Methyl
jasmonate (MJ, 10 µM), an elicitor of saponin biosynthesis, was treated for 3 days, and the total
ginsenoside contents (mg/g dry weight) (D) and gene expression (E) were analyzed. Vertical bars
indicate the mean value ± SE from five independent experiments. *: Significantly different from
the control at P <0.05. (F) PgHMGR1::GUS- or PgHMGR2::GUS-expressing seedlings grown
on ½ MS media for 4 days were treated with ethanol control (Cont), 1 or 10 µM of mevinolin,
and 10 µM of MJ for 3 hours. Bar= 100 µM.
Figure 5. Expression of PgHMGR1 is light-inhibited and possibly associated with shadowing
growth of ginseng. Three-year-old ginseng plants cultured hydroponically in perlite and
peatmoss were kept in darkness for 2 or 3 days. Control plants (Cont) were grown in a 16-h
light/8-h dark cycle, and samples were taken under light conditions. Parts of the leaf and root
were separately sampled and used for RT-PCR, HMGR activity assay, and ginsenoside analysis.
Total ginsenoside contents in the leaves (A) and roots (B) and individual major ginsenosides
were analyzed (Supplemental Figure 4). Vertical bars indicate the mean value ± SE from three
independent experiments. * and **, Significantly different from the control at P <0.05 and **
P<0.01, respectively. Relative HMGR activity in the leaves and roots was analyzed (C). Data
represent mean value ± SE (n=10). Gene expression of PgHMGR1 and PgHMGR2 in leaves (D)
and roots (E) was analyzed by real-time PCR. Data represent mean value ± SE (n=5). Putative
PIF3 binding motifs are found on the G-box or ACE element of the analyzed promoter of the
PgHMGR1 gene, which are not found in PgHMGR2, by Plantpan (F). Hypocotyl expression in
4-day-old etiolated seedlings of PgHMGR1::GUS was decreased by 2 h of light exposure (G).
Expression in true leaves of 10-day-light-grown PgHMGR1::GUS seedling was decreased by 9 h
of darkness (H).
Figure 6. Subcellular localization of PgHMGR1, PgHMGR2, and the catalytic domain of
PgHMGR1. Fluorescent images of PgHMGR1-CFP (A) and PgHMGR1-mRFP (B) were
visualized by a confocal laser scanning microscope. A fluorescent image of PgHMGR2-CFP was
visualized with green color in an ER-like vesicle (C). The catalytic domain of PgHMGR1 tagged
with CFP at the C-terminus was localized in the cytosol and nucleus (D). Fluorescent signals of
PgHMGR1-mRFP [Pro35S:PgHMGR1–mRFP in the ER-YK (ER marker), PX-YK (peroxisome
marker), PT-YK (plastid marker) transgenic background] overlap with ER-YK, PX-YK, and PT-
YK foci (E). All scale bars indicate 5 µm.
Figure 7. Overexpression of PgHMGR1 or the catalytic domain of PgHMGR1 results in higher
production of sterol and triterpene in plants. Quantification of endogenous sterols and triterpenes
in Arabidopsis HMGR1ox and H1CDox. Rosette leaves (200 mg) and inflorescences (25 mg) of
5-week-old plants were harvested and freeze-dried. The samples were then analyzed using GC-
MS. Values represent the mean ±SE of content (µg per g dry weight) calculated from three
independent sterol isolations and quantification. * and **: Significantly different from control at
P <0.05 and P<0.01, respectively. Quantitative analysis of triterpene ginsenoside contents in the
control and transgenic ginseng adventitious roots of Panax ginseng (cv. Yunpoong)
overexpressing PgHMGR1 were analyzed by HPLC (C). Vertical bars indicate the mean ± SE
from three independent experiments.
Figure 8. Conserved role of PgHMGR1. A phylogenetic tree of HMGRs from various organisms
including PgHMGR1 and PgHMGR2 (A). Eleutherococcus senticosus (EsHMGR, AFM77981),
Eucommia ulmoides (EuHMGR, AAV54051), Salvia miltiorrhiza (SmHMGR, ACD37361),
Panax quinquefolius (PqHMGR, ACV65036), Camptotheca acuminata (CaHMGR, AAB69727),
Solanum chacoense (ScHMGR, AEX26934), Arabidopsis thaliana (AtHMGR1S, At1g76490;
AtHMGR2, At2g17370), fungi Pichia jadinii (O74164), and animals such as Homo sapiens
(NP_000850), Drosophila melanogaster (P14773), and Mus musculus (NP_032281). The
neighbor-joining method was used, and the branch lengths are proportional to the divergence,
with the scale of 0.1 representing 10% changes. The conserved motifs among the members are
highlighted in colored boxes with an arranged number, and the sequences of the motifs are listed
in Supplemental Figure S7 (B). PgHMGR1 complemented hmgr1-1. Structure of T-DNA
insertion position in the Arabidopsis genome of SALK line (SALK_125435) (C). Closed boxes
are exons. The T-DNA insertion site is indicated by a flag. T-DNA sequences were inserted in the
first exon in hmg1-1. Expression analysis of the AtHMGR1, AtHMG2, AtSE1, and SAG12, a
senescence marker gene, in 3-week-old seedlings (F) of wild-type (WT), heterozygous (HZ), and
homozygous (HM) lines of hmgr1-1 (D). The expression level of an actin gene was used as an
internal control. Real-time quantitative RT-PCR analysis of AtHMGR1 confirmed the knock-out
of hmgr1-1. Data are normalized with β-actin and are relative to the levels in WT seedlings.
Fifty-day-old Arabidopsis plants (E, G-I). Heterozygous (HZ) hmgr1-1 shows a similar
phenotype to the wild-type (Col), whereas the homozygous (HM) mutant shows a severe dwarf
and sterile phenotype (E, F). Exogenous squalene treatment rescued the dwarf phenotype (E).
PgHMGR1ox (G) and PgHMGR1CDox (H) complemented the hmgr1-1 dwarf phenotype,
whereas PgHMGR2ox (I) did not.
Figure 9. Presumptive model for the HMGR-triggered ginsenoside biosynthesis. Sterols and
triterpenes produced via mevalonate (MVA) are catalyzed by 3-hydroxy-3-methylglutaryl-CoA
reductase (HMGR) in the endoplasmic reticulum (ER) or peroxisome of the vasculature of
ginseng. HMGR can be up-regulated by the light signaling factor, phytochrome-interacting factor
3 (PIF3), regulated by phyB (phytochrome B).
Supplemental Table 1. Primers used in confirmation of gDNA insertion and RT-PCR
No. Organism Gene Accession
No. Annotation Primers used (5'-3')
Annealing
temp. (°C)
1 Ginseng PgHMGR1
HMG-F TGCTGCCAATATCGTCTCTG 60
2
HMG-R CCAAGAAGGTTCAAGCAAGC
3 Ginseng PgHMGR1
H1_RT_F2 ATGTAGATCGCTCGCCGT 60
4
H1_RT_R2 GTGGAGGAGATAGTATGCGAC
5 Ginseng PgHMGR2
H2_RT_F AGGCCTTCGCCTGATGCCT 60
6
H2_RT_R GCAAATGCTGACTTGGGATT
7 Ginseng Pgactin DC03005B05 actin-left AGAGATTCCGCTGTCCAGAA 60
8
actin-right ATCAGCGATACCAGGGAACA
9 Arabidopsis AtHMGR1 At1G76490 Arabi_H1_F ATTGTCACCGAATCGCTTCC 59
10
Arabi_H1_R GATCTCCCGGTGACTCTCTG
11 Arabidopsis AtHMGR2 At2G17370 Arabi_H2_1300F TGAAGCTGAAGGCAACGACCT 59
12
hmgr2-2RP CTGTTGACTTGAGACGAAGGG
13 Arabidopsis AtSE1 At1g58440 AtSE_F AGCTGGTGTTGCTGGTTCT 57
14
AtSE_R CTTCCACACAATCTTCAATTCC
15 Arabidopsis SAG12 At5g45890 AtSAG12_F ATTCCTGCCGGAAGAACTTT 54
16
AtSAG12_R TATCCATTAAACCGCCTTCG
17 Arabidopsis Atactin At5g09810 At-actin2F GTGTGTCTTGTCTTATCTGGTTCG 58
18
At-actin2R AATAGCTGCATTGTCACCCGATACT
19 Vector CFP, GFP
GFP342rv CTCGACCAGGATGGGCAC
20
35S
promoter 35S GCACAATCCCACTATCCTTCG
21
NOS terminator p33NOS ACCGGCAACAGGATTCAATCT
37
Supplemental Data
Supplemental Figure S1. Genomic DNA gene sequence of PgHMGR1 with its promoter (-1317
to -1). The full-length genomic DNA sequence with its deduced amino acid sequence. A one-
letter code below the corresponding codons and intron sequence is shown. Thin-line boxes in the
promoter region=putative ‘TATA’ boxes; thin-line underlines=putative ‘CAAT’ boxes; shaded
bold box=putative ‘G-box’; bold underline=putative ‘ACE element’; shaded box in the N-
terminal=putative motif for ER retention; square boxes in the intron region=putative intron-exon
boundaries with the GT-AG rule.
Supplemental Figure S2. Genomic DNA gene sequence of PgHMGR2 with its promoter (-477
to -1). The full-length genomic DNA sequence with its deduced amino acid sequence. A one-
letter code below the corresponding codons and intron sequence is shown. Thin-line boxes in the
promoter region=putative ‘TATA’ boxes; thin-line underline=putative ‘CAAT’ boxes; shaded box
in the N-terminal=putative motif for ER retention; square boxes in the intron region=putative
intron-exon boundaries with the GT-AG rule.
Supplemental Figure S3. Ginsenoside contents by inhibitor or elicitor treatment. Individual
ginsenosides (A) were analyzed in 4-week-old adventitious roots (B) following 3 day treatment
with the chemicals; Mevinolin (10 µM), an inhibitor of HMGR and methyl jasmonate (MeJA, 10
µM), an elicitor of saponin biosynthesis. Vertical bars indicate the mean ± SE from three
independent experiments. *: Significantly different from control at P <0.05.
Supplemental Figure S4. Ginsenoside contents by dark treatment. Three-year-old ginseng
plants cultured hydroponically in perlite and peatmoss were kept in darkness for 2 or 3 days.
Control plants (Cont) were grown in a 16-h light/8-h dark cycle, and samples were taken under
light conditions. Individual major ginsenosides in leaves (A) and roots (B) were analyzed.
Vertical bars indicate the mean ± SE from three independent experiments. * and **, Significantly
different from control at P <0.05 and ** P<0.01, respectively.
Supplemental Figure S5. Multiple alignment of the deduced amino acid sequences of
38
PgHMGR1 and PgHMGR2 with homologous HMGRs from other plants (Fig. 8):
Eleutherococcus senticosus (EsHMGR, AFM77981), Eucommia ulmoides (EuHMGR,
AAV54051), Salvia miltiorrhiza (SmHMGR, ACD37361), Panax quinquefolius (PqHMGR,
ACV65036), Camptotheca acuminata (CaHMGR, AAB69727), Solanum chacoense (ScHMGR,
AEX26934), and Arabidopsis thaliana (AtHMGR1S, At1g76490; AtHMGR2, At2g17370).
Black boxes indicate identical residues; gray boxes indicate identical residues for at least two of
the sequences. Conserved domains are indicated with arrowed lines and three conserved motifs
are highlighted in colored boxes. Two putative HMGR-CoA-binding sites and two NADP(H)-
binding sites are indicated with a thin square box (Shen et al., 2006), and the motif responsible
for ER retention is highlighted with bold square box (Merret et al., 2007). The indicated triangle
represents the starting site for the H1CD cloning in this study.
Supplemental Figure S6. Conserved motifs among plant and animal HMGRs. Conserved
residues analyzed by MEME are represented by three motifs. The catalytically active residues
marked by a box (Abdin et al., 2012) were conserved in these three motifs.