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Qiao, J. Spermatophyte Sesquiterpene Synthases. Encyclopedia. Available online: (accessed on 14 April 2024).
Qiao J. Spermatophyte Sesquiterpene Synthases. Encyclopedia. Available at: Accessed April 14, 2024.
Qiao, Jianjun. "Spermatophyte Sesquiterpene Synthases" Encyclopedia, (accessed April 14, 2024).
Qiao, J. (2021, August 10). Spermatophyte Sesquiterpene Synthases. In Encyclopedia.
Qiao, Jianjun. "Spermatophyte Sesquiterpene Synthases." Encyclopedia. Web. 10 August, 2021.
Spermatophyte Sesquiterpene Synthases

Sesquiterpenes are important defense and signal molecules for plants to adapt to the environment, cope with stress, and communicate with the outside world, and their evolutionary history is closely related to physiological functions. In this study, the information of plant sesquiterpene synthases (STSs) with identified functions were collected and sorted to form a dataset containing about 500 members. The phylogeny of spermatophyte functional STSs was constructed based on the structural comparative analysis to reveal the sequence–structure–function relationships. We propose the evolutionary history of plant sesquiterpene skeletons, from chain structure to small rings, followed by large rings for the first time and put forward a more detailed function-driven hypothesis. Then, the evolutionary origins and history of spermatophyte STSs are also discussed. In addition, three newly identified STSs CaSTS2, CaSTS3, and CaSTS4 were analyzed in this functional evolutionary system, and their germacrene D products were consistent with the functional prediction. This demonstrates an application of the structure-based phylogeny in predicting STS function. 

spermatophyte sesquiterpene synthetase sesquiterpene phylogenetic analysis functional evolution

1. Introduction

Terpenes constitute a large class of chemically and structurally diverse natural products in the plant kingdom and serve multiple physiological and ecological functions. At present, more than 25,000 terpenoid structures and 80,000 compounds have been found [1]. Sesquiterpenes are the most complex group with structural diversity, and more than 300 kinds of basic skeletons have been found, which are widely distributed in plants and microorganisms [2]. Sesquiterpenes play important roles in interactions with pollinators and seed dispersers [3], direct defenses against herbivores [4] and pathogens [5], mediate plant–plant and plant–microbe interactions [6], and help acclimation to biotic and abiotic environmental stress [7].
The main structure of sesquiterpene is biosynthesized by STSs, which convert farnesyl diphosphate (FPP) into the sesquiterpene skeleton [8]. Common to all STSs is the formation of highly reactive carbocationic intermediates which can undergo a great variety of rearrangements resulting in a huge number of different sesquiterpene structures [9]. It is initiated by the divalent metal ion-dependent ionization of the substrate FPP, and the following cyclization reactions depending on which carbon−carbon double bond reacts with the initially formed allylic carbocation (Figure 1) [10]. One type involves cyclization of farnesyl cation to yield (E,E)-germacradienyl cation (C10-C1 closure) or (E)-humulyl cation (C11-C1 closure) rings. The other type is initiated by isomerization of the C2–C3 double bond of farnesyl cation to the tertiary nerolidyl cation. Then, the cisoid conformer of nerolidyl cation can undergo cyclization to either the central or distal double bond forming bisabolyl cation (C6-C1 closure), cycloheptanyl cation (C7-C1 closure), (Z,E)-germacradienyl cation (C10-C1 closure), or (Z)-humulyl cation (C11-C1 closure). The resulting cationic intermediate undergoes deprotonation or addition of water before termination sesquiterpene [1].
Figure 1. The reaction mechanism of sesquiterpene production starting with FPP. Farnesyl cation is formatted by losing the diphosphate moiety (OPP). Then, the farnesyl cation can be converted to the nerolidyl cation. Possible cyclizations for both cations are indicated in the figure.
The ancestral bifunctional diterpene synthases having γβα tri-domain architecture, presumably catalyzing production of gibberellin intermediate ent-kaurene, seem to the ancestor of all plant terpene synthases (TPS) [11]. The ancient gene duplication and sub-functionalization led to separate class II diterpene cyclases (γβα tri-domain) and subsequently class I TPSs (βα didomain) including the STSs [12]. At present, most studies on STS evolution focused on the phylogenetic location of STSs [13][14]. The TPS family is classified into seven subfamilies designated as TPS-a, TPS-b, TPS-c, TPS-d, TPS-e/f, TPS-g, and TPS-h [11]. Among these, the STSs are predominantly distributed in the angiosperm-specific TPS-a clade and the gymnosperm-specific TPS-d clade [12], and there are few STSs in the TPS-b, TPS-g, and TPS-e/f subfamilies [11]. However, functional evolution of plant STSs has not yet been clarified. It is difficult to predict enzyme function from STS sequences, because STSs represent a very diverse set of enzymes with a wide range of sequence similarities. Janani et al. gathered 262 plant STS sequences with experimentally characterized products, hoping to choose likely functional residues for mutagenesis studies; unfortunately, they did not reveal any general rules for STS function prediction [15].

2. Current Insights

Along with the species evolution, plants have evolved to produce a different collection of terpenenes to accommodate their biotic and abiotic environment [16]. The plant sesquiterpenes gradually formed and might be a potential result of escalating defense and counter-defense between plants and specialized herbivores [17][18]. Exploring the evolutionary origin and history of STSs can not only help to understand the evolutionary pattern and reaction mechanism, but also preliminarily predict the function of STSs. At present, there are many phylogenetic analyses of plant TPSs, most of which focus on taxonomic studies to indicate the evolutionary behavior of TPSs among and within species [19][20], without functional selection of sequences. In this study, all 394 spermatophyte STSs were divided into five distinct groups according to structure-based phylogenetic analysis to explore the evolutionary patterns of plant STSs and sesquiterpenes.
Many genes are involved in sesquiterpene biosynthesis in the genome of each plant species, which also provides a large platform for the evolution of new sesquiterpenes via gene duplication and subfunctionalization [21][22]. Intron loss, mutations, and coevolution with natural enemies are considered to be the most important evolutionary dynamics of STSs. Evolution in STSs is often the result of intron loss or mutations that lead to subfunctionalization or function loss [23]. For example, a large fragment loss of δ-selinene synthase 2 (Agsel2) from A. grandis was found around intron X that led to Agsel2 being transcribed as a pseudogene [24]. Single amino acid W279A switch converts δ-cadinene synthase (CAD1-A) into germacradien-4-ol synthase [25]. New sesquiterpenes keep arising in specific plant lineages, potentially as an outcome of coevolution with natural enemies [17]. Different plant lineages have evolved the ability to make additional “specialized” metabolites that are implicated in defense or the attraction of beneficial organisms, which indicates a dominant process dynamic evolution in STSs to the chemical diversity in plants.
STSs have various evolutionary forms. It can also be expected that STSs with altered subcellular localization and new substrate specificities would have evolved. Although TPSs often have broad substrate specificity and accept GPP, FPP, or GGPP in vitro, their function may be narrower in planta due to their subcellular localization [26]. Monoterpene synthases and diterpene synthases typically contain N-terminus signal peptides and are transported into plastids, STSs, however, are usually found in the cytosol [9]. There is increasing evidence for an exchange of TPS subcellular localization, especially under stress conditions [27][28]. Examples include the AmNES/LIS-1/2 and CsLIS/NES-1/2 analyzed above in the TPS-g subfamily. In this scenario, driven by adaptive evolution, ancestral monoterpene synthases losing the N-terminus signal peptide changed their substrate pool and gradually evolved into STSs. Models for gymnosperm TPS evolution proposed that STSs evolved from diterpene synthases through loss of introns, which resulted in, among other changes, the complete loss of the γ domain [24]. Based on this model, Abies grandis a-bisabolene synthase Ag1 (C6-C1 closure), a three-domain plant STSs, is potentially an intermediate in the evolutionary history from diterpene to sesquiterpene synthase [29].
Distinct catalytic features of the STSs arose early in spermatophyte evolution and the reactions have become more complex over time. In the evolution of STSs, it is easier to form acyclic sesquiterpenes than cyclic sesquiterpenes according to phylogenetic analysis. Acyclic sesquiterpenes were formed directly from farnesyl cation or nerolidyl cation by proton loss or addition of water [9]. For cyclic sesquiterpenes, they can be formed by typically catalyzing reaction cascades with additional steps, such as the isomerization of carbon–carbon double bond in the initial cation to allow alternate ring closures or additional cyclization [9]. Successive gene duplications and the subsequent accumulation of mutations led to the multitude of STSs, many of which catalyze more complex reactions than the ancestor. Although it is universally accepted that evolution of natural product biosynthesis has led to the formation of more and more complex structures, this process has rarely been documented at the level of a specific enzyme and plant group [30]. Overall, we speculated the early possible evolutionary process of spermatophyte STSs is from acyclic sesquiterpenes to cyclic sesquiterpenes, and the C6-C1 closure sesquiterpenes (small rings) may have formed earlier than C10-C1/C11-C1 closure sesquiterpenes. Interestingly, in some specific STSs, evolution may stop at a certain stage to form a series of characteristic metabolites under certain selective pressures. For example, Artemisia species’ STSs are clustered in the A1 clade of the TPS-a subfamily (Figure 2) and obviously originated from a common ancestor dedicated to producing 1,6-cyclized sesquiterpenes. Among these, AaADS from Artemisia annua produces the artemisinin specific intermediate amorpha-4,11-diene (C6-C1 cyclized bicyclic sesquiterpenes) [31]; however, other STSs highly homologous to ADS from Artemisia species cyclize FPP to (+)-a-bisabolol (C6-C1 monocyclic sesquiterpenes) [32].
Figure 2. The phylogeny of spermatophyte STSs in the TPS-a subfamily. Blue shadow represents the Rosanae STSs, yellow shadow represents the Asterids STSs, red shadow represents the monocot STSs, and orange shadow represents the magnoliid STSs. Red branches show the STSs with identical products, blue branches show the STSs with 1,11-cyclized functions and green branches show the probable HGT members from Santalum album. STSs producing 1,6-cyclized sesquiterpenes are marked with grey dots, and the neighbor STSs producing 1,10- or 1,11-cyclized products, respectively, are marked with black rectangles.
Systematic study on the evolutionary changes of STS structure is an effective way to elucidate its function. Over the last three decades, high-resolution crystal structures have become available for STSs, and the enzymatic structure–function relationships have revealed the evolutionary relationships of STSs [1][12]. It was recognized early that the structure of STS products depending on the initial substrate conformation imposed by the enzymatic active site cavity [33]. For example, SaSQS2 is the representative of C6-C1 cyclized STSs, and the shape of FPP conformation is close to natural straight chain in its active site cavity [34]. For C10-C1 cyclized STSs of TEAS [35] and XC1 [36], the shapes of FPP are obviously curved. For the formation of acyclic sesquiterpene, we choose the medium/long-chain-length prenyl pyrophosphate synthase as the representative because there is no crystal structure of acyclic STSs, in which the FPP conformation is almost natural straight chain [37] (Figure S2). Overall, in the evolution of STSs, the change of residues in the active site cavity made the straight chain FPP gradually bend, which made the C1 carbocation gradually approach the intramolecular double bond, and endowed STSs with the ability to form small ring and even large ring sesquiterpenes.
Horizontal gene transfer (HGT) also plays an important role in the evolution of STSs [38]. Sixteen Santalum STSs have been functionally charactered (Table S3), among which, 13 STSs are clustered in TPS-b and their products are mainly C6-C1 cyclized sesquiterpene β-bisabolene (Figure 3b), while three STSs are clustered in the R1 clade of the TPS-a subfamily and their products are mainly C10-C1 cyclized sesquiterpenes germacrene D-4-ol, hedycaryol and C11-C1 cyclized sesquiterpene a-humulene (Figure 2). This indicates that the evolutionary sources of STSs in these two parts are completely different. Sandalwood is a semi-parasitic plant, whose survival is inseparable from the host plant. Some studies have shown that sandalwood prefers to parasitize on nitrogen fixing woody plants [39], and HGT events between sandalwood and host plants have also been reported [40][41], which may indicate that STSs in sandalwood come from HGT. These parasitic plants are likely to obtain the synthesis ability of terpenenes and other secondary metabolites from the host through HGT, so as to better adapt to the environment or communicate with the host.
Figure 3. (a) The phylogeny of spermatophyte STSs in the TPS-g subfamily. STSs with diterpene synthases function in vitro are shown by red branches and uniformly distributed in the phylogenetic tree. (b) The phylogeny of spermatophyte STSs in the TPS-b subfamily. STSs with monoterpene synthases function in vitro are shown by red branches. STSs producing 1,6-cyclized sesquiterpenes and 1,10/1,11-cyclized sesquiterpenes are marked with grey dots and red dots, respectively. Products of STSs marked with black rectangles are acyclic farnesene or nerolidol.
In this comprehensive analysis, on the one hand, we collected the information of plant STSs with identified functions and constructed the phylogeny of plant functional STSs based on the structural comparative analysis, to reveal the sequence–structure–function relationships. On the other hand, we highlighted our incomplete understanding of the evolutionary pattern of sesquiterpenes in spermatophytes (Figure 4), from chain structure to small rings, followed by large rings for the first time, and discussed the evolutionary origins and history of STSs from spermatophyte plants. Then, we proved our evolutionary pattern is useful in predicting the function of STSs by three germacrene D synthases.
Figure 4. The proposed evolutionary history of spermatophyte STSs. Blue boxes represent ancestral TPSs, red boxes represent STSs, and purple boxes represent transition TPSs. The character “M” represents monoterpene synthases and the character “S” represents STSs. The red words represent the examples of STSs, the green words represent functional products of STSs.


  1. Christianson, D.W. Structural and Chemical Biology of Terpenoid Cyclases. Chem. Rev. 2017, 117, 11570–11648.
  2. Fraga, B.M. Natural sesquiterpenoids. Nat. Prod. Rep. 2013, 30, 1226–1264.
  3. Gershenzon, J.; Dudareva, N. The function of terpene natural products in the natural world. Nat. Chem. Biol. 2007, 3, 408–414.
  4. Rasmann, S.; Köllner, T.G.; Degenhardt, J.; Hiltpold, I.; Toepfer, S.; Kuhlmann, U.; Gershenzon, J.; Turlings, T.C.J. Recruitment of entomopathogenic nematodes by insect-damaged maize roots. Nature 2005, 434, 732–737.
  5. Dreher, D.; Baldermann, S.; Schreiner, M.; Hause, B. An arbuscular mycorrhizal fungus and a root pathogen induce different volatiles emitted by Medicago truncatula roots. J. Adv. Res. 2019, 19, 85–90.
  6. Agrawal, A.A.; Heil, M. Synthesizing specificity: Multiple approaches to understanding the attack and defense of plants. Trends Plant Sci. 2012, 17, 239–242.
  7. Erb, M. Volatiles as inducers and suppressors of plant defense and immunity-origins, specificity, perception and signaling. Curr. Opin. Plant Biol. 2018, 44, 117–121.
  8. Tholl, D.; Lee, S. Terpene Specialized Metabolism in Arabidopsis thaliana. Arab. Book 2011, 9, e0143.
  9. Degenhardt, J.; Kllner, T.G.; Gershenzon, J. Monoterpene and sesquiterpene synthases and the origin of terpene skeletal diversity in plants. Phytochemistry 2009, 69, 1621–1637.
  10. Davis, E.M.; Croteau, R. Cyclization enzymes in the biosynthesis of monoterpenes, sesquiterpenes, and diterpenes. In Biosynthesis: Aromatic Polyketides, Isoprenoids, Alkaloids; Leeper, F.J., Vederas, J.C., Eds.; Topics in Current Chemistry; Springer: Berlin/Heidelberg, Germany, 2000; Volume 209, pp. 53–95.
  11. Cao, R.; Zhang, Y.; Mann, F.M.; Huang, C.; Mukkamala, D.; Hudock, M.P.; Mead, M.E.; Prisic, S.; Wang, K.; Lin, F.-Y.; et al. Diterpene cyclases and the nature of the isoprene fold. Proteins-Struct. Funct. Bioinform. 2010, 78, 2417–2432.
  12. Gao, Y.; Honzatko, R.B.; Peters, R.J. Terpenoid synthase structures: A so far incomplete view of complex catalysis. Nat. Prod. Rep. 2012, 29, 1153–1175.
  13. Dudareva, N.; Martin, D.; Kish, C.M.; Kolosova, N.; Gorenstein, N.; Faldt, J.; Miller, B.; Bohlmann, J. (E)-beta-ocimene and myrcene synthase genes of floral scent biosynthesis in snapdragon: Function and expression of three terpene synthase genes of a new terpene synthase subfamily. Plant Cell 2003, 15, 1227–1241.
  14. Martin, D.M.; Faldt, J.; Bohlmann, J. Functional characterization of nine Norway spruce TPS genes and evolution of gymnosperm terpene synthases of the TPS-d subfamily. Plant Physiol. 2004, 135, 1908–1927.
  15. Durairaj, J.; Di Girolamo, A.; Bouwmeester, H.J.; de Ridder, D.; Beekwilder, J.; van Dijk, A.D.J. An analysis of characterized plant sesquiterpene synthases. Phytochemistry 2019, 158, 157–165.
  16. Pichersky, E.; Raguso, R.A. Why do plants produce so many terpenoid compounds? New Phytol. 2018, 220, 692–702.
  17. Richards, L.A.; Dyer, L.A.; Forister, M.L.; Smilanich, A.M.; Dodson, C.D.; Leonard, M.D.; Jeffrey, C.S. Phytochemical diversity drives plant-insect community diversity. Proc. Natl. Acad. Sci. USA 2015, 112, 10973–10978.
  18. Firn, R.D.; Jones, C.G. Natural products—A simple model to explain chemical diversity. Nat. Prod. Rep. 2003, 20, 382–391.
  19. Chen, F.; Tholl, D.; Bohlmann, J.; Pichersky, E. The family of terpene synthases in plants: A mid-size family of genes for specialized metabolism that is highly diversified throughout the kingdom. Plant J. 2011, 66, 212–229.
  20. Martin, D.M.; Bohlmann, J. Identification of Vitis vinifera (-)-alpha-terpineol synthase by in silico screening of full-length cDNA ESTs and functional characterization of recombinant terpene synthase. Phytochemistry 2004, 65, 1223–1229.
  21. Martin, D.M.; Aubourg, S.; Schouwey, M.B.; Daviet, L.; Schalk, M.; Toub, O.; Lund, S.T.; Bohlmann, J. Functional Annotation, Genome Organization and Phylogeny of the Grapevine (Vitis vinifera) Terpene Synthase Gene Family Based on Genome Assembly, FLcDNA Cloning, and Enzyme Assays. BMC Plant Biol. 2010, 10.
  22. Ma, L.-T.; Lee, Y.-R.; Liu, P.-L.; Cheng, Y.-T.; Shiu, T.-F.; Tsao, N.-W.; Wang, S.-Y.; Chu, F.-H. Phylogenetically distant group of terpene synthases participates in cadinene and cedrane-type sesquiterpenes accumulation in Taiwania cryptomerioides. Plant Sci. 2019, 289.
  23. Hillwig, M.L.; Xu, M.; Toyomasu, T.; Tiernan, M.S.; Wei, G.; Cui, G.; Huang, L.; Peters, R.J. Domain loss has independently occurred multiple times in plant terpene synthase evolution. Plant J. 2011, 68, 1051–1060.
  24. Trapp, S.C.; Croteau, R.B. Genomic organization of plant terpene synthases and molecular evolutionary implications. Genetics 2001, 158, 811–832.
  25. Loizzi, M.; Gonzalez, V.; Miller, D.J.; Allemann, R.K. Nucleophilic Water Capture or Proton Loss: Single Amino Acid Switch Converts -Cadinene Synthase into Germacradien-4-ol Synthase. Chembiochem 2018, 19, 100–105.
  26. Pazouki, L.; Niinemets, U. Multi-Substrate Terpene Synthases: Their Occurrence and Physiological Significance. Front. Plant Sci. 2016, 7.
  27. Gutensohn, M.; Orlova, I.; Nguyen, T.T.H.; Davidovich-Rikanati, R.; Ferruzzi, M.G.; Sitrit, Y.; Lewinsohn, E.; Pichersky, E.; Dudareva, N. Cytosolic monoterpene biosynthesis is supported by plastid-generated geranyl diphosphate substrate in transgenic tomato fruits. Plant J. 2013, 75, 351–363.
  28. Dong, L.; Jongedijk, E.; Bouwmeester, H.; Van Der Krol, A. Monoterpene biosynthesis potential of plant subcellular compartments. New Phytol. 2016, 209, 679–690.
  29. McAndrew, R.P.; Peralta-Yahya, P.P.; DeGiovanni, A.; Pereira, J.H.; Hadi, M.Z.; Keesling, J.D.; Adams, P.D. Structure of a Three-Domain Sesquiterpene Synthase: A Prospective Target for Advanced Biofuels Production. Structure 2011, 19, 1876–1884.
  30. Luck, K.; Chen, X.; Norris, A.M.; Chen, F.; Gershenzon, J.; Koellner, T.G. The reconstruction and biochemical characterization of ancestral genes furnish insights into the evolution of terpene synthase function in the Poaceae. Plant Mol. Biol. 2020, 104, 203–215.
  31. Chang, Y.J.; Song, S.H.; Park, S.H.; Kim, S.U. Amorpha-4,11-diene synthase of Artemisia annua: cDNA isolation and bacterial expression of a terpene synthase involved in artemisinin biosynthesis. Arch. Biochem. Biophys. 2000, 383, 178–184.
  32. Muangphrom, P.; Seki, H.; Suzuki, M.; Komori, A.; Nishiwaki, M.; Mikawa, R.; Fukushima, E.O.; Muranaka, T. Functional Analysis of Amorpha-4,11-Diene Synthase (ADS) Homologs from Non-Artemisinin-Producing Artemisia Species: The Discovery of Novel Koidzumiol and (+)-alpha-Bisabolol Synthases. Plant Cell Physiol. 2016, 57, 1678–1688.
  33. Tantillo, D.J. Biosynthesis via carbocations: Theoretical studies on terpene formation. Nat. Prod. Rep. 2011, 28, 1035–1053.
  34. Blank, P.N.; Shinsky, S.A.; Christianson, D.W. Structure of Sesquisabinene Synthase 1, a Terpenoid Cyclase That Generates a Strained 3.1.0 Bridged-Bicyclic Product. Acs Chem. Biol. 2019, 14, 1011–1019.
  35. Koo, H.J.; Vickery, C.R.; Xu, Y.; Louie, G.V.; O’Maille, P.E.; Bowman, M.; Nartey, C.M.; Burkart, M.D.; Noel, J.P. Biosynthetic potential of sesquiterpene synthases: Product profiles of Egyptian Henbane premnaspirodiene synthase and related mutants. J. Antibiot. 2016, 69, 524–533.
  36. Gennadios, H.A.; Gonzalez, V.; Di Costanzo, L.; Li, A.; Yu, F.; Miller, D.J.; Allemann, R.K.; Christianson, D.W. Crystal Structure of (+)-delta-Cadinene Synthase from Gossypium arboreum and Evolutionary Divergence of Metal Binding Motifs for Catalysis. Biochemistry 2009, 48, 6175–6183.
  37. Hsieh, F.-L.; Chang, T.-H.; Ko, T.-P.; Wang, A.H.J. Structure and Mechanism of an Arabidopsis Medium/Long-Chain-Length Prenyl Pyrophosphate Synthase. Plant Physiol. 2011, 155, 1079–1090.
  38. Jia, Q.; Li, G.; Kollner, T.G.; Fu, J.; Chen, X.; Xiong, W.; Crandall-Stotler, B.J.; Bowman, J.L.; Weston, D.J.; Zhang, Y.; et al. Microbial-type terpene synthase genes occur widely in nonseed land plants, but not in seed plants. Proc. Natl. Acad. Sci. USA 2016, 113, 12328–12333.
  39. Suetsugu, K.; Kawakita, A.; Kato, M. Host range and selectivity of the hemiparasitic plant Thesium chinense (Santalaceae). Ann. Bot. 2008, 102, 49–55.
  40. Rice, D.W.; Alverson, A.J.; Richardson, A.O.; Young, G.J.; Virginia Sanchez-Puerta, M.; Munzinger, J.; Barry, K.; Boore, J.L.; Zhang, Y.; dePamphilis, C.W.; et al. Horizontal Transfer of Entire Genomes via Mitochondrial Fusion in the Angiosperm Amborella. Science 2013, 342, 1468–1473.
  41. Barkman, T.J.; McNeal, J.R.; Lim, S.-H.; Coat, G.; Croom, H.B.; Young, N.D.; dePamphilis, C.W. Mitochondrial DNA suggests at least 11 origins of parasitism in angiosperms and reveals genomic chimerism in parasitic plants. Bmc Evol. Biol. 2007, 7.
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