Identification and characterisation of a mads box gene from rafflesia cantleyi solms laubach (rafflesiaceae)

IDENTIFICATION AND CHARACTERISATION OF A MADS-BOX GENE FROM RAFFLESIA CANTLEYI SOLMS-LAUBACH (RAFFLESIACEAE) PHUA EK KIAN, EDWIN (B.Sc., NUS) A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF SCIENCE DEPARTMENT OF BIOLOGICAL SCIENCES NATIONAL UNIVERSITY OF SINGAPORE 2010 Acknowledgements My deepest gratitude to my supervisor Associate Professor Hugh Tan Tiang Wah, and my co-supervisor Professor Prakash P. Kumar, for they have been most patient and understanding, and who believed in me and pushed me right to the end, despite a very long and tiring candidature. I need to thank the Economic Planning Unit, Prime Minister’s Office, Government of Malaysia for permission to collect Rafflesia cantleyi buds from Pulau Tioman, Pahang, Peninsular Malaysia; and Associate Professor Lim Saw Hoon, formerly of the Malaysia University of Science and Technology for her help in this project. I would like to thank Ang Kai Yang, Reuben Clements Gopalasamy, Norman Lim T-Lon, and Alvin Lok for their expertise in the field and in help with the collection of the Rafflesia flowers. I would also like to thank Dr. Rengasamy Ramamoorthy for his invaluable help and expertise in the laboratory. Last, but not least, I thank all my colleagues and labmates from the Plant Systematics Laboratory and the Plant Morphogenesis Laboratory, and friends in the Department of Biological Sciences for their support, advice, and help! I could not have done this without you! i Table of Contents Page Acknowledgements i Table of Contents ii Summary iv List of Abbreviations v List of Tables vii List of Figures viii Chapter 1: General Introduction 1 Chapter 2: Literature Review 2.1. Rafflesia R.Br. 5 2.1.1. Floral morphology Rafflesia 5 2.1.2. Rafflesia evolution and systematics 6 2.1.3 Molecular studies in Rafflesia 8 2.1.4. Rafflesia cantleyi Solms-Laubach 8 2.2. MADS-box genes 8 2.2.1. Floral organ identity genes 12 2.2.2. Flowering time genes 15 2.3. Heterologous expression system for functional analysis of genes 17 Chapter 3: Material and Methods 3.1. Plant Materials 18 3.2. RNA and DNA isolation 18 3.3. Reverse transcription 20 3.4. PCR amplification 20 3.5. Cloning of PCR products 22 3.6. Plasmid DNA purification 23 3.7. DNA sequencing 24 3.8. Sequence analysis 24 3.9. Phylogenetic analysis 25 3.10. Rapid amplification of cDNA ends 25 ii 3.11. Preparation of ectopic expression construct 25 3.12. Transformation of Agrobacterium tumefaciens 26 3.13. Genetic transformation of Arabidopsis thaliana 28 3.14. Quantitative real-time PCR analysis 29 3.15. Genomic Southern blot analysis 29 Chapter 4: Results and Discussion 4.1. Collection of Rafflesia cantleyi flower buds 32 4.2. RNA isolation 32 4.3. DNA isolation 35 4.4. Cloning MADS-box genes by RT-PCR 35 4.5. Phylogenetic analysis 40 4.6. Functional characterisation of 35S::RcMADS1 in A. thaliana 41 4.6.1. Construction of ectopic expression plasmid 41 4.6.2. Transgenic Arabidopsis thaliana T1 phenotypes 43 4.6.3. 35S::RcMADS1 effects on flowering time 45 4.6.4. Floral morphology in 35S::RcMADS1 transgenic lines 47 4.7. Molecular characterisation of selected transgenic lines 4.7.1. Genomic Southern blot analysis 53 4.7.2. Quantitative real-time PCR analysis 53 Chapter 5: General Discussion and Future Work 5.1. RcMADS1 may be involved in regulation of flowering time 60 5.2. Temporal and spatial expression of RcMADS1 in Rafflesia cantleyi 61 5.3. Future work 62 5.4. Conclusions 62 References 64 iii SUMMARY Rafflesia is a distinctive genus of holoparasitic endophytes found only in the Indo-Malayan region, with highly reduced vegetative morphology, and usually manifest only as large, fleshy flowers on their host plants. Despite its distinct ecology and morphology, very little is known about this taxon, including information on the molecular processes of floral development. The MADS-box genes, which encode transcription factors sharing a highly conserved MADS domain are known as the key regulatory genes that mediate flower development. Therefore, in an attempt to learn more about molecular floral development in Rafflesia, we cloned and characterised a MADS-box cDNA, RcMADS1, from Rafflesia cantleyi. Using RNA from flower buds of Rafflesia cantleyi, we performed RT-PCR with degenerate primers specific for the MADS domain, followed by 5′-RACE. This yielded a cDNA of 951 bp (named RcMADS1), encoding a polypeptide of 228 amino acids. Sequence analysis of this polypeptide revealed about 57% similarity to AGL24 and SVP, two proteins from Arabidopsis thaliana that are involved in mediating various flowering signals. Phylogenetic analysis showed RcMADS1 to be nested in the StMADS11 clade, together with AGL24 and SVP. Ectopic expression of RcMADS1 in Arabidopsis thaliana as a heterologous system produced several independent lines of transgenic plants that showed alterations in flowering time and floral morphology in a dose-dependent manner, similar to the phenotypes observed when AGL24 is overexpressed. Our data suggest that RcMADS1 may be functionally similar to AGL24. iv List of Abbreviations Chemicals and Reagents CTAB cetyltrimethylammonium bromide DEPC diethyl pyrocarbonate DTT dithiothreitol EDTA ethylenediaminetetraacetic acid HCl hydrochloric acid LB lysogeny broth ( = Luria Bertani) MgCl2 magnesium chloride NaCl sodium chloride NaOH sodium hydroxide PVPP polyvinyl polypyrrolidone SDS sodium dodecyl sulfate SSC standard saline citrate TBE Tris borate EDTA TE Tris EDTA Tris tris(hydroymethyl)aminomethane Units and Measurements bp base pairs cm centimeter cm2 square centimeter g gram g centrifugal force h hour l litre M molar ( = moles per litre) min minute mg milligram mJ millijoule ml millilitre mM millimolar v mm ng nanogram pmol picomole rpm revolutions per minute s second V volt v/v volume per volume w/v weight per volume °C degree Celsius µg microgram µl microlitre Others BLAST Basic Local Alignment Search Tool CaMV cauliflower mosaic virus cDNA complementary deoxyribonucleic acid DNA deoxyribonucleic acid dNTP deoxynucleoside triphosphate mRNA messenger ribonucleic acid OD optical density oligo(dT) oligodeoxythymidylic acid PCR polymerase chain reaction pH potential of hydrogen RACE rapid amplification of cDNA ends RNA ribonucleic acid RT-PCR reverse transcription polymerase chain reaction UTR untranslated region UV ultraviolet vi List of Tables Page Table 3.1 Degenerate primers used in cloning MADS-box genes from Rafflesia cantleyi. 21 Table 3.2 Primer pairs used in quantitative real-time PCR. 30 Table 4.1 Phenotype analysis of T1 transgenic plants generated. 44 Table 4.2 Comparison of flowering times of 35S::RcMADS1 T2 lines. 46 vii List of Figures Page Figure 2.1 Schematic representation of the structure of plant MIKC-type MADS-box genes. 11 Figure 3.1 Schematic diagram of expression construction. 27 Figure 4.1 Rafflesia cantleyi Solms-Laubach buds. 33 Figure 4.2 Gel electrophoresis of total RNA extracted from young Rafflesia cantleyi flower bud (~1 cm in diameter). 34 Figure 4.3 Cloning of MADS-box genes via degenerate PCR. 36 Figure 4.4 Structure of RcMADS1 cDNA. 38 Figure 4.5 Alignment of the derived amino acid sequences of RcMADS1 and other members of the StMADS11 clade. 39 Figure 4.6 Phylogenetic tree of MADS-box proteins. 42 Figure 4.7 Phenotype of wild-type-looking transgenic line ETL01. 48 Figure 4.8 Phenotype of strong transgenic line ETL12. 49 Figure 4.9 Phenotype of strong transgenic line ETL14. 50 Figure 4.10 Altered floral morphology due to ectopic expression of 35S::RcMADS1 in Arabidopsis thaliana. 51 Figure 4.11 Secondary inflorescence 35S::RcMADS1 plants. 52 Figure 4.12 Development of profuse branching. 35S::RcMADS1 development ectopic in 54 viii Figure 4.13 Genomic Southern blot analysis. 55 Figure 4.14 Comparison of expression levels of RcMADS1 across transgenic lines. 56 Figure 4.15 Effect of RcMADS1 ectopic expression on FLC and FT. 58 ix CHAPTER 1 GENERAL INTRODUCTION The parasitic plant genus Rafflesia is a distinctive flowering plant genus, highly unusual in the plant kingdom owing to its highly reduced vegetative morphology, prominent and large floral structures, and physiology. Rafflesia species are holoparasitic endophytes — plants that grow completely embedded within their host plants and completely dependent on them for nutrition. Unlike the majority of the flowering plants, they lack leaves, stems, and roots, and only manifest as flowers for sexual reproduction on host plants such as species of Tetrastigma (Kuijt, 1969). Rafflesia has particularly distinctive, large fleshy flowers that can grow up to a metre in diameter, producing the smell of rotting flesh which attracts carrion flies for pollination (Meijer, 1997). Besides being recognised as the largest individual flower among all extant angiosperms, Rafflesia flowers have some unusual structures, such as a modified perianth (perigone) enclosed by a diaphragm; a central column with an apical disk bearing long spike-like structures (processes), and the presence of ramenta, which are fine hairs, on the interior surface of the perigone tube and diaphragm (Meijer, 1997). The genus Rafflesia is confined to the Indo-Malayan region (Meijer, 1997), and has been little researched, with only a few studies (whether molecular or ecological) published in the past 20 years (e.g., Beaman et al., 1988; Nickrent and Starr, 1994; Barkman et al., 2004). There is a lack of extensive work on this genus partly owing to its rarity and the inaccessibility of its habitats. Holoparasitic plants like Rafflesia have many physiological and morphological adaptations as a result of their evolution and have lost many plant structures such as leaves, stems and roots, 1 thus making phylogenetic relationships with non-parasitic plants difficult. Barkman et al. (2004) sequenced the mitochondrial gene matR and produced a broad phylogenetic tree that showed a placement of Rafflesia in the Malpighiales. Rafflesia was later found to be nested in the Euphorbiaceae in a more restricted study of the Malpighiales using more mitochondrial genes and a chloroplastic gene (Davis et al., 2007). It is interesting to note that Rafflesia is considered to have evolved from a family with very small flowers. Because the flower is the only macroscopic structure of the plant that is visible, and that the flowers are highly unusual, Rafflesia can be studied from the molecular development perspective, which can help elucidate the evolutionary processes that Rafflesia has undergone. Flowering is a complex process that involves the regulation of various developmental programs by MADS-box genes, which encode transcription factors containing a highly-conserved MADS-box which is part of the DNA-binding domain (Becker and Theissen, 2003). Plant floral MADS-box genes also have three other domains in addition to the MADS (M) domain: an intervening (I) domain; a keratin-like coiled-coil (K) domain; and a C-terminal (C) domain. Together, these genes have an MIKC structure which is specific to plants (Nam et al., 2003). There are at least nine classes of MADS-box genes based on their function and expression patterns (Nam et al., 2003): classes A, B, C, D, E, F, G, Bs (B-sister), and T. Many of these MADS-box genes control flower formation and are known as floral MADS-box genes (Nam et al., 2003). The ‘ABC’ model of flower formation was originally proposed to explain how the genes (from classes A, B, and C) interact to produce the different organs (Weigel and Meyerowitz, 1994), and this model is being modified and updated as more information from continuing studies point to the involvement of other gene classes in floral development: such as class E genes acting 2 synergistically with combinations of A, B, and C genes to produce petals, stamens and carpels, as well as floral meristem formation (Honma and Goto, 2001); and class D genes are required for ovule development (Favaro et al., 2003). Such genes are being intensely studied using model organisms such as Arabidopsis thaliana (thale cress), Antirrhinum majus (snapdragon), Zea mays (maize), and Oryza sativa (rice). Changes in MADS-box genes are strongly correlated to the evolution of land plant reproductive structures (Theissen et al., 2000). However, model organisms represent only a small portion of the plant kingdom, and many more genes need to be identified before a thorough understanding of the control and evolution of flower development is achieved (Soltis et al., 2002, 2007). Work on many branches of plants have begun to fill in the gaps, such as from bryophytes (e.g., Physcomitrella), ferns (e.g., Ceratopteris), gymnosperms (e.g., Cycas, Gnetum, Ginkgo, Pinus) and basal angiosperms (e.g., Michelia, Piper). The identification and characterisation of MADS-box genes in Rafflesia could fill in some of these gaps in the genetic architecture of floral development, leading to the objectives of this study: 1) To clone one or more MADS-box genes from Rafflesia cantleyi, a species of Rafflesia from Pulau Tioman, Pahang, Malaysia, using a degerate PCR approach; 2) To identify and analyse the cloned MADS-box gene(s) through sequencing and phylogenetic analysis; 3) To characterise the function(s) of the cloned MADS-box gene(s) by studying the effects of ectopic expression in Arabidopsis thaliana plants generated via Agrobacterium-mediated transformation; 3 4) To characterise the molecular processes underlying the function(s) of cloned MADS-box genes using quantitative real-time PCR and other appropriate methods. 4 CHAPTER 2 LITERATURE REVIEW 2.1. Rafflesia R.Br. The genus Rafflesia R.Br. is well-known as a parasitic plant genus, with the largest known flowers in the world (Beaman et al., 1988). All species in Rafflesia are holoparasitic endophytes of the vine Tetrastigma (Vitaceae): the vegetative parts of the plant are wholly embedded inside the tissues of the host plants and completely dependent on their host plants for nutrition. These plants have no visible leaves, stems or roots; and only appear as flowers and fruits during sexual reproduction (Kuijt, 1969). The flowers are often large and can grow up to one metre in diameter, and produce a smell of rotten flesh during anthesis to attract carrion flies for pollination (Beaman et al. 1988). Besides these large flowers, and the unusual mode of animalaided pollination, Rafflesia flowers have unusual morphology. 2.1.1. Floral morphology of Rafflesia Rafflesia flowers are unisexual, where the female flowers possess rudimentary anthers (Meijer, 1997). The perianth is fused, forming a perigone partially closed by a diaphragm at the apex (leaving an aperture). There are five perigone lobes (the ‘petals’) which are reddish and often with white warts. A central column widens into a disk at the apex, which supports processes that are spike-like structures. The processes are hypothesised to radiate heat to aid in dispersal of the odour (that resembles decaying protein) as olfactory cues to attact carrion flies for pollination (Beaman et al., 1988). Underneath the disk is a groove, known as the sulcus; in the 5 male flowers, the anthers are situated under the rim of the disk adjacent to the sulcus (Meijer, 1997). The floral structures of Rafflesia have been thought to be possibly homologous to those in Passiflora (Kuijt, 1969; Barkman et al., 2004): the diaphragm of Rafflesia being homologous to the annular corona of the Passifloraceae; the Rafflesiaceous central column possibly homologous to the Passifloraceous androgynophore, and the Rafflesiaceous perigone tube possibly homologous to the Passifloraceous hypanthium. Phylogenetic data placing Passifloraceae as close relatives to Rafflesiaceae suggests a shared origin for these floral structures (Barkman et al., 2004). 2.1.2. Rafflesia evolution and systematics Because Rafflesia specimens are so rare and often found in remote habitats, and the ecology of Rafflesia is dependent on its host plants, the exact number of Rafflesia species is uncertain. Several species described in the 19th and early 20th centuries are not completely known, owing to incomplete descriptions, or the lack of type specimens (Meijer, 1997); these species include Rafflesia borneensis Koord.; Rafflesia ciliata Koord.; Rafflesia titan Jack; Rafflesia tuan-mudae Becc.; and Rafflesia witkampii Koord. In his treatment, Meijer (1997) accepted 13 species: Rafflesia arnoldii R.Br., with two varieties: Rafflesia arnoldii var. arnoldii R.Br., and Rafflesia arnoldii var. atjehensis (Koord.) Meijer; Rafflesia cantleyi Solms-Laubach; Rafflesia gadutensis Meijer; Rafflesia hasseltii Suringar; Rafflesia keithii Meijer; Rafflesia kerrii Meijer; Rafflesia manillana Teschemacher; Rafflesia micropylora Meijer; Rafflesia patma Blume; Rafflesia pricei Meijer; Rafflesia rochussenii Teijsm. & Binn.; Rafflesia schadenbergiana Göpp.; and Rafflesia tengku-adlinii Salleh & Latiff. Since 2002, 10 or 11 new species have been discovered in the Philippines, as 6 well as two others from outside the Philippines, bringing the number of currently recognised and described Rafflesia species to 27 (Barcelona et al., 2009). The new Philippine species are: Rafflesia baletei Barcelona & Cajano; Rafflesia leonardi Barcelona & Pelser; Rafflesia lobata R.Galang & Madulid; Rafflesia mira Fernando & Ong; Rafflesia philippensis Blanco; and Rafflesia speciosa Barcelona & Fernando. The other newly discovered species since the treatment by Meijer (1997) are Rafflesia azlanii Latiff & M.Wong from Peninsular Malaysia; and Rafflesia bengkuluensis Susatya, Arianto & Mat-Salleh from Sumatra, Indonesia. Owing to the highly unusual morphology and evolution as endophytic holoparasites, the taxonomy and phylogenetic affinities of Rafflesia were not clear. Rafflesia had been grouped together with other parasitic plants (such as Apodanthus, Pilostyles, Cytinus, Bdallophyton, and Mitrastema) in various taxonomic treatments (Meijer, 1997). More recent phylogenetic studies using molecular data had more precisely established the phylogenetic affinities of Rafflesiaceae sensu stricto (comprising Rafflesia, Rhizanthes, and Sapria). Using data from the mitochondrial gene matR from a wide analysis of 95 species of angiosperms and gymnosperms, Barkman et al. (2004) placed Rafflesia and Rhizanthes within the order Malpighiales, with sister families such as Passifloraceae, Salicaceae, and Violaceae. Rafflesiaceae was more confidently placed within the Malpighiales as nested in Euphorbiaceae using more data (five mitochondrial and one chloroplastic genes) from a focused sampling of species from all families of Malpighiales (Davis et al., 2007). These studies suggest a rapid evolution leading to highly specialised and unusual floral morphology. 7 2.1.3. Molecular studies in Rafflesia Apart from phylogenetic studies of Rafflesia (Nickrent et al., 1997; Barkman et al., 2004; Davis et al., 2007) mentioned above, there have been no other published studies of the molecular biology of Rafflesia, particularly the functional genomics and developmental biology. 2.1.4. Rafflesia cantleyi Solms-Laubach Rafflesia cantleyi Solms-Laubach is a species with relatively smaller flowers, compared to some of the better-known and large-flowered species such as Rafflesia arnoldii and Rafflesia keithii (Meijer, 1997). It is found in Malaysia, in the states of Perak, Kelantan, Pahang, and Kedah. Up to 1984, this species was considered to be identical with Rafflesia hasseltii by Meijer (1997) following identification by Ridley and other botanists, but was later re-identified as Rafflesia cantleyi as conceived by Solms-Laubach, owing to differences in the size and pattern of the warts on the perigone lobes. Meijer (1997) views this species to be closely related to Rafflesia hasseltii and that it seems to hybridise with it in the Malay Peninsula. 2.2. MADS-Box Genes Many key processes in growth and development are regulated by transcription factors, which are important proteins that bind to and affect the transcription of various target genes. Transcription factors can be classified into gene families according to the conserved DNA-binding domain present. In plants, the major transcription factor gene families include the basic-region leucine zipper (bZIP), MYB-related and MADS-box gene families (Pabo and Sauer, 1992; Martin and Paz-Ares, 1997; Liu et al., 1999). 8 MADS-box genes encode transcription factors involved in a variety of important developmental and signal transduction processes in eukaryotes (Messenguy and Dubois, 2003). The MADS-box encodes a DNA-binding domain comprising of approximately 60 amino acids, the MADS domain, which is highly conserved across plants, fungi, and animals (Theissen et al., 1996). “MADS” is an acronym for the four DNA-binding proteins whose similarity led to the definition of this gene family (Schwarz-Sommer et al., 1990): MINICHROMOSOME MAINTENANCE 1 (MCM1) from Saccharomyces cerevisiae (yeast) (Passmore et al., 1989), AGAMOUS (AG) from Arabidopsis thaliana (Yanofsky et al., 1990), DEFICIENS (DEF) from Antirrhinum majus (Sommer et al., 1990), and SERUM RESPONSE FACTOR (SRF) from Homo sapiens (Norman et al., 1988). This MADS domain folds into a structural motif for DNA interaction consisting of an antiparellel coiled coil of α-helices that lies flat on the DNA minor groove (Pellegrini et al., 1995). All known MADS-domain proteins are transcription factors which regulate target gene expression by binding to specific cis-acting DNA sequences, and have diverse biological roles primarily in development or cell differentiation such as celltype determination and pheromone response in yeast; trachea development in insects; muscle development in vertebrates and insects; and inflorescence and flower development in angiosperms (Shore and Sharrocks, 1995). Besides developmentrelated processes, MADS-domain proteins in yeast have also been found to control arginine metabolism (Messenguy and Dubois, 1993). MADS-domain proteins are proposed to be classed into two main groups of proteins comprising two lineages arising from an ancient duplication event: the Type I lineage which includes SRF-like proteins and the Type II lineage which includes MEF2-like (MYOCYTE-SPECIFIC ENHANCER FACTOR 2-like) proteins, both of 9 which are found in animals, fungi, and plants (Alvarez-Buylla et al., 2000). The two classes of MADS-domain proteins are further classified into subfamilies on the basis of sequence similarity of the C-terminal extensions (Theissen et al., 1996). In animals and fungi, the Type I (SRF-like) proteins contain an SAM domain in the C-terminal extension; this SAM domain (for SRF, ARG80, and MCM1) is based on the loose similarity shared between SRF, ARG80 and MCM1 (Shore and Sharrocks, 1995). Some plant MADS-box genes have been found to group with the animal and fungal SRF-like genes to form the Type I lineage, although the C-terminal domain extensions for these Type I plant MADS-domain proteins are not defined (AlvarezBuylla et al., 2000). In the Type II lineage, animal and fungal MEF2-like proteins contain an MEF2 domain, originally described for vertebrates (Yu et al., 1992). In plants, Type II proteins are of the MIKC structure characteristic of most known plant MADS-box genes (Alvarez-Buylla et al., 2000). MIKC-type proteins are found only in plants, and thus these proteins are thought to have evolved after plants have diverged from animals (and fungi) (Kaufmann et al., 2005). MIKC-type plant MADS-domain proteins characteristically have a modular structure comprising of four domains: the MADS (M), intervening (I), keratin-like (K), and C-terminal (C) domains (Figure 2.1) (Theissen et al., 1996, Alvarez-Buylla et al, 2000). The MADS-domain, which is highly conserved across organisms/plants, encodes a 60-amino-acid DNA-binding domain. This conserved domain binds DNA at a consensus recognition sequence known as the CarG box [CC(A/T)6GG] (Riechmann et al., 1996b). The K domain is a region approximately 70-amino-acidresidues long, with a sequence similar to the coiled-coil of keratin, and found only in plant Type II proteins (Theissen et al., 1996). This K domain is weakly conserved at the primary sequence level but the predicted potential to form amphipathic helices 10 MADS DNA binding K I Dimerisation C Multimerisation Figure 2.1. Schematic representation of the structure of plant MIKC-type MADS-box genes. From left (amino terminal): MADS domain, with DNA binding and dimerisation functions; I and K domains, which are involved in dimerisation; and C domain (at carboxyl terminal), which is variable in length, and postulated to be involved in transactivation and formation of multimeric protein complexes. (Modified from Alvarez-Buylla et al., 2000) 11 characterises this region. The weakly conserved I domain links the MADS domain to the K domain, and is predicted to form an α-helix similar to the MEF2S and SAM domains of non-plant MADS-domain proteins which are required for dimerisation (Huang et al., 2000), and thus influences the specificity of DNA-binding dimer formation (Riechmann et al., 1996a). The MADS+I domains have been found to be sufficient for the formation of DNA-binding dimers, although some class B proteins require part of the K domain as well (Huang et al., 1996; Riechmann et al., 1996a). The C-terminal domain is the least conserved region and is variable in length. However, there is differential conservation within subfamilies, and is particularly conserved in the DEF subfamily (Kaufmann et al., 2005). This domain has been postulated to function as a transactivation domain and contribute to the formation of multimeric protein complexes (Cho et al., 1999; Egea-Cortines et al., 1999; Honma and Goto, 2001). The MADS-box gene family is particularly important in controlling various aspects of plant development, such as floral transition, floral meristem identity, floral organ specification, and fruit and ovule development (Ng and Yanofsky, 2001). Plant MIKC-type MADS-box genes can be divided into at least nine classes based on their function and expression patterns (Nam et al., 2003): classes A, B, C, D, E, F, G, Bs (B-sister), and T. The classes of MADS-box genes which control flower formation are known as floral MADS-box genes (Nam et al., 2003). 2.2.1. Floral organ identity genes The best studied plant MADS-box transcription factors are those involved in floral organ identity determination. In the ‘ABC’ genetic model of determination of floral organ identity, combinatorial interactions between the three classes of floral 12 homeotic genes, A, B, and C, determine the identities of the four floral organs (Haughn and Somerville, 1988; Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Theissen, 2001). A typical flower consists of four different types of organs arranged in four whorls. The first and outermost whorl usually comprises green, leaflike sepals. The second whorl is composed of usually showy, colourful petals. The third whorl is the androecium, composed of the stamens, the reproductive organs that produce pollen. The fourth and innermost whorl is the gynoecium, consisting of the carpels, the reproductive organs that produce the ovules. The homeotic genes are active in two adjacent whorls in the flower: Class A genes alone in the first whorl specify sepals; both Class A and B genes in the second whorl specify petals; Class B and C genes in the third whorl specify stamens; and Class C genes alone in the fourth whorl specify carpels. In Arabidopsis thaliana, the Class A function is contributed by two different genes, APETALA1 (AP1)and APETALA2 (AP2), the B function also by two genes, APETALA3 (AP3) and PISTILLATA (PI), and the C function by just one gene, AG. All these genes, with the exception of AP2, are members of the MADS-box family. The ‘classical ABC model’ has been later extended to include D and E functions, yielding an ‘ABCDE model’ (Theissen, 2001; Krizek and Fletcher, 2005). The A, B, and C functions are the same as in the earlier ABC model, but a D function specifying ovules and an E function that is required for the specification of petal, stamen and carpel identity have been added. First described in Petunia (Angenent et al., 1995; Colombo et al., 1995), D-function genes act in concert with C-function genes to specify ovule development. Homologous genes in Arabidopsis, SEEDSTICK (STK), SHATTERPROOF1 (SHP1) and SHATTERPROOF2 (SHP2) were found to act redundantly and regulate each other’s expression. A stk shp1 shp2 triple mutant has 13 arrested ovule development, but each of the genes is sufficient for ovule development to proceed (Favaro et al., 2003). Class E genes are a new class of floral homeotic genes required for the specification of organ identity in the second, third, and fourth whorls (Jack, 2001). In Arabidopsis thaliana, the first E-function genes characterised were the three SEPATALLA genes (SEP1, 2, and 3). Loss of function of all three SEP genes caused the transformation of the second to fourth whorls of the flower into sepals (Pelaz et al., 2000). A fourth gene, SEP4, is required with the other three SEP genes to confer sepal identity and also contributes to the development of the other three floral organs (Ditta et al., 2004). A sep1 sep2 sep3 sep4 quadruple mutant shows a conversion of all four floral organ types into reiterating whorls of leaf-like structures, instead of sepals as is the case for the sep1 sep2 sep3 triple mutant. Using yeast two-hybrid screening, analyses of protein-protein interaction have shown that AG interacts with SEP1, SEP2, and SEP3, while AP1 interacts with SEP3 (Fan et al., 1997; Pelaz et al., 2001). Co-immunoprecipitation experiments suggest that the AP3–PI heterodimer can interact directly with SEP3 and AP1, as well as with SEP3 and AG to form ternary complexes in vitro (Honma and Goto, 2001). The ability of MADS-box proteins to form multimeric complexes may therefore provide the molecular basis for the combinatorial control of floral organ specification. In this hypothesis, different MADS homo- or hetero-dimer combinations interact with additional transcription factors, which then determine the functional specificity of the complexes formed (Riechmann et al., 1996b). This led to the formulation of a ‘quartet model’, which postulates that four different combinations of four different floral homeotic proteins determine the identities of the four different floral organs (Theissen, 2001; Theissen and Saedler, 2001). Specifically, tetramers of AP1–AP3–PI–SEP, 14 AP3–PI–AG-SEP, and AG–AG–SEP–SEP would specify petals in the second whorl, stamens in the third whorl, and carpels in the fourth whorl, respectively (Honma and Goto, 2001; Theissen and Saedler, 2001). These protein quartets represent one model of transactivation of genes for floral organ identity by MADS box protein complexes. Each dimer of a MADS-box tetramer recognises and binds to a single CArG box sequence; the C-terminal domains of the MADS-box proteins are involved in protein– protein interaction to form the tetramer (Jack, 2001). However, for this model to work, two closely linked CArG box sequences have to be present in the promoters of target genes. Two other models were hypothesised. One model is where multimeric MADSbox protein complexes bind to single CArG box sequences. Here, a single dimer binds to the CArG box sequence while other proteins which bind to this dimer via protein– protein interactions could provide either altered DNA-binding selection or affinity, or a transcriptional activation domain to the multimeric complex (Jack, 2001). Another, less likely, model is that dimers of MADS-box proteins cooperatively bind to adjacent CArG box sequences where there is no protein–protein interaction. This is however not well supported by existing data (Jack, 2001). 2.2.2. Flowering time genes Besides floral organ identity genes, there are MADS-box genes involved in flowering that have different functions, for example, in the control of flowering time, such as SHORT VEGETATIVE PHASE (SVP) and AGAMOUS-LIKE 24 (AGL24). SVP and AGL24 are members of the StMADS11 clade (Becker and Theissen, 2003) which are involved in the contrasting functions of repression and promotion of flowering, respectively. These genes have been categorised as Class T genes (Nam et al., 2003). 15 SVP forms a repressor complex of flowering time along with FLOWERING LOCUS C (FLC) (Liu et al., 2009a) which directly affects the expression of SUPPESSOR OF OVEREXPRESSION OF CONSTANS 1 (SOC1) and FLOWERING LOCUS T (FT) (Liu et al., 2009b). SVP has been shown to repress transcription of SOC1 in the shoot apex and leaves by binding directly to the SOC1 promoter (Li et al., 2008). In contrast, AGL24 promotes the expression of SOC1 by binding to the SOC1 promoter. These observations clearly show that SVP and AGL24 are key integrators of flowering signals, along with other floral transition signals (Liu et al., 2008). Overexpression of SVP results in the loss of carpels as well as the conversion of flowers into shoot-like structures with chimaeric characteristics of vegetative shoots and flowers. Similarly, overexpression of AGL24 results in the transformation of carpels into inflorescence-like structures, the sepals and petals into leaf-like structures, and initiation of secondary inflorescences in the axils of sepals (Liu et al., 2009a). Homologues of SVP and AGL24 have been isolated from a number of dicotyledonous and monocotyledonous species, and when they were ectopically expressed in Arabidopsis, phenotypes similar to those of 35S::SVP and 35S::AGL24, respectively, have been observed. This shows that they are likely to have conserved function in specifying floral meristem development (Liu et al., 2009a). The coregulator of LEAFY (LFY), namely SEPALLATA3 (SEP3), is repressed by SVP, AGL24 and SOC1 (Liu et al., 2009b). This is achieved by forming complexes with two chromatin regulators: TERMINAL FLOWER 2/LIKE HETEROCHROMATIN PROTEIN 1 (TFL2/LHP1) and SAP18. SVP interacts with TFL2/LHP1 to modulate histone H3 methylation while AGL24 and SOC1 interacts with SAP18 to modulate histone H3 acetylation in SEP3 chromatin. 16 2.2.3. Heterologous expression system for functional analysis of genes Arabidopsis thaliana has been used for understanding functions of genes cloned from species for which there is limited molecular and genetic information available. The ease of genetic transformation coupled with the availability of numerous mutants makes it a convenient heterologous system for such functional analyses. This is particularly useful for plants that are not amenable to transformation, for plants that lack mutants, and for plants with very long generation times. Examples include Eucalyptus grandis (Brill and Watson, 2004) , Cycas edentata (Zhang et al., 2004), and Paulownia kawakamii (Prakash and Kumar, 2002). From the foregoing review of literature, it is clear that molecular regulation of floral development in higher plants is understood in a fairly comprehensive manner. However, there is a paucity of information on the developmental regulation of parasitic plants. Despite having the world’s largest flowers, application of molecular tools were rarely used in studying floral development in Rafflesia species. In view of this, we initiated the current project of cloning MADS-box genes that might be involved in regulating flower development in Rafflesia cantleyi. It is hoped that our results will contribute to a better understanding of the development of highly specialised flowers of Rafflesia, and parasitic plants in general. 17 CHAPTER 3 MATERIALS AND METHODS 3.1. Plant materials Flower buds of various sizes of Rafflesia cantleyi Solms-Laubach were collected from a few localities along a trail between Tekek and Juara in Pulau Tioman, Pahang, Malaysia. Collection of Rafflesia cantleyi material in Peninsular Malaysia required a permit (permit number: UPE40/200/19 SJ. 1200) from the Economic Planning Unit, Prime Minister’s Department (Unit Perancang Ekonomi, Jabatan Perdana Menteri), Putrajaya, Malaysia. The buds were surface-sterilised using a 10% (v/v) Clorox® solution (1% sodium hypochlorite) for 5–10 min, followed by three rinses with sterile water. Tissues were cut and weighed, then flash-frozen in liquid nitrogen. All samples were stored at –80°C until further use. Transgenic and mutant Arabidopsis thaliana plants used in the experiments were of the same genetic background, Columbia ecotype. Arabidopsis thaliana seeds were sown on soil (Flora Fleur) and stratified for 3–4 days at 4°C to break seed dormancy and allow uniform germination, before being transferred to a growth chamber. The plants were grown at 23 ± 2°C under long-day photoperiod conditions (16 h of light / 8 h of darkness). 3.2. RNA and DNA isolation Total RNA from the Rafflesia cantleyi flower buds was isolated using a modified RNeasy® Plant Mini Kit (QIAGEN) method (Kim, 2004). The modification involves an initial CTAB extraction (Doyle and Doyle, 1987). 100 mg fresh weight of tissue was pulverised in liquid nitrogen and homogenised in 500 ml CTAB buffer 18 with 1 µl β-mercaptoethanol added by vigorously mixing using a vortex. The homogenate was incubated at 60°C for 10 min before 500 µl of chloroform–isoamyl alcohol (24:1) was added and then vigorously mixed. The homogenate was then centrifuged at 14,000 g for 15 min to pellet the insoluble cell debris. 360–400 µl of the aqueous phase was recovered and mixed with cold isopropanol (2/3 volume of the recovered supernatant) and incubated at −20°C for 1 h or more to precipitate the RNA. The preparation was then applied to an RNeasy® column and purification of the preparation was done following the manufacturer’s instructions. Total RNA from Arabidopsis thaliana plant tissues was isolated using the RNeasy® Plant Mini Kit (QIAGEN) following manufacturer’s instructions. Genomic DNA from Rafflesia cantleyi was isolated using a modified CTAB method (Lodhi et al., 1994). 100 mg fresh weight of tissue was pulverised in liquid nitrogen and homogenised in 500 ml CTAB buffer, with 1 µl β-mercaptoethanol and PVPP (100 mg/g plant tissue) added, by vigorously mixing using a vortex. The homogenate was incubated at 60°C for 25 min before 500 µl of chloroform–isoamyl alcohol (24:1) was added and then vigorously mixed. The homogenate was then centrifuged at 14,000 g for 15 min to pellet the insoluble cell debris. 360–400 µl of the aqueous phase was recovered, and 1/2 volume of 5 M NaCl was added to the supernatant. The resulting solution was then with cold isopropanol (2/3 volume of the recovered supernatant) and incubated at −4°C for 1 h or more to precipitate the DNA. The DNA was purified by repeated steps of centrifugation and washing with 76% ethanol, and then stored in deionised water or TE buffer. 19 3.3. Reverse transcription Analysis of RNA was performed in a two-step reverse transcription– polymerase chain reaction process (RT-PCR): cDNA was synthesised using poly(A)+ RNA primed with oligo(dT) in the first step; and PCR was performed using primers specific for the gene of interest in the second step. First-strand cDNA was synthesised using the SuperScript™ II RNase H–reverse transcriptase (Invitrogen), following manufacturer’s instructions. 50 ng to 5 µg of RNA was mixed with 1 µl of oligo(dT)12–18 (0.5 µg/µl) primer and 1 µl of 10 mM dNTP mix, and adjusted to a total volume of 10 µl with DEPC-treated water. The RNA and primer mix was denatured by incubation at 65°C for 5 min using a thermal cycler (Elmer Perkin) and subsequently placed on ice for 1 min. A 9 µl reaction mixture containing 2 µl of 10× RT buffer, 4 µl of 25 mM MgCl2 solution, 2 µl of 0.1 M DTT, and 1 µl of RNaseOUT™ Recombinant RNase Inhibitor, was added to the reaction tube containing the 10 µl mix of RNA and primer and the tube was incubated at 42°C for 2 min. 1 µl (50 units) of SuperScript™ II reverser transcriptase was then added, and the 20 µl total reaction mixture was incubated at 42°C for 50 min for cDNA synthesis to take place. The reaction was terminated by an incubation at 70°C for 15 min followed by chilling on ice for 5–10 min. 1 µl of RNase H was then added and the reaction mixture was incubated at 37°C for 20 min before being stored at −80°C. 3.4. PCR amplification PCR amplification of MADS-box genes from Rafflesia cantleyi was performed using degenerate primers and an oligo(dT)15 primer. These degenerate primers were designed based on the conserved MADS box of MADS box genes. The primers used were are listed in Table 3.1. PCR reactions were performed using 20 Table 3.1. Degenerate primers used in cloning MADS-box genes from Rafflesia cantleyi Name Sequence (5′→3′) Direction Reference MADS1 AARMGIMGIAAYGGIYTIYTIAARAARGC Forward N.A. MADS2 GGGGTACCAAYMGICARGTIACITAYTCIAAGMGIMG Forward N.A. MADS3 AARAARGCIYWYGARCTIKCKGTICT Forward N.A. MADS4 AAYMGRCARGTICAITAYTCRAARMG Forward Di Stilio et al., 2005 MADS5 GGIMGIAARATIGARATIAARRGIAT Forward Di Stilio et al., 2005 MADS6 AAYRGICARGTIACITTYTGYAARRGIRG Forward Di Stilio et al., 2005 MADS7 CAYTTRATGGGIGARGCICTIAGYTG Forward Di Stilio et al., 2005 MADS8 GGACGAGGACGDGTWCARCT Forward Jager et al., 2003 MADS9 SAGATCAAGMGIATHGARAAY Forward Jager et al., 2003 MADS10 GGGGTACCAAYMGICARGTIACITAYTCIAAGMGIMG Forward Kramer et al., 1998 oligo(dT) TTTTTTTTTTTTTTT Reverse N.A. 21 step-up conditions with the following cycling parameters: an initial denaturation at 95°C for 1 min; 10 cycles of denaturation at 95°C for 30 s, annealing at 35°C for 1 min, and extension at 72°C for 1 min; 25 cycles of denaturation at 95°C for 30 s, annealing at 40°C for 1 min, and extension at 72°C for 1 min; and a final extension at 72°C for 10 min. 1 µg of cDNA template was addeded to a reaction mixture consisting of 0.4 µl DyNAzyme polymerase, 0.2 mM dNTP mix, 1× DyNAzyme PCR buffer and 2 pmol each of forward and reverse primers. PCR reactions were visualised by performing gel electrophoresis in a 1.2% agarose gel. Amplified fragments over 400 bp in size were selected for cloning and sequencing. 3.5. Cloning of PCR products The PCR products were purified using the QIAquick® PCR purification kit (QIAGEN) following manufacturer’s instructions. Five volumes of buffer PB were added to each PCR sample and the mixture was applied to the QIAquick column and centrifuged at 14,000 g for 1 min. The flow-through was discarded and the column was washed with 0.75 ml of buffer PE diluted in ethanol. The column was centrifuged for 1 min to remove residual ethanol. To elute the purified DNA, 30 µl of buffer EB (10 mM Tris HCl, pH 8.5) was added to the column membrane and the column was allowed to stand for 1 min before centrifugation for 1 min. The purified PCR product was then cloned into the pGEM®-T Easy Vector (Promega). The PCR product was added to a reaction mix containing 1× Rapid Ligation buffer, 50 ng pGEM®-T Easy Vector and 3 units of T4 DNA ligase, and the mixture was incubated overnight at 4°C to maximise ligation products. The resulting recombinant plasmids were then introduced into competent Escherichia coli DH5α cells. 22 Cell transformation was performed by adding 10 µl of ligation products to 100 µl of competent cells. The mixture was incubated on ice for 30 min before a heat shock treatment of the cells at 42°C for 90 s, followed by incubation on ice for 5 min. 1 ml of LB medium was added to the mixture which was then incubated with shaking at37°C for 1 h. The mixture was then centrifuged gently at 4,000 rpm for 4 min to pellet the cells and the excess LB medium was removed. The cells were then plated onto a LB agarose plate containing ampicillin (10 mg/l) and incubated at 37°C overnight. Transformants were picked via blue/white colony selection and PCR was performed to check for presence of the insert. 3.6. Plasmid DNA purification Plasmid DNA was isolated from the Escherichia coli clones using the Wizard SV Miniprep Kit (Promega) following manufacturer’s instructions. Clones were picked from the agarose plates and grown overnight in 3 ml bacterial cultures using LB medium containing ampicillin. Each bacterial culture was centrifuged at 3,700 rpm for 5 min to pellet the cells. The cells were then resuspended in 250 µl resuspension buffer. The cells were lysed by addition of 250 µl of cell lysis solution followed by 10 µl of alkaline protease, a step which did not exceed 5 min, as recommended by the manufacturer, to prevent nicking of the plasmid DNA by the alkaline protease. The cell lysis solution was neutralised by addition of 350 µl of neutralisation solution. The cell debris was removed by centrifugation for 1 min at 14,000 rpm. The supernatant was then applied to the spin column, where the plasmid DNA would bind to the silica membrane. The lysate was passed through the column by centrifugation, and the column was rinsed with 750 µl of wash buffer. Residual 23 wash buffer was removed by an additional 2 min of centrifugation. The purified plasmid DNA was eluted using 30 µl deionised water. 3.7. DNA sequencing Selected clones were sequenced via an automated sequencing method using ABI PRISM™ Big Dye™ Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, USA). The sequencing reaction was prepared by mixing 150 ng of double-stranded DNA with 1.6 pmol of forward or reverse primer and 2 µl of Terminator Ready Reaction Mix and the final volume topped up to 5 µl with nuclease-free water. The sequencing reaction was performed using a thermal cycler (Elmer Perkin) for 25 cycles of denaturation at 96°C for 30 s, annealing at 52°C for 5 s, and extension at 60°C for 4 min. Sequence reactions were purified using the CleanSEQ® kit (Agencourt) following manufacturer’s instructions with slight modifications. 10 µl of CleanSEQ reagent containing magnetic beads and 31 µl of 85% ethanol were added into each sequence reaction tube, with thorough mixing. The tubes were then placed onto an Agencourt SPRIPlate, and incubated for 3 min, before the supernatant was removed. The sequencing products were washed with 100 µl of 85% ethanol, then air-dried. The sequence products were eluted with 40 µl of sterile water in each tube, and 12 µl of the elution was transferred out from each tube for automated sequencing using the ABI PRISM™ 3100 DNA Sequencer (Applied Biosystems, USA). 3.8. Sequence analysis Sequences obtained after automated sequencing were collated and compared with published sequences in the GenBank databases using the Basic Local Alignment 24 Search Tool (BLAST) program on the National Center for Biotechnology Information (NCBI) website. The algorithms used were blastn (to search the nucleotide database using a nucleotide query) and tblastx (to search the translated nucleotide database using a translated nucleotide query). 3.9. Phylogenetic analysis Multiple sequence alignments were carried out using the program CLUSTALW. Since the ~60 amino acid MADS domain is highly conserved and alignment is unproblematic, the data set for the phylogenetic analysis was based on this region. This data set was subjected to a parsimony analysis in TNT version 1.0 (Tree Analyses Using New Technology, Goloboff et al., 2000). 3.10. Rapid amplification of cDNA ends 5′-rapid amplification of cDNA ends (5′-RACE) was performend using BD SMART™ RACE cDNA Amplification Kit (Clontech) following manufacturer’s instructions. The 5′-region of the putative cDNA was amplified using UPM (forward primer provided by manufacturer) and a gene-specific reverse primer (5′-ACA GCT GCA GAC AAC AGT GG-3′). 3.11. Preparation of ectopic expression construct The full open reading frame of the putative gene RcMADS1 from Rafflesia cantleyi was amplified from the cDNA clone (obtained as above) using the following primers containing restriction enzyme sites: RcM1-F-HindIII (5′-CCC AAG CTT GGT CGT GCC GTA TTT GTT CT-3′, HindIII recognition site underlined) and RcM1-R-XbaI (5′-TGC TCT AGA CCT CTC TCT CCG TCA GCT TG-3′, XbaI 25 recognition site underlined). The amplified fragments were digested for 2 h with HindIII and XbaI restriction endonucleases to generate sticky ends, which were then inserted between the CaMV 35S promoter and CaMV terminator in a sense direction in the pGreen 0229 vector digested with the same restriction endonucleases (Figure 3.1). This ectopic expression construct was named 35S::RcMADS1. 3.12. Transformation of Agrobacterium tumefaciens The 35S::RcMADS1 construct was introduced into Agrobacterium tumefaciens strain GV3101 carrying the pSoup helper vector. A 2 ml culture LB medium containing 20 mg/l gentamycin and 10 mg/l tetracycline was inoculated with GV3101 cells and incubated at 28°C in for 2 days. 100 µl of the bacterial culture was then transferred to 50 ml of fresh LB medium and incubated at 28°C until an OD600 of 0.8– 1.0 was reached. This bacterial culture was transferred into a 50 ml polypropylene tube and incubated on ice for 5 min, before being centrifuged at 3,700 rpm for 10 min at 4°C, and the resulting pellet was resuspended in 5 ml of ice-cold water. The resuspended cells were centrifuged again at 3,700 rpm for 10 min at 4°C and the pellet was resuspended in 1 ml of ice-cold water and used immediately for transformation. 1 µl of recombinant DNA was added to 100 µl of freshly prepared competent cells and allowed to incubate on ice for 30 min. This mixture was transferred into an electroporation cuvette (Eppendorf, 1 mm gap width, 100 µl volume) and electroporation was carried out at 2,300 V (Eppendorf, Electroporator 2510). After electroporation, 500 µl of LB medium was added and the mixture was incubated with shaking at 28°C for 4 h before plating on an LB agar plate supplemented with antibiotics (50 mg/l kanamycin, 20 mg/l gentamycin, and 10 mg/l tetracycline). The 26 HindIII Noster LB bar Nospro 2× 35Spro XbaI RcMADS1 CaMVter RB Figure 3.1. Schematic diagram of 35S::RcMADS1 ectopic expression construct. The RcMADS1 cDNA fragment was ligated into a pGreen 0229 vector with the following features: LB, left border; Noster, nopaline synthase terminator; bar, bialaphos encoding sequencing, conferring resistance to glufosinate ammonium, a broad spectrum herbicide which is available as Basta®; Nospro, nopaline synthase promoter; 2× 35Spro, tandem copies of the cauliflower mosaic virus (CaMV) 35S promoter; CaMVter, CaMV terminator; RB, right border. Arrows indicate directions of transcription. HindIII and XbaI are restriction sites. 27 plate was incubated at 28°C for 2 days, and transformants were selected via PCR and confirmed by sequencing. 3.13. Genetic transformation of Arabidopsis thaliana Transformation of Arabidopsis thaliana plants was carried out using the floral dip method (Clough and Bent, 1998). Healthy Arabidopsis thaliana plants were grown on soil under long-day photoperiod conditions (16 h of light / 8 h of darkness), until flowering. An Agrobacterium tumefaciens transformant selected via PCR and confirmed by sequencing to carry the ectopic expression construct 35S::RcMADS1 was inoculated into a 3 ml culture of LB medium containing 50 mg/l kanamycin and the culture was incubated at 28°C for 2 days. 25 µl of the bacterial culture was then transferred to a fresh 25 ml culture and incubated at 28°C overnight. This bacterial culture was centrifuged at 3,700 rpm for 10 min and the resulting pellet was resuspended in a 5% sucrose solution. Before commencement of the floral dipping, Silwet L-77 was added to the Agrobacterium tumefaciens cell suspension to a final concentration of 0.03% (v/v). Arabidopsis thaliana inflorescences were immersed in the bacterial cell suspension for 5–10 s with gentle agitation. If possible, the rosette portions of the plants were immersed in the bacterial cell suspension as well, to maximise transgenic seed production. After dipping, the plants were covered with plastic bags for 16–24 h, to maintain high humidity. The plants were then allowed to grow under long-day conditions until siliques had developed. The seeds were harvested, germinated and the resulting seedlings were screened for herbicide resistance. 28 35S::RcMADS1 plants were grown under long-day conditions and sprayed with 250 mg/l Basta® solution (Finale, AgrEvo, California, USA) 3 days and 10 days after germination. After 2 weeks, the surviving seedlings were selected as putative transgenic plants and grown for the next generation prior to phenotypic characterisation. 3.14. Quantitative real-time PCR analysis Real-time PCR experiments were carried out using the Power SYBR® Green PCR Master Mix (Applied Biosystems, USA) on the ABI Prism 7000 Sequence Detection System (Applied Biosystems, USA). PCR was performed in 20 µl reactions containing 1 µl of the diluted first strand cDNA samples, 4 pmol of primers, and 10 µl of the SYBR Green PCR mix. The PCR thermocycling profile used was as follows: 1 cycle of 50°C for 2 min; 1 cycle of 95°C for 10 s; 40 cycles of 95°C for 15 s, and 60°C for 1 min. The gene-specific primer pairs used are listed in Table 3.2. Analysis of the results was carried out using the ABI Prism 7000 Sequence Detection System software. 3.15. Genomic Southern blot analysis Genomic DNA samples from Rafflesia cantleyi was prepared as described in the previous section (Section 3.2). The quality and quantity of genomic DNA were analysed by spectrophotometer (NanoDrop, Thermo Fisher Scientific, USA). A 1% agarose gel (w/v) containing 0.5× TBE buffer (45 mM Tris-boric acid, 1 mM EDTA, pH 8.0) was prepared. Rafflesia cantleyi genomic DNA was digested by the appropriate restriction endonuclease, and electrophoresis of the digested DNA was conducted using an agarose gel in 0.5× TBE buffer until the bromophenol blue 29 Table 3.2. Primer pairs used in quantitative real-time PCR Target Forward Primer (5′→3′) Reverse Primer (5′→3′) RcMADS1 5′- CCAAGCCAGCCATCTCTTGA-3′ 5′- GCTCAGTCGCACCCGATT-3′ AP1 5′-CATGGGTGGTCTGTATCAAGAAGAT-3′ 5′-CATGCGGCGAAGCAGCCAAGGTT-3′ AGL24 5′-GAGGCTTTGGAGACAGAGTCGGTGA-3′ 5′-AGATGGAAGCCCAAGCTTCAGGGAA-3′ CO 5′-TCAGGGACTCACTACAACGACAATGG-3′ 5′-TTGGGTGTGAAGCTGTTGTGACACAT-3′ FLC 5′-CCAAACGTCGCAACGGTCTC-3′ 5′-GTCCAGCAGGTGACATCTCC-3′ FT 5′-CTTGGCAGGCAAACAGTGTATGCAC-3′ 5′-GCCACTCTCCCTCTGACAATTGTAGA-3′ LFY 5′-ATCGCTTGTCGTCATGGCTG-3′ 5′-GCAACCGCATTGTTCCGCTC-3′ SEP3 5′-AGACTAAGGTTAGCTGATGGGTA-3′ 5′-ATGATGACGACCGTAGTGATC-3′ SOC1 5′-AGCTGCAGAAAACGAGAAGCTCTCTG-3′ 5′-GGGCTACTCTCTTCATCACCTCTTCC-3′ SVP 5′-CAAGGACTTGACATTGAAGAGCTTCA-3′ 5′-CTGATCTCACTCATAATCTTGTCAC-3′ TUB2 5′-GAGAATGCTGATGAGTGCATGG-3′ 5′-AGAGTTGAGTTGACCAGGGAACC-3′ 30 dye had indicated the sample had been separated for a sufficient distance. The gel was processed sequentially with depurination (in 250 mM HCl for 10 min), denaturation (in 1.5 M NaCl, 0.5 M NaOH for 30 min) and neutralization (in 1.5 M NaCl, 0.5 M Tris-HCl pH 7.5, for 30 min). Genomic DNA was then blotted onto a positively charged nylon membrane by capillary blocking overnight. DNA was fixed to the membrane by UV crosslinking for 20 s at 120 mJ/cm2. Blots were hybridised with denatured probes in hybridisation buffer (1% SDS (w/v), 1 M NaCl, 10% dextran sulfate (w/v) and 100 µg/ml denatured salmon sperm DNA) at 65°C overnight. After hybrisation, the blots were washed twice with 2× SSC (0.3 M NaCl and 0.03 M sodium citrate, pH 7.0) and 0.1% SDS (w/v) at 65°C for 15 min, once with 1× SSC and 0.1% SDS (w/v) at 65°C for 30 min, and then 0.1× SSC and 0.1% SDS (w/v) at 65°C for 5 min. The blots were then exposed to X-ray film. 31 CHAPTER 4 RESULTS AND DISCUSSION 4.1. Collection of Rafflesia cantleyi flower buds Flower buds of Rafflesia cantleyi were collected from Pulau Tioman, Malaysia (see Methods section for details). The size ranged from 1.0 cm to 11.4 cm in diameter (Figure 4.1). Our attempts to extract RNA from buds of various sizes yielded mixed results. The quality of RNA was generally poor in the extracts from the larger buds. However, we succeeded in optimising nucleic acid extraction from buds of 1.0 cm to 2.95 cm in diameter. These were used for PCR cloning of MADS-box genes. 4.2. RNA isolation Preliminary attempts at RNA extraction include the use of protocols such as TRIzol®, RNeasy® and CTAB. Tissues turned brown and highly viscous upon homogenisation, due to rapid polyphenol production, leading to poor quality RNA with all the methods tried. RNA recovery was negligible using TRIzol® and RNeasy® protocols, whereas it was low for the CTAB protocol, where the yield was less than 20 ng/µl. Optimisation of the CTAB protocol included the combination of the initial steps of the CTAB protocol with an additional series of purification steps using an RNeasy® column. This modification yield more appreciable amounts of total RNA, from 83.8 ng/µl to 614.2 ng/µl, as well as fairly good quality total RNA (A260/A280 ratio of about 1.5 to 1.9) (Figure 4.2). Young buds yielded better quality RNA than older buds. 32 A B D C 1 cm Figure 4.1. Rafflesia cantleyi Solms-Laubach buds. (A) Buds of a range of sizes collected from Pulau Tioman, Malaysia, from the largest on the left to the smallest on the right. Not all buds were viable for DNA and RNA extraction; some were aborted during development. (B) A bud of approximately 2.5 cm developing on a Tetrastigma sp. vine in the forest in Pulau Tioman, Malaysia. (C) A longitudinal section of a young bud (approximately 2 cm in diameter), showing generally undifferentiated tissue. The layers of tissue on the top would develop into petals and bracts, and while the layers of tissue in the centre would develop into the column. Browning due to polyphenol formation was rapid after the tissue was exposed to air; when cut, the bud is pale yellow and white in colour. (D) A longitudinal section of a bud close to anthesis (approximately 8 cm in diameter). 33 1 2 3 Figure 4.2. Gel electrophoresis of total RNA extracted from young Rafflesia cantleyi flower bud (~1 cm in diameter). The quality of RNA appeared to be good, due to the presence of the 28S and 18S rRNA bands. 34 4.3. DNA isolation Extraction of genomic DNA was achieved by using the CTAB method. Although the yield was appreciable and the quality of DNA was good, problems were encountered later during genomic DNA blot analysis. Digestion by restriction endonucleases was poor, indicating that polyphenolic compounds produced by the Rafflesia tissue may not have been thoroughly removed during extraction and genomic DNA was contaminated by such compounds, preventing cleaving of the genomic DNA. 4.4. Cloning MADS-box genes by RT-PCR The general approach to cloning the MADS-box genes from Rafflesia cantleyi is based on reverse-transcribing mRNA from the total RNA extracted from R. cantleyi flower buds and performing PCR using degenerate primers designed from consensus sequences of known MADS-box genes from various angiosperms species, as little nuclear gene data is available for Rafflesia. Using a step-up PCR strategy, long faint smeary bands were obtained after amplification with degenerate primers. As there were no particularly distinct bands (Figure 4.3A), these PCR products were purified and cloned into pGEM®-T, and the recombinant plasmids were used for another round of PCR amplification using SP6 and T7 primers, where distinct bands of 200 to 2000 bp were obtained. Plasmids containing fragments of 400 bp and longer were used for DNA sequencing (Figure 4.3B). One fragment of 750 bp, amplified from RNA from a young flower bud (~1 cm in diameter) using the primer MADS2, was found to be highly similar to MADS-box genes, with scores up to 148 bits using a tblastx search. The MADS-box genes with high similarity to this sequence include SVP, AGL24, 35 A B MADS1 Rc NC 1 2 MADS2 Rc NC 3 4 MADS3 Rc NC 5 6 PC Figure 4.3. Cloning of MADS-box genes via degenerate PCR. (A) Gel electrophoresis of PCR products after step-up PCR using three different pairs of degenerate primers (forward primers: MADS1, MADS2, and MADS3; reverse primer is an oligo-dT primer). All three sets of primers produced long smeary bands of different intensities. (B) Screening of fragments from degenerate PCR via pGEM-T cloning. Lanes 1–6 represent amplifcations from 6 different white colonies; PC: positive control. 36 STMADS11, and IbMADS4, which are all members of the STMADS11 clade of MADS-box genes. A sequence-specific primer was designed from this fragment sequence for use in Rapid Amplification of cDNA Ends (RACE) in sequencing the 5′ region of the mRNA of this putative MADS-box gene. 5′-RACE PCR using this new primer yielded an 800 bp fragment. A gene-specific primer meant for cloning the full-length cDNA was designed based on the 5′ untranslated region (UTR) from this fragment. We obtained a 951 base-pair-long sequence when we used the 5′ primer and the oligo(dT) primer. This cDNA sequence contains a 687 bp open reading frame (ORF) before a stop codon encoding a polypeptide of 228 amino acids, as well as 5′ and 3′ UTRs (Figure 4.4). The conserved MADS domain (69 amino acids) was identified using the NCBI conserved domain search, while comparison with known MADS-box genes allowed the identification of the K domain (79 amino acids). Results from a tblastx search using the online BLAST program revealed high similarities in protein sequence alignment to MADS-box proteins such as MADS1 from Populus tomentosa, MPF2 and MPF4 from Physalis pubescens, IbMADS4 from Ipomoea batatas, StMADS11 and StMADS16 from Solanum tuberosum, and SVP and AGL24 from Arabidopsis thaliana, all from the StMADS11 clade of MADSdomain proteins. An alignment of some of these proteins from the StMADS11 with RcMADS1 showed that the MADS domain and K domains are highly conserved amongst these members of the StMADS11 clade., with the I domain somewhat highly conserved as well (Figure 4.5). RcMADS1 showed 57.6% and 56.7% amino acid similarity respectively to AGL24 and SVP from Arabidopsis thaliana. At the nucleotide level, the RcMADS1 cDNA has a 64.6% and 61.2% similarity to AGL24 and SVP respectively. 37 1    GGGTTAGGTTTCCATATGCCCCACGGGCCACATATTCTTGAACCTGTCTAGCCTTCCCCA  60 61   ATTTATTGCCCGAAGCTGTACGTTCATATTTCGAAACTAAAGATTCGGTCGTGCCGTATT  120 121  TGTTCTGTTTCGGAGCAGATAATATAATGGCTCGAGAAAAGATCAAGATCAAAAAGATCG  180 1                               M  A  R  E  K  I  K  I  K  K I 11 181  ACAACATCACTGCAAGGCAAGTCACCTTCTCTAAGAGGAGACGAGGGCTCTTCAAGAAAG  240 12   D  N  I  T  A  R  Q  V  T  F  S  K  R  R R G  L  F  K  K 31 241  CAGAAGAGCTATCAGTTCTTTGTGATGCTCATGTGGCTGTCATTATCTTTTCTGCCACAG  300 32   A  E  E L  S  V  L  C  D  A  H  V  A  V  I  I F  S  A  T 51 301  GGAAGCTCTTCGACTATTCCAGCTCCAGCATGAAGGACATACTCTCGAGGTATGATGATC  360 52   G  K  L  F  D  Y  S  S S S M  K  D  I  L  S  R  Y D  D 71 361  TGCATTTCAATAACAAAGAGAAGCCAAGCCAGCCATCTCTTGAACTGCAGCTAGAAATTA  420 72   L  H  F  N  N K  E  K  P  S  Q  P  S  L  E  L  Q  L  E  I 91 421  GCAATCGGGTGCGACTGAGCAAAGAAGTTGCAGACAAGACTCGCCAACTAAGGCAAATGA  480 111 92   S  N  R  V  R  L  S  K  E  V  A  D  K  T  R  Q  L  R  Q  M 481  GAGGAGAAGATCTGAACGAATTAAATGTGGAGGAACTGCAGCAACTGGAGAACTTGCTTG  540 112  R  G  E  D  L  N  E  L  N  V  E  E L  Q  Q L  E  N  L  L 131 541  AGGTGGGCCTCCAGCGCGTTACTGATGCCAAGGGCAAGCGCATCACAAATGAGATATCTG  600 151 132  E  V  G  L  Q  R  V  T  D  A  K  G  K  R  I  T  N  E  I  S 601  AACTCGAAAGGAAGGGAGCGCAGCTGATGGAAGAAAATAAGCAACTAAAGCAAAAAATGG  660 152  E  L  E  R  K  G  A  Q  L  M  E  E N  K  Q  L  K  Q  K M 171 661  TGATGATGTGCAGTGGAACAAGACCTGCCATCCTGGAGTCTGATATCACAACCCATGAAG  720 191 172  V  M  M C  S  G  T  R  P  A  I  L  E  S  D  I  T  T H  E 721  AGGGCATGTCCTCTGATTCTGCCACTGTTGTCTGCAGCTGTAGCAATGGCCCCCCCGTGG  780 192  E  G  M  S  S D  S  A  T  V  V C  S  C  S  N  G  P  P V 211 781  AAGATGATATCTCCGATACGTATCTTAAATTGGGATTGTCCTTCTCAAGCTGACGGAGAG  840 212  E  D  D I  S  D  T  Y  L  K  L  G  L  S  F  S  S *          229 841  AGAGGTGCAGGATTCTTCAATGGCGGTGCAATGAATGAACACATACAAATCTTTACACG   899 Figure 4.4. Structure of RcMADS1 cDNA. The upper row indicates the nucleotide sequence, and the lower row the deduced amino acid sequence. The termination (TGA) codon is shown by an asterisk (*). The MADS-box and K domains are shown in bold type. 38 MADS domain I domain RcM1 MPF2 MPP3 AGL24 SVP STMADS16 STMADS11 MAREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSVLCDA VAVIIFSATGKLFDYSSSSMKDILSRY LH N K MAREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSVLCDAHVAVIIFSATGKLFDYSSSSMKDILSRYDDLHFNNKEKP MMAREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSVLCDADVALIIFSSTGKLFDFSSSSMKDILGKYK.LQSANLDKV AREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSVLCDADVALIIFS TGKLFDFSSSSMKDILGKYK LQS NL K MMAREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSILCDADVALIIFSSTGKLFDFSSSSMKDILGKYK.LQSANLDKV AREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSILCDADVALIIFS TGKLFDFSSSSMKDILGKYK LQS NL K MMAREKIRIKKIDNITARQVTFSKRRRGIFKKADELSVLCDADVALIIFSATGKLFEFSSSRMRDILGRYS.LHASNINKL AREKIRIKKIDNITARQVTFSKRRRGIFKKADELSVLCDADVALIIFSATGKLFEFSSS MRDILGRY LH NI K MMAREKIQIRKIDNATARQVTFSKRRRGLFKKAEELSVLCDADVALIIFSSTGKLFEFCSSSMKEVLERHN.LQSKNLEKL AREKI IRKIDN TARQVTFSKRRRGLFKKAEELSVLCDADVALIIFS TGKLFEF SSSMKEVLER LQS NL K MMAREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSVLCDADVALIIFSATGKLFDFASTSMKDILGKYK.LQSASLEKV AREKIKIKKIDNITARQVTFSKRRRGLFKKAEELSVLCDADVALIIFSATGKLFDF S SMKDILGKYK LQS L K MMVRQKIQIKKIDNLTARQVTFSKRRRGLFKKAQELSTLCDADIGLIVFSATGKLFEYSSSSMMQLIEKHK.MQSERDSMD RQKI IKKIDNLTARQVTFSKRRRGLFKKAQELS LCDADIGLIVFSATGKLFEYSSSSM QLIEK K MQS RcM1 MPF2 MPP3 AGL24 SVP STMADS16 STMADS11 ..SQPSLELQL.EISNRVRLSKEVADKTRQLRQMRGEDLNELNVEELQQLENLLEVGLQRVTDAKGKRITNEISELERKG QPSL LQL E S RLSK VADKTRQLRQMRGEDL EL VEELQQLE LE G RV D KG RI EI LERKG ..DQPSLDLQL.ENSLNVRLRKQVADKTRELRQMKGEELEGLSLEELQQIEKRLEAGFNRVLEIKGTRIMDEIANLQRKG DQPSL LQL ENS RL K VADKTRELRQMKGEEL GL LEELQQIEK LE G RVLEIKG RIM EI LQRKG ..DQPFLDLQL.ENSLNVRLRKQVADKTRELRQMKGEELEGLSLEELQQIEKRLEAGFNRVLEIKGTRIMDEIANLQRKG DQP L LQL ENS RL K VADKTRELRQMKGEEL GL LEELQQIEK LE G RVLEIKG RIM EI LQRKG M.DPPSTHLRL.ENCNLSRLSKEVEDKTKQLRKLRGEDLDGLNLEELQRLEKLLESGLSRVSEKKGECVMSQIFSLEKRG D PS L L EN RLSK V DKTKQLR LRGEDL GL LEELQ LEK LE G RV E KG VM QI LEKRG ..DQPSLELQLVENSDHARMSKEIADKSHRLRQMRGEELQGLDIEELQQLEKALETGLTRVIETKSDKIMSEISELQKKG DQPSL LQL ENS RMSK IADK H LRQMRGEEL GL IEELQQLEK LE G RVIE KS KIM EI LQKKG ..DEPSLDLQL.ENSLNMRLSKQVADKTRELRQMRGEELEGLSLEELQQIEKRLEAGFNRVLEIKGTRIMDEITNLQRKG DEPSL LQL ENS RLSK VADKTRELRQMRGEEL GL LEELQQIEK LE G RVLEIKG RIM EI LQRKG NPEQLHSSNLLSEKKTHAMLSRDFVEKNRELRQLHGEELQGLGLDDLMKLEKLVEGGISRVLRIKGDKFMKEISSLKKKE EQ L E LSR EK RELRQLHGEEL GL LDDL LEK VE G RVL IKG KFM EI L KKE 80 79 79 79 79 79 79 K domain 157 156 156 157 157 156 159 C-terminal RcM1 MPF2 MPP3 AGL24 SVP STMADS16 STMADS11 AQLMEENK LKQKM M G P I AQLMEENKQLKQKMVMMCSGTRPAILESDITTHEEGMSSDSATV...VCSCSNGPPVEDDISDTYLKLGLSFSS...... EEG SS S V S G P EDD S LKLG AAELMEENKKLKQKMEMMKLGKFPLLTDMDCMVIEEGQSSDSIITTNNVCSSNSGPPPEDDSSNASLKLGCNNGLAAVDDD ELMEENK LKQKME M GK PLL V EEG SS S T NV S G P EDDSS SLKLG VD AAELMEENKKLKQKMEMMKLGKLPLLTDMDCMVMEEGQSSDSIITTNNVCSSNTGPPPEDDSSNASLKLGCNNGLAPVDDD ELMEENK LKQKME M GK PLL V EEG SS S T NV S G P EDDSS SLKLG VD SELVDENKRLRDKLETLERAKLTTLKEALETESV..........TTNVSSYDSGTPLEDDSDT.SLKLGLPSWE...... ELVDENK LRDKLE L AK L T NV S G P EDDS SLKLG MQLMDENKRLRQQGTQLTEENERLGMQICNNVHAHGGAE...........SENAAVYEEGQSSESITNAGNSTGAPVDSE QLMDENK LRQ L E L V G A EEG S SI A VD AAELMEENKQLKHKMEIMKKGKFPLLTD...MVMEEGQSSESIITTNN........PDQDDSSNASLKLG...GTTAVEDE ELMEENK LKHKME M GK PLL V EEG SS S T N P QDDSS SLKLG VE AAQLQEENSQLKQQSQARLNE...............................EGQNVIEQGHSADSITNNRSLVNSHQDYN QL EEN LKQ Q E EQG S SI D 228 236 236 220 226 222 208 RcM1 MPF2 MPP3 AGL24 SVP STMADS16 STMADS11 ............. CSITSLKLGLPLS S TSLKLGLP CSITSLKLGLPLS S TSLKLGLP ............. SSDTSLRLGLPYG S TSLRLGLP CSITSLKLGLPFS S TSLKLGLP DSDTSLKLCLAFP S TSLKL LA 228 249 249 220 239 235 221 Figure 4.5. Alignment of the derived amino acid sequences of RcMADS1 and other members of the StMADS11 clade. The MADS, I and K domains are all relatively conserved across the various proteins. Identical residues are coloured dark blue, conserved residues Key to sequences included: RcM1 = RcMADS1 from Rafflesia cantleyi; MPF2 and MPF3 from Physalis pubescens; AGL24 and SVP from Arabidopsis thaliana; and StMADS16 and StMADS11 from Solanum tuberosum. 39 As this cDNA had been amplified from a young and small flower bud, perhaps just past the floral transition stage and which had not quite started developing distinct floral organs (see Figure 4.1C), it seems highly probable that RcMADS1 would have a function in promoting or repressing flowering. Also, with relatively high amino acid sequence similarity to AGL24 and SVP, two proteins known to promote and repress flowering in Arabidopsis thaliana respectively, RcMADS1 could be a functional homologue of AGL24 or SVP in Rafflesia cantleyi. 4.5. Phylogenetic analysis Phylogenetic analysis of the conserved MADS-box domain using proteins from the StMADS11 clade as well as representative members of other MADS-box protein clades showed that RcMADS1 is nested with the StMADS11 clade (Figure 4.6), and it appears to be more related to AGL24 than SVP based on the subclades they are nested in. RcMADS1 is grouped with PtMADS1, a protein from Populus tomentosa, and both these two proteins are sister to a clade containing AGL24 and StMADS16. Together, RcMADS1 and AGL24 form a sister clade to a clade containing SVP and JOINTLESS (from Solanum lycopersicum). The StMADS11 clade comprises of proteins with diverse functions from a wide variety of plants from both gymnosperms and angiosperms (Becker and Theissen, 2003). The presence of homologues from gymnosperms seems to be evidence that StMADS11-like proteins have ancestral functions in controlling aspects of vegetative development, and the present functions of StMADS11 proteins in angiosperms are newly evolved after the split from gymnosperms (Becker and Theissen, 2003). This phylogenetic analysis shows that RcMADS1 is more closely related to AGL24 than to SVP, despite the higher alignment similarity between RcMADS1 and 40 SVP as shown by a BLAST search. Since AGL24 and SVP have contrasting functions in regulating flowering time, it appears likely that RcMADS1 would have a function more similar to AGL24 than to SVP. 4.6. Functional characterisation of 35S::RcMADS1 in Arabidopsis thaliana As genetic transformation of Rafflesia is not possible, due to the inherent difficulty of cultivating the endophytic and holoparasitic Rafflesia, the functions of identified genes from Rafflesia will have to be studied using some other plant model organism such as Arabidopsis thaliana. Although this would be a heterologous plant system involving genes foreign to the model organism, there are benefits: many mutants are available for complementation studies, and there is a lot of published information on ectopic expression of putative homologues in Arabidopsis thaliana. In the case of RcMADS1, there are many studies already published for AGL24 and SVP, both StMADS11-like genes. Therefore, in order to elucidate the function of the gene, we used an Arabidopsis thaliana transgenic plant system to study the expression of the gene in comparison with AGL24 and SVP. Note: RcMADS1 cDNA was recently (in January 2010) independently cloned and sequenced by Dr. Rengasamy Ramamoorthy in our laboratory. The independent sequence verification showed a 100% similarity to the original clone. 4.6.1. Construction of ectopic expression plasmid A plasmid for ectopic expression of RcMADS1 was constructed using a pGreen 0229 vector containing a BAR gene conferring resistance to Basta® (a herbicide containing glufosinate ammonium) (Figure 4.7). Arabidopsis thaliana plants transformed with the pGreen vector would survive Basta® application. The 41 LAMB1 ABS GGM2 GGM12 ANR1 FLC AGL19 SOC1 © M rEnt DAL3 OsMADS1 AGL6 AGL13 AGL3 SEP3 SEP1 SEP2 OsMADS14 SQUA FUL AP1 CAL FBP11 ZAG2 AG CyAG GGM3 DAL2 FARINELLI SHP1 SHP2 AGL12 GGM10 AGL16 AGL17 AGL21 AP3 DEF GLO PI STMADS11 AGL24 IbMADS4 STMADS16 IbMADS3 MPF2 MPP3 MPP4 PtMADS1 STMADS11 clade RcMADS1 JOINTLESS PkMADS1 SVP BM1 OsMADS47 OsMADS55 BM10 OsMADS22 Figure 4.6. Phylogenetic tree of MADS-box proteins. This consensus phylogenetic tree was generated via parsimony analysis using TNT version 1.0, with a data set based on the conserved MADS-box domain of approximately 60 amino acids. RcMADS1 is found to be nested within the StMADS11 clade (shown by arrow). 42 completed construct was sequenced to verify that the RcMADS1 gene was correctly inserted into the vector. 4.6.2. Transgenic Arabidopsis thaliana T1 phenotypes Transgenic Arabidopsis thaliana plants were obtained using the floral dip method (see Section 3.13) (Clough and Bent, 1998). Seeds produced after floral dipping were sown and germinated, and seedlings were subjected to 205 mg/l Basta® selection. For the T1 generation, over 30 transgenic plants were generated after Basta® screening (Table 4.1), with three distinctive phenotypes observed. Seven of the T1 plants had a ‘strong’ phenotype where some homeotic conversion of floral organs were observed: sepals and petals were converted to leaflike structures bearing conspicuous trichomes. Additionally, the development of the carpel was abnormal. Instead of developing into a silique after self-fertilisation, the gynophore elongated and the carpel itself usually split and developed an inflorescence. Most of the time, siliques were not formed, and the plants did not produce viable seed. Six plants had a ‘weak’ phenotype where full homeotic conversion of floral organs did not occur, unlike in the ‘strong’ phenotype, but there were some alterations to the development of the floral organs. The sepals became more leaf-like. Both sepals and petals were persistent, not abscising when the flower grew older. Similar to those in the ‘strong’ phenotype, the carpels did not develop normally into siliques most of the time, and viable seed was not produced. This phenotype is similar to what was observed for the attenuated phenotype with only bract-like sepals in 35S::AGL24 hemizygotes (Yu et al., 2004). The remaining 18 plants generated after Basta® screening of the T1 seeds had a ‘normal’ phenotype where the phenotype was similar to that of wild-type plants. 43 Table 4.1. Phenotype analysis of T1 transgenic plants generated Phenotype Morphology Number of plants generated Mean number of rosette leaves formed before bolting Normal Flowers look like wild type. 18 11.7 ± 1.7 Weak Sepals become leaf-like, persistent, not abscising when silique matures. 6 10.8 ± 2.6 Strong Sepals and petals become leaf-like, with presence of trichomes; abnormal carpel development; iterative inflorescence formation 7 11.7 ± 1.7 44 Most of the plants showing altered phenotype (especially with severely deformed carpels) did not produce viable seeds. Hence, seeds from only 17 independent transgenic lines harbouring 35S::RcMADS1 were recovered. Due to the ‘strong’ phenotype exhibited by some of the T1 plants which bear some similarities to AGL24 overexpression in Arabidopsis thaliana plants (Yu et al., 2002), some lines were chosen for further analysis in the T2 generation and beyond. 4.6.3. 35S::RcMADS1 effects on flowering time Some transgenic Arabidopsis lines harbouring 35S::RcMADS1 were selected for an analysis of the effect of the transgene on flowering time (Table 4.2). Lines ETL01, ETL04, ETL05, and ETL09 had earlier exhibited no significant changes in flower morphology in the T1 generation, while ETL06 has shown some mild changes in floral morphology (i.e. the ‘weak’ phenotype), and ETL12 and ETL14 are transgenic lines with total conversion of floral organs into leaf-like structures (i.e. the ‘strong’ phenotype). The T2 plants show a correlation in terms of flowering time. Lines ETL01, ETL04, ETL05, and ETL09 had similar flowering time to wild-type plants, with 10–11 rosette leaves before bolting. ETL06 (a ‘weak’ phenotype line) segregated into a normal-looking phenotype, and a ‘weak’ phenotype with mild floral organ homeosis, and the flowering time was lowered to a mean of 8.2 ± 1.2 and 5.5 ± 0.7 rosette leaves, respectively, before bolting. ETL12 (a ‘strong’ phenotype line) segregated into a normal-looking phenotype, and a ‘strong’ phenotype exhibiting total strong floral organ homeosis, and the flowering time was a mean of 10.4 ± 1.9 and 7.4 ± 1.4 rosette leaves before bolting. ETL14 (another ‘strong’ phenotype line) only produced ‘strong’ phenotype plants in the T2 generation, and the flowering time was 6.5 ± 0.9 leaves before bolting. These results suggest that an earlier flowering 45 Table 4.2. Comparison of flowering times of 35S::RcMADS1 T2 lines Line ‘Normal’ phenotype ‘Weak’ phenotype no. of plants mean no. of leaves no. of plants Col WT 48 15.9 ± 1.2 48 ETL01 9 11.7 ± 1.1 32 ETL04 25 11.4 ± 1.0 31 ETL05 24 11.6 ± 1.2 28 ETL06 14 8.2 ± 1.2 ETL09 23 10.1 ± 0.8 ETL12 23 10.4 ± 1.9 ETL14 2 ‘Strong’ phenotype mean no. no. of of leaves plants mean no. of leaves 5.5 ± 0.7 Total number of plants 24 28 7 7.4 ± 1.4 34 18 6.5 ± 0.9 22 Notes: Flowering time is presented as the number of rosette leaves on the main shoot when the inflorescence was ≈ 3 cm in height. “Total number of plants” refer to plants sown before Basta® screening (but not applicable to Columbia wild-type plants, which acted as controls). 46 time was associated with floral organ homeosis. Lines that had a transgene insert (conferring resistance to Basta®) but no discernible difference in phenotype had flowering times similar to that of wild-type plants. Lines that showed observable changes in floral morphology had distinctly earlier flowering times (as seen in ETL06, ETL12, and ETL14). 4.6.4. Floral morphology in 35S::RcMADS1 transgenic lines The morphology of the transgenic lines that showed the ‘strong’ phenotype were further studied in the T2 and T3 generations, alongside two lines that expressed the ‘strong’ phenotype only in the T2 generation and beyond. In the T1 generation, the lines ETL10 and ETL17 did not express any discernible changes in floral morphology, but expressed floral homeosis only in the T2 generation. As a comparison, ETL01, a wild-type-looking line was also studied (Figure 4.7). The habit, floral morphology, and silique development of ETL01 all closely resembled that of wild-type plants. In the ‘strong’ phenotype, changes in floral morphology were drastic in the lines observed, namely ETL10, ETL12 (Figure 4.8), ETL14 (Figure 4.9), and ETL17. Both sepals and petals were converted into leaf-like structures, with the second whorl losing the petalloid appearance, and developed conspicuous trichomes typically seen in leaves (Figure 4.10). Secondary inflorescences developed from the axils of the whorls. The carpel developed abnormally, with an elongation of the gynophore, and silique formation was aborted. Secondary inflorescences were observed developing from the aborted silique (Figure 4.11). In some plants of ETL14, the development of the inflorescence proceeded directly from the fourth whorl, and bypassed the intermediate step of carpel formation and development (see Figure 4.8B). 47 A B C D E F Figure 4.7. Phenotype of wild-type-looking transgenic line ETL01. (A) Flower, showing typical whorls, resembling wild-type flower. (B) Habit. (C) Comparison of the habits of wild-type plant (left) and ETL01 transgenic plant (right). (D) and (E) Branch of plant showing an inflorescence with wild-typelooking flowers. (F) Comparison of the inflorescences and flowers of wild-type plant (left) and ETL01 transgenic plant (right). 48 A B C D E F Figure 4.8. Phenotype of strong transgenic line ETL12. (A) Flower, showing conversion of sepals and petals into leaf-like structures. (B) Habit. (C) Comparison of the habits of wild-type plant (left) and ETL12 transgenic plant (right). ETL12 plants display more profuse branching. (D) Branch of ETL12 transgenic plant. (E) Comparison of branches of wild-type plant (left) and ETL12 transgenic plant (right). (F) A secondary inflorescence developing from an aborted carpel. 49 A B D E C F Figure 4.9. Phenotype of strong transgenic line ETL14. (A) Flower, showing conversion of sepals and petals into leaf-like structures with conspicuous trichomes, and conversion of the fourth whorl into an inflorescence. (B) Flower showing a secondary inflorescence developing directly from the fourth whorl. (C) Habit. (D) Branch of ETL14 transgenic plant. (E) Comparison of the branches of wild-type plant (left) and ETL14 transgenic plant (right). (F) Comparison of the habits of wild-type plant (left) and ETL14 transgenic plant (right). 50 A B C D Figure 4.10. Altered floral morphology due to ectopic expression of 35S::RcMADS1 in Arabidopsis thaliana. (A) Arabiodosis thaliana wild-type flower showing all four whorls of floral organs. (B) 35S::RcMADS1 flower showing a deformed carpel on an elongated gynophore, and conversion of sepals and petals to leaf-like structures bearing conspicuous trichomes. (C) 35S::RcMADS1 flower with a secondary inflorescence developing from the axil of a sepal (red arrowhead). (D) 35S::RcMADS1 flower bearing multiple secondary inflorescences (red arrowheads). 51 A B C D Figure 4.11. Secondary inflorescence development in 35S::RcMADS1 plants. (A) In T2 lines observed (plant here is from ETL14), the secondary inflorescence develops from the aborted silique through a split in the carpel. (B) 35S::RcMADS1 flower from a T3 line (ETL14) whose fourth whorl directly develops into an inflorescence, indicated by the red arrowhead. (C) A more developed 35S::RcMADS1 flower from a T3 line (ETL14), showing the elongation of the secondary inflorescence which developed from the fourth whorl. There is no intermediate step of carpel/silique development. (D) Closer view of the secondary inflorescence developed directly from the fourth whorl of the flower. 52 Due to the iterative inflorescence development, transgenic lines harbouring the 35S::RcMADS1 displayed rather profuse branching habits (Figure 4.12). Secondary inflorescences would develop from the axils of the fourth whorl or from carpels/gynophores, which would in turn give rise to further inflorescences developing from the abnormal flowers on the secondary inflorescences. 4.7. Molecular characterisation of selected transgenic lines 4.7.1. Genomic Southern blot analysis All T2 transgenic Arabidopsis thaliana lines analysed in the genomic Southern blot were found to have two insertions of the 35S::RcMADS1 transgene (Figure 4.13). The use of two different restriction endonucleases in the analysis revealed that the insertions were in different loci and that the transgenic lines were independently generated. This indicates that the changes in flowering time and morphology observed were due to the effect of the transgene and not likely to be due to a direct disruption of native gene function by the insertions. 4.7.2. Quantitative real-time PCR analysis To show that the transgene RcMADS1 is expressed in the various transgenic lines, we performed quantitative real-time PCR and compared its expression across the different transgenic lines (Figure 4.14). Five transgenic lines (ETL01, ETL10, ETL12, ETL14, and ETL17) were selected. ETL01 and ETL10 are transgenic lines that are wild-type-looking, with no discernible changes in floral morphology or in flowering time. ETL12, ETL14, and ETL17 are transgenic lines showing a ‘strong’ phenotype, with severe altered floral morphology and early flowering. All lines examined showed increases in expression levels of RcMADS1, indicating successful 53 A B C Figure 4.12. Development of profuse branching. (A) Wild-type Arabidopsis thaliana on the left, 35S::RcMADS1 transgenic line on the right. Note the more profuse branching in the transgenic plant. (B) General habit of wild-type Arabidopsis thaliana (left) and 35S::RcMADS1 transgenic line (right). (C) Branch from a 35S::RcMADS1 transgenic plant, showing development of secondary inflorescences from an aborted carpel (indicated by red arrowhead), and subsequent abnormal development of the carpels in the flowers on the secondary inflorescences. 54 HindIII L 01 10 12 14 17 XbaI 01 10 12 14 17 L Figure 4.13. Genomic Southern blot analysis. Genomic DNA from various transgenic lines was digested with either HindIII or XbaI, as indicated above the lanes, before hybridisation with the RcMADS1-specific probe. Transgenic lines examined include ETL01 and ETL10 (‘normal’ phenotype); ETL12, ETL14, and ETL17 (‘strong’ phenotype). ‘L’ indicates DNA Marker III. 55 Figure 4.14. Comparison of expression levels of RcMADS1 across transgenic lines. Leaves of selected transgenic lines were collected for quantitative real-time PCR analysis of RcMADS1 expression in the Arabidopsis thaliana plants harbouring 35S::RcMADS1. ETL01 and ETL10 are wild-type-looking lines; while ETL12, ETL14, and ETL17 show a ‘strong’ phenotype with severely altered floral morphology. All transgenic lines showed increased expression levels. Relative to ETL01, the other lines had expression levels 4 to 16 folds higher. Error bars represent SD. 56 insertion and expression in these transgenic lines. Using ETL01 as the control, the other transgenic lines showed expression levels 4 to 16 folds higher, after normalisation. This shows that increased levels of RcMADS1 may have an effect on floral development. In Arabidopsis thaliana, AGL24 and SVP regulate and are regulated by a host of genes in the control of floral meristem identity. These genes include AP1, CO, FLC, FT, LFY, SOC1, and SEP3. To examine the effect of ectopic expression of RcMADS1 in Arabidopsis thaliana and elucidate the putative function of this gene, we performed quantitative real-time PCR to compare the expression of all these genes that RcMADS1 may interact with: AGL24, AP1, CO, FLC, FT, LFY, SEP3, SOC1, and SVP. This preliminary survey showed that FLC was upregulated (Figure 4.15A) while FT was downregulated (Figure 4.15B) in transgenic Arabidopsis lines displaying the ‘strong’ phenotype (i.e. ETL12, ETL14, and ETL17), whereas gene expression of FLC and FT in wild-type-looking transgenic lines (ETL01 and ETL10) were not significantly different from that in Columbia wild-type plants. All other genes studied did not show any changes in expression levels. A decrease in FT expression levels would most probably be due to an increase in FLC expression levels, since FLC acts as a repressor of FT (Michaels et al., 2003). In Arabidopsis thaliana, FLC acts in tandem with SVP to repress FT expression, which would otherwise activate SOC1 and AP1 expression (Liu et al., 2009a), and FLC itself also directly represses SOC1 expression. SOC1 and AGL24 directly upregulate the expression of one another, and together form a protein complex which activates LFY. AGL24 therefore directly and indirectly upregulates SOC1 and LFY respectively, but the data did not show increases in SOC1 and LFY expression levels, indicating that RcMADS1 does not quite behave like AGL24 in regulating native 57 A B Figure 4.15. Effect of RcMADS1 ectopic expression on FLC and FT. Leaves of selected transgenic lines were collected for quantitative real-time PCR analysis of genes that may be affected by RcMADS1 ectopic expression. (A) FLC expression had increased 8 to 12 folds in transgenic lines displaying severe alterations in floral morphology (ETL12, ETL14, and ETL17), while remaining relatively unchanged in transgenic lines that are wild-type-looking (ETL01 and ETL10). (B) FT expression was contrastingly decreased, about 2 to 5 folds compared to Columbia wild type, in ETL12, ETL14, and ETL17; FT expression levels were relatively unchanged in ETL01 and ETL10. Error bars represent SD. 58 Arabidopsis thaliana genes downstream of AGL24, as might be expected of a putative homologue of AGL24. While an increased level of SVP (or its homologue) may explain the increase of FT through the action of the SVP–FLC complex, but it does not explain the increased expression of FLC itself. The respective changes in gene expression of FLC and FT were both correlated to increased expression of RcMADS1 in the ‘strong’ transgenic lines, indicating that RcMADS1 acts in a dosage-dependent manner, much like AGL24. However, since the material used were leaves harvested after bolting had occurred, not much conclusive data can be gained. Almost all of the pertinent interactions occur in the shoot apical meristems during floral transition (Liu et al., 2009a). AGL24 does not appear to upregulate FLC natively in Arabidopsis thaliana (Michaels et al., 2003), whereas RcMADS1 does, as shown in the quantitative realtime PCR data (Figure 4.15). While RcMADS1 and AGL24 are similar in amino acid sequence, and in phenotypic effect during ectopic expression or overexpression in Arabidopsis thaliana, it seems possible that RcMADS1 may interact with protein partners in a manner rather different from AGL24. This could also indicate a different native function in Rafflesia, despite the similarities in phenotypes between 35S::AGL24 and 35S::RcMADS1 when ectopically expressed in Arabidopsis thaliana. 59 CHAPTER 5 GENERAL DISCUSSION AND FUTURE WORK 5.1. RcMADS1 may be involved in regulation of flowering time In Arabidopsis thaliana, AGL24 is an integrator of flowering signals leading to a precise regulation of floral meristem specification (Liu et al., 2009a). RcMADS1, a putative homologue of AGL24 from Rafflesia cantleyi may have such a function, similar to genes from other species in the StMADS11 clade, such as INCO from Antirrhinum majus (Masiero et al., 2004), BM1 from Hordeum vulgare (Trevaskis et al., 2007), and OsMADS22 and OsMADS47 from Oryza sativa (Fornara et al., 2008). Ectopic expression of RcMADS1 resulted in early flowering and floral reversion, with altered floral morphology that are largely similar to results obtained in similar studies overexpressing AGL24 in Arabidopsis thaliana (Yu et al., 2002; Yu et al., 2004). Moreover, the effects of ectopic expression of RcMADS1 is dosagedependent, as is the case for AGL24 (Yu et al., 2002). This suggests RcMADS1 is a functional homologue of AGL24 in Rafflesia cantleyi. However, the preliminary quantitative real-time PCR data suggests that there are some differences between RcMADS1 and AGL24 when interacting with intermediates in the regulatory network of AGL24 in Arabidopsis thaliana. AGL24 has not been known to upregulate FLC. Other than that, there is no enough data for quantitative real-time PCR at this stage to draw more definite conclusions. StMADS11-like genes are quite diverse in function. Besides regulating flowering time and specification and maintenance of floral meristems in Arabidopsis thaliana (Liu et al., 2009a), there are StMADS11-like genes with other functions in development. For example, PkMADS1 is involved in shoot induction and phyllotaxy 60 in Paulownia kawakamii (Prakash and Kumar, 2002); MPF2-like genes are involved in development of the inflated calyx syndrome in Solanaceae (Hu and Saedler, 2007); OsMADS22, OsMADS47, and OsMADS55 are involved in regulation of brassinosteroid responses in Oryza sativa (Fornara et al., 2008). The study of the rice StMADS11-like genes suggested that heterologous overexpression of StMADS11-like genes in Arabidopsis thaliana causes the same phenotypes as that of overexpression of AGL24 and SVP genes. This could be due to inappropriate protein–protein interactions that disturb normal flower development (Fornara et al., 2008), and is no indication of the real function of such genes in the native non-Arabidopsis thaliana plants. This finding was underscored by the fact that the StMADS11-like genes from rice did not complement agl24 and svp mutants (Fornara et al., 2008). While the phenotypes observed from transgenic plants harbouring 35S::RcMADS1 are shown to be similar to those overexpressing AGL24, and shows that RcMADS1 is a StMADS11like gene with possibly similar function to AGL24, it is not conclusive evidence for the actual native function in Rafflesia cantleyi since all StMADS11-like genes cause similar floral reversions when overexpressed in Arabidopsis thaliana. 5.2. Temporal and spatial expression of RcMADS1 in Rafflesia cantleyi Ideally, there has to be more information on when, where, and how RcMADS1 is expressed natively. This would allow us to have a better understanding of the possible mechanisms of regulation RcMADS1 may be controlling in Rafflesia cantleyi. Due to technical difficulties in isolating good quality RNA and DNA in appreciable quantities, it was not possible to obtain data on temporal and spatial expression patterns of RcMADS1. Additionally, it would be difficult to study the stage before floral transition, given that Rafflesia is holoparasitic — the vegetative tissues are 61 threadlike and embedded within the host plant tissues, and therefore are difficult to isolate. 5.3. Future work The quantitative real-time PCR data for the effects of ectopically expressing RcMADS1 in Arabidopsis thaliana is quite preliminary in nature, using leaf tissue after bolting had occurred. For a clearer picture of how RcMADS1 interacts with other flowering time genes and floral development genes, changes in gene expression levels of target genes should be studied during the floral transition stage. This would allow more appropriate examination of the respective changes of various members of the regulatory network involved in integrating flowering signals, forming the inflorescence meristems, and in acquiring and maintaining floral meristem identity. Protein–protein interaction studies can also be carried out to elucidate the differences (in interaction with protein partners) between AGL24 and RcMADS1. More importantly, temporal and spatial expression patterns of RcMADS1 (and possibly other MADS-box genes identified from Rafflesia) should be studied, in order to better understand the role and function in Rafflesia itself. 5.4. Conclusions Here, we report the cloning and characterisation of a MADS-box gene from Rafflesia cantleyi. This cDNA has been named RcMADS1. The full-length cDNA was cloned using a degenerate PCR approach. RcMADS1 shows high sequence similarity to several MADS-box genes, in particular SVP and AGL24, all of which belong to the StMADS11 clade. Ectopic expression of this gene (35S::RcMADS1) in a heterologous system, namely Arabidopsis thaliana, resulted in a floral phenotype similar to that of 62 35S::AGL24. Quantitative real-time PCR analysis of selected target genes in the regulatory network of AGL24 and SVP were performed in transgenic Arabidopsis thaliana harbouring 35S::RcMADS1. To the best of our knowledge, this is the first report of cloning and partial functional characterisation of a floral pathway gene from Rafflesia. 63 References Alvarez-Buylla, E. R., S. Pelaz, S. J. Liljegren, S. E. Gold, G. Burgeff, G. S. Ditta, L. Ribas de Pouplana, L. Martinez-Castilla, and M. F. Yanofsky. 2000. An ancestral MADS-box gene duplication occurred before the divergence of plants and animals. 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The Plant Journal 37: 566–577. 70 [...]... reactions were performed using 20 Table 3.1 Degenerate primers used in cloning MADS- box genes from Rafflesia cantleyi Name Sequence (5′→3′) Direction Reference MADS1 AARMGIMGIAAYGGIYTIYTIAARAARGC Forward N .A MADS2 GGGGTACCAAYMGICARGTIACITAYTCIAAGMGIMG Forward N .A MADS3 AARAARGCIYWYGARCTIKCKGTICT Forward N .A MADS4 AAYMGRCARGTICAITAYTCRAARMG Forward Di Stilio et al., 2005 MADS5 GGIMGIAARATIGARATIAARRGIAT... GGIMGIAARATIGARATIAARRGIAT Forward Di Stilio et al., 2005 MADS6 AAYRGICARGTIACITTYTGYAARRGIRG Forward Di Stilio et al., 2005 MADS7 CAYTTRATGGGIGARGCICTIAGYTG Forward Di Stilio et al., 2005 MADS8 GGACGAGGACGDGTWCARCT Forward Jager et al., 2003 MADS9 SAGATCAAGMGIATHGARAAY Forward Jager et al., 2003 MADS1 0 GGGGTACCAAYMGICARGTIACITAYTCIAAGMGIMG Forward Kramer et al., 1998 oligo(dT) TTTTTTTTTTTTTTT Reverse N .A 21 step-up... cantleyi Solms- Laubach; Rafflesia gadutensis Meijer; Rafflesia hasseltii Suringar; Rafflesia keithii Meijer; Rafflesia kerrii Meijer; Rafflesia manillana Teschemacher; Rafflesia micropylora Meijer; Rafflesia patma Blume; Rafflesia pricei Meijer; Rafflesia rochussenii Teijsm & Binn.; Rafflesia schadenbergiana Göpp.; and Rafflesia tengku-adlinii Salleh & Latiff Since 2002, 10 or 11 new species have been... species of angiosperms and gymnosperms, Barkman et al (2004) placed Rafflesia and Rhizanthes within the order Malpighiales, with sister families such as Passifloraceae, Salicaceae, and Violaceae Rafflesiaceae was more confidently placed within the Malpighiales as nested in Euphorbiaceae using more data (five mitochondrial and one chloroplastic genes) from a focused sampling of species from all families of. .. functional genomics and developmental biology 2.1.4 Rafflesia cantleyi Solms- Laubach Rafflesia cantleyi Solms- Laubach is a species with relatively smaller flowers, compared to some of the better-known and large-flowered species such as Rafflesia arnoldii and Rafflesia keithii (Meijer, 1997) It is found in Malaysia, in the states of Perak, Kelantan, Pahang, and Kedah Up to 1984, this species was considered... as 6 well as two others from outside the Philippines, bringing the number of currently recognised and described Rafflesia species to 27 (Barcelona et al., 2009) The new Philippine species are: Rafflesia baletei Barcelona & Cajano; Rafflesia leonardi Barcelona & Pelser; Rafflesia lobata R.Galang & Madulid; Rafflesia mira Fernando & Ong; Rafflesia philippensis Blanco; and Rafflesia speciosa Barcelona... CHAPTER 3 MATERIALS AND METHODS 3.1 Plant materials Flower buds of various sizes of Rafflesia cantleyi Solms- Laubach were collected from a few localities along a trail between Tekek and Juara in Pulau Tioman, Pahang, Malaysia Collection of Rafflesia cantleyi material in Peninsular Malaysia required a permit (permit number: UPE40/200/19 SJ 1200) from the Economic Planning Unit, Prime Minister’s Department... lack of type specimens (Meijer, 1997); these species include Rafflesia borneensis Koord.; Rafflesia ciliata Koord.; Rafflesia titan Jack; Rafflesia tuan-mudae Becc.; and Rafflesia witkampii Koord In his treatment, Meijer (1997) accepted 13 species: Rafflesia arnoldii R.Br., with two varieties: Rafflesia arnoldii var arnoldii R.Br., and Rafflesia arnoldii var atjehensis (Koord.) Meijer; Rafflesia cantleyi. .. Rafflesia have many physiological and morphological adaptations as a result of their evolution and have lost many plant structures such as leaves, stems and roots, 1 thus making phylogenetic relationships with non-parasitic plants difficult Barkman et al (2004) sequenced the mitochondrial gene matR and produced a broad phylogenetic tree that showed a placement of Rafflesia in the Malpighiales Rafflesia was... some of these gaps in the genetic architecture of floral development, leading to the objectives of this study: 1) To clone one or more MADS- box genes from Rafflesia cantleyi, a species of Rafflesia from Pulau Tioman, Pahang, Malaysia, using a degerate PCR approach; 2) To identify and analyse the cloned MADS- box gene( s) through sequencing and phylogenetic analysis; 3) To characterise the function(s) of ... 361  TGCATTTCAATAACAAAGAGAAGCCAAGCCAGCCATCTCTTGAACTGCAGCTAGAAATTA  420 72   L  H  F  N  N K  E  K  P  S  Q  P  S  L  E  L  Q  L  E  I 91 421  GCAATCGGGTGCGACTGAGCAAAGAAGTTGCAGACAAGACTCGCCAACTAAGGCAAATGA  480... 541  AGGTGGGCCTCCAGCGCGTTACTGATGCCAAGGGCAAGCGCATCACAAATGAGATATCTG  600 151 132  E  V  G  L  Q  R  V  T  D  A  K  G  K  R  I  T  N  E  I  S 601  AACTCGAAAGGAAGGGAGCGCAGCTGATGGAAGAAAATAAGCAACTAAAGCAAAAAATGG  660... 1    GGGTTAGGTTTCCATATGCCCCACGGGCCACATATTCTTGAACCTGTCTAGCCTTCCCCA  60 61   ATTTATTGCCCGAAGCTGTACGTTCATATTTCGAAACTAAAGATTCGGTCGTGCCGTATT  120 121  TGTTCTGTTTCGGAGCAGATAATATAATGGCTCGAGAAAAGATCAAGATCAAAAAGATCG  180 1                               M  A  R  E  K  I  K  I  K  K