Plant Cells as Pharmaceutical Factories

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Current Pharmaceutical Design, 2013, 19, 5640-5660

Plant Cells as Pharmaceutical Factories Heiko Rischer1,*, Suvi T. Häkkinen1, Anneli Ritala1, Tuulikki Seppänen-Laakso1, Bruna Miralpeix2, Teresa Capell2, Paul Christou2,3 and Kirsi-Marja Oksman-Caldentey1 1

VTT Technical Research Centre of Finland, P.O. Box 1000, 02044 VTT (Espoo), Finland; 2Departament de Producció Vegetal i Ciència Forestal (PVCF), Universitat de Lleida-Agrotecnio Center, Av. Alcalde Rovira Roure, 177, Lleida, E-25198, Spain; 3 Institució Catalana de Recerca i Estudis Avançats (ICREA) Barcelona, Spain Abstract: Molecules derived from plants make up a sizeable proportion of the drugs currently available on the market. These include a number of secondary metabolite compounds the monetary value of which is very high. New pharmaceuticals often originate in nature. Approximately 50% of new drug entities against cancer or microbial infections are derived from plants or micro-organisms. However, these compounds are structurally often too complex to be economically manufactured by chemical synthesis, and frequently isolation from naturally grown or cultivated plants is not a sustainable option. Therefore the biotechnological production of high-value plant secondary metabolites in cultivated cells is potentially an attractive alternative. Compared to microbial systems eukaryotic organisms such as plants are far more complex, and our understanding of the metabolic pathways in plants and their regulation at the systems level has been rather poor until recently. However, metabolic engineering including advanced multigene transformation techniques and state-of-art metabolomics platforms has given us entirely new tools to exploit plants as Green Factories. Single step engineering may be successful on occasion but in complex pathways, intermediate gene interventions most often do not affect the end product accumulation. In this review we discuss recent developments towards elucidation of complex plant biosynthetic pathways and the production of a number of highvalue pharmaceuticals including paclitaxel, tropane, morphine and terpenoid indole alkaloids in plants and cell cultures.

Keywords: Plant cell culture, medicinal plants, natural products, secondary metabolites, pharmaceuticals, genetic engineering. INTRODUCTION Eukaryotic organisms such as higher plants are exquisite designers and producers of a variety of small compounds - referred to as secondary metabolites - that are beneficial as foods, flavours, fragrances, fine chemicals and pharmaceuticals [1]. The first written document about the use of medicinal plants can be found in Papyrus Ebers (1550 BC). Even though the use of certain medicinal plants to treat diseases was known for millenia, only about 200 years ago the first active chemical constituent responsible for a pharmacological effect was isolated. This compound was morphine which was isolated from opium poppy (Papaver somniferum L.) by the German pharmacist Friedrich Sertürner. Morphine is still today being isolated from plant material. It is an interesting natural compound which is in wide use because of its superior effects as an analgesic drug. Many plant-derived compounds are used in the pharmaceutical industry, and plants also serve as an important source for new lead compounds. Many plants containing high-value secondary metabolites are difficult to cultivate or are becoming endangered because of overharvesting. Furthermore, the chemical synthesis of plant-derived compounds is often uneconomical due to their highly complex structures and the specific stereochemical features (see Miralpeix et al., this issue). The biotechnological production of valuable secondary metabolites in plant cell or organ cultures is an attractive alternative to the extraction of whole plant material. However, the use of plant cell or organ cultures has had only limited commercial success thus far. This can be explained by the empirical nature of selecting high-yielding, stable cultures and the lack of our understanding of how secondary metabolites are synthesized and how their synthesis is regulated [2]. More recently spectacular advances in plant genomics and metabolite profiling have offered unprecedented opportunities to explore the extraordinary complexity of the plant’s biochemical capacities. State-of-the-art genomics tools can be used *Address correspondence to this author at the VTT Technical Research Centre of Finland, P.O. Box 1000, 02044 VTT (Espoo), Finland; Tel: +358207224461; Fax: +358207227071; E-mail: [email protected] 18-8/13 $58.00+.00

to improve the production of known natural compounds or to synthesize entirely novel plant constituents by “combinatorial biochemistry” in cultivated plant cells. Therefore the utilization of plants cells as green chemical factories is becoming more realistic [3]. Although there is a large number of important natural products on the market we have selected here only some representative examples among terpenes and alkaloids. We discuss below the importance of plant-derived natural compounds as pharmaceutical compounds, the challenges in their chemical analysis, and their role in modern drug discovery. Furthermore we present the current knowledge and rapid development in the elucidation of the biosynthetic pathways leading to the desired end products. The state of the art in utilizing biotechnological processes in the production of specific plant secondary metabolites is also discussed. NATURAL PRODUCTS AS PHARMACEUTICALS Plant secondary metabolites or plant natural products can be defined as small molecules that have no recognized role in the maintenance of fundamental life processes in the plants that synthesize them, but they certainly have an important role in the interaction of the plant with its environment. Each plant has its particular spectrum of secondary metabolites and their synthesis is often strongly correlated to the vegetative stage of the plant. Many secondary metabolites have a very complex and unique structure thus introducing analytical challenges. Often their production is enhanced by both biotic and abiotic stresses. Based on their biosynthetic origins plant secondary metabolites can be structurally divided into five major groups: polyketides, isoprenoids (e.g. terpenoids), alkaloids, phenylpropanoids and flavonoids. Typically these compounds occur only in very low amounts in plant tissues [3]. The nutritional value of edible plants and their constituents has been studied for decades. In addition to edible plants there is an immense variety of toxic and medicinal plants in the plant kingdom [4], (Table 1). According to recent estimations there are about 422 000 higher plant species (K. Saito, Chiba University, Japan, personal communication) of which less than 10% are characterized © 2013 Bentham Science Publishers

Plant Cells as Pharmaceutical Factories

Table 1.

Current Pharmaceutical Design, 2013, Vol. 19, No. 31

5641

Number of plant species used for food and medication worldwide (according to [4]). Use of plants

Existing higher plants

Number of plant species 250000 – 500000*

Used for foods

3000

Commercially cultivated species

150

Highest caloric intake

20

Used for medication Used in western medicine

10000 150-200

*estimates vary according to different taxonomic concepts

chemically at least to some extent [5]. The majority of plant species occur in tropical rain forests. Natural products preparations or plant extracts and infusions have historically been the major source of the pharmaceutical agents. After the isolation of the first natural plant compound, morphine, 200 years ago, pure isolated natural compounds have become important pharmaceuticals in modern society. Often it has been difficult to find a new chemical substitute to these old natural compounds that possess the same efficacy and specificity. These include many alkaloids or terpenes derived from medicinal plants such as Atropa belladonna, Camptotheca acuminata, Capsicum annuum, Catharanthus roseus, Erythroxylum coca, Papaver somniferum, Cannabis sativa, Artemisia annua and Taxus species – just to name a few. These plants have been and still are an important source of pharmaceuticals with high monetary value. According to the WHO 11% of the current 252 drugs considered essential for humans are exclusively derived from flowering plants [6]. In industrialized countries ca. 25% of prescription medicines contain at least one compound that is directly or indirectly (via semi-synthesis) derived from natural origin [7]. Despite the huge efforts which was put on drug discovery using combinatorial chemistry the success rate has been disappointingly low. To date, only one de novo new chemical entity, sorafenib (Bayer) for treatment of renal cell carcinoma (2005) and hepatocellular carcinoma (2007) was reported in the public domain as resulting from this technology. Undoubtedly the chemical diversity of plants is much larger than any chemical library made by humans and thus the plant kingdom represents an enormous reservoir of pharmacologically valuable molecules just waiting to be discovered. Therefore many pharmaceutical companies have now renewed their interest in plant-derived compounds. More than 100 000 secondary metabolites have been discovered in higher plants but only half their structures have been elucidated. Plants are thus excellent organic chemists and constantly respond to environmental challenges by adjusting their capacity to produce secondary metabolites. Natural products have continued over the centuries to be significant sources of drugs and lead compounds [7]. Their dominant role is evident as over 50% of new anticancer drugs have been discovered in nature (Table 2). During the past 30 years 1355 new drug entities (NDEs) were introduced to the market; 27% were either natural products or were derived from natural products as semi-synthetic derivatives. In addition, 20% of the drugs were synthesized after the molecule was first discovered from natural resources. Table 2 shows the origin of the 458 NDEs representing the four major therapy groups with anti-infectives (antibacterial, antiviral, antifungal and antiparasitic), anticancer, anti-hypertensive or anti-inflammatory activities discovered between 1981 and 2010. It is remarkable that over ca: 70% of all antibacterial compounds and 64% of antiparasitic compounds for example were directly or indirectly derived from natural resources (Table 2). Natural sources

will undoubtedly continue to play a prominent role in the discovery of pharmaceuticals also in the future. The production of a secondary metabolite of interest, either a known molecule or a promising lead compound, for industrial use is often a big challenge. As discussed earlier these compounds accumulate in plants in small quantities and the production rates often vary from year to year. Some substances can only be isolated from extremely rare plants which is not ideal for sustainable production. The biotechnological production of high-value plant secondary metabolites therefore is a viable option as compared to isolation from intact native plants or total chemical synthesis. However, the biosynthetic pathways leading to their formation in plants are often long, with complex multiple steps catalyzed by various enzymes, and are still largely unknown at the enzyme and genetic level. The best characterized pathways after decades of intensive classical biochemical research are those of opium and terpenoid indole alkaloids. Plant molecular biology has developed enormously during the past decades but the full utilization of plant metabolic engineering is still lacking compared to microorganisms. However, new sequencing technologies, multigene transformation techniques and development of metabolomics tools have enabled the rapid elucidation of biosynthetic pathways in plants. Spectacular advances in plant genomics combined with metabolite profiling offer unprecedented possibilities to explore the extraordinary complexity of the plant biochemical capacity [3]. In the future, better advantage of plant biotechnology for improving production rates of known compounds will be taken and new tools for drug discovery will be developed. The examples discussed in subsequent sections of this article further highlight more recent developments in the field. PACLITAXEL Background The diterpene paclitaxel (Taxol®) is found in Taxus species (yew) which are small to medium-size evergreen trees. The trees are relatively slow growing, typically 10-30 m high and can live hundreds of years [8]. Historical use of Taxus other than medical applications encompasses also weapon- and furniture making as well as gardening. At least eight Taxus species are known [9]: T. baccata (European yew), T. brevifolia (Pacific yew), T. canadensis (Canadian yew), T. cuspidate (Japanese yew), T. chinensis (Chinese yew), T. wallichiana (Himalayan yew), T. floridana (Floridian yew), T. globosa (Mexican yew), and two hybrids Taxus x media (T. baccata x T. cuspidata) and Taxus x hunnewelliana (T. cuspidata x T. canadensis). Paclitaxel (Fig. 1) was discovered in the 1960’s from the inner bark of T. brevifolia and its complex structure was elucidated in 1971. Contents, however, is very low and typically less than 0.01% of the dry weight of the inner bark. When its medical importance

5642 Current Pharmaceutical Design, 2013, Vol. 19, No. 31

Table 2.

Rischer et al.

Number of new drug entities (NDEs) discovered during 1981–2010 belonging to the four most important therapy groups. The figures are modified according to [7] N

ND

NS

B

S

Total

N+ND+NS (%)

12

74

34

13

60

230

52

Antibacterial

10

64

1

0

23

109

69

Antifungal

0

3

0

1

25

29

10

Antiviral

0

2

31

12

8

78

42

Antiparasitic

2

5

2

0

4

14

64

Anticancer

9

25

17

17

30

100

51

Antihypertensive

0

2

34

0

41

77

47

Anti-inflammatory

0

13

0

1

37

51

26

Total

21

114

69

31

168

458

48

Antimicrobial

Abbreviations: N, natural product; ND, derived from natural product, semisynthetic product; NS, synthetic but pharmacophore is from a natural product; B, biological origin, usually a large protein or peptide either isolated from an organism or cell line or produced by biotechnological means in a host; S, derived through total chemical synthesis

was recognized in a screening program by the National Cancer Institute (Frederic, Maryland, USA) in the 1970’s very early estimations indicated that the demand for paclitaxel might exceed 300 kg per year, which would amount to 750 000 yew trees. Soon it became obvious that the yew trees would be endangered and investigators started to look for an alternative source of paclitaxel such as micro-organisms or Taxus endophytic fungi [10]; [11]. Despite strong postulations of endophytes being able to synthesize paclitaxel independently no conclusive confirmation has been obtained thus far [12]. Paclitaxel can be synthetized chemically but the method is far from being economically feasible. Paclitaxel is now obtained by semi-synthesis from 10-deacetylbaccatin III which is isolated from the needles of T. baccata. Another large-scale production system developed by the company Phyton Biotech uses cultivated cell cultures of Taxus. Similarly docetaxel (Taxotere®) (Fig. 1), a semi-synthetic taxane, is also synthesized from 10-deacetylbaccatin III. The sales of Taxol® from Bristol-Myers Squibb and Taxotere® from Sanofis Aventis amounted to more than  2.3 billion (data from the 2009 Annual Reports of the two companies). Due to expiration of the Taxol® patent in 2010 a number of generic products have been entering the market very recently. Pharmacology Microtubule-binding natural products, such as vincristine, vinblastine and paclitaxel play important roles in the war against cancer. The compounds exerting this mechanism of action have contributed to the substantial development of current cancer therapy. Paclitaxel’s mode of action was discovered in 1979 [13] but only in 1992 the compound entered clinical trials. Paclitaxel was then ap-

AcO O NH

O

proved by the FDA for the treatment of refractory metastatic ovarian cancer. Two years later the FDA approved it for metastatic breast cancer (refractory or insensitive to anthracyclines), non-small cell lung cancer and AIDS related Kaposi’s sarcoma [14]. Docetaxel was approved in 1996 for locally advanced or metastatic breast cancer or non-small cell lung cancer refractory to anthracycline-based chemotherapy. Together with paclitaxel it has become the most used anticancer compound for treatment of solid tumours in the breast, lung and ovaries. Peripheral neuropathy, one of the side effects of taxanes, appears less frequently and less severely with docetaxel [15]. The clinical use of the taxanes is limited mainly by drug resistance and toxicity. Resistance to taxanes involves a multifactorial mechanism and might include the overexpression of the membrane efflux pump P
gp, tubulin mutations and increased microtubule dynamics associated with altered microtubule-associated protein (MAP) expression [16]. The use of taxanes has been investigated also for treatment of stomach and upper GI tract adenocarcinoma, urothelial cancer and metastatic melanoma and myeloma [17] as well as other diseases such as Parkinson’s, Alzheimer’s and Multiple Sclerosis which also require microtubule stabilization and avoidance of cell proliferation and angiogenesis [18]; [19]. The cytoskeleton is composed of microtubules, actin microfilaments and intermediate filaments. Microtubules are hollow rods with an outer diameter of about 25 nm and an inner diameter of about 14 nm. Microtubules are composed of two heterodimer molecules, - and -tubulin [20], and they are highly dynamic structures, in continuous state of assembly/disassembly. At least six isotypes of -tubulin and seven isotypes of -tubulin have been identified.

AcO

OH

O

(CH3 )3 C—O

O

HO

OBz OAc

Paclitaxel Fig. (1). Chemical structures of paclitaxel (TaxolR) and docetaxel (TaxotereR).

OH

O

O

O OH

NH

O

O

O HO

OH

Docetaxel

OBz OAc

Plant Cells as Pharmaceutical Factories

Current Pharmaceutical Design, 2013, Vol. 19, No. 31

They show tissue-, cell- and tumour-specific patterns of expression as well as differences in drug binding [21]. They have thus an important role in controlling mitosis and the mitotic spindle. Therefore microtubules constitute a highly attractive target for anticancer drug design. An increasing number of antitumor drugs target microtubules and exhibit their anticancer activities by altering their polymerization dynamics, arresting mitosis, and thus inhibit cell proliferation and induce apoptosis. Although paclitaxel is targeted at microtubules its mode of action is completely different from that of vincristine which prevents tubuline polymerization. Low concentrations of paclitaxel (
0.05 M) stabilize microtubules by binding specifically to the  subunit of the tubulin heterodimers, which induces cell blockage in the mitosis phase and leads to cell death by apoptosis, especially in the proliferating tumour cells. However, high concentrations (5 M) stabilizes microtubules regardless of the cell cycle stage and inhibits cell progression to the S phase, thus inducing cell death by necrosis [22]. Since docetaxel has a higher affinity for the taxane-binding site than paclitaxel, its effect on cancer cells also differs from that of paclitaxel. For example, paclitaxel induces cell cycle arrest at the G2-M phase while docetaxel exerts its maximum cell-killing effect on cells in the S phase. It has been suggested that microtubules represent the single best cancer target identified to date and microtubule-targeted drugs undoubtedly will continue to be an important chemotherapeutic class

Table 3.

5643

of drugs [21]. Moreover, it is important to discover new targets and new sources of microtubule-binding natural products as well as novel formulations for improved cancer therapy. Biosynthesis of Paclitaxel Diterpenes, such as paclitaxel, are formed exclusively from geranylgeranyl diphosphate (GGPP) which is synthetized from three isopentenyl diphosphate (IPP) and one dimethyl diphosphate (DMAPP) molecules by the enzyme GGPP synthase. Both the cytosolic and plastidic pathways for IPP and DMAPP exist in plants and it is not entirely clear from which one paclitaxel is derived. Cusidó and co-workers [23] demonstrated that in cell cultures of T. baccata paclitaxel biosynthesis is blocked by fosmidomycin and mevinolin, well-known inhibitors of plastidic and cytosolic pathways, respectively. It has been postulated that 19 enzymes and 13 corresponding genes are involved in the biosynthesis of paclitaxel after the formation of GGPP [24]-[26] (Fig. 2). Once GGPP undergoes cyclization into taxadiene, eight hydroxylations, five acyl/aroyl transferases, one epoxidation, one aminomutase reactions, two CoA esterifications and a N-benzoylation are required to obtain paclitaxel (see Table 3). The first committed step in paclitaxel biosynthesis is the cyclization of GGPP to the taxadiene/taxoid skeleton by taxadiene synthase (TXS) (Fig. 2). Apparently this step is not rate-limiting

Known and postulated enzymes involved in paclitaxel biosynthesis and their corresponding known genes (modified from [26]) Biosynthetic Enzyme

Abbreviation

Gene Known

Publication

GGPPS

yes

[29]

TXS

yes

[190]

T5OH

yes

[191]

TAT

yes

[30]

Taxadiene-13-hydroxylase

T13OH

yes

[31]

Taxane-10-hydroxylase

T10OH

yes

[32]

Taxane-1-hydroxylase

T1OH

no

Taxane-2-hydroxylase

T2OH

yes

[192]

Taxane-7-hydroxylase

T7OH

yes

[193]

taxane-9- hydroxylase

T9OH

no

TBT

yes

Geranylgeranyl diphosphate synthase Taxadiene synthase Taxadiene-5-hydroxylase Taxadiene-5-ol-O-acetyl transferase

Taxane-2-O-benzoyl transferase C4,C20-epoxidase

no

Oxomutase

no

C9 oxidase

no

[33]

10-deacetylbaccatin III-10-O acetyltransferase

DBAT

yes

[34]

Phenylalanine aminomutase

PAM

yes

[194]

- phenylalanoyl-CoA Baccatin III: 3-amino, 13-

no BAPT

yes

Taxane 2’-hydroxylase

T2’OH

no

N-benzoyl transferase

DBTNBT

yes

[36]

phenylpropanoyltransferase

[37]

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Rischer et al.

OPP

9

GGPPS

6

7

9

TXS

11

12 1

3

5

4

3 1

20

OPP

OPP

T5αOH

OH

T13αOH OH HO

OH

HO

OH

OH

OH

TAT (T1βOH) T2αOH T7βOH (T9αOH)

OAc

HO OH

OH

OAc

HO

HO

T10βOH

OAc

OAc

TBT

HO

OH

HO

OH

OAc

HO OH

Epoxydase Oxomutase

OH

HO

OH

O

AcO

OH

C9 Oxydase O

HO

OBz

HO

NH2

HO

OBz OAc

Baccatin III

NH2

O

O

HO

OBz OAc

10-Deacetylbaccatin III

O

OH

DBAT O

HO

OBz OAc

OH

O

O

3' 2'

OH

OH

PAM

O-CoA

CoA transf. BAPT

β-Phenylalanine

AcO O NH

O

AcO

OH

O

DBTNBT HO

OH

NH2

O

O OBz OAc

O

AcO

OH

O

(T2’αOH) O

O OH

HO

OBz OAc

NH2

O

OH

O O

O HO

OBz OAc

Paclitaxel

Fig. (2). Schematic biosynthesis of paclitaxel starting from GGPP involves 19 enzymes and 13 corresponding genes (adopted from [26]). Abbreviations as indicated in Table 3. Hypothetical enzymes are marked in brackets.

although it is a slow reaction [27]-[28]. Taxadiene has not been found in cell suspension cultures of Taxus species. The next step is the hydroxylation at C5 and the migration of the double bond from 4 (5) to 4 (20). This reaction is cytochrome P450 catalysed, and also not rate-limiting [29]. The biosynthesis continues to the formation of taxadiene5
,10,13
-triol acetate catalyzed either by taxadiene-5
-ol-Oacetyl transferase (TAT) and taxadiene-10-hydroxylase (T10OH) or alternatively by only taxadiene- 13
-hydroxylase (T13
OH) [30]-[32]. The next steps involve several hydroxylations at C1, C2, C7 and C9 positions as well as oxidation of C9 and epoxidation at the C4C5 double bond (Fig. 2). The order of the individual steps is unclear [25].

The next important compound in the pathway is 10-deacetylbaccatin III (DABIII) which is currently used commercially as a starting molecule for the semi-synthetic production of paclitaxel. This enzymatic step is catalysed by taxane 2
-O-benzoyl transferase (TBT) [33]. DABIII is converted to baccatin III by 10deacetyl-baccatin III-10-O-acetyl transferase (DBAT) [34]. An essential step in the last part of paclitaxel biosynthesis is the esterification of the C13 hydroxyl group of baccatin III with the -phenylalanoyl CoA side chain. The side chain is derived from
phenylalanine by phenylalanine aminomutase (PAM) [35]. An unknown ester CoA ligase probably activates the compound so that it can bind to baccatin III. The enzyme involved in the conjugation of the -phenylalanoyl-CoA side chain to baccatin III is C-13-

Plant Cells as Pharmaceutical Factories

phenylpropanoyl-CoA transferase (BAPT), yielding the compound 3’-N-debenzoyl-2’-deoxytaxol [36]. Although the role of taxane 2’
-hydroxylase has not been confirmed it appears that N-benzoyl transferase (DBTNBT) [37] is responsible for the very last step of the biosynthetic pathway of paclitaxel. Wheeler and co-workers [38] have suggested that the taxane core is produced in plastids or in the cytosol whereas most of the hydroxylation steps occur in the endoplastic reticulum and are cytochrome P450-mediated events [38]. All transferases seem to be cytosolic. Obviously there must be considerable intracellular trafficicking of taxanes. Much insight has been obtained into the complex biosynthetic pathway of paclitaxel during the past two decades. However, the pathway has not been completely elucidated. In particular a more in depth understanding is needed on the regulation of the specific biosynthetic steps and corresponding gene function. Biotechnological Production of Paclitaxel During the past decades much effort has been devoted to the production of paclitaxel and its precursors in plant cell and tissue cultures. A biotechnological process has been developed through an empirical approach (Phyton Biotech) by optimizing and selecting a high-producing cell line. This approach is usually not sustainable since undifferentiated cells are often unstable and lose their ability to produce the desired compound in the longer term [3]. Cell cultures are reported to produce paclitaxel up to 0.2 mg per kg of the dry weight and about double the amount of the precursor baccatin III [39]. Classical methods have been also used to enhance the production of taxanes in cell cultures. These include e.g. two-stage growth and use of elicitors [40]. The well-established elicitor methyl jasmonate had a pronounced effect on taxane production [41]-[42]. The highest amount of paclitaxel reported so far is 2.71 mg per liter in immobilized cells of T. baccata [42]. An interesting approach would be to express the entire biosythetic pathway in microorganisms [43]. Extensive studies on various strategies for increasing biotechnological production of taxanes are discussed in recent reviews [44]-[45]. To be able to take full advantages of biotechnological production a more in depth understanding of paclitaxel biosynthesis is require at the genetic, enzymatic and regulation levels. Our current knowledge is still limited although a lot of studies on paclitaxel biosynthesis have been carried out for decades. Several hundreds of taxanes have been identified and they differ from each other in the side chain, hydroxylation patterns in the core or the side chain and substitutions with acyl or aryl groups [24]. TROPANE ALKALOIDS Background Through the ages humans have performed experiments using extracts made of plants and animals surrounding them. Some of the extracts have been poisonous, some hallucinogenic and some have possessed medicinal properties. Many of the current pharmaceuticals originate from these vehicles of murder, magic and medicine [46]. Of all plant species known, few are as closely connected to these ancient crafts as Atropa and Hyoscyamus, well-known plants producing tropane alkaloids. The family Solanaceae is considered as the home of tropane alkaloids. Although this family includes important food species, it also became well known for its toxic and medicinal counterparts. Since tropane alkaloids are chemically relatively simple structures and easy to extract, they were widely used in e.g. witchcraft and tribal ceremonies for their hallucinogenic properties. Nowadays, tropane alkaloids, such as atropine, hyoscyamine and scopolamine have an important role as medicinal compounds acting as parasympatolytes. Tropane alkaloids form a subclass of a diverse and impressive class of natural compounds called alkaloids, which are found mainly in plants. The number of tropane alkaloids known from natural sources exceeds 200 [47].

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There are several subfamilies of Solanaceae in which typical Solanaceae-specific tropane alkaloids, such as hyoscyamine, atropine and scopolamine have been found, including Anisodus, Anthocercis, Atropa, Brugmansia, Cyphanthera, Datura, Duboisia, Hyoscyamus, Latua, Mandragora, Physochlaina, Przewalskia, Scopolia, Solandra and Symonanthus [48]. The most important species for commercial cultivation are Datura, Duboisia, Atropa, Hyoscyamus, Scopolia and Brugmansia [49]. Besides Solanaceae, tropane alkaloids have also been found from other families such as Proteaceae, Euphorbiaceae, Rhizophoraceae, Cruciferae, Convolvulaceae and Erythroxylaceae. Pharmacology Muscarinic receptors mediate the effects of acetylcholine in target tissues. Drugs which arrest the muscarinic receptors are called anticholiergic drugs or parasympatholytes. Hyoscyamine and scopolamine are classical examples of anticholinergic drugs, affecting on parasympathetic nervous system [50]. The natural and pharmacologically active form of hyoscyamine is its (-)-form. However, hyoscyamine is very easily racemized leading to the formation of atropine, which is a mixture of (+) and (-)-forms. Atropine binds to all subclasses of muscarinic receptors (M1-M5) and acts as competing antagonist. Both hyoscyamine and atropine are used as skeletal muscle relaxants, against excessive hydrochloric acid formation in stomach and they are also used as antidotes for cholinesterase inhibitor poisoning. Atropine increases the heart rate and prevents bradycardia and the muscarinagonistic dilation of blood vessels. In the eye, atropine paralyzes muscles in the pupil leading to its dilation. It also decreases saliva formation. Scopolamine (hyoscine), in contrast to hyoscyamine, is a depressant for the central nervous system, and is used in the treatment of motion sickness and as a sedative before operations. Anisodamine, an intermediate in the biosynthesis of scopolamine from hyoscyamine, also has potential in the treatment of e.g. septic shock, respiratory disorders, migraine and eclampsia [51]. Biosynthesis of Tropane Alkaloids Tropane alkaloids, as other alkaloids, derive from amino acids. Putrescine, a key intermediate in alkaloid biosynthesis (Fig. 3), can be synthesized directly from ornithine in a reaction catalyzed by ornithine decarboxylase (ODC, EC 4.1.1.17), or formed indirectly from arginine in a reaction sequence initiated by arginine decarboxylase (ADC, EC 4.1.1.19). Putrescine metabolism is directed to higher polyamines in one branch and to alkaloids in another branch, in a reaction catalyzed by putrescine-N-methyltransferase (PMT, EC 2.1.1.53). N-methylputrescine, the product of the PMTcatalyzed reaction, is further converted to N-methylamino butanal by a diamine oxidase DAO, or to N-methylputrescine oxidase MPO (EC 1.4.3.6). This enzyme was first described from tobacco roots [52] and has been further purified and characterised [53]. Later, the gene encoding MPO was cloned [54]; [55]. The pyrrolidine structure of tropane alkaloids is biosynthesized from methyl-1pyrrolinium cation. This same structure serves as a backbone for nicotine biosynthesis in tobacco and further stands as a branching point in the common pathway of tropane and nicotine-related alkaloids. The rest of the tropane skeleton is built similarly in hyoscyamine-, scopolamine- and cocaine-forming species, i.e. via addition of acetoacetate into N-methyl-1-pyrrolinium and formation of 4-(1-methyl-2-pyrrolidinyl)-3-oxobutenoate as a precursor [56]. The final step before formation of tropane skeleton is the cyclization of 4-(1-methyl-2-pyrrolidinyl)-3-oxobutenoate into carboxytropinone. The role of tropinone in tropane alkaloid biosynthesis was controversial for a long time, until in 1990 Landgrebe and Leete showed it to be an intermediate in the biosynthesis of tropine [57]. Tropinone is further converted into tropine and pseudotropine by the reactions catalysed by two distinct enzymes, tropinone reductase I (TRI; [58]) and tropinone reductase II (TRII; [58]; [59]), respectively. Condensation of tropine and phenyl lactic acid leads

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Rischer et al.

AS Arginine

Ornithine

ADC

ODC Putrescine PMT

N-Methylputrescine MPO N-Methylaminobutanal N-Methylpyrrolidinyl hygrine N-Methylpyrrolinium

Nicotine

Hygrine

Cocaine

Cuscohygrine

CH 3 N

Norlittorine O H3C

Tropinone TRI

N

TRII

CH 3

Phenyllactic acid

CH 3 N

N O

TPAT OH

H

OH

3β−Tigloyloxytropane

O

CYP80F1

OH

H

Tropine

Pseudotropine

ATAT

Hyoscyamine aldehyde

Littorine

APAT TTAT

3α-Acetoxytropane

3α-Tigloyloxytropane

ADH

3β−Acetoxytropane

Other esters Calystegines

H3C N

O

O

Norhyoscyamine

OH

Hyoscyamine H6H CH3

CH3 N

N O

HO

O

O

H6H O

6β-Hydroxyhyoscyamine

OH

O

OH

Scopolamine

Fig. (3). Biosynthetic pathway of tropane alkaloids. One arrow may represent more than one step. Hypothetical steps are indicated with dashed lines. Abbreviations: ADC, arginine decarboxylase; ODC, ornithine decarboxylase; PMT, putrescine N-methyltransferase; MPO, N-methylputrescine oxidase; TRI, tropinone reductase I; TRII, tropinone reductase II; ATAT, acetyl-CoA:tropine acyl transferase; TTAT, tigloyl-CoA:tropine acyl transferase; TPAT, tigloylCoA:pseudotropine acyl transferase; APAT, acetyl-CoA:pseudotropine acyl transferase; H6H, hyoscyamine 6
-hydroxylase.

to formation of littorine, a positional isomer of hyoscyamine (Fig. 3). Li and co-workers exploited virus-induced gene silencing (VIGS) to discover the rearrangement of littorine to hyoscyamine [60]. It was suggested that the (R)-littorine is converted to hyoscyamine in a two-step process, first by formation of hyoscyamine aldehyde catalyzed by a cytochrome P450 enzyme, proceeding to the formation of hyoscyamine by an alcohol dehydrogenasecatalysed reaction. Recently, Al Balkhi et al. [61] reported on the basis of isotopic labelling and elicitation studies made with Datura

innoxia hairy roots that norlittorine and norhyoscyamine are conversion products of littorine and hyoscyamine, respectively. An important enzyme with dual functions in the tropane pathway is hyoscyamine-6-hydroxylase (H6H; EC 1.14.11.11), which catalyses the hydroxylation of hyoscyamine to 6-hydroxyhyoscyamine (anisodamine) and further epoxidation of the latter leading to scopolamine [62]; [63]. The hydroxylase activity of H6H has commonly been observed to be much higher than the epoxidase activity, leading to the formation of scopolamine. The primary site

Plant Cells as Pharmaceutical Factories

for hyoscyamine and scopolamine biosynthesis is the root tissue. Hyoscyamine is transported from roots to aerial parts for storage and for possible conversion to other products. The translocation mechanism is not completely known. Based on grafting experiments it has been suggested that specialized cells are not required [64], whereas other reports indicate that transport proceeds via xylem [65]. Several studies have indicated that root morphology is required for tropane alkaloid biosynthesis in vitro, and generally low levels of tropane alkaloids are obtained in undifferentiated cell and tissue cultures. When it comes to medicinal plants, the first breakthrough example of metabolic engineering was performed by Yun and co-workers [65]. They cloned the h6h gene from Hyoscyamus niger and introduced it into Atropa belladonna. As a result, the majority of the hyoscyamine was converted into scopolamine. Later, this finding has resulted in the engineering of this final step in tropane alkaloid pathway in various other Solanaceae species [66]-[69]. Among the genes known in the tropane alkaloid pathway, pmt and h6h have been considered to be rate-limitingenzyme genes. Overexpression of pmt in D. metel and H. muticus hairy roots cultures were shown to be crucial for increasing alkaloid production [70]. However, overexpression of pmt alone in A. belladonna plants and Duboisia hybrid root cultures was not sufficient in increasing the end product formation [71]; [72]. On the other hand, it can be concluded that overexpression of h6h increases scopolamine biosynthesis without exception in the reported tropane alkaloid-producing species. By overexpressing both of the rate-limitingenzyme genes, significantly higher scopolamine production rates have been achieved in hairy root cultures of A. belladonna [73] and H. niger [68]. Over nine-fold higher scopolamine concentration was achieved in transgenic hairy root lines of H. niger expressing both pmt and h6h compared to the wild type, and more than twice the amount compared to the highest production achieved with only h6h expression [68]. The best line produced up to 411 mg/l scopolamine, which is by far the highest production hitherto reported. Calystegines, which are synthesized in the other branch of the tropane pathway (Fig. 3), are polyhydroxynortropane alkaloids, possessing strong glycosidase inhibitory activity [74]. The compounds were originally found in Calystegia sepium (L.) [75], but they have also been detected in many Solanaceae and other families, even in some species which were not thought to possess the tropane alkaloid pathway, such as Solanum tuberosum (L.) [76]. Calystegines are formed from tropinone in a reaction catalysed by TRII [77]. Besides tropane alkaloids and calystegines, Solanaceous plants produce other tropine- and pseudotropine-derived esters, namely 3
- and 3-acetoxytropane [78], and 3
- and 3-tigloyloxytropane [79] (Fig. 3). These compounds are present in low amounts in Datura whole plants and roots [78]; [80]-[81] and cell suspension cultures [82], Atropa root and suspension cultures [80]; [83], and in Hyoscyamus hairy root cultures [80]; [84]-[85]. Biotechnology of Tropane Alkaloids Bioconversion of hyoscyamine to 6-hydroxyhyoscyamine and scopolamine was accomplished by expressing h6h from B. candida in Saccharomyces cerevisiae [86]. The specific activity obtained from H6H was similar or even higher than those reported from plant-derived preparations. Even though scopolamine production remained low, up to 83% of the added hyoscyamine was converted into 6-hydroxyhyoscyamine. Later, Kai et al. [87] expressed h6h from Anisodus acutangulus in E. coli and the majority of the added hyoscyamine was converted into scopolamine, whereas only a minor amount remained as 6-hydroxyhyoscyamine; this indicated much higher epoxidase than hydroxylase activity of heterologously expressed H6H. In our studies the uptake of hyoscyamine as well as bioconversion of hyoscyamine to scopolamine was followed with N. tabacum and Hyoscyamus muticus hairy roots carrying heterologous H6H

Current Pharmaceutical Design, 2013, Vol. 19, No. 31

5647

from H. niger [88]. Hyoscyamine was efficiently taken up by N. tabacum hairy roots; on average 95% of the hyoscyamine added to the medium was taken up, leading to 45% bioconversion of hyoscyamine to scopolamine. Interestingly, the produced scopolamine in N. tabacum hairy roots was efficiently secreted out of the cells. Up to 85% of the total scopolamine produced was secreted into the culture medium, compared to only 12% for H. muticus. This is noteworthy, since as observed in this study, tropane alkaloids are commonly retained in the root tissues in hairy roots of endogenous tropane alkaloid producers [66]; [67]. In a similar manner, in a study by Hallard et al. [89], tobacco cells overexpressing terpenoid indole alkaloid genes secreted the products into the medium, whereas in the endogenous producer Catharanthus roseus the alkaloids were stored inside the cell vacuoles. It was suggested that scopolamine-producing plants possess a regulatory mechanism for the cell-specific expression of H6H. Earlier studies indicate that in Nicotiana roots the promoter of the H. niger H6H gene is only expressed in meristem cells, whereas in Hyoscyamus expression is also demonstrated in the pericycle [90]. Further experiments performed with the immobilized cell cultures deriving from hairy roots described here showed that a marked secretion of scopolamine (74-81%) also took place in bioreactor cultivations of these undifferentiated cells [91]. The transport mechanism of scopolamine in Hyoscyamus and Nicotiana is not known, although it is possible that different transporter proteins are involved in the two species because scopolamine is an endogenous product in the former but a foreign metabolite in the latter. Earlier it has been reported that berberine is transported by an MDR-type transporter in Coptis japonica [92], for which berberine is an endogenous metabolite, whereas in Arabidopsis another type of transporter MATE is involved [93]. These results provide additional insights concerning the divergent role of H6H in different Solanaceae species, as varying hydroxylase and epoxidase activities of H6H have been reported in endogenous and non-hyoscyamine producers [94]; [95]. Subroto and co-workers [96] tested another approach for scopolamine production via bioconversion of hyoscyamine using coculture of hairy roots and shooty teratomas of A. belladonna and Duboisia hybrids. Although A. belladonna shooty teratomas were able to take up exogenously added hyoscyamine efficiently, about 72% of the hyoscyamine added to the medium was stored in shoots unconverted. The overall yield of scopolamine remained very low, only 0.12% (w/w). However, of the scopolamine produced by the shoots, up to 70% was found in the medium. By contrast, Duboisia hybrid shooty teratomas produced up to 38% (w/w) scopolamine and 37% (w/w) 6
-hydroxyhyoscyamine from the exogenously added hyoscyamine. None of the produced scopolamine was found in the culture medium in Duboisia hybrid, indicating species specific transport and accumulation processes even between species within the same family. Co-culture of hairy roots and shooty teratomas increased scopolamine and scopolamine:hyoscyamine ratios significantly: up to 0.84 mg/g DW scopolamine was produced in A. belladonna shooty teratomas [96]. This is comparable to that achieved by Hashimoto et al. [66] with genetically engineered A. belladonna hairy roots overexpressing H6H. Besides overexpressing pathway genes, product biosynthesis can be increased by more traditional ways e.g. by modulating the environmental cultivation conditions or by elicitation. In our recent studies, we observed that by altering the ploidy level of H. muticus plants and hairy roots, the ratio of hyoscyamine:scopolamine was changed efficiently towards scopolamine [97]. Commercial Exploitation of Tropane Alkaloids Atropa belladonna and Duboisia spp. are among the principal world sources of tropane alkaloids. Due to the difficult synthesis of these rather complex molecules, both alkaloids are still extracted from plants. Contemporary medicine exploits tons of atropine and

5648 Current Pharmaceutical Design, 2013, Vol. 19, No. 31

Table 4.

Rischer et al.

Yields of high scopolamine contents obtained from cultivated species and engineered hairy root cultures. Species

Tissue

Yield (% d.w.)

Reference

Brugmansia sanguinea

leaves

0.80

[49]

Duboisia leichhardtii x D. myoporoides

leaves

1.54

[195]

Duboisia hybrids

leaves

2.0-4.0

[98]

Hyoscyamus muticus

leaves

4.41

[196]

Atropa belladonna

hairy roots

0.30

[66]

Datura candida

hairy roots

0.57

[197]

Duboisia leichhardtii

hairy roots

1.80

[198]

Duboisia myoporoides

hairy roots

3.20

[199]

Hyoscyamus niger

hairy roots

4.11*

[68]

* estimated from the figure

scopolamine extracted from field-grown cultivars, while the constantly growing demand for tropane alkaloids pushes science and industry to develop new efficient ways for their production. Yields of scopolamine obtained from cultivated crops are listed in (Table 4). For example, over 400 tonnes of dried leaves are collected from Brugmansia sanguinea farms in Equador annually. In Australia, Duboisia hybrids are cultivated in large scale and approximately 10-15 tons of fresh leaves per hectare are collected per harvest three times a year for global supply to the pharmaceutical industry [98]. Today, one of the leading producers of scopolamine is Boehringer Ingelheim [99]. Duboisia plantations located in Australia provide the leaf material, which is then dried and shipped to Germany for scopolamine extraction. After isolation and purification, scopolamine is chemically converted to the active ingredient scopolamine butyl bromide (Buscopan®). It is manufactured in ca. 21 tons annually [98]. Annual global production of scopolamine is 600 kg, with the sales over 77 million $, for atropine 1000 kg (67 million $) and for hyoscyamine 140 kg (67 million $) [98]. So far, there is no synthetic process in sight which could compete with the natural sources for tropane alkaloid supply. Advanced biotechnological methods for alternative and sustainable production platforms for tropane alkaloids have shown their potential, especially those consisting of genetically engineered hairy root cultivations (Table 4), opening possibilities for optimized and controlled production as well as consistent yields.

[101]. Over the years several syntheses have been reported for these substances, but we are still lacking a commercially feasible total chemical synthesis. Thus a biotechnological approach for the production of these highly valuable benzylisoquinoline alkaloids has attracted considerable interest.

MORPHINE ALKALOIDS Background The opium poppy, Papaver somniferum L. (Papaveraceae), is an annual herb, up to 1-1.5 meters in height, and the source of the drug opium which contains a large number of isoquinoline alkaloids, more specifically benzylisoquinoline alkaloids. These alkaloids are also found in other Papaveraceae plants, in the genus Eschscholzia (Californian poppy; Eschscholzia californica Cham.) and in the genus Papaver (P. bracteatum (Lindl.) [100]. Opium is the inspissated latex from unripe fruit capsules of opium poppy. The production of opium is very laborious and can therefore be practised only in countries with cheap labour. The alkaloids are extracted either from crude opium or from the opium poppy straw to produce a concentrate of poppy straw (CPS), which is sold as a narcotic raw material or refined into active pharmaceutical ingredients. Total chemical synthesis of codeine and morphine was achieved by Gates and coworkers in 1952 and 1956, respectively

Biosynthesis of Morphine Alkaloids The biosynthesis of morphine occurs in all major plant organs of P. somniferum starting within the first seven days after seed germination. The biosynthesis continues throughout the lifecycle of this annual plant, with the highest biosynthetic activity taking place in the capsule after petal fall [103]. Morphine biosynthesis is well characterized at the enzymatic level, especially the first part to Sreticuline which is shared by the pathway leading to berberine. The biosynthesis of berberine was the first example of a plant secondary metabolite which was completely characterized at the enzyme level (Zenk and coworkers in the early 1990s). Today, most of the encoding genes in the benzylisoquinoline alkaloid pathway have been cloned as a composite work done on several species and by different research teams (Fig. 4). Benzylisoquinoline alkaloid biosynthesis initiates from tyrosine, with tyrosine decarboxylase (TYDC) as the first enzyme in the pathway. It is encoded by a small gene family composed of about 15 members. The TyDC gene family is divided into two subfami-

Pharmacology Four of the opium alkaloids, papaverine, morphine, codeine and noscapine have attained wide medicinal use and morphine can be considered as the most well-known drug in the world used to control moderately severe to severe pain. Morphine belongs to the group of opioids that work at opioid receptors. Morphine is a Mu () receptor agonist. Mu receptors are found primarily in the brainstem and medial thalamus, and are responsible for supraspinal analgesia, respiratory depression, euphoria, sedation, decreased gastrointestinal motility, and physical dependence. Thus morphine exerts its principal pharmacological effect on the central nervous system and gastrointestinal tract. Its primary actions of therapeutic value are analgesia and sedation. Morphine is a rapid-acting narcotic, and it is known to bind very strongly to the -opioid receptors, and for this reason it often has a high incidence of euphoria/dysphoria, respiratory depression, sedation, pruritus, tolerance, and physical and psychological dependence. Morphine is also characterized as a relatively long-acting opioid and the elimination half-life is approximately two hours. The effects of morphine can be countered with opioid antagonists such as naloxone and naltrexone [102].

Plant Cells as Pharmaceutical Factories

Current Pharmaceutical Design, 2013, Vol. 19, No. 31

COOH

HO

NH 2

HO

HO TYDC

5649

HO NH 2

HO

NCS

NH

HO

H

Dopamine +

L-Dopa

CHO HO HO 4-Hydroxyphenylacetaldehyd

(S)-Norcoclaurine 6OMT CNMT CYP80B1 4'OMT H3CO

H3CO N

HO

NCH3

HO

H OH

OC H

H

HO

H3CO

3

NCH3

HO

H

HO

BBE

(S)-Scoulerine

H3CO

H3CO

(S)-Reticuline

(R)-Reticuline STS

H3CO

H3CO

H3CO

HO

HO

HO

H H3CO H3COOC

SAT

N C H3

H3CO

H

HO

SalR

N C H3

H

H

H3CO

H

O Salutaridine

Salutaridinol

Salutaridinol-7-O-acetate

N C H3

spont.

HO

H3CO

H3CO

O

O

O T6ODM H

N C H3

H

N C H3

CODM H H3CO

H3CO

O

N C H3

Neopinone

Oripavine

Thebaine

T6ODM spont. H3CO

H3CO

O

HO

O H

H

O

N C H3

O

COR H

H

N C H3

O

CODM

HO Codeinone

HO

H

H

HO Codeine

N C H3

COR H

H

N C H3

O Morphine

Morphinone

Fig. (4). Biosynthetic pathway of benzylisoquinoline alkaloids in Papaver somniferum. Abbreviations: tyrosine decarboxylase (TYDC), norcoclaurine synthase (NCS), 6-O-methyltransferase (6OMT), N-methyltransferase (CNMT), (S)-N-methycoclaurine 3´-hydroxylase (CYP80B1), 4-O-methyltransferase (4OMT), berberine bridge enzyme (BBE), salutaridine synthase (STS), salutaridine reductase (SalR), salutaridine 7-O-acetyltransferase (SAT), thebaine 6-Odemethylase (T6ODM). codeine O-demethylase (CODM).

lies, based on sequence identity, represented by TyDC1 and TyDC2 [104], [105]. TyDC1-like subgroup is expressed abundantly in roots and TyDC2-like subgroup in roots and stem. The two subgroups also show differences in developmental, tissue-specific and inducible expression patterns [106]. The first committed step of the benzylisoquinoline pathway is catalyzed by the enzyme norcoclaurine synthase (NCS, EC 4.2.1.78), which condenses dopamine and 4hydroxyphenylacetaldehyde to give (S)-norcoclaurine. The enzyme has been isolated, purified and characterized [107] [108] and the encoding NCS was finally isolated from a meadow rue (Thalictrum flavum ssp. glaucum) cell suspension culture cDNA library. The

enzyme was shown to have highest sequence homology to the pathogenesis-related (PR)-10 and Bet v 1 protein families but the biochemical context of this homology is still unclear. However, the enzyme contains a putative 19 amino acid signal peptide, suggesting that it is associated with a subcellular compartment other than the cytosol, e.g. endoplasmic reticulum. The recombinant enzyme was overexpressed E. coli and the kinetics of the enzymatic reaction have now been evaluated [109], [110]. (S)-Coclaurine is produced from (S)-norcoclaurine by a 6-Omethyltransferase (6OMT; [111], and converted to (S)-Nmethylcoclaurine by a N-methyltransferase (CNMT; [112]. 4-O-

5650 Current Pharmaceutical Design, 2013, Vol. 19, No. 31

methyltransferase (4OMT; [111] in turn forms S-reticuline from (S)-3-hydroxy-N-methylcoclaurine. Facchini and Park [113] isolated molecular clones for 6OMT, 4OMT and CNMT from opium poppy that were found to have high homology with those of the Japanese goldthread (Coptis japonica). By introducing tyrosine/dopa decarboxylase (TYDC), (S)-N-methylcoclaurine-3´hydroxylase (CYP80B1), berberine bridge enzyme, (BBE), (7S)salutaridinol 7-O-acetyltransferase (SAT), and codeinone reductase (COR) together with 6OMT, 4OMT and CNMT to opium poppy they observed that transcript levels generally increased in developing seedlings, were high in stems and flower buds and variable in roots and leaves of mature plants. The transcript accumulation was also observed to respond on elicitation, except in the case of COR. The cytochrome P-450-dependent mono-oxygenase (S)-Nmethylcoclaurine-3-hydroxylase (CYP80B1, CYP80B3) lies on the pathway to the benzylisoquinoline alkaloid branch point intermediate (S)-reticuline. It hydroxylates (S)-N-methylcoclaurine to form (S)-3-hydroxy-N-methylcoclaurine [114], [115]. The overexpression of cyp80b3 cDNA in opium poppy resulted in up to 450% increase in the amount of total alkaloid in latex, indicating that this is one of the key regulation steps in the biosynthesis of morphine. This carries great potential for improving the opium poppy elite lines for pharmaceutical interest [116]. S-reticuline serves as a branch-point intermediate in the biosynthesis of numerous isoquinoline alkaloids. The berberine bridge enzyme (BBE, EC.1.5.3.9) represents a key branch point in the biosynthesis of benzylisoquinoline alkaloids. BBE catalyzes the conversion of S-reticuline to S-scoulerine, which represents the first step in the pathway leading to the antimicrobial alkaloids sanguirine and berberine. Several genomic clones (e.g. bb1, bb2, bb3) are isolated from P. somniferum and Eschscholzia californica [117], [118], [119]. By blocking the BBE, some groups have aimed at higher accumulation of morphine. Park and co-workers [120] introduced antisense constructs of the genes encoding BBE and Nmethylcoclaurine-3´-hydroxylase (CYP80B1) separately into Californian poppy cell culture. The results were similar in both transgenic lines. About 50 % of the transgenic cell cultures showed reduced accumulation of benzophenantridine alkaloids when compared to cell lines transformed with a marker gene. Both antisense suppressions caused an overall reduction in benzophenanthridine alkaloid accumulation but they did not cause accumulation of pathway intermediates. This was suggested to involve feedback inhibition of early biosynthetic enzymes by one or more alkaloid intermediates, a putative degradation of alkaloid intermediates, or possibly that benzylisoquinoline alkaloid biosynthetic enzymes operate as part of a metabolon or metabolic channel. The growth rate of the transgenic lines was also decreased. This might have resulted from the low-level accumulation of cytotoxic pathway intermediates. The transgenic lines also contained large cellular pools of several amino acids. The relative abundance of tyrosine was only twofold higher in antisense-suppressed cells than in controls. However, the elevated tyrosine pool cannot account for the decrease in the accumulation of tyrosine-derived benzylisoquinoline alkaloids in transgenic cell lines. The team concluded that the proposed model suggesting that plant cells can be engineered to accumulate valuable benzylisoquinoline alkaloids using an mRNA-mediated approach [121] does not appear to be feasible in some cases because of the complex and poorly understood overall metabolic regulation of the pathway. Especially better understanding of the subcellular localization of the biosynthetic enzymes, the trafficking of intermediates and the mechanisms that control flux is essential for rational design of metabolic engineering of cell cultures. Salutaridinol 7-O-acetyltransferase (SAT, EC 2.3.1.150) catalyzes the conversion of salutaridinol to salutaridinol-7-O-acetate, the immediate precursor of thebaine along the morphine biosynthetic pathway [100]. A cDNA corresponding to the internal amino acid sequences of the native enzyme was isolated and a genomic

Rischer et al.

DNA blot analysis indicated that there is probably a single copy of this gene in the genome of Papaver somniferum. In addition, the gene transcript was detected in extracts from Papaver orientale and Papaver bracteatum [122]. Kempe et al. [123] introduced an RNAi construct designed to reduce transcript levels of SAT into opium poppy and observed an accumulation of intermediates (salutaridine and salutaridinol) up to 2- to 56-fold. The morphine levels remained constant but amounts of thebaine and codeine in latex were lower. On the other hand over-expression of SAT resulted in an increase in capsule morphine, codeine and thebaine as reported by Allen and co-workers [124]. Codeinone reductase (COR) catalyzes the NADPH-dependent reduction of codeinone to codeine (Fig. 4). This enzyme is highly substrate specific and stereoselective. Unterlinner et al. [103] isolated and characterized four full-length and two partial cDNAs encoding codeinone reductase isoforms. These represent six alleles from a gene family that may have at least 10 members. The four isoforms have a high amino acid sequence identity (95-96%). These codeinone reductase isoforms are 53% identical to 6´-deoxychalcone synthase of soybean, suggesting an evolutionary link between phenylpropanoid and alkaloid biosynthesis. Larkin et al. [125] successfully over-expressed COR in opium poppy resulting in even up to 10-fold increased Cor transcript levels in transgenic leaves. Furthermore, the morphinan alkaloid contents were between 15% and 30% higher than in control non-transgenic samples. No other significant changes in alkaloid profiles or quantities were observed in leaf, roots, pollen or seed. The demethylation of thebaine and codeine was thought to be catalyzed by cytochrome P450-dependent enzymes [122]. This hypothesis was proved to be incorrect by Hagel and Facchini [126]. They isolated thebaine 6-O-demethylase (T6ODM) and codeine Odemethylase (CODM) by functional genomics. These non-heme dioxygenases were demonstrated to catalyze O-demethylation, and by virus-induced gene silencing (VIGS) of T6ODM and CODM the metabolism at thebaine and codeine was blocked. The authors postulated that with these genes the production of codeine and morphine in scalable microbial systems might be feasible, thus offering an alternative to conventional systems with respect to production costs and the regulation of controlled substances. By using VIGS, Vijekoon and Facchini [127] investigated the regulation the final six steps in morphine biosynthesis, including the steps of salutaridine synthase (STS), salutaridine reductase (SalR), salutaridine 7-O-acetyltransferase (SAT), thebaine 6-Odemethylase (T6ODM), codeinone reductase (COR), and codeine O-demethylase (CODM). Silencing and reduced levels of STS, SalR, T6ODM and CODM correlated with lower morphine levels and substantial increase in the accumulation of reticuline, salutaridine, thebaine and codeine, respectively. The silencing of genes encoding SalAT and COR resulted in higher levels of salutaridine and reticuline, which are not the actual substrates for these enzymes. These observations confirmed the function of some of the previously characterized enzymes and provided some further insight into the biochemical regulation and complexity of the morphine pathway. Genes encoding regulatory factors isolated from Arabidopsis, soybean and corn have been screened to identify those that modulate the expression of genes encoding for enzymes involved in the biosynthesis of morphinan alkaloids in opium poppy (Papaver somniferum) and benzophenanthridine alkaloids in California poppy (Eschscholzia californica). In opium poppy, the overexpression of selected regulatory factors increased the levels of PsCOR (codeinone reductase), Ps4'OMT (S-adenosyl-l-methionine: 3'-hydroxy-N-methylcoclaurine 4'-O-methyltransferase) and Ps6OMT [(R,S)-norcoclaurine 6-O-methyltransferase] transcripts by 10- to more than 100-fold. These transcriptional activations translated into an enhancement of alkaloid production in opium poppy of up to at least 10-fold. In California poppy, the transactiva-

Plant Cells as Pharmaceutical Factories

tion effect of regulatory factor WRKY1 resulted in an increase of up to 60-fold in the level of EcCYP80B1 [(S)-N-methylcoclaurine 3'-hydroxylase] and EcBBE (berberine bridge enzyme) transcripts. As a result, the accumulations of selected alkaloid intermediates were enhanced up to 30-fold. The transactivation effects of other regulatory factors led to the accumulation of the same intermediates. These regulatory factors also led to the production of new alkaloids in California poppy callus culture. The average weight percentage of total alkaloid in the dry capsules of the control lines (i.e. regenerated non-transgenic wild-type) was 0.04%, with an average ratio of morphine : codeine : thebaine of 85 : 10 : 5. The notable transgenes (ERX, HMGB5 and Myblike) increased the total weight percentage of alkaloids to between 0.16% (fourfold increase relative to the control) and 0.21% (fivefold increase), with substantial increases in thebaine yield representing 74% and 81% of the morphine–codeine–thebaine total, respectively. This means that these transgenes changed the thebaine content from the wild-type level of 0.002% to between 0.12% (60-fold) and 0.17% (85-fold) [128]. Biotechnological Production of Morphine Alkaloids The option to utilize plant cell cultures for production of morphine-type alkaloids has been of interest since the early 1970s. Misawa [129] reviewed the works carried out during 1970s and 1980s and at that time morphogenetic differentiation was found to be a prerequisite for production of thebaine by cell cultures. However, Hsu and Park [130] observed that callus obtained from hypocotyls of P. somniferum contained thebaine and a low level of codeine. In addition, codeine (0.016% of DW) and thebaine (0.004% of DW) but no morphine have been detected in rhizogenous but not in embryogenic callus of P. somniferum album [131]. Remarkably, suspension cultures of P. somniferum accumulated not only codeine (3 mg/g DW -> 0.3% of dw) but also morphine (2.5 mg/g DW -> 0.25% of dw) after removal of the hormones from the culture medium. In general, improved alkaloid production was correlated with overall slower growth rate [132]. Alkhimova et al. [133] also found morphine from the de-differantiated callus initiated from mature seeds of P. somniferum. Total alkaloid content of this dedifferentiated P. somniferum callus line was 5.4±0.32 mg/g DW, of which 70% was morphine (3.8 mg/g DW -> 0.38% of DW). Hairy root induction of P. somniferum (var. album) by A. rhizogenes (LBA 9402) was published in 2004 [134]. Total alkaloid content in hairy roots was slightly higher than in the control (0.46% DW >< untransformed roots 0.32% DW). However, these hairy roots accumulated three times more codeine (0.18% DW) than intact roots (0.05% DW) and sanguinarine was only detected in transgenic hairy roots (0.02% DW). The morphine levels were similar in hairy roots and in untransformed control roots (0.26% and 0.27% DW, respectively). In addition, it was shown that the liquid culture medium contained morphine (0.26% DW) and sanguinarine (0.014% DW). On the other hand, Facchini and Bird [135] stated that only sanguinarine has been found at significant levels from poppy cell cultures. Moreover, even production of sanguinarine is not constitutive but is typically induced only after treatment with a fungusderived elicitor. The absence of morphine, codeine and papaverine in de-differantiated opium poppy cell cultures suggests that benzylisoquinoline alkaloid biosynthesis is developmentally regulated and requires the differentiation of specific tissues. A requirement of specialized laticifer cells is suggested before morphine and codeine can accumulate. Farrow and co-workers [200] carried out a very comprehensive transcriptomic and metabolite profiling study of cell cultures originating from 18 plant species and four families (Papaveraceae, Berberidaceae, Ranunculaceae, Menispermaceae). They constructed EST libraries, each containing approximately 3500 unigenes per species for a total of 58,787 unigenes. Sequence resources were analyzed in the context of the targeted metabolite profiles obtained for each cell culture, making the prediction of

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enzyme functions possible. Their results suggested that speciesspecific metabolite accumulation was influenced by the presence and absence of the key enzymes and by the substrate specificities of these enzymes. This data can be seen as an important tool for the discovery of benzylisoquinoline alkaloid pathway genes. Elicitation is one important way of boosting secondary metabolite accumulation, as was mentioned earlier. Transient and rapid accumulation of the berberine bridge enzyme (BBE) mRNA has been observed with fungal elicitor and methyl jasmonate (MeJA). However, sanguinarine was found only in cell cultures of Papaver somniferum treated with fungal elicitor (Botrytis, Pythium) but not in MeJA-elicitated [117] cultures. The elicitation of P. somniferum cell culture by the fungal mycelium of Botrytis sp. led to 26-fold increased amounts of sanguinarine (29% of DW) when compared to non-elicitated culture (reviewed by Misawa [129]). Bobak et al. [136] also reported a 30-fold increase in sanguinarine contents after treatment of elicitor prepared from Botrytis cinerea. Archambault et al. [137] in turn reported that the elicitation of P. somniferum cell suspension culture by chitin resulted in sanguinarine production at 5% of DW (final concentration 385 mg/l and productivity 26 mg/l per day throughout the culture). They also optimized the nutrient solution and feeding system needed for best sanguinarine production [138]. The production level in bioreactors was 3- to 14-times lower than in shake flask cultures [137]. Addition of MeJA to the culture medium of opium poppy cells induced accumulation of cyp80b1 and bbe1, but not of cor1 (gene encoding for codeinone reductase) transcripts. This is consistent with the fact that sanguinarine accumulation, but not morphine, can be induced in opium poppy cell suspension culture by MeJA elicitation [115]. TERPENOID INDOLE ALKALOIDS Background Terpenoid indole alkaloids (TIAs) comprise more than 3000 known structures, which makes them one of the major classes of alkaloids in plants [139]. Their distribution is restricted to only a few plant families. The main producers are species of the Apocynaceae, Loganiaceae and Rubiaceae. Although the chemical structures can be quite complex, a tryptamine portion i.e. the indole part is generally easily discerned. The joined C9 or C10 fragment is of terpenoid origin [140] and further classified according to structural rearrangements, i.e. Corynanthe type, Aspidosperma type and Iboga type. Pharmacology Preparations including TIAs have been used since ancient times. Two well-known examples are Rauwolfia serpentina (Apocynaceae), the use of which goes back thousands of years in India as a snake-bite remedy and Tabernanthe iboga (Apocynaceae), which has long been used for ceremonial rites by indigenous people in tropical Africa due to its psychoactive ingredients. In more recent times individual compounds have been isolated, starting with the purification of strychnine from Strychnos ignatii (Loganiaceae) [141]. Although this powerful neurotoxin is nowadays mainly employed for pest control, there were even pharmaceutical claims such as increasing appetite, memory enhancement and toning skeletal musculature [142]. Really successful TIA drugs isolated from plants were only introduced in the 1950s. Especially Rauwolfia proved a very rich source of medically important compounds. The anti-hypertensive and anti-psychotic drug reserpine was first introduced in 1954 by the CIBA Pharmaceutical Company, followed by the antiarrhythmic agent ajmaline and the anti-hypertonic compound ajmalicine. The latter is also found from Madagascar periwinkle (Catharanthus roseus). This pantropic shrub, that is widely grown as an ornamental species, came to fame almost by coincidence [143]. Due to its traditional medical use as an anti-diabetic agent in Jamaica, pure compounds isolated from the plant ended up in drug tests.

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Surprisingly it was not the anti-diabetes but the anti-cancer activity that led to the isolation and characterisation of two new dimeric TIAs: vinblastine (vincaleucoblastine, VLB, Velban®) [144] and vincristine (VCR, Oncovin®) [145]. Vinblastine and vincristine were approved for clinical use in 1961 and 1963, respectively. Initially it was observed in animal studies that injection of the compounds decreased the cell count of white blood cells in bone marrow. This led further to the establishment of these natural compounds and the related anhydrovinblastine [146] as well as semisynthetic agents including vindesine, vinorelbine and vinflunine as potent chemotherapeutic agents for the treatment of haematological and lymphatic neoplasms, acute leukaemia, Hodgkin’s disease, non-Hodgkin lymphomas as well as solid tumours such as breast, testicular and non-small cell lung cancers. The dimeric alkaloids are mitotic inhibitors through their ability to prevent the polymerization of tubulin into microtubules in dividing cells and thereby inducing programmed cell death in cancerous cells [147]. The use of TIAs is restricted by drug resistance mediated by P-gp and side effects including myelosuppression and neurotoxicity. The development of new halogenated analogs such vinflunine (Javlor®) has led to less serious side effects and drug resistance. Even though monoclonal antibodies now dominate the global cancer market in terms of market share, treatment is superior in combination with chemotherapy. A good example is Rituximab (trade name MabThera with sales of $5.1bn in 2010), which is usually administered as R-CHOP (i.e. in combination with cyclophosphamide, hydroxydaunorubicin, oncovin i.e. vincristine and prednisone) for the treatment of B-cell lymphoma. Similarly, Velban® i.e. vinblastine is still in clinical use for the treatment of advanced Hodgkin’s disease, lymphocytic lymphoma, histiocytic lymphoma, advanced testicular cancer, Kaposi’s sarcoma and various cancers that have stopped responding to previous chemotherapy. Biosynthesis of Catharanthus Alkaloids Altogether more than 120 TIAs have been isolated from Catharanthus during the past decades [148] and the numbers are still rising even in recent years [149]; [150]. As indicated by their name, TIAs are fusion products of precursors derived from two pathways: tryptamine from the shikimate pathway and secologanin from the non-mevalonate (MEP) pathway [151] (Fig. 5). Chorismate supplied by the plastid-localised shikimate pathway constitutes the first building block for tryptophane biosynthesis. The pathway to this amino acid is well described and involves reactions catalyzed by anthranilate synthase (AS), phosophoribosylanthranilate synthase, phosophoribosylanthranilate isomerase, indole3-glycerolphosphate synthase and tryptophan synthase. Finally cytosolic tryptophane decarboxylase (TDC) converts tryptophan to tryptamine. In plants terpenoids are biosynthetically derived from two precursors: isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP). These in turn can in principal arise both from the cytosolic mevalonate (MVA) pathway or the plastidic nonmevalonate (MEP) pathway (also called DOXP pathway). It has been established that the major channel in Catharanthus is the MEP pathway, although some crosstalk between the pathways appears to exist [152]. All the seven steps leading to IPP/DMAPP have been identified, including the corresponding enzymes and encoding nuclear genes [153]. The first reaction catalyzed by deoxyxylulose 5phosphate synthase (DXS) is the condensation of (hydroxyethyl)thiamin derived from pyruvate with glyceraldehyde 3phosphate to build deoxyxylulose 5-phosphate (DXP). 1-deoxy-Dxylulose 5-phosphate reductoisomerase (DXR) initiates intramolecular rearrangement and reduction to form 2-C-methyl-Derythritol 4-phosphate (MEP). Four successive steps involving 2-Cmethyl-D-erythritol 4-phosphate cytidyltransferase (MCT), 4(cytidine 5’-diphospho)-2-C-methyl-D-erythritol kinase (CMK), 2C-methyl-D-erythritol 2,4-cyclodiphosphate synthase (MDS) and

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(E)-4-hydroxy-3-methylbut-2-enyl diphosphate synthase (HDS) lead to the formation of hydroxymethylbutenyl diphosphate (HMBPP) that is finally converted by (E)-4-hydroxy-3-methylbut2-enyl diphosphate reductase HDR to a mixture of IPP and DMAPP (Fig. 5). The monoterpene-secoiridoid pathway commences by the formation of geranylpyrophosphate and then geraniol through the action of the corresponding synthases GPPS and GES, respectively. The cytochrome P450 mono-oxygenase geraniol-10-hydroxylase (G10H) has been isolated from Catharanthus and transfers geraniol jointly with a NADPH:cytochrome P450 reductase (CPR) [154] to 10-hydroxygeraniol. 10-hydroxygeraniol undergoes a reversible dehydrogenation to produce 10-oxogeraniol which is oxidized further to give 10-oxogeranial [155]. The following sequence of reactions leading eventually to loganic acid is mainly postulated on the basis of retrosynthetic considerations and labeling experiments [151]. Accordingly it is postulated that 10-oxogeranial is first cyclized and reduced to form iridodial, which in turn undergoes hydroxylation, dehydrogenation and again a cyclisation. Loganic acid would then be synthesized from the resulting iridotrial by sequential hydroxylation, glucosylation and hydroxylation. Two further reactions, i.e. a methylation and an acyclisation catalyzed by S-adenosyl-L-methionine:loganic acid methyltransferase (LAMT) [156] and (SLS) [157], respectiviely, lead to the formation of secologanin. The condensation of tryptamine and secologanin catalysed by strictosidine synthase (STR) results in the formation of strictosidine, the common precursor for TIAs [158] which is then converted into cathenamine [151]. From here multiple routes most of which are only partly elucidated lead to the great diversity of alkaloids. A notable exception is the formation of ajmalicine, which has been demonstrated with enzyme preparations [159]. Vindoline and catharanthine constitute the building blocks for the formation of the pharmaceutically important bisindoles. Vindoline biosynthesis starts from the hydroxylation of tabersonine catalysed by the cytochrome P450-dependent tabersonine 16-hydroxylase (T16H) [160]. Then 16-hydroxytabersonine is O-methylated by 16-hydroxytabersonine 16-O-methyltransferase (OMT) [161]. The enzyme performing the hydratation of the 2,3 double bond of 16methoxytabersonine is unknown and the N-methyltransferase generating desacetoxyvindoline has only been partly characterised [162]. Desacetoxyvindoline-4-hydroxylase (D4H) that has been characterised and the corresponding gene cloned [163] provides the substrate for deacetylvindoline-4-O-acetyltransferase (DAT) to eventually form vindoline [164]. Ultimately peroxidase-like
-3’,4’-anhydrovinblastine synthase (AVLBS) joins vindoline and catharanthine in a coupling reaction [165].
-3’,4’-Anhydrovinblastine constitutes a ‘dimeric’ monoterpenoid indole alkaloid and is presumably the precursor of the anticancer drugs VLB and VCR. In addition to the pathway elucidation at the level of chemical reactions and involved enzymes including their transcriptional regulation [166], much progress has been made concerning the spatial and temporal regulation of the TIA biosynthesis in intact plants. Especially in situ hybridisation and immunocytochemical localisation have helped to reveal compartmentalization and transportation of the highly complex and dynamic TIA metabolism in C. roseus [167]. In the aerial part transcripts for enzymes catalyzing early pathway steps (including DXS, DXR, MECS and G10H) were localised to the internal phloem-associated parenchyma [168], whereas downstream enzymes such as SLS were restricted to epidermal cells or more specifically to laticifers [169]. In the roots, TDC enzymatic activity was not found in the maturation zone but was located in the cytosol and in the apoplastic region of the meristematic cells [170]. At the subcellular level parts of the TIA biosynthesis involve chloroplasts, cytosol, vacuoles and endoplasmatic

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MEP pathway

G3P

Pyr DXR MCT CMK MDS HDS HDR

IPP/DMAPP

Shikimate pathway

GPPS GES G10H/CPR 10HGO LAMT SLS

Chorismate

AS TDC

H NH2

N

H

STR

OGlc H

H Tryptamine

O CH 3OO C Secologanin NH

N

H

H

OGlc H O

CH 3OO C Strictosidine

SGD

N

N H

H H O CH 3OO C Cathenamine

N H

N

N H

N

H

H N

H O

H

CH 3OOC

MAT

COOC H 3

H

T16H OMT NMT D4H DAT

H

AVLBS

N

O N

N

COOC H 3

Catharanthine

Tabersonine

Ajmalicine

N

H

N H

H

O C OC H3

H

C OO C H 3

19-O-Acetylhörhammericine

N

H

CH 3O

H OH COOC H 3 N

OCOCH 3 N H OH CH 3 COOCH 3 Vindoline CH 3O

H

OCOC H 3 N OH H CH 3 COOC H 3 Vinblastine

Fig. (5). Biosynthetic pathway of TIAs in Catharanthus roseus. Abbreviations: AS (anthranilate synthase), CMK (4-diphosphocytidyl-2C-methyl-D-erythrol kinase), CMS (4-diphosphocytidyl-2C-methyl-D-erythrol-4phosphate synthase), CPR (cytochrome P450 reductase), DAT (deacetylvindoline 4-O-acetyltransferase), D4H (desacetoxyvindoline 4-hydroxylase), DXR (1deoxy-D-xylulose-5-phosphate reductoisomerase), DXS (1-deoxy-D-xylulose-5-phosphate synthase), G10H (geraniol 10-hydroxylase), GPPS (geranyl diphosphate synthase), HDR (1-hydroxy-2-methyl-2-(E)-butenyl-4-diphosphate reductase), HDS (1-hydroxy-2-methyl-2-(E)-butenyl-4-diphosphate synthase), 10HGO (10-hydroxygeraniol oxidoreductase), IPPI (isopentenylpyrophosphate isomerase), MAT (acetyl-CoA:minovincine-O-acetyltransferase), MECS (2Cmethyl-D-erythrol-2,4-cyclodiphosphate synthase), NMT (N-methyltransferase), OMT (O-methyltransferase), SGD (strictosidine ß-D-glucosidase), SLS (secologanin synthase), STR (strictosidine synthase), TDC (tryptophan decarboxylase), T16H (tabersonine 16-hydroxylase).

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reticulum as illustrated by the spatial organization of the vindoline pathway [171]. Interestingly, catharanthine is even excreted out of the cells to the cuticula of the leaves, where it deters insect predators [172]. Biotechnological Production of Catharanthus Alkaloids Over the years many efforts have been made to improve production of the various therapeutically valuable alkaloids of C. roseus. Enormous progress has been made in elucidating the biosynthetic pathway at the level of chemical reactions, structural genes, enzymes, transport and regulation. Functional genomics tools have also helped to considerably accelerate the pace of discoveries in non-model medicinal plants [173]. Undifferentiated cell suspensions would be most suited for virtually unlimited upscaling in containment, but they only accumulate intermediate TIAs or side branch products such as ajmalicine and not the most valuable heterodimeric TIAs vinblastine and vincristine. Approaches including feeding experiments, chemical treatments and genetic transformation have been used to overcome this problem [148]. The influence of phytohormones on alkaloid production has been specifically noted. Auxins suppress alkaloid accumulation whereas cytokinins enhance it [174], and jasmonates trigger extensive transcriptional reprogramming leading to the concerted activation of pathways through the action of master regulators [175]. However, it must be admitted that to date the sustainable improvement of TIA production has not nearly been achieved either in Catharanthus cells or in differentiated hairy roots or whole plants. Further unravelling of the genetic, catalytic and transport processes is needed in order to fully understand the whole complexity of the involved mechanisms and to develop suitable engineering strategies. Recently, resources from the activities of several large research consortia have become available, e.g. http://www.smartcell.org/index.php and http://medicinalplantgenomics.msu.edu/index.shtml that will be helpful for the whole research community. Commercial Exploitation of Catharanthus Alkaloids Since the concentration of alkaloids in the plant is extremely low, i.e. 0.001-0.002% dw in leaves [143], their production is costly. Initially one gram cost $100000 at the pharmacy level [176]. The difficult supply prompted attempts to chemically synthesise the compounds but to date there is no commercially viable route. However, meanwhile a whole list of derivatives semi-synthesized for example from the more abundant monomers vindoline and catharanthine are available, such as vinorelbine (Navelbine®) and vinflunine (Javlor®), that show superior anti-tumour activities and less side effects (http://www.pierre-fabre.com). Despite all these developments the natural compounds as such are still indispensable for cancer chemotherapy after decades [177], maintaining a continued interest in Catharanthus research. ANALYTICAL CHALLENGES IN ALKALOID AND TERPENOID ANALYSES Extraction Taxanes Extraction of taxanes from Taxus species (yew) by 80% ethanol using ultrasonication at 40°C for 60 minutes has been shown to give best yields. The recoveries were higher than 93% for paclitaxel and six other toxoids [178]. Alkaloids Alkaloids are usually isolated from plant material by successive liquid-liquid extraction. Lyophilized and ground plant or cell material is extracted with dilute mineral acid, the separated extract containing alkaloid salts is made alkaline with ammonia and the free alkaloids are extracted with dichloromethane. The final extract is largely free from other molecules or impurities.

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Other methods include extraction with ethanol [174], methanol [179] or methanol-water mixtures, after which the evaporated extract can be treated with alkaline buffer and the alkaloids extracted into ethyl acetate. The extraction efficiency of major morphine alkaloids from Papaver plants was improved by using ultrasoundassisted extraction with 20% methanol at low temperature instead of a long maceration process [180]. Analytical Methods Taxanes Trace analysis of paclitaxel and other taxoids can be performed by reversed phase HPLC-tandem mass spectrometry using ESI or APCI in positive ion mode and quantitated by MRM mode using typical transition masses [178]. The limit of quantitation (LOQ) for paclitaxel and six other taxanes extracted from yew needles ranged from 14 to 32 ng/ml. Tropane Alkaloids Tropane alkaloids are most traditionally analyzed by gas chromatography (GC) with flame ionization detection (FID) or by GC combined with mass spectrometry (GC-MS). Typically, the final alkaloid extract is dissolved in an organic solvent such as dichloromethane, and can thus be directly subjected to GC-MS [181] in which the identification is based on mass fragmentation, commonly in electrospray ionization (ESI) in positive mode. Trimethylsilylation is further used to improve volatility, stability and reliable quantification. However, thermolabile compounds are problematical for GC analysis. The sensitivity has been limited when using high performance liquid chromatography (HPLC) with UV detection, but when coupled to MS, especially tandem MS, this problem is overcome. Fast separations also suitable for sample throughput are obtained by using ultra performance liquid chromatography (UPLC) operating at high pressures (up to 1000 bar) with small particle size columns and normal flows. Quantification of alkaloids is performed by selected ion (SIM) or multiple reaction monitoring (MRM) techniques in GC- and LC-MS, respectively. The use of labeled internal standards is preferred but, unfortunately, can be performed only when a few well-known analytes are quantified. As in the case of tropane alkaloids which do not contain strong chromophores, evaporative light scattering detection (ELSD) becomes a good alternative and complementary detection method for UV [182]; [183]. Novel column technology, e.g. new monolithic and chiral columns, also enables better separation and quantification of tropane alkaloids and their enantiomers. In capillary electrophoresis (CE), although it is a fast method, the minute sample amounts (nanoliter scale) restrict the sensitivity at hardly below micromolar range, compared to nano- or picomolar range in GCand LC-MS. Morphine Alkaloids Extraction of major opium alkaloids, related opiates and metabolites from opium latex with 2.5% acetic acid gives similar recovery to that obtained with methanol or a mixture of 2.5% acetic acid and methanol [184]. The extracts were analysed by liquid chromatography-atmospheric pressure chemical ionization mass spectrometry (LC-APCI-MS) in positive ion mode. When using the selected ion monitoring technique for the analysis of rat urine after inhalation exposure of opium, it was found that only trace levels of thebaine was detected in urine despite the fact that it is one of the major alkaloids in opium. Meconin, in turn, existed in relatively large amounts suggesting that it is indicative of opium ingestion by inhalation. Terpenoid Indole Alkaloids Ion pair techniques in reversed phase HPLC have been used in the analysis of indole alkaloids [179]. Hexane sulphonic acid improved the peak shape and response of the alkaloids at an optimum

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Fig. (6). General strategy for the metabolic engineering of microorganisms for drug production (modified from Lee et al [185]).

concentration of 2.5 mM. The pH of the eluent was 2.5 and by using this method it was possible to separate indole alkaloids which were structurally very closely related, such as isomers and even epimers. In order to understand the regulatory system governing total indole alkaloid (TIA) metabolism in elicited Catharanthus roseus cells, targeted and non-targeted LC-MS analyses at pH 10 were used in combination with the cDNA-amplified fragment length polymorphism (AFLP) technology. This provided insights not only into TIA metabolism but also into secondary metabolism in C. roseus in general [174]. Metabolic Profiling Omics tools can be exploited at different stages of drug development (Fig. 6), [185]. They are used for global profiling of metabolites or for quantifying and controlling the levels of known medicinal compounds and potential metabolic intermediates in plant tissues or cell cultures. Combining the data between the metabolome and genome of medicinal plants provides novel information about the genes and markers of medicinal compound synthesis [186]. Metabolomic fingerprinting is most commonly performed by using HPLC or UPLC coupled to negative or positive electrospray ionization mass spectrometry employing accurate mass time of flight (TOF) or ion trap multistage tandem mass spectrometry

[187]; [188]. GC
GC-TOF techniques for volatile compounds, and NMR (nuclear magnetic resonance) spectroscopy are also more frequently used. Data analysis is based on multivariate statistical methods, e.g. principal component analysis (PCA) or partial least squares discriminant analysis (PLSD). Sophisticated methods are needed for data-mining and filtering of the huge amount of data and novel imaging techniques have in turn considerably aided interpretation of the complex data. However, the metabolic profiling of higher plants biosynthesizing approximately 30000 low molecular weight products, as well as the efforts in the combinatorial biosynthesis of secondary metabolites such as terpenoids (paclitaxel, artemisinin) and alkaloids (morphine, Vinca alkaloids) and their by-products [189], necessitate more and more efficient and sensitive analytical and computational tools. CONFLICT OF INTEREST The authors confirm that this article content has no conflicts of interest. ACKNOWLEDGEMENTS VTT and UdL gratefully acknowledge support from European Union Framework 7 Program Integrated Project 222716 SmartCell; European Union Framework 7 European Research Council IDEAS Advanced Grant (to PC) Program-BIOFORCE; COST Actions

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FA0804 Molecular farming: plants as a production platform for high value proteins and FA1006 Plant Metabolic Engineering for High Value Products.

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[27]

REFERENCES [1] [2]

[3] [4]

[5] [6] [7] [8] [9] [10]

[11] [12]

[13] [14] [15]

[16] [17]

[18]

[19] [20] [21] [22]

[23]

[24] [25]

Verpoorte R. Secondary metabolism. In: Verpoorte R, Alfermann AW. Eds. Metabolic engineering of plant secondary metabolism. Kluwer Academic Publishers 2000; pp. 1-29. Weid M, Ziefler J, Kutchan TM. The roles of latex and the vascular bundle in morphine biosynthesis in the opium poppy, Papaver somniferum. Proc Natl Acad Sci USA 2004; 101: 13957-62. Oksman-Caldentey K-M, Inzé D. Plant cell factories in the postgenomic era: new ways to produce designer secondary metabolites. Trends Plant Sci 2004; 9: 433-40. McChesney JD, Venkataraman SK, Henri JT. Plant natural products: Back to the future or into extinction? Phytochemistry 2007; 68: 2015-22. Hostettmann K, Terreaux C. Search for new lead compounds from higher plants. Chimia (Aarau) 2000; 54:652-7. Raskin I, Ribnicky DM, Komarnytsky S, et al. Plants and human health in the twenty-first century. Trends Biotechnol 2002; 20: 52231. Neuman DJ, Cragg GM. Natural products as sources of new drugs over 30 years from 1981-2010. J Nat Prod 2012; 75: 311-35. Lewington A, Parker E. Ancient trees: Trees that live for a thousand years. Collins & Brown Limited, London 1999; pp.192. Cope EA. Taxaceae: The genera and cultivated species. Bot Rev 1998; 64: 291-322. Stierle A, Strobel GA, Stierle DB, Grothaus P, Bignami G. The search for a taxol-producing microorganism among the endophytic fungi of the Pacific yew, Taxus brevifolia. J Nat Prod 1995; 58: 1315-24. Stone R. Surprise! A fungus factory for taxol? Science 1993; 260: 154-5. Staniek A, Woerdenbag HJ, Kayser O. Taxomyces andreanae: A presumed paclitaxel producer demystified? Planta Med 2009; 75: 1561-6. Schiff PB, Fant J, Horwich SB. Promotion of microtubule assembly by taxol. Science 1979; 227: 665-7. Bristol-Myers Squipp. (http://packageinserts.bms.com/pi/pi_taxol.pdf) update July 2007; rd Access 3 November 2008. Montero A, Fossella F, Hortobagyi G, Valero V. Docetaxel for treatment of solid tumours: a systematic review of clinical data. Lancet Oncol 2005; 6: 229-39. McGrogan BT, Gilmartin B, Carney DN, McCann A. Taxanes, microtubules and chemoresistant breast cancer. Biochim Biophys Acta 2008; 1785: 96-132. Brown DT. Pre-clinical and clinical studies of taxanes. In: Itokawa H, Lee KH. Eds. Taxus, the genus Taxus, Taylor and Francis Group, London, New York 2003; 387-420. Zhang B, Maiti A, Shively S, Lakhani F, et al. Microtubule-binding drugs offset tau sequestration by stabilizing microtubules and reversing fast axonal transport deficits in a tauopathy model. Proc Nat Acad Sci USA 2005; 102: 227-31. Mastronardi FG, Tsui H, Winer S, et al. Synergy between paclitaxel plus an exogenous methyl donor in the suppression of murine demyelinating diseases. Multiple Sclerosis 2007; 13: 596-609. Risinger AL, Giles FJ, Mooberry SL. Microtubule dynamics as a target in oncology. Cancer Treat Rev 2009; 35: 255-61. Jordan MA, Kamath K. How do microtubule-targeted drugs work? An overview. Curr Cancer Drug Targets 2007; 7: 730-42. Yeung TK, Germond C, Chen X, Wang Z. The mode of action of taxol: Apoptosis at low concentration and necrosis at high concentration. Biochem Biophys Res Comm 1999; 263: 398-404. Cusidó RM, Palazón J, Bonfill M, Expósito O, Moyano E, Piñol MT. Source of isopentenyl diphosphate for taxol and baccatin III biosynthesis in cell cultures of Taxus baccata. Biochem Eng J 2007; 33: 159-67. Croteau R, Ketchum REB, Long RM, Kaspera R, Wildung MR. Taxol biosynthesis and molecular genetics. Phytochem Rev 2006; 5: 75-97. Vongpaseuth K, Roberts SC. Advance in the understanding of paclitaxel metabolism in tissue culture. Curr Pharm Biotech 2007; 8: 219-36.

[28]

[29]

[30]

[31]

[32]

[33]

[34]

[35] [36]

[37]

[38]

[39] [40]

[41] [42]

[43] [44]

[45] [46] [47] [48]

Onrubia Ibáñez M. A molecular approach to taxol biosynthesis. Doctoral dissertation. Universitat Pompeu Fabra, Barcelona, Spain. 2012; p. 173 + app. Koepp AE, Hezari M, Zajicek J, et al. Cyclization of geranylgeranyl diphosphate to taxa-4(5),11(12)-diene is the committed step of taxol biosynthesis in Pacific yew. J Biol Chem 2005; 270: 8686-90. Hezari M, Ketchum REB, Gibson DM, Croteau R. Taxol production and taxadiene synthase activity in Taxus canadensis cell suspension cultures. Arch Biochem Biophys 1997; 337: 185-90. Hefner J, Ketchum REB, Croteau R. Cloning and functional expression of a cDNA encoding geranylgeranyl diphosphate synthase from Taxus canadensis and assessment of the role of this prenyltransferase in cells induced for taxol production. Arch Biochem Biophys 1998; 360: 62-74. Walker K, Schoendorf A, Croteau R. Molecular cloning of a taxa4(20),11(12)-dien-5
-ol-O-acetyl transferase cDNA from Taxus and functional expression in Escherichia coli. Arch Biochem Biophys 2000; 374: 371-80. Jennewein S, Rithner CD, Williams RM, Croteau R. Taxol biosynthesis: taxane 13
hydroxylase is a cytochrome P450-dependent monooxygenase. Proc Nat Acad Sci USA 2001; 98: 13595-600. Schoendorf A, Rithner CD, Williams RM, Croteau R. Molecular cloning of a cytochrome P450 taxane-10-hydroxylase cDNA from Taxus and functional expression in yeast. Proc Nat Acad Sci USA 2001; 98: 1501-6. Walker K, Croteau R. Taxol biosynthesis: molecular cloning of a benzoyl CoA:taxane-2
-O-benzoyl transferase cDNA from Taxus and functional expression in Escherichia coli. Proc Nat Acad Sci USA 2000a; 97: 13591-6. Walker K, Croteau R. Molecular cloning of a 10-deacetylbaccatin III-10-O-acetyl transferase cDNA from Taxus and functional expression in Escherichia coli. Proc Nat Acad Sci USA 2000b; 97: 583-7. Floss HG, Mocek U. Biosynthesis of taxol In: Suffness M. Ed. Taxol – Science and applications. CRC Press, Boca Raton. FL, USA 1995; pp 191-208. Walker K, Fujisaki S, Long R, Croteau R. Molecular cloning and heterologous expression of the C-13 phenylpropanoid side chainCoA acyltransferase that functions in taxol biosynthesis. Proc Nat Acad Sci USA 2002a; 99: 12715-20. Walker K, Long R, Croteau R. The final acylation step in taxol biosynthesis: cloning of the taxoid C13-side-chain Nbenzoyltransferase from Taxus. Proc Nat Acad Sci USA 2002b; 99: 9166-71. Wheeler LA, Long RM, Ketchum REB, Rithner CD, Williams RM, Croteau R. Taxol biosynthesis: differential transformation of taxadien-5
-ol and its acetate ester by cytochrome P450 hydroxylases from Taxus suspension cells. Arch Biochem Biophys 2001; 390: 265-78. Fett-Neto AG, DiCosmo F, Reynolds WF, Sakata K. Cell culture of Taxus as a source of the antineoplastic drug taxol and related taxanes. Nature Biotech 1992; 10: 1572-175. Cusidó RM, Palazón J, Bonfill M, Navia-Osorio A, Morales C, Piñol MT. Improved paclitaxel and baccatin III production in suspension cultures of Taxus media. Biotechn Prog 2002; 18: 418-23. Yukimune Y, Tabata H, Higashi Y, Hara Y. Methyl jasmonate induced overproduction of paclitaxel and baccatin III in Taxus cell suspension cultures. Nature Biotech 1996; 14: 1129-32. Bentebibel S, Moyano E, Palazón J, et al. Effects of immobilization by entrapment in alginate and scale-up on paclitaxel and baccatin III production in cell suspension cultures of Taxus baccata. Biotech Bioeng 2005; 89: 647-55. Liu T, Koshla C. A balancing act for taxol precursor pathways in E. coli. Science 2010; 330: 44-5. Malik S, Cusidó RM, Mirjalili MH, Moyano E, Palazón J, Bonfill M. Production of the anticancer drug taxol in Taxus baccata suspension cultures: A review. Process Biochem 2001; 46: 23-34. Sabater-Jarra AB, Tudela LR, López-Pérez AJ. In vitro culture of Taxus sp.: strategies to increase cell growth and taxoid production. Phytochem Rev 2010; 9: 343-56. Mann J. Murder, magic and medicine. Oxford University Press, 1992; 240 p. Lounasmaa M, Tamminen T. In: G.A. Cordell (Ed.), The alkaloids. Vol. 44. Academic Press, San Diego, USA 1993; p. 1. Eich E. Solanaceae and Convolvulaceae: Secondary metabolites. Springer- Verlag, Berlin, Heidelberg, 2008; pp.33-212.

Plant Cells as Pharmaceutical Factories [49] [50] [51] [52] [53]

[54]

[55] [56]

[57] [58]

[59]

[60]

[61]

[62] [63]

[64]

[65]

[66] [67]

[68]

[69]

[70]

[71]

Griffin WJ, Lin GD. Chemotaxonomy and geographical distribution of tropane alkaloids. Phytochemistry 2000; 53: 623-37. Eger, E. Artopine, scopolamine and related drugs. Anesthesiology 1962; 23: 365-383. Poupko JM, Baskin SI, Moore E. The pharmacological properties of anisodamine. J Appl Toxicol 2006; 27: 116-21. Mizusaki S, Tanabe Y, Noguchi M, Tamaki E. N-methylputrescine oxidase from tobacco roots. Phytochemistry 1972; 11: 2757-62. McLauchlan WR, McKee RA, Evans DM. The purification and immunocharacterisation of N-methylputrescine oxidase from transformed root cultures of Nicotiana tabacum l. cv SC58. Planta 1993; 191: 440-5. Heim WG, Sykes KA, Hildreth SB, Sun J, Lu RH, Jelesko JG. Cloning and characterization of a Nicotiana tabacum methylputrsecine oxidase transcript. Phytochemistry 2007; 68: 454-63. Katoh A, Shoji T, Hashimoto T. Molecular cloning of Nmethylputrescine oxidase from tobacco. Plant Cell Physiol 2007; 48: 550-4. Samuelsson G, Bohlin L. Drugs of Natural Origin. 6th revised edition. Apotekarsocieteten – Swedish Academy of Pharmaceutical Sciences, Kristianstad, Sweden. 2009; 776 p. Landgrebe ME, Leete E. The metabolism of tropinone in intact Datura innoxia plants. Phytochemistry 1990; 29: 2521-4. Nakajima K, Hashimoto T, Yamada Y. Two tropinone reductases with different stereospecificities are short-chain dehydrogenases evolved from a common ancestor. Proc Natl Acad Sci USA 1993; 90: 9591-5. Keiner R, Kaiser H, Nakajima K, Hashimoto T, Dräger B. Molecular cloning, expression and characterization of tropinone reductase II, an enzyme of the SDR family in Solanum tuberosum (L.). Plant Mol Biol 2002; 48: 299-308. Li R, Reed DW, Liu E, et al. Functional genomic analysis of alkaloid biosynthesis in Hyoscyamus niger reveals a cytochrome P450 involved in littorine rearrangement. Chemistry and Biol 2006; 13: 513-20. Al Balkhi MH, Schiltz S, Lesur D, Lanoue A, Wadouachi A, Boitel-Conti M. Norlittorine and norhyoscyamine identified as products of littorine and hyoscyamine metabolism by 13C-labeling in Datura innoxia hairy roots. Phytochemistry 2012; 74: 105-14. Hashimoto T, Yamada Y. Hyoscyamine 6-hydroxylase, a 2oxoglutarate-dependent dioxygenase, in alkaloid-producing root cultures. Plant Physiol 1986; 81: 619-25. Matsuda J, Okabe S, Hashimoto T, Yamada Y. Molecular cloning of hyoscyamine 6ß-hydroxylase, a 2-oxoglutarate-dependent dioxygenase, from cultured roots of Hyoscyamus niger. J. Biol. Chem 1991; 266: 9460-4. Hashimoto T, Hayashi A, Amano Y, et al. Hyoscyamine 6hydroxylase, an enzyme involved in tropane alkaloid biosynthesis, is localized at the pericycle of the root. J Biol Chem 1991; 266: 4648-53. Yun D-J, Hashimoto T, Yamada Y. Metabolic engineering of medicinal plants: transgenic Atropa belladonna with an improved alkaloid composition. Proc Natl Acad Sci USA 1992; 89: 11799-803. Hashimoto T, Yun D-J, Yamada Y. Production of tropane alkaloids in genetically engineered root cultures. Phytochemistry 1993; 32: 713-8. Jouhikainen K, Lindgren L, Jokelainen T, Hiltunen R, Teeri T, Oksman-Caldentey KM. Enhancement of scopolamine production in Hyoscyamus muticus L. hairy root cultures by genetic engineering. Planta 1999; 208: 545-51. Zhang L, Ding R, Chai Y, et al. Engineering tropane biosynthetic pathway in Hyoscyamus niger hairy root cultures. Proc Natl Acad Sci USA 2004; 101: 6786-91. Oksman-Caldentey K-M, Häkkinen ST, Rischer H. Metabolic engineering of the alkaloid biosynthesis in plants: functional genomics approaches. In: Applications of plant metabolic engineering. Verpoorte R, Alfermann AW & Johnson TS (eds.), Springer, Heidelberg 2007; 109-27. Moyano E, Jouhikainen K, Tammela P, et al. Effect of pmt gene overexpression on tropane alkaloid production in transformed root cultures of Datura metel and Hyoscyamus muticus. J Exp Bot 2003; 54: 203-11. Rothe G, Hachiya A, Yamada Y, Hashimoto T, Dräger B. Alkaloids in plants and root cultures of Atropa belladonna overexpressing putrescine N-methyltransferase. J Exp Bot 2003; 54: 2065-70.

Current Pharmaceutical Design, 2013, Vol. 19, No. 31 [72]

[73] [74]

[75] [76] [77]

[78]

[79] [80]

[81] [82] [83]

[84]

[85]

[86]

[87]

[88]

[89]

[90]

[91] [92]

[93]

[94]

5657

Moyano E, Fornalé S, Palazón J, Cusidó RM, Bagni N, Piñol MT. Alkaloid production in Duboisia hybrid hairy root cultures overexpressing the pmt gene. Phytochemistry 2002; 59: 697-702. Yang C, Chen M, Zeng L, et al. Improvement of tropane alkaloid production in hairy root cultures of Atropa belladonna by overexpressing pmt and h6h genes. Plant Omics J 2011; 4: 29-33. Asano N, Nash RJ, Molyneux RJ, Fleet GW. Sugar-mimic glycosidase inhibitors: natural occurrence, biological activity and prospects for therapeutic application. Tetrahedron Asymmetry 2000; 11: 1645-80. Goldmann A, Milat ML, Ducrot PH, et al. Tropane derivatives from Calystegia sepium. Phytochemistry 1990; 29: 2125-8. Dräger B, van Almsick A, Mrachatz G. Distribution of calystegines in several Solanaceae. Planta Med 1995; 61: 577-9. Dräger B, Funck C, Hoehler A, et al. Calystegines as a new group of tropane alkaloids in Solanaceae. Plant Cell Tissue Org Cult 1994; 38: 235-40. Robins RJ, Bachmann P, Robinson T, Rhodes MJC, Yamada Y. The formation of 3
- and 3-acetoxytropanes by Datura stramonium transformed root cultures involves two acetyl-CoA-dependent acyltransferases. FEBS 1991; 292: 293-7. Rabot S, Peerless AC, Robins RJ. Tigloy-CoA:pseudotropine acyl transferase – an enzyme of troane alkaloid biosynthesis. Phytochemistry 1995; 39: 315-22. Robins RJ, Bachmann P, Peerless ACJ, Rabot S. Esterification reactions in the biosynthesis of tropane alkaloids in transformed root cultures. Plant Cell Tissue Organ Cult 1994; 38: 241-7. Witte L, Müller K, Arfmann H-A. Investigation of the alkaloid pattern of Datura innoxia plants by capillary gas-liquidchromatography-mass-spectrometry. Planta Med 1987; 53: 192-7. Hiraoka N, Tabata M. Acetylation of tropane derivatives by Datura innoxia cell cultures. Phytochemistry 1983; 22: 409-12. Hartmann T, Witte L, Oprach F, Toppel G. Reinvestigation of the alkaloid composition of Atropa belladonna plants, root cultures, and cell suspension cultures. Planta Med 1986; 52: 390-395. Wilhelmson A, Häkkinen ST, Kallio P, Oksman-Caldentey KM, Nuutila AM. Heterologous expression of Vitreoscilla hemoglobin (VHb) and cultivation conditions affect the alkaloid profile of Hyoscyamus muticus hairy roots. Biotechnol Prog 2006; 22: 350-8. Häkkinen ST. A functional genomics approach to the study of alkaloid biosynthesis and metabolism in Nicotiana tabacum and Hyoscyamus muticus cell cultures. VTT Publications 696 (http://www.vtt.fi/publications/index.jsp) 2008; 90 p. Cardillo AB, Rodríguez Talou J, Giulietti AM. Expression of Brugmansia candida hyoscyamine-6- hydroxylase gene in Saccharomyces cerevisiae and its potential use as biocatalyst. Microbial Cell Fact 2008; 7: 17. Kai G, Liu Y, Wang X, Yang S, Fu X, Luo X, Liao P. Functional identification of hyoscaymine 6 ß-hydroxylase from Anisodus acutangulus and overproduction of scopolamine in geneticallyengineered Escherichia coli. Biotechnol Lett 2011; 33: 1361-5. Häkkinen ST, Moyano E, Cusidó RM, Palazón J, Piñol MT, Oksman-Caldentey K-M. Enhanced secretion of tropane alkaloids in Nicotiana tabacum hairy roots expressing heterologous hyoscyamine-6ß-hydroxylase. J Exp Bot 2005; 56: 2611-8. Hallard D, van der Heijden R, Verpoorte R. Suspension cultured transgenic cells of Nicotiana tabacum expressing tryptophan decarboxylase and strictosidine synthase cDNAs from Catharanthus roseus produce strictosidine upon feeding of secologanin. Plant Cell Rep 1997; 17: 50-4. Kanegae T, Kajiya H, Amano Y, Hashimoto T, Yamada Y. Species-dependent expression of the hyoscyamine 6-hydroxylase gene in the pericycle. Plant Physiol 1994; 105: 483-90. Moyano E, Palazón J, Bonfill M, et al. Biotransformation of hyoscyamine into scopolamine in transgenic tobacco cell cultures. J Plant Physiol 2007; 164: 521-524. Shitan N, Bazin I, Dan K, et al. Involvement of CjMDR1, a plant multidrug-resistance-type ATP-binding cassette protein, in alkaloid transport in Coptis japonica. Proc Natl Acad Sci USA 2003; 100: 751-6. Li L, He Z, Pandey GK, Tsuchiya T, Luan S. Functional cloning and characterization of a plant efflux carrier for multidrug and heavy metal detoxification. J Biol Chem 2002; 277: 5360-8. Yun D-J, Hashimoto T, Yamada Y. Transgenic tobacco plants with two consecutive oxidation reactions catalyzed by hyoscyamine-6hydroxylase. Biosci Biotech Biochem 1993; 57: 502-3.

5658 Current Pharmaceutical Design, 2013, Vol. 19, No. 31 [95]

[96]

[97]

[98]

[99] [100] [101]

[102] [103]

[104]

[105]

[106] [107]

[108]

[109]

[110]

[111]

[112]

[113] [114]

[115]

Rocha P, Stenzel O, Parr A, et al. Functional expression of tropinone reductase I (trI) and hyoscyamine-6
-hydroxylase (h6h) from Hyoscyamus niger in Nicotiana tabacum. Plant Sci 2002; 162: 90513. Subroto MA, Kwok KH, Hamill JD, Doran PM. Coculture of genetically transformed roots and shoots for synthesis, translocation, and biotransformation of secondary metabolites. Biotechnol Bioeng 1996; 49: 481-94. Dehghan E, Häkkinen ST, Oksman-Caldentey KM, Shariari F. Production of tropane alkaloids in diploid and tetraploid plants and in vitro hairy root cultures of Egyptian henbane (Hyoscyamus muticus L.). Plant Cell Tissue Org Cult 2012; 110: 35-44. Grynkiewicz G, Gadzikowska M. Tropane alkaloids as medicinally useful natural products and their synthetic derivatives as new drugs. Pharmacol. Rep. 2008; 60: 439-63. Boehringer Ingelheim Annual report 2011. (www.boehringeringelheim.com). Lenz R, Zenk MH. Acetyl coenzyme A:salutaridinol-7-Oacetyltransferase from Papaver somniferum plant cell cultures. J Biol Chem 1995; 280: 31091-6. Rice KC. Synthetic opium alkaloids and derivatives. A short total synthesis of (±)-dihydrothebainone, (±)-dihydrocodeinone and (±)nordihydrocodeinone as an approach to a practical synthesis of morphine, codeine and congeners. J Org Chem 1980; 45: 3135-7. Trescot AM, Datta S, Lee M, Hansen H. Opioid Pharmacology. Pain Physician 2008: Opioid Special Issue: 11:S133-53. Unterlinner B, Lenz R, Kutchan TM. Molecular cloning and functional expression of codeinone reductase: the penultimate enzyme in morphine biosynthesis in the opium poppy Papaver somniferum. Plant J 1999; 18:465-75. Facchini PJ, De Luca V. Differential and tissue spesific expression of a gene family for tyrosine/dopa decarboxylase in opium poppy. J Biol Chem 1994; 269: 26684-90. Maldonado-Mendoza IE, López-Meyer M, Galef JR, Burnett RJ, Nessler CL. Molecular analysis of a new member of the Opium poppy tyrosine/3,4-dihydrohyphenylalanine decarboxylase gene family. Plant Physiol 1996; 110:43-9. El-Ahmady SH, Nessler CL. Cellular localozation of tyrosine decarboxylase expression in transgenic opium poppy and tobacco. Plant Cell Rep 2001; 20: 313-7. Samanani N, Facchini PJ. Isolation and partial characterization of norcolaurine synthase, the first committed step in benzylisoquinoline alkaloid biosynthesis, from opium poppy. Planta 2001; 213:898-906. Samanani N, Facchini PJ. Purification and characterization of norcolaurine synthase - the first committed enzyme in benzylisoquinoline alkaloid biosynthesis in plants. J Biol Chem 2002; 277: 3387883. Samanani N, Liscombe DK, Facchini PJ. Molecular cloning and characterization of norcoclaurine synthase, an enzyme catalyzing the first committed step in benzylisoquinoline alkaloid biosynthesis. Plant J 2004; 40: 302-13. Luk LYP, Bunn S, Liscombe DK, Facchini PJ, Martin E Tanner. Mechanistic studies on norcoclaurine synthase of benzylisoquinoline alkaloid biosynthesis: An enzymatic Pictet-Spengler reaction. Biochem 2007; 46:10153-61. Morishige T, Tsujita T, Yamada Y, Sato F. Molecular characterization of the S-adenosyl-L-methionine: 30-hydroxy-Nmethylcoclaurine-40-O-methyltransferase of isoquinoline alkaloid biosynthesis in Coptis japonica. J Biol Chem 2000; 275: 23398405. Choi K-B, Morisige T, Shitani N, Yazaki K, Sato F. Molecular cloning and characterization of coclaurine N-methyltransferase from cultured cells of Coptis japonica. J Biol Chem 2002; 277: 830-5. Facchini PJ, Park SU. Developmental and inducible accumulation of gene transcripts involved in alkaloid biosynthesis in opium poppy. Phytochemistry 2003; 64: 177-86. Pauli HH, Kutchan TM. Molecular cloning and functional expression of two alleles encoding (S)-N-methylcoclaurine 3`hydroxylase (CYP80B1), a new methyl jasmonate-inducible cytochrome P-450-dependent mono-oxygenase of benzylisoquinoline alkaloid biosynthesis. Plant J 1998; 13: 793-801. Huang F-C, Kutchan TM. Distribution of morphinan and benzophenanthridine alkaloid gene transcript accumulation in Papaver somniferum. Phytochemistry 2000; 53: 555-64.

Rischer et al. [116]

[117] [118]

[119]

[120]

[121] [122]

[123]

[124]

[125] [126]

[127] [128]

[129] [130] [131]

[132] [133]

[134]

[135] [136]

[137] [138]

Frick S, Kramella R, Kutchan TM. Metabolic engineering with a morphine biosynthetic P450 in opium poppy surpasses breeding. Metabolic Eng 2007; 9: 169-76. Facchini PJ, Penzes C, Johnson AG, Bull D. Molecular characterization of berberine enzymes genes from opium poppy. Plant Physiol 1996; 112: 1669-77. Hauschild K, Pauli HH., Kutchan TM. Isolation and analysis of a gene bb1 encoding the berberine bridge enzyme from the Californian poppy Eschscholzia californica. Plant Mol Biol 1998; 36: 4738. Park SU, Johnson AG, Penzes-Yost C, Facchini PJ. Analysis of promoters from tyrosine/dihydroxyphenylalanine decarboxylase and berberine bridge enzyme genes ivolved in benzylisoquinole alkloid biosynthesis in opium poppy. Plant Mol Biol 1999; 40: 12131. Park SU, Yu M, Facchini PJ. Antisense RNA-mediated suppressiom of benzophenanthridine alkaloid biosynthesis in transgenic cell cultures of Californian poppy. Plant Physiol. 2002; 128: 696706. Kutchan TM. Alkaloid biosynthesis - The basis for metabolic engineering of medicinal plants. Plant Cell 1995; 7: 1059-70. Grothe T, Lenz R, Kutchan TM. Molecular characterization of the salutardidinol 7-O-acetyltransferase involved in morphine biosynthesis in opium poppy Papaver somniferum. J Biol Chem 2001; 276: 30717-23. Kempe K, Higashi Y, Frick S, Sabarna K, Kutchan TM. RNAi suppression of the morphine biosynthetic gene salAT and evidence of association of pathway enzymes. Phytochemistry 2009; 70: 57989. Allen RS, Miller JAC, Chitty JA, Fist AJ, Gerlach WL, Larkin PJ. Metabolic engineering of morphinan alkaloids by over-expression and RNAi suppression of salutaridinol 7-O-acetyltransferase in opium poppy. Plant Biotech J 2008; 6: 22-30. Larkin PJ, Miller JAC, Allen RS, et al. Increasing morphinan alkaloid production by over-expressing codeinone reductase in transgenic Papaver somniferum. Plant Biotech. J. 2007; 5: 26-37. Farrow SC, Hagel JM, Facchini PJ. Transcript and metabolite profiling in cell cultures of 18 plant species that produce benzylisoquinoline alkaloids Phytochemistry 2010; 77: 79-88. Wijekoon CP, Facchini PJ. Systematic knockdown of morphine pathway enzymes in opium poppy using virus-induced gene silencing. Plant J 2012; 69: 1052-63. Apuya NR, Park JH, Zhang L, et al. Enhancement of alkaloid production in opium and California poppy by transactivation using heterologous regulatory factors. Plant. Biotech J 2008; 6: 160-75. Misawa M. Plant tissue culture: An alternative for production of useful metabolite. FAO Agricultural services bulletin 1994; No 108. ISBN 92-5-103391-9. Hsu AF, Park J. Metabolism of 14C-codein in cell culture of Papaver somniferum. Phytochemistry 1989; 28: 1879-81. Kassem MA, Jacquin A. Somatic embryogenesis, rhizogenesis and morphinan alkaloids production in two species of opium poppy. J Biomed Biotech 2001; 1:70-8. Siah CL, Doran PM. Enhanced codeine and morphine production in suspended Papaver somniferum cultures after removal of exogenous hormones. Plant Cell Rep 1991; 10: 349-53. Alkhimova OG, Kyrylenko TK, Vagyn Yu V, Heslop-Harrison JS. Alkaloid biosynthesis in Papaver sp. cells in culture and during organogenesis. Ukrainskii Biokhimicheskii Zhurnal 2001; 73: 141-6. Le Flem-Bonhomm V, Laurain-Mattar D, Fliniaux MA. Hairy root induction of Papaver somniferum var. album, a difficult-totransform plant, by A. rhizogenes LBA 9402. Planta 2004; 218: 890-3. Facchini PJ, Bird DA. Developmental regulation of benzylisoquinoline alkaloid biosynthesis in opium poppy plants and tissue cultures. In vitro Cell Dev Biol Plant 1998; 34: 69-79. Bobak M, Nadaska M, Smaj J, et al. The influence of elicitation on the subcellular localization and content of sanguinarine in callus cells of Papaver somniferum L. Biol Plant 1995; 37: 501-6. Archambault J, Williams RD, Bedard C, Chavarie C. Production of sanguinarine by elicitated plant cell cultures: I Shake flask suspension cultures. J Biotechnol 1996; 46: 95-105. Williams RD, Bedard C, Chavarie C, Archambault J. Production of sanguinarine by elicitated plant cell cultures: II Further nutritional aspects. J Biotechnol 1996; 46:107-20.

Plant Cells as Pharmaceutical Factories [139]

[140]

[141] [142] [143] [144] [145] [146] [147]

[148]

[149] [150]

[151]

[152]

[153] [154]

[155]

[156] [157]

[158]

[159]

[160] [161]

[162]

Saxton JE (ed.). Monoterpenoid indole alkaloids. Supplement to Part 4, The Chemistry of Heterocyclic Compounds, Wiley, New York, 1994; 860 p. Yang L, Hill M, Wang M, Panjikar, Stöckigt J. Structural basis and enzymatic mechanism of the biosynthesis of C9- from C10monoterpenoid indole alkaloids. Angew Chem Int Ed 2009; 48: 5211-3. Pelletier PJ, Caventou JB. Note sur un nouvel alcali. Ann Chim Phys 1818; 8: 323. Barron SE, Guth PS. Uses and limitations of strychnine as a probe in neurotransmission. Trends Pharmacol Sci 1987; 8: 204-6. Noble RL. The discovery of the Vinca alkaloids – chemotherapeutic agents against cancer. Biochem Cell Biol 1990; 68: 1344-51. Noble RL. Role of chance observation in chemotherapy: Vinca rosea. Ann NY Acad Sci 1958; 76: 882-94. Johnson IS, Wright HF, Svoboda GH. Experimental basis for clinical evaluation of anti-tumour principles derived from Vinca rosea. Linn J Lab Clin Med 1959; 54: 830. Schmidt B, Kutney J, Mayer L. Anhydrovinblastine for the treatment of cancer. PCT Int Appl 1998 WO 9839004. Takano Y, Okudeira M, Harmon BV. Apoptosis induced by microtubule disrupting drugs in cultured human lymphoma cells – inhibitory effects of phorbol ester and zinc sulphate. Pathol Res Pract 1993; 189: 197-203. Van der Heijden R, Jacobs DI, Snoeijer W, Hallard D, Verpoorte R. The Catharanthus alkaloids: pharmacognosy and biotechnology. Curr Med Chem 2004; 11: 607-28. Wang L, He HP, Di YT, Zhang Y, Hao XJ. Catharoseuramine, a new monoterpenoid indole alkaloid possessing a peroxy bridge from Catharanthus roseus. Tetrahedron Lett 2012; 53: 1576-8. Wang Y, Liu Z, Song F, Liu S. Electrospray ionization tandem mass spectrometric study of the aconitines in the roots of aconite. Rapid Communications in Mass Spectrometry 2002; 16; 2075-82. Verpoorte R, van der Heijden, Moreno PRH. Biosynthesis of terpenoid indole alkaloids in Catharanthus roseus cells. In: The alkaloids – chemistry and pharmacology Volume 49. Edited by: Cordell GA. Academic Press, San Diego, 1997. Arigoni D, Sanger S, Latzel C, Eisenreich W, Bacher A, Zenk MH. Terpenoid biosynthesis from 1-deoxy-D-xylulose in higher plants by intramolecular skeletal rearrangement. Proc Natl Acad Sci USA 1997; 94: 10600-5. Phillips MA, Leon P, Boronat A, Rodriguez-Concepcion M. The plastidal MEP pathway: unified nomenclature and resources. Trends Plant Sci 2008; 13: 619-23. Meijer AH, de Waal A, Verpoorte R. Purification of the cytochrome P450 enzyme geraniol 10-hydroxylase from cell cultures of Catharanthus roseus. J Chromatogr 1993; 635: 237-49. Ikeda H, Esaki N, Nakai S, et al. Acyclic monoterpene primary alcohol NADP+ oxidoreductase of Rauwolfia serpentina cells – the key enzyme in biosynthesis of monoterpene alcohols. J Biochem 1991; 109: 341-7. Madyastha KM, Guarnaccia R, Baxter C, Coscia CJ. S-adenosyl – L-methionine:loganic acid methyltransferase. J Biol Chem 1973; 248: 2497-501. Irmler S, Schröder G, St-Pierre B, et al. Indole alkaloid biosynthesis in Catharanthus roseus: new enzyme activities and identification of cytochrome P450 CYP72A1 as secologanin synthase. Plant J 2000; 24: 797-804. Stöckigt J, Ruppert M. Strictosidine: the biosynthetic key to monoterpenoid indole alkaloids. In Comprehensive Natural Products Chemistry Volume 4. Edited by: Kelly JW. Elsevier Science Ltd. Amsterdam, 1999. Stöckigt J, Hemscheidt T, Höfle G, Heinstein P, Formacek V. Steric course of hydrogen transfer during enzymatic formation of 3-heteroyohimbine alkaloids. Biochem 1983; 22: 3448-52. St Pierre B, De Luca V. A cytochrome P-450 monooxygenase catalyzes the first step in the conversion of tabersonine to vindoline in Catharanthus roseus. Plant Phys 1995; 109: 131-9. Levac D, Murata J, Kim WS, De Luca V. Application of corundum abrasion for investigating the leaf epidermis: molecular cloning of Catharanthus roseus 16-hydroxytabersonine 16-Omethyltransferase. Plant J 2008; 53: 225-36. De Luca V, Balsevic J, Tyler RT, Kurz WGW. Characterisation of a novel N-methyltransferase (NMT) from Catharanthus roseus plants. Plant Cell Rep 1987; 6: 458-61.

Current Pharmaceutical Design, 2013, Vol. 19, No. 31 [163]

[164] [165]

[166]

[167] [168]

[169]

[170]

[171] [172]

[173] [174]

[175]

[176] [177] [178]

[179]

[180]

[181]

[182]

[183]

5659

Vazques-Flota F, De Carolis E, Alarco AM, De Luca V. Molecular cloning and characterization of desacetoxyvindoline-4hydroxylase, a 2-oxoglutarate dependent dioxygenase involvled in the biosynthesis of vindoline in Catharanthus roseus (L) G. Don. Plant Mol Biol 1997; 34: 935-48. De Luca V, Balsevic J, Kurz WGW. Acetyl coenzyme A:deacetylvindoline O-acetyltransferase, a novel enzyme from Catharanthus. J Plant Physiol 1985; 121:417-28. Sottomayor M, Lopez-Serano M, DiCosmo F, Barcelo A. Purification and characterization of -3’,4’-anhydrovinblastine synthase (peroxidase-like) from Catharanthus roseus (L.) G. Don. FEBS Lett 1998; 428: 299-303. Memelink J, Verpoorte R, Kijne W. ORCAnization of jasmonateresponsive gene expression in alkaloid metabolism. Trends Plant Sci 2001; 6: 212-9. Mahroug S, Burlat V, St Pierre B. Cellular and sub-cellular organisation of the monoterpenoid indole alkaloid pathway in Catharanthus roseus. Phytochem Rev 2007; 6: 363-81. Burlat V, Oudin A, Courtois M, Rideau M, St Pierre B. Coexpression of three MEP pathway genes and geraniol 10-hydroxylase in internal phloem parenchyma of Catharanthus roseus implicates multicellular translocation of intermediates during the biosynthesis of monoterpene indole alkaloids and isoprenoid-derived primary metabolites. Plant J 2004; 38: 131-41. St-Pierre B, Vazquez-Flota FA, De Luca V. Multicellular compartmentation of Catharanthus roseus alkaloid biosynthesis predicts intercellular translocation of a pathway intermediate. Plant Cell 1999; 11: 887-900. Moreno-Valenzuela OA, Minero-Garcia Y, Brito-Argaez L, et al. Immunocytolocalisation of tryptophan decarboxylase in C. roseus hairy roots. Mol Biotechnol 2003; 23: 11-8. Guirimand G, Guihur A, Poutrain P, et al. Spatial organisation of the vindoline biosynthetic pathway in Catharanthus roseus. J Plant Phys 2011; 168: 549-57. Roepke J, Salim V, Wu M, et al. Vinca drug components accumulate exclusively in leaf exudates of Madagascar periwinkle. Proc Natl Acad Sci USA 2010; 107: 15287-92. Goossens A, Rischer H. Implementation of functional genomics for gene discovery in alkaloid producing plants. Phytochemistry Reviews 2007; 6: 35-49. Rischer H, Orei M, Seppänen-Laakso T, et al. Gene-tometabolite networks for terpenoid indole alkaloid biosynthesis in Catharanthus roseus cells. Proc Natl Acad Sci USA 2006; 103; 5614-9. De Geyter N, Gholami A, Goormachtig S, Goossens A. Transcriptional machineries in jasmonate-elicited plant secondary metabolism. Trends Plant Sci 2012; 17: 349-59. Zenk MH, Juenger M. Evolution and current status of the phytochemistry of nitrogenous compounds. Phytochemistry 2007; 68: 2757-72. Son JK, Rosazza JPN, Duffel MW. Vinblastine and vincristine are inhibitors of monoamine oxidase B. J Med Chem 1990; 33: 1845-8. Li S, Fu Y, Zu Y, et al. Determination of paclitaxel and other six taxoids in Taxus species by high-performance liquid chromatography-tandem mass spectrometry. J Pharm Biomed Anal 2009; 49: 81-9 Gerasimenko I, Sheludko Y, Unger M, Stöckigt J. Development of an efficient system for the separation of indole alkaloids by high performance liquid chromatography and its applications. Phytochem Anal 2001; 12; 96-103. Fakhari AR, Nojavan S, Ebrahimi SN, Evenhuis CJ. Optimized ultrasound-assisted extraction procedure for the analysis of opium alkaloids in papaver plants by cyclodextrin-modified capillary electrophoresis. J Sep Sci 2010; 33: 2153-9. Aehle E, Dräger B. Tropane alkaloid analysis by chromatographic and electrophoretic techniques: An update. J Chromatogr B 2010; 878: 1391-406. Russo R, Guillarme D, Rudaz S, Bicchi C, Veuthey JL. Evaluation of the coupling between ultra performance liquid chromatography and evaporative light scattering detector for selected phytochemical applications. J Sep Sci 2008; 31: 2377-87. Peng CA, Ferreira JFS, Wood AJ. Direct analysis of artemisinin from Artemisia annua L. using high-performance liquid chromatography with evaporative light scattering detector, and gas chromatography with flame ionization detector. J Chromatogr A 2006; 1133: 254-8.

5660 Current Pharmaceutical Design, 2013, Vol. 19, No. 31 [184]

[185]

[186] [187]

[188]

[189]

[190] [191]

Kikura-Hanajiri R, Kaniwa N, Ishibashia M, Makinob Y, Kojima S. Liquid chromatographic-atmospheric pressure chemical ionization mass spectrometric analysis of opiates and metabolites in rat urine after inhalation of opium. J Chromatogr B 2003; 789; 139-50. Lee SY, Kim HU, Park JH, Park JM, Kim TY. Metabolic engineering of microorganisms: general strategies and drug production. Drug Discovery Today 2009; 14: 78-88. Ulrich-Merzenich G, Panek D, Zeitler H, Vetter H, Wagner H. Drug development from natural products: Exploiting synergistic effects. Indian J Exp Biol 2010; 48; 208-19. Wang L, Zhang Y, He HP, Zhang Q, Li SF, Hao XJ. Three new terpenoid indole alkaloids from Catharanthus roseus. Planta Med 2011; 77: 754-8. Gu D, Yang Y, Abdulla R, Aisa HA. Characterization and identification of chemical compositions in the extract of Artemisia rupestris L. by liquid chromatography coupled to quadrupole time-offlight tandem mass spectrometry. Rapid Communications in Mass Spectrometry 2011; 26: 83-100. Julsing MK, Koulman A, Woerdenbag HJ, Quax WJ, Kayser O. Combinatorial biosynthesis of medicinal plant secondary metabolites. Biomol Eng 2006; 23: 265-79. Wildung MR, Croteau R. A cDNA clone for taxadiene synthase, the diterpene cyclase that catalyzes the committed step of Taxol biosynthesis. J Biol Chem 1996; 271: 9210-04. Jennewein S, Long RM, Williams RM, Croteau R. Cytochrome P450 taxadiene 5
-hydroxylase, a mechanistically unusual monooxygenase catalyzing the first oxygenation step of taxol biosynthesis. Chem Biol 2004; 11: 379-87.

Received: November 30, 2012

Accepted: January 31, 2013

Rischer et al. [192]

[193] [194]

[195] [196]

[197] [198]

[199] [200]

Chau M, Croteau R. Molecular cloning and characterization of a cytochrome P450 taxoid 2
-hydroxylase involved in taxol biosynthesis. Arch Biochem Biophys 2004; 427: 48-57. Chau M, Jennewein S, Walker K, Croteau R. Taxol biosynthesis: molecular cloning and characterization of a cytochrome P450 taxoid 7-hydroxylase. Chem Biol 2004; 11: 663-72. Walker K, Klettke K, Akiyama T, Croteau R. Cloning, heterologous expression and characterization of a phenylalanine aminomutase involved in taxol biosynthesis. J Biol Chem 2004; 279: 5394754. Gritsanapan W, Griffin WJ Alkaloids and metabolism in Duboisia hybrid. Phytochemistry 1992; 31: 471-7. Oksman-Caldentey K-M, Vuorela H, Isenegger M, Strauss A, Hiltunen R. Selection for high tropane alkaloid content in Hyoscyamus muticus. Plant Breed 1987; 99: 318-26. Christen P, Roberts MF, Phillipson JD, Evans WC. High yield production of tropane alkaloids by hairy root cultures of a Datura candida hybrid. Plant Cell Rep 1989; 8: 75-7. Mano Y, Okhawa H, Yamada Y. Production of tropane alkaloids by hairy root cultures of Duboisia leichhardtii transformed by Agrobacterium rhizogenes. Plant Sci 1989; 59: 191-201. Yukimune Y, Yasuhiro H, Yasuyuki Y. Tropane alkaloid production in root cultures of Duboisia myoporoides obtained by repeated selection. Biosci Biotechnol Biochem 1994; 58: 1443-6. Farrow SC, Hagel JM, Facchini PJ. Transcript and metabolite profiling in cell cultures of 18 plant species that produce benzylisoquinoline alkaloids. Phytochemistry 2012; 77: 79-88.