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Investigation of the genomics of gender regulation in Populus trichocarpa Temmel, Nyssa A. 2011

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INVESTIGATION OF THE GENOMICS OF GENDER REGULATION IN POPULUS TRICHOCARPA  by Nyssa A. Temmel  B.Sc. (Hons) University of Victoria, 2002  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  December 2011  © Nyssa A. Temmel, 2011  Abstract This thesis reports the findings of four projects conducted to study the genomics of gender regulation in Populus trichocarpa. Sex-linked markers previously discovered in Salix vimilanis were tested to determine if they were also sex-linked in other Salix species and P. trichocarpa. It was found that the DNA sequence of the SCAR 354 marker, and its position at the 5’ end of a gene encoding an Ssu72-like protein, was conserved with some SFP variability in species of Salix and P. trichocarpa. While this marker may be useful for phylogenetic or population studies in Salix, this marker was not sex-linked in the species investigated in this study. An investigation of genes located on the telomeric end of chromosome 19, the putative sex chromosome in P. trichocarpa, was conducted to look for gender-biased SNPs that would indicate recombination suppression in the region on a sex locus.  A large  variability in the number of SNPs was observed in the gene sequences studied, but no SNPs that segregated with gender were discovered so a genetic marker that could be used to sex P. trichocarpa individuals of unknown gender could not be developed. Using a microarray approach, gender-biased gene-expression was studied in leaf tissue of P. trichocarpa.  While some gender-biased gene-expression was observed in  vegetative tissues the differences observed were statistically insignificant due to biological variation in the samples tested, the small sample size used in this study, and changes in the genome annotation between version 1.1 and 2.0 of the poplar genome. This study could not verify the microarray results using rtPCR in a larger sample of male and female leaf tissue. MADS-box genes involved in floral development were identified as having genderbiased gene-expression using a microarray approach. Thirteen putative MADS-box genes  ii  that showed gender-biased expression in male and female inflorescences were discovered. Novel expression patterns for nine floral MADS-box genes were identified with this microarray data, and the expression patterns of three of these genes were investigated in further detail using reverse-transcription PCR.  iii  Preface A version of chapter two has been published. Temmel, Nyssa A., Rai, Hardeep S., and Cronk, Quentin C.B. (2007) Sequence characterization of the putative sex-linked Ssu72like locus in willow and its homologue in poplar. Canadian Journal of Botany 85(11):10921097 I wrote the majority of the manuscript and was responsible for all the laboratory work, while Dr. Rai contributed the DNA sequence editing and alignment, and Dr. Cronk assisted with the writing and the research was conducted in his laboratory. In chapter three and four, Dr. Rick White, the Managing Director of the Statistical Consulting and Research Laboratory at the University of British Columbia, did the statistical analysis of the microarray data. Collection of the P. trichocarpa leaf and floral buds from trees on the UBC Vancouver campus that were used for RNA, DNA and cDNA preparation was assisted by Collin Varner, the Horticulturist/Arborculturist with UBC Plant Operations and Gregg Doughty, the UBC Plant Operations arborist.  iv  Table of Contents  Abstract .............................................................................................................................. ii Preface ................................................................................................................................iv Table of Contents ................................................................................................................v List of Tables ................................................................................................................... xiii List of Figures...................................................................................................................xiv List of Abbreviations........................................................................................................xvi Acknowledgements..........................................................................................................xvii Dedication.........................................................................................................................xix Chapter 1: Introduction.....................................................................................................1 1.1  Sexual systems in plants ...........................................................................................1  1.2  Evolution of dioecy ..................................................................................................3  1.2.1 Gynodioecious pathway .....................................................................................3 1.2.2 Heterostyly pathway ..........................................................................................4 1.2.3 Monoecious pathway .........................................................................................5 1.2.4 Evolution of dioecy in Populus ..........................................................................5 1.3  Sex chromosomes in plants.......................................................................................6  1.3.1 Identification and description of sex chromosomes ............................................7 1.3.2 A brief history of the discovery of sex chromosomes .........................................7 1.3.3 Discovery of sex chromosomes in plants............................................................8 1.3.4 Evolution of sex chromosomes...........................................................................9 1.3.5 Proposed mechanism for the evolution of sex chromosomes ............................10  v  1.3.6 Degeneration of the Y chromosome .................................................................12 1.3.7 The absence of sex chromosomes in Populus ...................................................14 1.4  The genetics of gender............................................................................................14  1.4.1 Gender-determining genes ...............................................................................15 1.4.2 Gender-differentiation and development in plants ............................................16 1.4.3 Gender-differentiation in Zea mays ..................................................................17 1.4.4 Gender-differentiation in Cucumis sativus and C. melo ....................................18 1.4.5 The role of MADS box genes in gender differentiation ....................................21 1.4.6 Gender-associated genes ..................................................................................23 1.4.7 Differences in genetic control of gender-determination between monoecy and dioecy …………………………………………………………………………………. 24 1.5  Populus trichocarpa as a model organism for studying gender-determination in  plants...............................................................................................................................25 1.5.1 Life history of Populus trichocarpa .................................................................25 1.5.2 Characteristics of Populus trichocarpa as a model species ...............................26 1.5.3 Genomic markers linked to sex-differentiation in Populus and Salix ................27 1.5.4 Gender-determination and chromosome 19 in Populus.....................................28 1.6  Research objectives ................................................................................................30  Chapter 2: Characterization of sex-linked genetic markers developed in Salix viminalis in other Salix species and Populus trichocarpa.................................................................31 2.1  Introduction............................................................................................................31  2.1.1 Sex-linked genetic markers in Salix..................................................................32 2.2  Objectives...............................................................................................................33  vi  2.3  Methods..................................................................................................................34  2.3.1 Collection and preparation of plant materials ...................................................34 2.3.2 PCR conditions and sequencing of samples......................................................35 2.3.3 Sequencing alignment for the SCAR sequences amplified in Salix ...................36 2.3.4 Novel primer design based on SCAR sequence and associated coding gene regions in P. trichocarpa .............................................................................................36 2.3.5 Cloning, sequencing and phylogenetic analysis of S. arctica and S. reticulata samples .......................................................................................................................37 2.4  Results....................................................................................................................38  2.4.1 Amplification and characterization of the SCAR 354 marker ...........................38 2.4.2 Gene-anchored amplification in Salix and P. trichocarpa.................................40 2.4.3 Identifying SNP differences that may explain why this SCAR marker appears to be sex-linked ...............................................................................................................41 2.5  Discussion ..............................................................................................................47  2.5.1 SCAR 354 amplification in Salix......................................................................47 2.5.2 Amplification of the Ssu72-like gene region in Salix and P. trichocarpa ..........48 2.5.3 Identifying single feature polymorphisms (SFPs) that may explain why this region is sex-linked in some Salix species....................................................................49 2.5.4 Potential utility of this region...........................................................................50 2.5.5 Considering this research and version 2.0 of the Populus trichocarpa genome. 51 2.6  Conclusions ............................................................................................................54  Chapter 3: Investigation of chromosome 19 of the Populus trichocarpa genome to identify molecular genetic markers that can be used to identify gender.........................55  vii  3.1  Introduction............................................................................................................55  3.1.1 Investigation of sex chromosomes and sex loci to identify gender-determining genes …………………………………………………………………………………. 55 3.1.2 Sex-determination in the genus Populus...........................................................56 3.1.3 Features of chromosome 19 that may be involved in gender-determination ......59 3.2  Objectives...............................................................................................................60  3.3  Methods..................................................................................................................61  3.3.1 Collection and preparation of biological materials............................................61 3.3.2 In silico investigation of the gene content of scaffold 117 ................................61 3.3.3 PCR conditions and sequencing of samples......................................................62 3.3.4 Sequence and polymorphism data analysis.......................................................63 3.4  Results....................................................................................................................64  3.4.1 Investigation of gene content of scaffold 117 and the telomeric end of chromosome 19 ...........................................................................................................64 3.4.2 Analysis of sequence and polymorphism data obtained for eleven genes located on the telomeric region of chromosome 19 ..................................................................66 3.5  Discussion ..............................................................................................................71  3.5.1 Identifying if the homogametic gender of Populus trichocarpa ........................71 3.5.2 Version 1.1 versus version 2.0 of the poplar genome........................................72 3.5.3 Developing a genetic marker on chromosome 19 that segregates with gender in Populus trichocarpa ....................................................................................................73 3.5.4 Investigation of recombination rates in genes located in the telomeric region of chromosome 19 in P. trichocarpa................................................................................73  viii  3.5.5 Sex-linked markers on chromosome 19 of P. trichocarpa ................................74 3.5.6 Nucleotide diversity of genes located on the telomeric end of chromosome 19 in P. trichocarpa .............................................................................................................75 3.5.7 Nucleotide diversity observed in other species of Populus................................77 3.6  Conclusions ............................................................................................................78  Chapter 4: Exploration of genome-wide gender-biased gene-expression patterns to identify genes involved in gender-determination in Populus trichocarpa........................80 4.1  Introduction............................................................................................................80  4.1.1 The biology of gender-specific differences in dioecious species .......................81 4.1.2 Secondary sexual characteristics in plants ........................................................82 4.1.3 Clustering of differentially expressed genes near the sex locus.........................83 4.1.4 Investigating gender-biased gene-expression using genome wide survey approaches...................................................................................................................84 4.2  Objectives...............................................................................................................86  4.3  Methods..................................................................................................................87  4.3.1 Collection of and preparation of biological materials........................................87 4.3.2 cDNA preparation and PCR conditions ............................................................87 4.3.3 Microarray gene-expression profiling experiment and statistical analysis of the microarray data............................................................................................................88 4.3.4 Investigation of gender-biased gene-expression patterns on chromosome 19....91 4.4  Results....................................................................................................................92  4.4.1 Results from the statistical analysis of the P. trichocarpa leaf tissue data from both microarray experiments .......................................................................................92  ix  4.4.2 Investigation of gender-biased gene-expression patterns on chromosome 19....96 4.5  Discussion ..............................................................................................................99  4.5.1 Sample size on statistical power of microarray experiments .............................99 4.5.2 Effect of biological variation on interpreting microarray results .....................100 4.5.3 Interpreting gender-biased gene-expression using genome wide survey approaches.................................................................................................................103 4.5.4 Considerations when working with the developing model system of Populus trichocarpa................................................................................................................104 4.6  Conclusions ..........................................................................................................106  Chapter 5: MADS-box genes in the genus Populus and their role in floral development ..........................................................................................................................................109 5.1  Introduction..........................................................................................................109  5.1.1 Evolution of MADS-box genes in plants ........................................................110 5.1.2 MADS-box genes and their role in floral development ...................................110 5.1.3 MADS-box gene research in the genus Populus. ............................................111 5.2  Objectives.............................................................................................................113  5.3  Methods................................................................................................................114  5.3.1 Collection of and preparation of biological materials......................................114 5.3.2 cDNA preparation and PCR conditions ..........................................................115 5.3.3 Microarray gene-expression profiling experiment ..........................................115 5.3.4 Updating information on MADS-box genes in P. trichocarpa from version 1.1 to version 2.0 of the poplar genome...............................................................................116 5.3.5 Phylogenetic analysis of MADS-box genes in P. trichocarpa........................117  x  5.4  Results..................................................................................................................118  5.4.1 Updating available MADS-box gene research from version 1.1 to version 2.0 of the poplar genomes....................................................................................................118 5.4.2 Identifying P. trichocarpa MADS-box genes showing gender-biased expression using a microarray approach......................................................................................121 5.4.3 Identification of putative gene function based on protein sequence homology for P. trichocarpa MADS-box genes...............................................................................124 5.4.4 Confirming the MADS-box gene microarray results using cDNA expression experiments ...............................................................................................................128 5.5  Discussion ............................................................................................................131  5.5.1 MADS-box genes in the second version of the P. trichocarpa genome...........131 5.5.2 Gender development in poplar flowers...........................................................132 5.5.3 Comparing the MADS-box genes involved in floral development in A. thaliana and P. trichocarpa .....................................................................................................133 5.5.4 A-class gene-expression in P. trichocarpa inflorescences...............................135 5.5.5 B-class gene-expression in P. trichocarpa inflorescences...............................136 5.5.6 C-class gene-expression in P. trichocarpa inflorescences...............................138 5.5.7 E-class gene-expression in P. trichocarpa inflorescences ...............................139 5.6  Conclusions ..........................................................................................................141  Chapter 6: Conclusions..................................................................................................142 6.1  Developing a genetic marker to identify the gender of an individual using vegetative  tissues ...........................................................................................................................143 6.2  Investigation of the genetics of floral development in P. trichocarpa....................146  xi  6.3  Future directions and applications of this research ................................................147  References........................................................................................................................150 Appendices.......................................................................................................................166 Appendix A List of PCR primers designed to amplify genes studied in chapters three, four and five .........................................................................................................................166 Appendix B List of gene names, genomic locations, and functional annotations for genes studied in chapters two, three and four...........................................................................176 Appendix C List of gene names, genomic locations, and functional annotations for genes studied in chapter five ...................................................................................................179 Appendix D List of all MADS-box genes reported in Leseberg et al. (2006) updated to version 2.0 of the poplar genome...................................................................................181 Appendix E Protein alignment of MADS-box protein domain for Populus trichocarpa MADS-box genes used to construct the phylogenetic tree in chapter five ......................194  xii  List of Tables Table 2.1 Species tested for amplification with the SCAR 354 primers...............................40 Table 2.2 List of putative genes in the region surrounding the location of the Populus trichocarpa sequence on chromosome 15 that show homology to the SCAR 354 Salix sequence. ............................................................................................................................43 Table 2.3 Gene-anchored primers designed to amplify conserved regions 5' of the poplar Ssu72-like gene (fgenesh1_pg.C_LG_XV000380) and the equivalent region in Salix..........43 Table 2.4 Species tested for amplification with Ssu72-like primers.....................................44 Table 2.5 Comparison of the position of the list of putative genes in the region surrounding the location of the Populus trichocarpa sequence on chromosome 15 that shows homology to the SCAR 354 Salix sequence in version 1.1 and version 2.0 of the poplar genome. ............53 Table 3.1 Gene models located on scaffold 117 and chromosome 19 annotated as putative chloroplast terpene synthase in P. trichocarpa.....................................................................67 Table 3.2 Gene models unique to scaffold 117, updated with names and positions on version 2.0 of the P. trichocarpa genome. .......................................................................................67 Table 3.3 Overview of polymorphism data for genes located on the telomeric end of chromosome 19...................................................................................................................70 Table 4.1 Genes showing a statistically significant difference in the gene-expression between males and females in P. trichocarpa, on version 1.1 of the genome. ......................95 Table 5.1 Relating available information on the expression of MADS-box genes in Populus species to MADS-box genes showing gender-biased expression patterns...........................123 Table 5.2 Populus MADS-box floral genes that have been characterized with expression patterns in floral tissues and their A. thaliana homologs. ...................................................126  xiii  List of Figures Figure 1.1 Sexual systems in plants. .....................................................................................2 Figure 1.2 Three functional groups of genes are involved in gender-determination in plants. ............................................................................................................................................15 Figure 1.3 Schematic comparing how gender-determining, gender-differentiating and gender-associated genes act in monoecious versus dioecious mating systems. .....................25 Figure 2.1 Structure of SCAR 354 marker regions in Salix species, and homologous region found on the Populus trichocarpa genome. .........................................................................42 Figure 2.2 Sequence alignment of the SCAR 354 sequences obtained from Salix caprea and Populus trichocarpa (for the region alignable between the two genera). ..............................44 Figure 2.3 Consensus sequence (402bp long) for S. reticulata obtained from gene-anchored primers amplifying the SCAR 354 conserved region and a portion of the coding region of the Ssu72-like gene...................................................................................................................45 Figure 2.4 Consensus sequence (522bp long) for S. arctica obtained from gene-anchored primers amplifying the SCAR 354 conserved region and a portion of the coding region of the Ssu72-like gene...................................................................................................................46 Figure 3.1 Summary of the data available on the position of the putative sex locus on chromosome 19 in the genus Populus..................................................................................58 Figure 3.2 Genes sampled along the length of scaffold 117 (version 1.1) and their corresponding positions on chromosome 19 (version 2.0)....................................................68 Figure 3.3 Haplotype diagram for gene SCA_117_10 (gene model POPTR_0019s01790) based on 772bp sequenced in ten females and four males. ...................................................69  xiv  Figure 4.1 Collecting leaf and floral buds from P. trichocarpa individuals. ........................88 Figure 4.2 Arrangement of P. trichocarpa mRNA samples on the NimbleGen Affymatrix oligonucleotide microarray chips.........................................................................................90 Figure 4.3 Histogram of the parametric p-values plotting gene-expression in male leaf tissue (x axis) against the frequency of gene-expression in female leaf tissue (y-axis). ..................93 Figure 4.4 Histogram of the parametric p-values comparing gene-expression differences in leaf tissue between the two microarray experiments. ...........................................................95 Figure 4.5 Comparison of the gene-expression values from the microarray data for chromosome 19, based on version 1.1 of the poplar genome, to the physical map chromosome 19 from version 2.0 of the poplar genome.......................................................97 Figure 5.1 Distribution of 88 MADS-box genes on P.trichocarpa chromosomes. .............120 Figure 5.2 Un-rooted phylogenetic tree produced using a Maxmimum Likelihood (ML) analysis to show the homology in protein sequence data between the MADS-box domain from the poplar floral MADS-box genes and A.thaliana floral MADS-box genes..............127 Figure 5.3 Electrophoresis gel photo showing differential expression patterns measured by PCR of cDNA from male and female P. trichocarpa flower tissues...................................129 Figure 5.4 Electrophoresis gel photo showing no detectible expression in male and female P. trichocarpa leaf tissue of the MADS box genes that had differential expression patterns in male and female floral tissues............................................................................................130 Figure 5.5 Comparing differences in MADS-box gene-expression in floral tissues between A. thaliana and P. trichocarpa...........................................................................................134  xv  List of Abbreviations AFLP Amplified Fragment Length Polymorphisms CAGE Cap Analysis of Gene-expression MPSS Massively Parallel Signature Sequencing MSY Male Specific Region PAR Pseudoautosomal Region RAPD Random Amplified Polymorphic DNA SAGE Serial Analysis of Gene-expression SCAR Sequence Chracterized Amplified Region SNPs Single Nucleotide Polymorphisms SSR Simple Sequence Repeat  xvi  Acknowledgements I would like to thank my supervisor, Dr. Quentin Cronk, for his support and guidance throughout my graduate studies, and for encouraging me to explore the field of molecular evolution and development in plants. I would also like to thank my thesis committee, Drs. Sally Aitken, Carl Douglas, and Brian Ellis, for their thoughtful feedback on my research and help in crafting my thesis. I was fortunate to have NSERC support during my degree, so I would like to thank the Natural Sciences and Engineering Research Council of Canada for awarding me grant funding that supported me for four years of my studies. The UBC Department of Botany was a wonderful place to work and study so I would like to thank the faculty and staff of this department for the many helpful informal conversations that provided assistance, both with my research and with preparing grant applications, applying for TAships and getting all the paperwork of studying in on time. I was fortunate to work with a great group of laboratory colleagues and fellow graduate students during my studies, so I would like to thank the following people for the assistance they gave me during my degree. Dorothy Cheung and Dr. Hardeep Rai provided excellent technical assistance as I learned new laboratory techniques and worked out research problems, Cindy Sayre and Julia Nowak assisted in collecting leaf samples used for the molecular work in my research, and Hardy Hall helped with the initial statistical analysis of the microarray experiments. Sæmundur Sveinsson helped me perform the phylogenetic analysis of the MADS-box protein sequences, and Dr. Armando Geraldes was instrumental in assisting me with learning to edit DNA sequences and analyze them.  Drs. Athena  McKown, Ji Yong Yang, Robin Young and Robyn Seipp provided advice and encouragement throughout my degree but especially during the writing of my thesis.  xvii  Finally I would like to thank my friends and family, and especially my husband Jason Evans, for reminding me to laugh when my research was not going as well as hoped for, and allowing me to focus on it when it was.  xviii  Dedication This thesis is dedicated to the memory of three people who influenced my decision to pursue a graduate degree. First, to my grandmother, Dr. Ingrid Maria Mathilda Willhelmina Hamburger Temmel, who completed her doctorate in Botany from the University of Graz in July of 1948, and was an inspiration to my pursuing studies in the field of Botany. Also, to my Gr. 9 and 10 science teacher Mr. David Stacey, who taught me that science was fun, and that, if you were lucky, learning was something you never stopped doing. And finally, I’m dedicating my thesis to the memory of my good friend Sandra Patricia Hunter, who passed away too soon, but lived her life with passion, courage and wisdom, and always encouraged me to follow my dreams.  xix  Chapter 1: Introduction 1.1  Sexual systems in plants Gender-determination in animals and plants is a developmental process that  results in the physical separation of female and male gamete production, on the same or on separate individuals.  Approximately 90% of the world’s angiosperms have  hermaphroditic or “perfect” flowers, which contain both male and female reproductive parts (Ainsworth, 2000). The remaining 10% of the flowering plants produce various combinations of male, female or hermaphroditic flowers on the same or separate plants. The most common combinations are, first, monoecy, where male and female flowers are present on the same plant and, secondly, dioecy, which occurs when male and female flowers are produced on separate plants (Ainsworth, 2000). Dioecious species account for 6%, or 14 000 species of all flowering plants (Renner and Won, 2001). Many plant species have evolved other combinations of flower types as well, such as gynodioecy, gynomonoecy, trimonoecy, androdioecy, andromonoecy and trioecy, but these combinations are less common and are usually considered to be intermediate forms in the evolution of unisexual flowers, which contain either male or female reproductive organs (Ainsworth, 2000) (Figure 1.1). These combinations of sexual systems in plants have been well described morphologically and physiologically, and there is some understanding of the possible ecological reasons for their evolution, such as avoidance of inbreeding depression caused by self-pollination (Charlesworth and Charlesworth, 1979). However, how gender is determined in plants at the molecular level or what the evolutionary pathway(s) for the development of separate genders might have been is a growing field of research  1  facilitated by the advent of genome sequencing. Research into gender-determination in dioecious plants has generally been focused around plant species that have welldeveloped heteromorphic sex chromosomes such as Silene or Rumex (Ainsworth et al., 1995; Filatov et al., 2000).  Figure 1.1 Sexual systems in plants. Circles represent floral forms, and plants that produce only hermaphroditic flowers are not shown.  2  1.2 1.2.1  Evolution of dioecy Gynodioecious pathway Approximately 160 plant families include dioecious species and it is estimated  that dioecy has evolved over 100 separate times (Charlesworth and Guttman, 1999). Dioecy is thought to have evolved through several evolutionary pathways.  In the  gynodioecy pathway, it is theorized that female plants arise by mutations that produce male sterility in hermaphroditic plants.  These spread through populations of  hermaphroditic plants because they produce out-crossed, and therefore fitter, offspring (Barrett, 2002). Selection then acts on the remaining hermaphroditic plants to reduce female function so that male plants are produced. It is most likely that dioecy evolved via gynodioecy or androdioecy as an intermediate stage because the initial step in this process is a single mutation resulting in either female or male sterility (Charlesworth and Charlesworth, 1978). The probability of dioecy evolving directly from a hermaphrodite is small because it is unlikely that two independent mutations, one for female and one for male sterility, would arise simultaneously and become tightly linked on the genome to ensure a stable dioecious system (Ainsworth, 2000).  A species that provides an  opportunity to study this is Carica papaya, where comparisons of silent mutation rates observed on the Y and Yh chromosomes indicate that the dioecious and gynodioecious breeding systems in this plant diverged approximately 73 000 years ago (Yu et al., 2008). Recent research with Fragaria virginiana Mill., which has both subdioecious and gynodioecious sexual systems, may also provide insight into how the transition from gynodioecy to dioecy occurred (Spigler et al., 2008).  3  1.2.2  Heterostyly pathway In a number of species dioecy has probably evolved from heterostyly, whereby  male flowers are derived from short-styled flowers and female flowers from long-styled flowers (Bawa, 1980). There are no fewer than 13 plant families where distyly has been observed (Charlesworth and Charlesworth, 1979), and plant species exhibiting this are characterized by pin flowers with long styles and short stamens and thrum flowers having short styles and long stamens (Beach and Bawa, 1980). The two flower types are produced in a population as if they are controlled by alleles at a single locus, where the pin flower phenotype is recessive (Charlesworth and Charlesworth, 1979). The genetic model put forward to explain the evolution of dioecy from heterostyly proposes the existence of a two-gene linkage model whereby a dominant gene for producing thrum flowers is linked to a recessive gene that causes the abortion of female organs (Muenchow and Grebus, 1989). However, the most probable force driving the change from distyly to dioecy is a change in pollinators. Reciprocal pollen transfer ensures the gene flow between the two floral types, so if the pollination system changes, gene flow ceases (Beach and Bawa, 1980). Unidirectional pollen flow (pin to thrum, and thrum to pin only) then facilitates the change from functional unisexuality to structural unisexuality as mutations from male sterility (pin flowers) and female sterility (thrum flowers) occur and spread through the population (Beach and Bawa, 1980). It is unlikely that dioecy evolved from heterostyly in P. trichocarpa because the flowers of this species show no evidence of heterostyly, and the genus is wind-pollinated.  4  1.2.3  Monoecious pathway Dioecy has also evolved through monoecy where it is thought that disruptive  selection acted on existing genetic variation in floral gender ratios in monoecious plants, which resulted in more specialization of the genders (Barrett, 2002). First, male fertility could be reduced through mutations that either reduce pollen production or convert male flowers to female ones (Charlesworth and Charlesworth, 1978).  These kinds of  mutations would reduce selfing in a population and increase female fertility. Secondly, a mutation that results in reduced fertility in the female plants would have to take place. A second wave of mutations resulting in reduced female fertility is unlikely to spread through the population of remaining monoecious plants unless it is tightly linked to the male fertility locus because such a mutation would be disadvantageous to the females (Charlesworth and Charlesworth, 1978).  Dioecy could therefore have evolved from  monoecious species through a process of alternating mutations that reduced male and female fertility over time (Charlesworth and Charlesworth, 1978). 1.2.4  Evolution of dioecy in Populus There is a strong association between the occurrence of dioecy and the presence  of monoecious species in related genera. This may be because once a group of species has acquired the ability to genetically suppress male or female function in flowers within an individual, the jump to dioecy can occur through changes in floral gender ratios on individual plants (Renner and Ricklefs, 1995). Eventually these genetic differences that cause differentiated expression of gender in individuals become fixed between males and females at “sex loci”.  5  From the distribution of sexual systems in related genera, it appears that dioecy in Populus evolved via the monoecious pathway. The genus Populus is classified in the angiosperm Eurosid I clade, in the order Malpighiales (Jansson and Douglas, 2007a). The order Malpighiales contains approximately 16 000 species divided into 39 families (Stevens, 2001), including the family Salicaceae, where Populus is placed. The family Salicaceae  contains  nine  genera  (Bennettiodendron,  Carrierea,  Idesia,  Itoa,  Macrohasseltia, Olmediella, Poliothyris, Populus and Salix), which are predominantly dioecious  or  monoecious  and  insect-pollinated  except  the  hermaphroditic  Macrohasseltia, and the wind-pollinated Populus (Cronk, 2005). 1.3  Sex chromosomes in plants When it comes to gender-determination, gonochorism, or sexual reproduction  between unisexual individuals, is the most prevalent system in animals, and is regulated by the presence of sex chromosomes in almost all cases (Janousek and Mrackova, 2010). Sex chromosomes are chromosomes that are involved in the genetic determination of gender in an organism. In the majority of plant species that are dioecious, the genetic mechanism that determines gender is not well understood, and not all dioecious species have identified sex chromosomes. Sex chromosomes in plants appear to be less evolved than those observed in animals, so studying them in plants provides an opportunity to understand the processes involved in their evolution from autosomes (Moore, 2009), as well as the exploring of how gender-determination is controlled genetically in the absence of sex chromosomes.  6  1.3.1  Identification and description of sex chromosomes Although sex chromosomes were discovered around the same time in both plants  and animals early in the 20th century, knowledge about sex chromosome structure and function in plants has lagged behind our understanding of the role of sex chromosomes in animals such as Caenorhabditis elegans, Drosophila sp., Homo sapiens and Mus musculus (Vyskot and Hobza, 2004).  As noted above, the fact that many sex  chromosomes are heteromorphic from autosomes, and from each other, facilitated their discovery. This heteromorphy is related to the two sex-determination systems that have been observed in most sexually reproducing organisms.  In heterogametic sex-  determining systems, the gender of an individual is determined by either the presence of a Y chromosome (XY system), or the W chromosome when females are heterogametic (ZW system) (Vyskot and Hobza, 2004). In the second system sex is determined by the ratio between the number of sex chromosomes and the number of autosomes, the X/A ratio (Jamilena et al., 2008). Since sex chromosomes were first described, the basic definition of what constitutes a sex chromosome has expanded to state that sex chromosomes do not recombine during meiosis, along at least a portion, if not the entirety of their length, and that sex-determining genes that control male and female fertility are located on them (Vyskot and Hobza, 2004). 1.3.2  A brief history of the discovery of sex chromosomes The genetic basis for sex-determination remained a mystery until the early years  of the 20th century. One of the first indications that genetic control of sex-determination was associated with chromosomes came in 1891 when Henking, while studying the insect Pyrrhocorus apterus, noticed a chromatin body in the spermatocyte that was larger than  7  the other chromatin bodies, and stained more darkly than the others (Anderson, 2000). These first observations were confirmed by subsequent researchers but no one was yet prepared to state that the sex of progeny was determined exclusively by chromosomes at the time of fertilization, and it wasn’t until 1902 that McClung postulated that the “accessory chromosome” identified by Henking could play a part in sex-determination (Brush, 1978). Conclusive evidence for the connection between chromosomes and sexdetermination came in 1905 when research on Tenebrio molitor (meal worm, n = 20) done by Dr. Nettie Stevens found that sex-determination must be controlled by chromosomes, as female somatic cells contained 20 large chromosomes, and the same cells in males contained one small, and 19 large chromosomes (Brush, 1978). The same year that Stevens discovered the sex-determination system that would become known as XY, where the males (Y) are the heterogametic gender, work on a different insect species, Anasa tristis, also inferred that chromosomes were responsible for sexdetermination, through a XO sexual determination system, where an X chromosome is present in the females, and males do not have this extra X chromosome (Brush, 1978). Between 1905 and 1909 the terms “heterchromosome”, “autosome”, “sex chromosomes”, and “X” and “Y” chromosomes came into use to describe the newly established link between “accessory chromosomes” and sex-determination in many organisms (Anderson, 2000). 1.3.3  Discovery of sex chromosomes in plants Conclusive evidence of the existence of sex chromosomes and their role in sex-  determination in plants was established in 1917 from research done on two species of liverwort, Sphaerocarpos donnellii Aust. and S. texanus Aust. (Anderson, 2000). The  8  haploid gametophyte of these species have eight chromosomes; in females the karyotype consists of seven autosomes and an X chromosome that determines sex, and in males there are seven autosomes and a Y chromosome (Allen, 1917). Heteromorphic sex chromosomes in dicotyledonous plants were first observed in the dioecious species Lychnis alba (now named Silene latifolia) where evidence for XX females and XY males was reported (Blackburn, 1923). However, plants with separate genders do not usually have sex chromosomes and the mechanism and control of gender identity in the majority of plants remains obscure, despite the numerous studies that have investigated the role of sex chromosomes in determining sex in haploid, diploid and triploid plants (Sakamoto et al., 2005). 1.3.4  Evolution of sex chromosomes The evolution of sex chromosomes has probably been driven by the need to limit  recombination between genes that determine gender (Ainsworth, 2000). In those plants with sex chromosomes, the X-Y system is thought to have evolved more recently than in animals. Evidence suggests that the X-Y system only arose approximately ten million years ago in Silene spp., whereas mammals developed sex chromosomes around 166 million years ago (Veyrunes et al., 2008). Birds have a Z-W system of sex chromosomes that evolved independently (Hake and O'Conner, 2008).  Of the estimated 14 620  dioecious plant species only 13 species in five families are known to have heteromorphic sex chromosomes (Jamilena et al., 2008; Ming et al., 2007), and an additional 13 plant families containing approximately 16 additional species exhibit homomorphic sex chromosomes (Ming et al., 2007). In mammals the degeneration of gene content and size of sex chromosomes through accumulation of deleterious mutations and subsequent gene  9  loss due to their long evolutionary history is characteristic, smaller size does not appear to be a genomic feature of young sex chromosomes in plants dues to local duplications and transposable element insertions (Ming and Moore, 2007).  However, where they  exist, the genetic mechanisms by which sex chromosomes control sex-determination, either via an active Y chromosome, or by the balance of the number of X chromosomes with the number of autosomes, are the same in plants and in animals (Jamilena et al., 2008). 1.3.5  Proposed mechanism for the evolution of sex chromosomes Sex chromosome evolution is a multiple step process, one of the first processes  involved being the suppression of genetic recombination surrounding one or more sexdetermining genes at a sex locus (Liu et al., 2004). At some point in the evolutionary history of the plant, a male or female sterility mutation occurs on a nascent sex chromosome, leading to suppression of recombination happens at the locus and its immediate neighbouring region. This process can be observed in Asparagus officinalis L. (2n = 20), in which the chromosome pair L5 have been identified as the homomorphic sex chromosomes (Reamon-Büttner et al., 1998). In this species sexual dimorphism is controlled by a dominant gene M, with female plants being homozygous for the recessive allele (mm), and male plants being either homozygous (MM) or heterozygous (Mm) at the sex locus (Jamsari et al., 2004). In many organisms that have a X-Y mating system, the sex chromosomes contain a pseudoautosomal region (PAR) where X and Y still recombine (Nicolas et al., 2005). This is the case in the dioecious crop species Spinacia oleracea (spinach) (Khattak et al., 2006), and asparagus (Telgmann-Rauber et al., 2007), where both species have homomorphic sex chromosomes that recombine along most of  10  their length with little or no measurable degeneration of the Y sex chromosome. In Carica papaya where a sex locus has also been identified, gender is determined by one chromosome region with three allelic forms, combinations of which result in the male, female and hermaphroditic phenotypes (Liu et al., 2004). The sex locus is distinguished on the homologous autosomes as a region of increased polymorphism with a low level of recombination, known as the male-specific region, or MSY (Liu et al., 2004). A single locus with three alleles also determines gender in Ecballium elaterium (Ainsworth, 2000). Species that have heteromorphic chromosomes can also exhibit this kind of sexdetermining system, as is the case in Humulus lupulus (European cultivated hops), where analysis of sex-linked markers has revealed that suppression of recombination between the X and Y chromosomes is localized to the region around the sex-determining locus (Seefelder et al., 2000). Once a sex-determining locus has formed on an autosome, the suppression of recombination observed around the sex locus can spread along the length of the autosome, and the proto Y chromosome will slowly accumulate deleterious mutations, along with favorable alleles, which will simultaneously reduce the size and functional gene content of the Y while selecting for male advantageous mutations via “Müller’s Ratchet” and genetic hitchhiking (Rice, 1987). In the genus Silene three species, S. latifolia, S. dioica and S. diclinis, have a heteromorphic X-Y sex-determination system where the Y chromosome determines maleness (Nicolas et al., 2005).  In these Silene  species the correlation between the synonymous divergence between the X and Y chromosomes and the distance from the PAR on the X chromosome genetic map indicates that suppression of recombination between the sex chromosomes started at an  11  ancient sex-determining locus and moved towards the current PAR (Nicolas et al., 2005). Now with the identification of eleven Y-linked loci with homologs on the X chromosome, researchers have confirmed that the X and Y sex chromosomes in these species stopped recombining at between ten and twenty million years ago (Bergero et al., 2007).  The accumulation of chloroplast DNA sequences on the Y chromosome in S.  latifolia has also contributed to the degeneration of genes (Kejnovsky et al., 2006). Rumex acetosa has heteromorphic sex chromosomes, but in this dioecious species sex-determination is controlled by the X/A ratio, where a ratio between numbers of X chromosomes and autosomes controls flower gender (Jamilena et al., 2008). In Rumex acetosa female plants have two X chromosomes (2n = 14, XX) and male plants have one X and two different Y chromosomes (2n = 15, XY1Y2) (Ming et al., 2007). In this species the non-recombining regions of the Y chromosomes are heterochromatic and may have become specialized due to an accumulation of repetitive DNA sequences (Shibata et al., 1999). Therefore, the Y chromosomes in Rumex acetosa represent an advanced stage of sex chromosome evolution (Mariotti et al., 2006). 1.3.6  Degeneration of the Y chromosome The current theory of sex chromosome evolution is that Y chromosomes shrink in  size, and genetic information is lost as they evolve (Ming and Moore, 2007). However, given that sex chromosomes in plants have evolved more recently than those in animals, the smaller size of the Y chromosome is not a universal genomic feature.  The  accumulation of repetitive DNA sequences, either tandem repeats like those seen in Rumex sp., or transposable elements, could account for the increased size of the Y chromosome in plants that have heteromorphic sex chromosomes (Jamilena et al., 2008).  12  In Silene latifolia and Rumex acetosa, where sex chromosomes are most highly evolved, the Y chromosomes have not degenerated to a point where non-functional DNA sequences would be lost.  Neither of these species have accumulated transposable  elements on their respective Y chromosomes (Jamilena et al., 2008), unlike the Y chromosomes of Cannabis sativa and Marchantia polymorpha. In C. sativa the Y is twice the size of the X due in large part to the accumulation of a specific Long INterspersed Element-like (LINE-like) retrotransposon on the terminal end of the long arm of the Y chromosome, which may play a role in suppressing pairing or recombination with the X chromosome (Sakamoto et al., 2000). Unlike in animals where a considerable amount of information on the gene organization on the Y chromosome is available, due in large part to work in some primates and Drosophila, data on gene sequence and organization on the Y chromosomes of plants is not as available.  In the  liverwort Marchantia polymorpha the haploid genome consists of eight autosomes and either an X chromosome in females (n = 8 + X) or a Y in males (n = 8 + Y) (Yamato et al., 2007). The X and Y chromosomes do not recombine along their entire length, and 64 genes have been identified on the Y chromosome, 14 of which are expressed in reproductive organs and are only found in the male genome (Yamato et al., 2007). Active Y chromosome specific genes were first reported in this plant species, and while repetitive DNA sequences are a common feature of Y chromosomes in the plant listed above and in animals, the nucleotide sequences that make up these repeats appear to be unique to each species studied, indicating that Y chromosomes have arisen independently in multiple taxa as a sex-determining system (Okada et al., 2001).  13  1.3.7  The absence of sex chromosomes in Populus There have been no sex chromosomes identified in Populus trichocarpa, although  a putative sex locus has been mapped by using sex-linked genetic markers on chromosome 19 in the genomes of six poplar species (including P. trichocarpa) (Paolucci et al., 2010). The genus Populus may have a similar sex-determination system as that found in Fragaria sp. (strawberry) as species in this genus do not share the same sexdetermination system, they may share a single origin for a sex-determining region in a common ancestor (Goldberg et al., 2010). 1.4  The genetics of gender The developmental processes that drive the differentiation between males and  females are controlled through gene-expression patterns in response to internal or environmental cues, and in plants this results in a large variety of sexual phenotypes. Given the myriad of sexual systems that exist in plant families, and the observation that these systems have evolved multiple times, there appear to be many ways that plants have evolved genetic controls for the development of sexual organs and mating types. Until recently, the lack of genetic data for plant species with diverse sexual systems has prevented a better understanding of how genetics relates to sexual phenotypes. Now that the genes involved in gender-determination have been identified in multiple species, it appears that these genes can be divided into three functional groups: gender-determining genes, gender-differentiating genes, and gender-associated genes (Figure 1.2). Gender-determining genes are located at a sex locus and are fixed as homozygous in one sex and heterozygous in the other so that they segregate with gender. Genderdifferentiating genes can be located throughout the genome and are responsible for the  14  development of sexual organs and reproductive functions in the plant - primary sexual characteristics. The products of gender-determining genes act upon this group of genes. Gender-associated genes play no direct role in gender function, but may result in secondary sexual characteristics that are associated with one or the other gender.  Figure 1.2 Three functional groups of genes are involved in gender-determination in plants. Solid arrows represent direct relationships between gene groups, and dashed arrows indicate indirect relationships between the gene groups. 1.4.1  Gender-determining genes Gender-determining genes are regulatory genes, which exist as homozygotes in  one sex and heterozygotes in the other sex, segregating at one or more loci in the 1:1 sex ratio observed in Populus (Comtois et al., 1986; Farmer, 1964). These genes are distinct from the gender-differentiating genes that result in the differences observed between male and female floral organs. Gender-determining genes presumably initiate a cascade of gender-regulated gene-expression that determines the observed male and female phenotypes.  In the liverwort, Marchantia polymorpha, the gene content of the sex  chromosomes has been investigated, and Y specific genes have been identified, as well as  15  a gene M2D3.4, located on an autosome, which is expressed only in male gametic cells (Tanurdzic and Banks, 2004). However, none of these genes have been characterized as being gender-determining.  Orthologs of the Arabidopsis thaliana genes SHOOT-  MERISTEMLESS (STMS) and CUPSHAPED COTYLEDON (CUC) have been shown to control the establishment and development of the floral meristems in Silene latifolia, and therefore may play a role in the gender-determination pathway in this species (Zluvova et al., 2006). However, to date no gene directly involved in gender-determination has been characterized in a dioecious species, while genes that respond to physiological or environmental cues to give rise to unisexual flowers in the monoecious species Zea mays and Cucumis sativus have been characterized (Jamilena et al., 2008). 1.4.2  Gender-differentiation and development in plants Genes may be gender-differentiating because their transcriptional regulation is  downstream of a gender-determining gene, or because they are gender-linked and therefore differ in copy number between sexes, for instance they may be present in only one sex. An example of this is found in the fly Drosophila melanogaster, where sexdetermination is controlled by the ratio of numbers of X chromosomes to sets of autosomes, as the X chromosome contains almost one third of the genes in this fly, an extra copy of this chromosome leads to aneuploid conditions and disruption of cellular equilibrium (Hake and O'Conner, 2008).  DNA methylation is also important in  regulating X-chromosome inactivation (Feng et al., 2010), and in Carica papaya methylated heterochromatin structures have been observed associated with the malespecific region of the Y chromosome (MSY), but not with the corresponding region on the X chromosome (Zhang et al., 2008).  16  An example of gender-differentiating genes can be found in the fern Ceratopteris richardii, where two genes are thought to be responsible for producing male (gene TRA) or female (gene FEM1) gametophyte traits depending on the presence of the hormone ACE (antheridiogen Ceratopteris) which activates a set of genes, HER, NOT1 and MAN1, which in turn influence the expression of TRA and FEM1 (Tanurdzic and Banks, 2004). The size and gender composition of the gametophyte population surrounding a germinating spore, and therefore the level of ACE, ultimately control the gender of the developing gametophyte, so this is a good example of hormonal mediation of gender development.  A similar situation has been observed in the androdioecious species  Mercurialis annua, where three unlinked loci (A, B1 and B2) determine gender via changes in levels of the plant hormones auxin and cytokinin (Khadka et al., 2005), but the gender ratio of flowers in hermaphroditic individuals is influenced by the frequency of male plants in the surrounding population (Dorken and Pannell, 2009). 1.4.3  Gender-differentiation in Zea mays One of the species that has been developed as a model system for studying how  gender-differentiation occurs in a monoecious plant is Zea mays. In this species an interaction of gender-differentiating genes, hormones and environmental factors combine to produce unisexual flowers in the same plant by acting on the immature floral meristem and causing the selective abortion of the inappropriate sex organs (Dellaporta and Calderon-Urrea, 1994). Z. mays produces flowers in inflorescences that consist of many individual spikelets, each subtended by two glumes that enclose two florets. At the beginning of development, the florets are bisexual, containing both male and female sexual organs, but when the florets transition to being unisexual, their location in the  17  plant, being either in a tassel (male) or ear (female) flower, and their position within the spikelet decides the gender that the floret will assume (Calderon-Urrea and Dellaporta, 1999). The abortion of either the stamens in female florets, or pistils in male florets, is regulated by the plant hormones gibberellin (GA) (Banks, 2008), and jasmonic acid (JA) (Acosta et al., 2009). The model pathway for the differentiation of male florets in Z. mays contains three genes (tasselseed1, tasselseed2, and silkless1) that appear to regulate the production of jasmonic acid which plays a role in the programmed cell death in pistil primordia (Browse, 2009; Calderon-Urrea and Dellaporta, 1999). For the development of female florets, it appears that two genes that are cell-cycle regulators (CYCLIN B and WEE1) block the cell-cycle from continuing in stamens, thereby producing pistillate florets (Kim et al., 2007). A second class of mutations that causes dwarf (d) plants also results in stamen abortion.  d1, d2, d3, and d5 mutations disrupt steps in the gibberellin  biosynthesis pathway which results in lower concentrations of endogenous gibberellin and causes an almost complete conversion of staminate to pistillate florets (Dellaporta and Calderon-Urrea, 1994). Environmental factors such as short day-length and low light levels have been shown to increase gibberellin levels in Z. mays, thereby causing the plant to produce predominantly female florets (Dellaporta and Calderon-Urrea, 1994). 1.4.4  Gender-differentiation in Cucumis sativus and C. melo The role that environmentally induced changes in hormone levels play in  affecting gender-differentiation has been looked into in more detail in two species of the Cucurbitaceae, Cucumis sativus L. and C. melo. Species in this family exhibit unisexual flowers, within either a dioecious, monoecious, gynoecious, or androdioecious mating,  18  and phylogenetic data indicates that dioecy is the ancestral mating system (Zhang et al., 2006). In C. sativus and C. melo gender expression is controlled by a combination of environmental, hormonal and genetic factors (Boualem et al., 2008; Trebitsh et al., 1997). In these two species gibberellin, long days and high temperatures cause the plants to produce a higher proportion of staminate flowers, whereas more pistillate flowers are produced under conditions of low temperature and short days, and in the presence of ethylene and auxin (Knopf and Trebitsh, 2006).  Genes involved in the ethylene  biosynthesis pathway mediate gender differentiation, but there are differences at the genetic level between the two species. Interestingly, though C. sativus and C. melo differ in geographic origin and chromosome number, and their most recent common ancestor is estimated to have occurred over 40 MYA, it appears that mutations resulting in reduced activity of 1-aminocyclopropane-1-carboxylic acid synthase (ACS) in both species have produced hermaphroditic, or perfect, flowers in both species independently (Boualem et al., 2009). In C. melo gender-differentiation is governed by two genes, andromonoecious (a) and gynoecious (g), and different sexual types arise from different combinations of alleles of these two genes. Andromonoecious (aaG-) and monoecious (A-G-) genotypes produce male flowers on the main stems with perfect or female flowers on axillary branches, respectively, while gynoecious (AAgg) and hermaphroditic (aagg) individuals produce only female or perfect flowers (Boualem et al., 2008). The a gene has been characterized and codes for a 1-aminocyclopropane-1-carboxylic acid synthase (ACS), now named CmACS-7, based on homology to the Arabidopsis ACS-7 gene (Boualem et al., 2008). CmACS-7 is an enzyme thought to catalyze the first rate-limiting step in the ethylene  19  biosynthesis pathway and in C. melo is required for the development of female flowers in monoecious plants, and a reduction in the activity of CmACS-7 produces perfect flowers on andromonoecious plants (Boualem et al., 2008). C. sativus L. var sativus, the wild-type cucumber, is monoecious and flowers are produced along the stem in a predetermined developmental sequence with staminate flowers occurring first, followed by a region where staminate and pistillate flowers are interspersed, and then at the end of the stems only pistillate flowers occur (Trebitsh et al., 1997). However, in domesticated C. sativus there are four different mating types and three major genes determine these: Female (F/f), Monoecious (M/m) and Androecious (A/a). The F gene promotes femaleness throughout the entire plant whereas the M gene determines if flowers are unisexual (M-) or perfect (mm), and when the F gene is homozygous recessive the A gene influences sex expression (Mibus and Tatlioglu, 2004). The genotypes of the four mating systems in C. sativus are as follows: gynoecious (MFF), monoecious (MMffA-), hermaphrodite (mmF-) and andromonoecious (mmffaa) (Mibus and Tatlioglu, 2004). Due to incomplete dominance of the F gene, female sex expression can be enhanced by interacting with two additional loci, Intensifier of female sex expression (In-F) and gynoecious (gy), though these interactions have not been well characterized (Knopf and Trebitsh, 2006).  Two alleles of the F gene have been  characterized as CS-ACS1G (dominant allele) and CS-ASC1 (recessive allele) and genotypes that are homozygous for the dominant allele (gynoecious and hermaphroditic plants) show increased ethylene concentrations, which results in more pistillate flowers (Mibus and Tatlioglu, 2004).  20  The C. sativus gene CsACS2 is highly homologous to the C. melo gene CmACS-7, therefore the C. sativus locus M may be orthologous to the C. melo a locus (Boualem et al., 2009). It appears that in C. sativus CsACS2 functions in a localized way to sense inhibiting ethylene in stamens and is therefore dedicated to arresting stamen development in floral primordia (Boualem et al., 2009). An ETHYLENE-INSENSITIVE3-like genetic sequence is also associated with the M locus in C. sativus, providing further evidence that gender-differentiation in this species is ethylene mediated (Liu et al., 2008). While further research is needed to identify the roles that the loci A, gy, and In-F play in genderdifferentiation in C. sativus, it seems evident that the F locus controls ethylene concentrations, while the M locus mediates differences in ethylene sensitivity in male and female floral primordia (Liu et al., 2008). How gibberellin and auxin influence genderdifferentiation in these species, and what relationship temperature and day length have to plant hormone levels that cause floral differentiation, has yet to be determined. 1.4.5  The role of MADS box genes in gender differentiation Plant genes have been found to be fairly compact and usually grouped together in  clusters, surrounded by repetitive DNA sequences, even in large genomes (Kellogg and Bennetzen, 2004).  Floral development in hermaphroditic, or perfect flowers, is a  complex, multistep process that begins with the establishment of a floral meristem, followed by the differentiation of floral organs that then develop into the structures observed in flowers (Zik and Irish, 2003). The current model for describing the genetic interactions that control flower development is the ABC model. This model defines three regions of the floral meristem, each of which is controlled by A-class of genes – A, B, or C (Coen and Meyerowitz, 1991). Region A comprises the first and second whorls of the  21  floral meristem, region B includes the second and third whorls and region C contains the third and forth whorls (Coen and Meyerowitz, 1991). Expression of A-class genes alone specifies sepal identity, A + B-class genes specify petal development in the second whorl, B + C-class genes expression results in stamen formation in the third whorl and C-class genes alone specify carpel development in the forth whorl (Zik and Irish, 2003). Many of the ABC model genes that control reproductive development in flowering plants belong to the MADS-box gene family of transcription factors, so called because these genes share a common motif found in the MCM1, AGAMOUS, DEFICIENS, and SRF genes (Coen and Meyerowitz, 1991). Further research has indicated that two other classes of genes are also involved in the floral development pathway. D-class genes have been found to be crucial for the determination of ovule identity (Vandenbussche et al., 2003), and E-class genes are required for specifying stamen, petal and carpel identity (Pelaz et al., 2000). The floral dichotomy observed in dioecious species results from developmental modifications of a perfect flower to a female or male flower via organ suppression (Ainsworth, 2000). In Silene latifolia the genes that are differentially expressed in female and male flowers are only involved in male floral development, and neither they, nor the characterized Y-linked genes in this species, control gender-determination (Jamilena et al., 2008). It has been found that gender-determination in Thalictrum dioicum is likely to be regulated by genes upstream from organ identity genes because stamen or carpel primordia are not initiated in the unisexual female and male flowers, respectively (Di Stilio et al., 2005). The authors of this study postulated that differentially regulated alleles of B-class genes might be linked to a sex-determining factor. A similar system  22  may exist in Populus trichocarpa whereby a genetic switch at a sex locus controls expression of B and C-class genes, thereby controlling the development of male or female flowers. MADS-box genes have not been shown to be associated with gender in either Silene latifolia, or Cucumis sativus, both of which have sex chromosomes (Hardenack et al., 1994; Perl-Treves et al., 1998). However, the C function genes have shown some gender-specific expression in Rumex acetosa (Ainsworth et al., 1995) and Liquidambar styraciflua (Liu et al., 1999). While the expression of B and C-class floral homeotic genes has been shown to decrease in the aborted organs in unisexual flowers in all species that have been studied, it is not known if this is causal or a consequence of the organ abortion (Tanurdzic and Banks, 2004). 1.4.6  Gender-associated genes Unlike animals, where secondary sexual characteristics are usually very evident  and important in mating habits and the life history of the species, in plants generally little is known about the evolution of sexual dimorphism that would result from the development of secondary sexual characters associated with one or the other gender (Meagher, 1984).  Consequently, to date gender-associated genes are not well  characterized, though observations in Chamaelirium luteum (Meagher, 1984), Asparagus officinalis L. (Bracale et al., 1991), and Populus trichocarpa (Brunner, 2010), have indicated that male plants can exhibit slightly different growth patterns or flowering times than females.  In the homosporous fern Ceratopteris richardii the male and  hermaphroditic gametophytes can be distinguished from the female gametophyte by the absence of a multicellular meristem (Tanurdzic and Banks, 2004), but it is uncertain that  23  this difference could be considered a secondary sexual characteristic controlled by gender-associated genes.  One limitation to the development of secondary sexual  characteristics in plants may be that in order to continue to reproduce, males and females of a species must maintain overlap in their habits and ecological requirements so that they can breed effectively (Meagher, 1984). 1.4.7  Differences in genetic control of gender-determination between monoecy  and dioecy One of the indications that the genetic control of gender-determination differs between monoecious and dioecious mating systems is that the ratio of males and females in a population is determined differently in each system. In dioecious species like Populus trichocarpa, the female to male ratio of individuals in a population is thought to be determined by genetic segregation of alleles at one or more loci, whereas in monoecious species the proportion of female to male gametes is influenced by environmental or epigenetic cues (Irish and Nelson, 1989). This would seem to indicate that while both monoecious and dioecious species have gender-differentiation and gender-associated genes, only dioecious species have gender-determining genes (Figure 1.3).  24  Figure 1.3 Schematic comparing how gender-determining, gender-differentiating and gender-associated genes act in monoecious versus dioecious mating systems. 1.5  Populus trichocarpa as a model organism for studying gender-determination in  plants 1.5.1  Life history of Populus trichocarpa Populus trichocarpa is a deciduous, paleopolyploid tree native to Western North  America where its range extends from California to Alaska (Brunner et al., 2004a). Though it has a broad environmental range, P. trichocarpa has fairly specific ecological requirements in that it is generally confined to river flood plains and the riparian zones of alluvial streams (Rood and Polzin, 2003). Populus trichocarpa has a juvenile stage that  25  lasts 5-10 years, during which time the trees are unable to flower and reproduce sexually (Brunner, 2010). The female and male flowers of this species are highly reduced in form, consisting of a floral disk with either 40- 50 stamens or a single pistil (Eckenwalder, 1996). The inflorescences are pendulous catkins that flush three to four weeks before leaves appear in the spring (Boes and Strauss, 1994). P. trichocarpa flowers are wind pollinated and after fertilization occurs in the spring, 20-40 capsules/female inflorescence are produced. Each capsule contains approximately 40 seeds that are released and wind dispersed in the early summer (Boes and Strauss, 1994). 1.5.2  Characteristics of Populus trichocarpa as a model species As a member of the genus Populus, P. trichocarpa is a good model species for the  study of gender in plants because of its small genome (approximately 500 million base pairs and 45 000 genes), high growth rates, economic importance of the genus, and the ease with which it is clonally propagated (Wullschleger et al., 2002a). It is already used as a model to study tree morphology, physiology, biochemistry and genetics. Arabidopsis thaliana, already a model species for studying most aspects of plant biology, is more related to Populus than the majority of dicot species for which genomic resources exist, making it possible to make informative comparisons between the two species’ genomes (Jansson and Douglas, 2007). Much of the interest in developing P. trichocarpa as a model organism has centered on the importance of secondary xylem formation and the application of this research in the forest industry (Wullschleger et al., 2002b). However, there has been some research done on P. trichocarpa genes that could be engineered to render the tree reproductively sterile, which is of interest because of the  26  importance in limiting gene flow between transgenic and wild populations (Skinner et al., 2003). The recent sequencing of the female P. trichocarpa clone “Nisqually 1” genome (version 1.1 released September 2004, version 2.0 released in January 2010), in concert with developments in gender-determination research in other model species provides exciting opportunities to understand patterns in the genomic architecture relative to gender. P. trichocarpa is the first dioecious plant to have had its genome sequenced. 1.5.3  Genomic markers linked to sex-differentiation in Populus and Salix Fixed heterozygosity should occur in the region of the sex locus to ensure that  there is no recombination, otherwise separate sexes would not be conserved. Research on Salix viminalis L. has determined the existence of a sex locus in this species (Gunter et al., 2003b), and as species in the closely related genera Salix and Populus exhibit stable dioecious sexual systems (Gunter et al., 2003b), it is probable that Populus species have a similar system. A marker co-segregating with gender has been found in S. viminalis L. (Gunter et al., 2003b) and it has been postulated that several loci interacting via epistasis may be responsible for sex-determination in this species (Alström-Rapaport et al., 1998). When genetic markers from chromosome 19 for a female P. alba and a male P. nigra were aligned the linkage maps indicated that the sex morphological trait mapped to two different regions of chromosome 19, providing evidence that at least two loci are involved in sex-determination in Populus (Gaudet et al., 2008). If a multilocus sexdetermination system is at work in the Salicaceae, I hypothesized that I should be able to use fine-scale mapping of genetic markers linked to gender that, when combined with the 27  sequenced genome, would enable me to pinpoint the exact location(s) of the P. trichocarpa sex locus/loci.  RAPD markers associated with sex-determination in P.  tomentosa have been reported (Hou et al., 2009), however, so far attempts to find markers that reliably co-segregate with gender in P. trichocarpa have failed (G. Tuskan, pers. comm.). 1.5.4  Gender-determination and chromosome 19 in Populus In the genus Populus, studies suggest that gender is controlled via a genetic  mechanism, and that the sex locus is located on chromosome 19 of the linkage map or sequenced genome (Paolucci et al., 2010). Initially it was thought that in Populus the females may be the heterogametic gender, and that the males are homogametic at the sex locus. This is because in the closely related genus Salix, sex specific markers have been found for females but not for males (Gunter et al., 2003b). However, this is opposite to what has been found in Asparagus and many other genera containing dioecious species (Semerikov et al., 2003). The association between gender-determination and chromosome 19 in the genus Populus was first reported in Populus nigra (Gaudet et al., 2008) and P. trichocarpa (Yin et al., 2008), where detailed genetic maps constructed for these species indicated that a genetic marker segregating with gender existed on chromosome 19. Several features of chromosome 19 in P. trichocarpa, such as a region of recombination suppression and segregation distortion extending over a large portion of the chromosome, as well as haplotype divergence observed for this chromosome suggest that it is involved in sexdetermination (Yin et al., 2008).  28  It appears that there is considerable variability across the six species in which gender has been investigated in Populus as to the position of the putative sex locus on chromosome 19, and research indicates that either males or females can be the heterogametic gender, depending on species. In P. trichocarpa, the sex locus is reported to be located on the peritelomeric portion of chromosome 19, and evidence from distorted segregation of microsatellite markers, suppression of recombination, and haplotype divergence observed only in the maternal parent in pedigreed crosses indicates that the female is the heterogametic gender in this species (Yin et al., 2008). A putative sex locus in P. nigra is also located on the terminal end of chromosome 19, though in this species the mapping of AFLP, SSR and SNP genetic markers on the genome indicated that the male was the heterogametic gender (Gaudet et al., 2008). In two species of Populus the sex locus has been mapped to a non-telomeric position on chromosome 19. In P. tremuloides, sex as a morphological trait was mapped to a central position on chromosome 19 in the male parent using an interspecific pedigreed of 61 full sibling hybrids of P. tremula L. x P. tremuloides Michx. (Pakull et al., 2009). The sex locus also maps to the middle of chromosome 19 in P. alba, though in this case the heterogametic sex is reported to be the female (Paolucci et al., 2010). In P. tomentosa, a RAPD marker has been identified that produces a DNA fragment in the male individuals that were sampled, but not the female individuals, though the reason for this difference in DNA amplification between the genders has not been identified (Hou et al., 2009).  29  1.6  Research objectives To further explore the genetics of gender in dioecious species I have used P.  trichocarpa as a model species to investigate the development of sex-linked markers for Populus based on those reported in Salix viminalis (chapter two). I conducted a survey of the genes located on the telomeric end of chromosome 19 looking for gender-biased SNPs in gene sequences (chapter three), as this region of the genome in numerous Populus species has been reported to be involved in gender-determination in this genus. I also investigated gender-bias in genome wide gene-expression using a microarray approach (chapter four). Finally I identified three MADS box genes that show a genderbiased expression pattern in male and female floral tissues (chapter five).  30  Chapter 2: Characterization of sex-linked genetic markers developed in Salix viminalis in other Salix species and Populus trichocarpa 2.1  Introduction The majority of flowering plants produce hermaphroditic flowers, which contain  both male (stamens and pollen) and female (style, stigma and ovules) reproductive parts (Ainsworth, 2000). Only approximately ten percent of the world’s angiosperms have mating systems that divide male and female function between separate flowers on the same individual (monoecy), or in some cases, separate individuals (dioecy). Of this ten percent, approximately six percent (14 000 species of flowering plants) are dioecious (Renner and Won, 2001). Gender systems in plants have been characterized morphologically and physiologically, and possible explanations for the ecological basis for their evolution have been studied as well (Ainsworth, 2000; Ming et al., 2007). However, until recently there has been little research on how gender is determined at the molecular level, or what evolutionary pathway(s) development of separate genders could have taken.  Many  studies on dioecious species have focused on plants that, unlike poplar and willows, have well-developed heteromorphic sex chromosomes such as plants in the genera Rumex or Silene (Ainsworth et al., 1995; Filatov et al., 2000). The publication of the Populus trichocarpa genome provides an exciting opportunity to explore the genetic underpinning of gender in plants, as it is the first dioecious species to have a sequenced genome. The genus Populus is closely related to Salix, in the family Salicaceae, and both these genera have 19 chromosomes with no evidence of heteromorphic sex chromosomes (Semerikov et al., 2003) Almost all species 31  in these two genera are dioecious, but two species (Populus lasiocarpa and Salix martiana) have been reported to be monoecious (Semerikov et al., 2003), and the mechanism by which sex is determined in this family is not well understood. Due to the strong association between the occurrence of dioecy and monoecy in related genera, it is thought that dioecy evolved via the intermediate step of monoecy in Salicaceae. Determining how gender is regulated in Salicaceae could increase the understanding of the evolution of sexual systems in plants, but this information also has important economic uses. Many Populus and Salix species are used in short-rotation plantations for energy production and it has been shown that male Populus clones have higher dry weight fiber yields on average than females (Tschaplinski et al., 1994). Gender in dioecious tree species therefore influences their economic value and can affect breeding schemes (Alström-Rapaport et al., 1998). Although Populus clones can achieve reproductive maturity in four to six years under specific conditions (Tuskan et al., 2006), the majority of individuals start flowering between five and ten years after germination (Braatne et al., 1996). Because of this, the process of selecting trees with traits that are beneficial to the wood fiber industry, or for use in ecological reclamation or phytoremediation can be a long one. The development of a sex-linked marker that could be used to determine the gender of clones prior to sexual maturity could aid selective breeding programs. 2.1.1  Sex-linked genetic markers in Salix Previous work in Salix viminalis has identified a genetic marker that appears to  segregate with gender. Alstrom-Rapaport et al. (1998) discovered a decamer primer (UBC 354) which generated a randomly amplified polymorphic DNA (RAPD) product  32  consisting of a single 560bp band. This marker was shown to be biparentally inherited, and is associated with femaleness in S. viminalis. The pattern of gender segregation led the authors to propose that a two-locus epistatic genetic model of sex-determination exists in this species.  However, a later study using amplified fragment length  polymorphism (AFLP) fragments suggested a single locus determinant of gender in S. viminalis (Semerikov et al., 2003).  This same study found four AFLP fragments  associated with sex in S. viminalis, all of which were predominantly present in females, but absent in males (Semerikov et al., 2003). The results from these two studies suggest that females of this species may possess some chromosomal regions not found in the males. However, the sex-linked markers developed in S. viminalis did not co-segregate with gender in Salix caprea (Semerikov et al., 2003), which suggests that species of the genus Salix do not have complete synteny with respect to their sex-determination system. Sequence characterized amplified region (SCAR) markers developed from the original S. viminalis RAPD markers identified by Alstrom-Rapaport et al. (1998) resulted in identification of two sex-linked markers associated with femaleness in this species (Gunter et al., 2003a). These two DNA SCAR markers, SCAR 354 and SCAR AE08, were tested in Salix eriocephala Michx. and it was found that the presence or absence of both these markers differed significantly between male and female plants (Gunter et al., 2003b). 2.2  Objectives It is not known how conserved these SCAR markers for gender are across a  broader range of willow species, or if the markers will be associated with sex in other genera in Salicaceae, such as Populus. In order to widen the survey of willow species  33  tested with these markers I identified three objectives for my research. For my first objective I attempted to amplify the SCAR 354 and AE08 markers in 12 willow samples (comprising different species and sexes), and in Populus trichocarpa males and females, and sequenced the resulting products. My second objective was to design primers based on the poplar genome sequence in order to expand the amplified SCAR sequence into the associated coding gene regions identified in poplar, to further characterize the sex-linked SCAR marker. The third objective was to sequence the gene-anchored sequences that primers were designed for, and see if differences in the DNA sequences in this region could be identified that would explain why this SCAR marker appears to be sex-linked. 2.3 2.3.1  Methods Collection and preparation of plant materials I collected leaf samples from twelve species of Salix from the UBC Botanical  Gardens, and VanDusen Botanical Gardens, Vancouver B.C., in the fall of 2006, and stored them in silica gel at room temperature to preserve and desiccate the tissues until DNA was extracted from them. The sex of the Salix species collected was determined if possible. I collected the P. trichocarpa leaf samples used in this work in April of 2006, from trees in natural stands located on the UBC campus that I had previously identified as males or females. The P. trichocarpa leaf tissue was collected fresh and stored at 80°C. Collections of Salix reticulata and S. arctica were made in the spring of 2006 from natural populations near Prince George B.C., and stored in silica gel at room temperature. Genomic DNA was extracted from leaf tissue of twelve Salix species, and two Populus trichocarpa individuals using a modified CTAB procedure (Doyle and Doyle,  34  1987). DNA samples were cleaned up using the QIAquick PCR purification kit (ON, Canada) and stored at minus 20ºC. 2.3.2  PCR conditions and sequencing of samples SCAR primers were ordered from Integrated DNA Technologies, Inc (Coralville,  IA, USA).  The fifteen genomic DNA samples were tested for marker presence or  absence with SCAR 354 (forward primer: 5’-GAGAGGGAGGGAGATTTAAG-3’; reverse primer: 5’-GCCGTAGCAGATTGTTAATCAC-3’) under the following reaction conditions: Each 50 !l reaction contained 5 !l of 10X Taq buffer (100mM Tris-HCl (pH 8.8), 500mM KCl, 0.8% Nonidet P40), 5 !l of 5µM forward primer, 5 !l of 5µM reverse primer, 5 !l of 2mM dNTPs (Fermentas Life Sciences, CA, USA), 3 !l of 25 mM MgCl2, 0.25 !l Taq DNA polymerase (Fermentas Life Sciences, CA, USA), 40 ng of genomic DNA and ddH2O made up to 50 !l. Polymerase chain reactions (PCRs) were run on an Eppendorf Mastercycler gradient thermocycler for 35 cycles under the following reaction conditions: 94˚C 1 min, 55˚C 1 min, and 72˚C 2 mins, with an initial 3 min 95˚C denaturation step and a final 5 min 72˚C polymerization step. Amplification products were visually scored for the presence or absence of marker bands. Amplified PCR products were then purified using the QIAquick PCR purification kit (ON, Canada), resuspended in ddH2O, and prepared for sequencing. Each 10 !l sequencing reaction contained 1 !l of 5µM primer (either forward or reverse), 40ng amplified DNA suspended in ddH2O, 0.5 !l BigDye v3.1 (Applied Biosystems, CA, USA), 2.5 !l 5x BigDye v3.1 buffer, and ddH2O to 10 !l. Sequencing reactions were run under the following conditions: 96˚C 10 seconds, 55˚C 5 seconds, 60˚C 4 mins, with a 72˚C 1 min extension, and an initial 96˚C denaturation step.  The sequenced products  35  were purified using a Sephadex column procedure (Graham and Olmstead, 2000), and run out on an ABI 377XL machine. 2.3.3  Sequencing alignment for the SCAR sequences amplified in Salix Resulting sequences were aligned using Se-Al 1.d1 Sequence Alignment Editor,  available from http://evolve.zoo.ox.ac.uk (Rambaut, 1996) and Sequencher 4.2.2 (Gene Codes  Corp.,  Ann  Arbor,  (http://www.ch.embnet.org).  MI),  and  displayed  using  Boxshade  3.21  The sequence for the Salix viminalis SCAR amplified  fragment sequence (Gunter et al. 2003b; accession number = AY192565) was obtained from the NCBI website (www.ncbi.nlm.nih.gov/Genbank/index.html) and added to the alignment as well.  Primer sequences were removed from both ends of the SCAR  sequences prior to BLASTing them against the poplar genome on the JGI website (http://genome.jgi-psf.org/Poptr1_1/Poptr1_1.home.html), and the resulting Populus sequence that had sequence similarity with the Salix sequences was added to the alignment. Sequences were deposited with GenBank (accession numbers EF206296EF206307). The Salix SCAR marker sequences were aligned and compared to the sequenced Populus trichocarpa genome using a BLAST search. All the work reported here was completed prior to the publication of version 2.0 of the poplar genome, so gene names and annotations are with respect to version 1.1 of the poplar genome. 2.3.4  Novel primer design based on SCAR sequence and associated coding gene  regions in P. trichocarpa Following the comparison of the SCAR sequences to the P. trichocarpa genome, I designed primers based on the poplar genome sequence in order to expand the amplified SCAR sequence into the associated coding gene regions identified in poplar. Four new  36  PCR primers for this project were designed based on the P. trichocarpa genome sequence that showed similarity to the Salix SCAR sequences (Table 2.2, Figure 2.1). Primers were designed using Primer3 (v. 0.4.0, http://frodo.wi.mit.edu/primer3/), and tested in silico to determine the probability of primer dimers or unspecific primer binding with Amplify 3X (v.3.1.4, http://engels.genetics.wisc.edu/amplify). The twelve willow and two poplar genomic DNA samples were then tested, as well as thirteen individuals of S. arctica (nine female, and four males), and twelve individuals of S. reticulata (eight females and four males), for marker presence or absence with the new primers using the same PCR conditions that were used for SCAR 354 amplification, but with an annealing temperature of 50˚C.  PCR products from these reactions were then purified and  sequenced following the same protocol as above in section 3.2. 2.3.5  Cloning, sequencing and phylogenetic analysis of S. arctica and S. reticulata  samples In order to further investigate the relationship between SCAR 354 and gender, I sequenced 13 individuals of S. arctica (9 females and 4 males) and 12 individuals of S. reticulata (8 females and 4 males). Because S. arctica is a tetraploid and S. reticulata is a diploid, to get good quality sequences for these willow samples, I cloned the samples using the TOPO TA Cloning Kit Sequ 20 rxn (cat. # K457501, Invitrogen Canada Inc., Burlington, ON), and then confirmed the presence of my target sequence in the plasmid DNA by performing an Eco R1 digestion. The plasmid DNA was cleaned up using the QIAprep Spin Miniprep Kit (50) (cat. # 27104, Qiagen Inc., Toronto, Ont., Canada), and then samples were sequenced following the same protocol outlined above in section 3.2. Once I obtained good quality sequences for all the individuals, they were aligned  37  following the same method outlined above. I used PAUP (Swofford, 2001) to construct a consensus tree, and then looked at SNP variation using parsimony (DELTRAN), as implemented in MacClade 4.0 (Maddison and Maddison, 2000). 2.4  Results  2.4.1  Amplification and characterization of the SCAR 354 marker No amplification products were obtained with SCAR AE08 so all the work  reported here concerns SCAR 354, and the primers designed are based on SCAR 354 and Populus trichocarpa sequences. The SCAR 354 marker was amplified in only five of the Salix species tested (S. caprea, S. fargesii, S. gracilistyla, S. glauca, and S. nakamura) and was not amplified in Populus trichocarpa. To the extent that the sex of this material was known, I was able to amplify the SCAR 354 marker in both male and female willow samples (Table 2.1).  The sequences appeared to be homologous to the previously  determined sequence for SCAR 354 in S. viminalis (GenBank AY192565). When the amplified SCAR 354 sequence was BLASTed against the poplar genome, the part of the sequence that was conserved across Salix species had very high similarity to a sequence in Populus trichocarpa that maps to chromosome 15 (LG_XV) in  the  poplar  genome  (Figure  2.1),  very  close  to  a  Ssu72-like  gene  (fgenesh1_pg.C_LG_XV000380). A putative homologue of this poplar gene also occurs on chromosome 1 of Arabidopsis (AT1G73820.1, Ssu72-like family protein) with 82% similarity to the gene in poplar. However the upstream region in Arabidopsis shows no substantial similarity with the upstream region in P. trichocarpa and Salix species. Based on the comparison of the various willow sequences, three distinct regions were identified in the amplified region (Figure 2.1). At the 5’ end is a microsatellite-like  38  region (MSR), which consists of approximately 177bp, and varies considerably in length among the five Salix species it was amplified in (Table 2.1) but is not found in poplar. Adjacent to the MSR is a semi-conserved region (SCR) which shows good similarity among the Salix species but which does not completely align with the P. trichocarpa sequence. The majority of the SCAR 354 sequence consists of a conserved region that shows a 92% similarity between the Salix species and P. trichocarpa. This conserved region is 298bp long in P. trichocarpa, and 313bp in S. viminalis; the discrepancy in sequence length in this region is due to three insertions totaling 15bp in Salix. The conserved region is immediately 5’ to the Ssu72-like gene (85 bp from the putative  start  site  of  the  gene).  The  close  proximity  to  the  gene  fgenesh1_pg.C_LG_XV000380 (a putative protein involved in transcriptional start site  selection) is a possible reason for the high level of sequence conservation across species. Table 2.2 lists the other putative genes that are located in the vicinity of the homologous sequence to SCAR 354 in the poplar genome.  39  Table 2.1 Species tested for amplification with the SCAR 354 primers. Species  Accession/Collection number  Sex  Amplification  Salix (Chosenia) arbutifolia UBC-BG 036493-0126-2002  N/D*  no  S. caprea  VanDusen 0822 1993  female  yes  S. eleagnos  UBC-BG 013854-0013-76  female  no  S. fargesii  22112-027-1982  male  yes  S. glauca  UBC-BG 14540-284-77  N/D  yes  S. gracilistyla  UBC-BG 034625-5555-1999  male  yes  S. hookeriana  UBC-BG 037096-1003-2004  N/D  no  S. lapponum  UBC-BG 013859-0013-1976  female  no  S. lucida  UBC-BG 10512-268-74  N/D  no  S. nakamura  UBC-BG 033409-653-97  male  yes  S. purpurea  UBC-BG 034628-5555-1999  female  no  S. sitchensis  UBC-BG 2538-099-71  N/D  no  Populus trichocarpa  54-524  female  no  P.trichocarpa  50-533  male  no  .  *N/D indicates that the gender was undetermined. 2.4.2  Gene-anchored amplification in Salix and P. trichocarpa Three primer pairs successfully amplified part of the coding region, and the  conserved region immediately to the 5' end of the Ssu72-like gene in Salix and poplar (Table 2.3). The amplification using 1F/2R was consistent irrespective of species or gender (Table 2.4). Amplification using 1F/5R was less consistent but gave longer sequences for S. caprea and P. trichocarpa (Figure 2.2). These results confirm that the SCAR marker sequence is adjacent to the Ssu72-like gene in Salix species just as in P. trichocarpa.  40  2.4.3  Identifying SNP differences that may explain why this SCAR marker  appears to be sex-linked The SCAR 354 sequence consisted of a length of variable purine-rich repeats, and a region highly conserved between species (Figure 2.1).  The conserved region  corresponds to the putative promoter region of an Ssu72-like gene (involved in transcriptional start site regulation) on chromosome 15 in poplar. Gene-anchored primers were designed to amplify the conserved region and part of the Ssu72-like coding region in willows and poplar (all genders and species). The conserved region the gene-anchored primers amplified contained numerous single feature polymorphisms (SFPs) both within and between species. To investigate the relationship between SCAR 354 and gender, 12 individuals of S. reticulata (eight females and four males), a diploid willow species (Figure 2.3), and 13 individuals of S. arctica (nine females and four males), a tetraploid willow species (Figure 2.4), were sequenced. A consensus tree attained for S. reticulata did not indicate that the SNPs segregated with gender, but a bootstrap analysis supported the existence of haplotypes. A consensus tree attained for S. arctica did not indicate that the SNPs segregated with gender, but a bootstrap analysis supported the existence of three haplotypes. I detected female biased SNPs in both S. reticulata (one SNP) and S. arctica (three SNPs) consensus sequences, though the majority of SNPs in both species did not show a gender-bias in their distribution.  41  Figure 2.1 Structure of SCAR 354 marker regions in Salix species, and homologous region found on the Populus trichocarpa genome. MSR = microsatellite-like region, SCR = semi-conserved region. The numbers above the SCAR indicate the base pair number, 5’ to 3’. The numbers below the poplar region are the base pair positions along chromosome 15.  42  Table 2.2 List of putative genes in the region surrounding the location of the Populus trichocarpa sequence on chromosome 15 that show homology to the SCAR 354 Salix sequence. Gene Name Location on LG_XV -5 fgenesh1_pg.C_LG_XV000375 4162512-4165784 -4 fgenesh1_pg.C_LG_XV000376 4172930-4174370 -3 estExt_fgenesh4_pg.C_LG_XV0383 4176495-4177658 -2 eugene3.00150420 4181734-4182456 -1 estExt_Genewise1_v1.C_LG_XV1107 4186527-4190414 0  fgenesh1_pg.C_LG_XV000380  4199692-4203247  +1 +2 +3 +4 +5  eugene3.00150423 fgenesh1_pg.C_LG_XV000390 grail3.0043012001 grail3.0043012101 grail3.0043012201  4207845-4210979 4214615-4217179 4229045-4229320 4229463-4229628 4229950-4233882  Putative gene function C-type lectin Predicted transposase Not known Not known Ribonucleoprotein complex SRP, Srpl 1 component Ssu72-like (Protein involved in transcription start site selection) Not known Aldehyde dehydrogenase Leucine-rich repeat proteins Leucine-rich repeat proteins Leucine-rich repeat proteins  Table 2.3 Gene-anchored primers designed to amplify conserved regions 5' of the poplar Ssu72-like gene (fgenesh1_pg.C_LG_XV000380) and the equivalent region in Salix. Primer name Ssu72-like 1F Ssu72-like 1R Ssu72-like 2R Ssu72-like 5R  Primer sequence (5'-3') GAA CAA TTC TCA ATC AAG TTC ATC ACC GGC ACY GTA CAG TGT TCT GAT TSG ACG AAC TAT AAG GGG TGC CAA AGT CG  Tm 49.3 56.0 49.6 54.6  .  43  Table 2.4 Species tested for amplification with Ssu72-like primers. Species Sex Salix (Chosenia) arbutifolia N/D* S. caprea female S. eleagnos female S. fargesii male S. glauca N/D S. gracilistyla male S. hookeriana N/D S. lucida N/D S. nakamura male S. purpurea female Populus trichocarpa female P. trichocarpa male *N/D indicates that the gender was undetermined.  1F + 1R yes yes yes no yes no yes no yes yes yes yes  1F + 2R yes yes yes yes yes yes yes yes yes yes yes yes  .  Figure 2.2 Sequence alignment of the SCAR 354 sequences obtained from Salix caprea and Populus trichocarpa (for the region alignable between the two genera). JGI indicates the sequence from the genome project and UBC is another poplar sequence (50-533). There is an 8 bp deletion in the genome sequence (JGI) relative to the other sequence (UBC). The relative positions of the SCAR sequence and the coding region are indicated.  44  Figure 2.3 Consensus sequence (402bp long) for S. reticulata obtained from geneanchored primers amplifying the SCAR 354 conserved region and a portion of the coding region of the Ssu72-like gene. Insertions are indicated by red, deletions are indicated by X, SNPs are indicated by green. Female biased SNPs are indicated in purple. In cases where there are two alternate bases for a SNP, the most frequent change is above the line.  45  Figure 2.4 Consensus sequence (522bp long) for S. arctica obtained from gene-anchored primers amplifying the SCAR 354 conserved region and a portion of the coding region of the Ssu72-like gene. Insertions are indicated by red, deletions are indicated by X, SNPs are indicated by green. Female biased SNPs are indicated in purple. In cases where there are two alternate bases for a SNP, the most frequent change is above the line.  46  2.5 2.5.1  Discussion SCAR 354 amplification in Salix. My study, using species other than S. viminalis, found that amplification using the  original SCAR 354 primers is inconsistent with respect to species and gender. Salix nakamura, S. gracilistyla, S. glauca and S. caprea yielded discrete amplification products when tested with the SCAR 354 primers, and of these four species, S. nakamura and S. gracilistyla have been confirmed as male, and S. caprea is a female. For S. glauca, the sex is as yet undetermined due to poor growth of the plant. This inconsistency is to be expected, as there is considerable nucleotide divergence between the species sequenced. All the species of willow that successfully amplified are in subgenera Vetrix or Chamaetia, which are separate from subgenus Salix (Azuma et al., 2000). Comparative sequence  alignment  reveals  different  levels of  sequence  conservation in the SCAR region. The microsatellite-like region (MSR) at the 5’ end of the SCAR 354 sequence does not have a counterpart in the poplar genome (Figure 2.1), but is present and variable in length in all Salix species tested. The MSR consists of an AG (purine)-rich repeat region with many AGAGG and similar repeats. In the sequences the MSR varies in length, shortest in S. gracilistyla and S. nakamura, and longest in S. viminalis (GenBank AY192565). The conserved region identified between the SCAR 354 sequences in the Salix species and the P. trichocarpa sequence is located 84 bp from the 5’ end of a putative gene on the poplar genome (Figure 2.1). The conserved region also contains a candidate TATA box. Promoters are generally located 50-300 bps away from the beginning of a gene (Cooper et al., 2006), which is consistent with the location of this conserved region.  47  2.5.2  Amplification of the Ssu72-like gene region in Salix and P. trichocarpa Given that there is 92% similarity between the Salix and P. trichocarpa sequences  in the conserved region (Figure 2.2), and that this region exhibits many of the characteristics of a promoter region, it is plausible that, as in the poplar genome, the SCAR marker in Salix is also adjacent to this gene. This hypothesis was tested by designing primers based on putatively conserved regions of the poplar gene and the conserved part of the Salix SCAR. The designed primer pairs consistently amplified this region, and the resulting sequences indicate that the association between the SCAR and Ssu72-like gene in the Salix species sampled is homologous to P. trichocarpa (Figure 2.2).  This is not  unexpected as Salix and Populus are closely related, have the same chromosome number and share a whole genome duplication event prior to 65 million years ago (Tuskan et al., 2006). The Ssu72 protein in yeast consists of 206 amino acids and functions to decrease the elongation rate of RNA polymerase II by balancing elongation and termination of this polymerase activity (Dichtl et al., 2002). In yeast it is thought to be a phosphatase that interacts with CTD kinase Kin28 and CTD phosphatase Fcp1 to terminate the transcription of pre-snoRNA and some pre-mRNAs, which in turn regulates cis and trans-acting signals in the cell (Ganem et al., 2003). The poplar Ssu72-like protein has a single-copy homologue in Arabidopsis, the gene At1g73820 (chr. 1: bp 27675718-27781654). The putative Arabidopsis protein is the same length and highly conserved relative to poplar. The main differences between the genes are (1) a greatly expanded second intron in the poplar gene, and (2) the absence 48  in Arabidopsis of the 87 bp intron between the third and fourth exons (Arabidopsis therefore has a three exon gene structure rather than the four exon gene structure of poplar). 2.5.3  Identifying single feature polymorphisms (SFPs) that may explain why this  region is sex-linked in some Salix species. Consensus trees attained for S. arctica indicated that while the individuals sampled did not segregate with gender, three distinct haplotypes supported by bootstrap analysis seem to exist within this species for the SCAR 354 marker sequence. For S. reticulata, two haplotypes were supported by bootstrap analysis of the data. These results could also be explained by sequencing errors or pseudo gene clones, but the existence of difference haplotypes within the individuals sequenced is the best explanation for these results. It appears that these SFPs could be the reason for SCAR marker 354 segregating with gender, as the phylogenetic analysis of the S. arctica and S. reticulata populations seems to indicate that male and female individuals are segregating according to gender, though the gender-bias is not statistically significant. Studies of Arctic willow species have suggested that the gender-bias observed in populations of some species could be explained by niche partitioning or gender-biased mortality due to variation in physiology between males and females. In populations of S. polaris and S. herbacea a 60:40 female to male gender ratio has been documented, and in S. polaris it has been observed that females have greater stomatal resistance than males, which perhaps makes females less susceptible to desiccation during the short Arctic growing season (Crawford and Balfour, 1983). In S. arctica, 2:1 female to male genderbiases have been observed, and many physiological differences, such as leaf size and  49  water use efficiency, have been documented between males and females of this species (Dawson and Bliss, 1989).  Growth chamber experiments using S. arctica seem to  indicate that there is a genetic basis for the sex-specific differences between the genders in this species (Dawson and Bliss, 1989), and therefore that the gender-biased distribution of females in wetter habitats and males in dryer habitat is not caused solely by gender specific mortality. It appears that once the genetic mechanism for determining gender in Salix is discovered, it will be possible in some species to link the genetics of secondary sexual characteristics such as leaf size or drought tolerance to gender. In a harsh environment such as the arctic, with short growing seasons, it may be important for the ecological requirements of the genders, and therefore their physiology, to diverge to minimize competition between the genders for limited resources (Cox, 1981). 2.5.4  Potential utility of this region The consistent amplification of the conserved Ssu72-like gene-associated region  in the Salix species sampled indicates that this marker may be useful for population and phylogenetic studies in willow. There are numerous SFPs found in the sequences both within and between species. For instance there is an 8 bp deletion in the genome project sequence relative to the two other individuals of Populus trichocarpa sequenced (Figure 2.2). Sequence divergence in the conserved region is 0-5.4% within Salix, and 7.7-9.9% between Salix and Populus. In addition to sequence variation between Salix species, there are also indications of heterozygous nucleotide positions (from double peaks on the sequencing electropherograms) that are consistent when sequenced in both directions.  50  This raises the possibility that in the future association studies might be performed between individual SFPs and gender (or other characters). The conserved region has an apparent homologue in the poplar genome, where it corresponds to the putative promoter region of an Ssu72-like gene (involved in transcriptional start site regulation) on chromosome 15. The poplar genome sequence was used to design gene-anchored primers that consistently amplify this region and part of the Ssu72-like coding region in willows as well as poplars, irrespective of species and gender.  The gene-anchored primers amplify a region that, while conserved, has  numerous single feature polymorphisms (SFPs) both within and between species. This region could thus be used for population and phylogenetic studies. 2.5.5  Considering this research and version 2.0 of the Populus trichocarpa  genome. I completed this research prior to the release of the second version of the Populus trichocarpa genome, but as this resource is now available, I revisited the results in order to include the most current gene annotation information available for the sequences I worked with for this project. Notably, the content of Table 2.2 has changed significantly due to the changes in genome assembly and gene model names used between version 1.1 and version 2.0 of the poplar genome. I performed a BLAST search of the sequences for the genes listed in Table 2.2 against version 2.0 of the poplar genome (http://www.phytozome.net). I was unable to find the gene model fgenesh1_pg.C_LG_XV000375 on version 2.0 of the genome, but all of the other genes had their best BLAST hits on chromosome 15, the same chromosome on which they were located on in version 1.1. The order of the genes was more or less  51  conserved  as  well,  except  that  the  poplar  Ssu72-like  gene  (fgenesh1_pg.C_LG_XV000380, POPTR_0015s04700 on version 2.0) is not at the centre of the list of putative genes in the region surrounding the location of the Populus trichocarpa sequence on chromosome fifteen that shows homology to the SCAR 354 Salix sequence (Table 2.5). The positional order of the nine genes is conserved in version 2.0 of the genome, as are the gene functional annotations for the most part (Appendix B). Three of the genes listed on version 1.1 of the genome (grail3.0043012001, grail3.0043012101, grail3.0043012201) are now considered to be one single gene on version 2.0 of the genome (POPTR_0015s04830).  52  Table 2.5 Comparison of the position of the list of putative genes in the region surrounding the location of the Populus trichocarpa sequence on chromosome 15 that shows homology to the SCAR 354 Salix sequence in version 1.1 and version 2.0 of the poplar genome. Genes are listed in the 5’ to 3’ direction on chromosome 15, and the gene fgenesh1_pg.C_LG_XV000380 / POPTR_0015s04700 encodes for the Ssu72-like protein. Gene order -5 -4 -3 -2 -1  Location 4162512 - 4165784 4172930 - 4174370 4176495 - 4177658 4181734 - 4182456 4186527 - 4190414  Version 1.1 Gene name Version 2.0 gene name fgenesh1_pg.C_LG_XV000375 Not Found fgenesh1_pg.C_LG_XV000376 POPTR_0015s04750 estExt_fgenesh4_pg.C_LG_XV0383 POPTR_0015s04760 eugene3.00150420 POPTR_0015s04770 estExt_Genewise1_v1.C_LG_XV1107 POPTR_0015s04780  Location Gene order N/A N/A 4964515 – 4966108 -4 4968103 – 4969266 -3 4973382 – 4974227 -2 4978598 – 4982760 -1  0  4199692 - 4203247  fgenesh1_pg.C_LG_XV000380  POPTR_0015s04700  4923219 – 4926558  -5  +1 +2 +3 +4 +5  4207845 - 4210979 4214615 - 4217179 4229045 - 4229320 4229463 - 4229628 4229950 - 4233882  eugene3.00150423 fgenesh1_pg.C_LG_XV000390 grail3.0043012001 grail3.0043012101 grail3.0043012201  POPTR_0015s04810 POPTR_0015s04820 POPTR_0015s04830 POPTR_0015s04830 POPTR_0015s04830  4999929 – 5003025 5011059 – 5013623 5022992 – 5030320 5022992 – 5030320 5022992 – 5030320  +1 +2 +3 +3 +3  53  2.6  Conclusions I was able to identify the position of the SCAR 354 marker on chromosome 15 on  the P. trichocarpa genome, and amplified the SCAR 354 marker sequence as well as the adjacent gene sequence for a Ssu72-like protein. This indicated that the position of this marker with respect to this gene is conserved in P. trichocarpa and the Salix spp. I was unable to confirm that the SCAR 354 marker that segregated with sex in S. viminalis was also a sex-linked marker in other Salix species or in P. trichocarpa. By sequencing the PCR products amplified by the gene-anchored primers, I characterized the SCAR marker sequence, and identified distinct regions of it that varied in the amount that they were conserved between the species I sampled.  When I  investigated the gene-anchored sequence obtained in males and females of S. arctica and S. reticulata I found some evidence that gender-biased SNPs do exist in this sequence, which may explain why the SCAR 354 marker segregates with gender in pedigreed families of S. viminalis (Alström-Rapaport et al., 1998), though the gender-bias I observed was not statistically significant. Given that the conserved Ssu72-like gene associated region consistently amplified in the Salix species I sampled, this marker may be useful for phylogenetic and population studies of willows.  It may be possible to discover species specific single feature  polymorphisms in the gene-anchored region my primers amplified, and these SFPs could prove to be useful genetic tools for identifying Salix species, which can be difficult to classify based on morphological traits alone.  54  Chapter 3: Investigation of chromosome 19 of the Populus trichocarpa genome to identify molecular genetic markers that can be used to identify gender 3.1 3.1.1  Introduction Investigation of sex chromosomes and sex loci to identify gender-  determining genes Gender-determining genes initiate the developmental process that results in male and female phenotypes. Plant genes have been found to be fairly compact and usually grouped together in clusters, surrounded by repetitive DNA sequences, even in large genomes (Kellogg and Bennetzen, 2004). When heteromorphic sex chromosomes were first observed in dioecious plant species such as Silene latifolia by Blackburn (1923), researchers began investigating these genomic regions looking for gender-determining genes using gene cloning techniques to enrich DNA libraries with sex-linked transcripts or sex chromosome sequences (Shibata et al., 1999). This approach has been used in dioecious species that have well defined sex chromosomes like Rumex acetosa and Silene latifolia, and it resulted in the identification of a few genes that may be involved in the gender-determination pathway in S. latifolia (Zluvova et al., 2006).  However, the  majority of dioecious plants do not have sex chromosomes, so this approach to investigating gender-determination has not identified the genetic mechanism involved in the majority of plant species studied, despite the numerous studies that have investigated the role of sex chromosomes in determining sex in haploid, diploid and triploid plants (Sakamoto et al., 2005).  55  3.1.2  Sex-determination in the genus Populus Given the 1:1 sex ratios that have been observed in Populus populations, gender-  determining genes probably segregate at one locus, or as several closely linked loci on an autosome. In the genus Populus, previous studies have indicated that chromosome 19 appears to be consistently involved in the genetics of gender, though there is some discussion as to which gender is homogametic and which is heterogametic, depending on the species studied. There have been four main studies on the genetics of gender in Populus. In the first study of this kind, Yin et al. (2008) identified a gender-associated locus that consistently mapped to the peritelomeric region of chromosome 19 in P. trichocarpa using microsatellite markers, and their results suggested that the female is the heterogametic gender. In 2008 Gaudet et al. also produced genetic linkage maps of Populus nigra L. using AFLPs, SSRs, and SNPs, and found a sex-linked marker that mapped to a terminal position on chromosome 19 in the male parent, suggesting that the male is the heterogametic gender. Genetic linkage maps in aspen (P. tremula x P. tremuloides) constructed using AFLP and SSR markers showed that a sex-linked marker could be mapped to a non-terminal position on chromosome 19 in the male P. tremuloides (Pakull et al., 2009). An AFLP and SSR marker-based genetic map of Populus alba L. also located the sex-determining locus at a non-terminal position on chromosome 19 of the female parental map (Paolucci et al., 2010). The results of these four studies are summarized in Figure 3.1. There have also been RAPD markers related to a sex locus reported in P. tomentosa by a Chinese research group, but no genetic location has been assigned to these markers (Hou et al., 2009).  56  As seen in Figure 3.1, while the sex locus in all four species investigated is located on chromosome 19, its position on chromosome 19 (either terminally or centrally located), and which gender is heterogametic seems to vary with species, with evidence that females are heterogametic in two species, and males are heterogametic in the other two species (Paolucci et al., 2010). While these observations may seem contradictory, there is evidence in other species that hybrid zones may play a role in plasticity of sexdetermination and the dynamics of how sex chromosomes evolve. It is well known that many members of the genus Populus interbreed and form viable hybrids easily (Hamzeh et al., 2006), so it may be that this ability has allowed Populus species to maintain a certain amount of flexibility in their sex-determination mechanism. The Japanese frog, Rana rugosa, forms population groups depending on the type of heterogamety and sex chromosome morphology (Hamzeh and Dayanandan, 2004; Janousek and Mrackova, 2010). In this species of frog there are five population groups, three that have a XX/XY sex-determination system, and two where the female is the heterogametic sex (ZZ/WZ) (Ogata et al., 2007). From the phylogenetic data on the species, it appears that the Y and Z sex chromosomes arose in populations from Western Japan, and the X and W sex chromosomes in central Japan in geographical isolation, and subsequently came together again and reciprocally hybridized, resulting in the different sex-determination systems observed within the same species (Janousek and Mrackova, 2010). Experimental breeding programs (Ogata et al., 2003) and theoretical studies (van Doorn and Kirkpatrick, 2007), support the idea that in populations that have a strong sex bias, strong positive selection pressure for the minor sex-favoring gene could reestablish a 1:1 sex ratio in a population. This is because a strong sexual bias will promote sexually  57  antagonistic genes to accumulate, resulting in heteromorphic sex chromosome formation or even the acquisition of a novel dominant sex-determining locus, and the evolution of a novel sex-determination system (van Doorn and Kirkpatrick, 2007). Given that the first  Figure 3.1 Summary of the data available on the position of the putative sex locus on chromosome 19 in the genus Populus. The putative sex locus of each chromosome is indicated by a red box. Markers indicated in red are genomic markers that are shared across all four Populus species studied. The numbers on the left of each chromosome indicate the marker positions in centiMorgans. The numbers in black on the right of each chromosome indicate the SSR markers used in mapping the sex loci. (Figure originally published by Paolucci et al., 2010 Copyright Springer-Verlag).  58  poplar fossils date back to the Eocene period (Cronk, 2005) it is quite plausible that a similar process could have affected the evolution of sex-determination in Populus, resulting in the different male or female herterogamy observed among the species studied thus far in the genus. 3.1.3  Features of chromosome 19 that may be involved in gender-determination Features of chromosome 19 on version 1.1 of the poplar genome such as a marker  associated with gender-determination mapped to this chromosome, and a region of recombination suppression on chromosome 19, as well as a haplotype divergence observed for this chromosome, suggest that it is involved in sex-determination in Populus trichocarpa (Yin et al., 2008). This research also indicated that a number of previously unassembled scaffolds, principally scaffold 117, seem to overlap with the peritelomeric region of chromosome 19 that appears to be associated with sex-determination in P. trichocarpa. One of the lines of reasoning put forward by Yin et al. (2008) to support the existence and position of a sex locus in P. trichicarpa was the evidence for recombination suppression on the telomeric end of chromosome 19. Scaffold 117, a 1MB segment of the P. trichocarpa genome containing ~80 gene models, was mapped to the peritelomeric region of chromosome 19 using 27 microsatellite markers in version 1.1 of the P. trichocarpa genome, in the region proposed to contain the sex locus (Yin et al., 2008). Fine scale mapping for scaffold 117 indicated no recombination within the upper 706kb region of chromosome 19, while the lower 257Kb region of this scaffold included several recombination positions with chromosome 19. Significant segregation distortion for alleles located on maternal haplotypes was observed between scaffold 117 and  59  chromosome 19, whereas no distortion within paternal haplotypes was observed between these two genomic regions (Yin et al., 2008). Based on these results, scaffold 117 potentially represents a divergent haplotype of the telomeric end of chromosome 19, and the distortion of the alleles located on the maternal haplotype provides evidence that the female is the heterogametic sex in P. trichocarpa. Following the reasoning that the telomeric end of chromosome 19 of the poplar genome may contain a sex locus, I decided to investigate gene sequences in the region of chromosome 19 that now incorporates the 1MB scaffold 117, the putative heterogametic haplotype of chromosome 19, looking for SNPs that segregate with gender. 3.2  Objectives In order to investigate how gender is regulated in Populus trichocarpa, I  identified two objectives based on the availability of the sequenced genome for this species, and previous work done that identified chromosome 19 of this genome as being involved with the genetics of gender differentiation in this species (Yin et al., 2008). My first objective was to identify if the homogametic gender contains two copies of telomeric region of chromosome 19, or two copies of a putatively sex-linked contiguous sequence (contig) labelled scaffold 117. My second objective was to develop a genetic marker that segregates with gender that may be associated with the genetic mechanism of genderdetermination, by looking at SNP variation in gene sequences located on the 5’ telomeric region of chromosome 19.  60  3.3 3.3.1  Methods Collection and preparation of biological materials I collected plant tissue from leaf buds of male and female P. trichocarpa trees in  March of 2006 and 2007 from natural stands located on the UBC campus previously identified as males or females by Dr. Cronk. These tissues were stored at –80ºC to ensure that they would be suitable for RNA and DNA extraction. I extracted DNA from 22 individuals (13 female and 9 male) using a modified CTAB extraction protocol (Doyle and Doyle, 1987), and DNA samples were diluted to a concentration of approximately 50ng/µl in nuclease free water, and then stored at -20 ºC. 3.3.2  In silico investigation of the gene content of scaffold 117 Given that it is thought that the female is the heterogametic gender in P.  trichocarpa (Yin et al., 2008), I attempted to determine if the male chromosome 19 consists of two copies of chromosome 19 as it appears in the sequenced genome, or two copies of scaffold 117. The gene content of scaffold 117 was investigated to develop a marker that segregates with gender, and to identify if the homogamete contains two copies of chromosome 19 sequence or two copies of scaffold 117. A series of in silico searches were performed to identify all the gene models located on scaffold 117, then all the scaffold 117 gene model sequences were BLASTed against the assembled version 1.1 P. trichocarpa genome to look for gene models that could be used to identify scaffold 117 versus chromosome 19, and therefore identify which of these scaffolds was present in two copies in the homogametic gender. Once the genes that were unique to scaffold 117 were identified, primers were designed based on the gene sequence reported in  61  version 1.1 of the poplar genome, and I attempted to amplify these genes using DNA extracted from the collected leaf tissue samples. 3.3.3  PCR conditions and sequencing of samples PCR primers for this project were designed using Primer3 (v. 0.4.0,  http://frodo.wi.mit.edu/primer3/), and tested in silico to determine the probability of primer  dimers  or  unspecific  primer  http://engels.genetics.wisc.edu/amplify).  binding  with  Amplify  3X  (v.3.1.4,  Primers were ordered from Integrated DNA  Technologies Inc (Coralville, Iowa), and a list of the primers used in this project is included in Appendix A. DNA amplification for this project was performed under the following reaction conditions: Each 25 !l reaction contained 2.5 !l of 10X Taq buffer (100mM Tris-HCl (pH 8.8), 500mM KCl, 0.8% Nonidet P40), 1 !l of 5µM forward primer, 1 !l of 5µM reverse primer, 2.5 !l of 2mM dNTPs (Fermentas Life Sciences, CA, USA), 2.5 !l of 25 mM MgCl2, 0.25 !l Taq DNA polymerase (Fermentas Life Sciences, CA, USA), 100 ng of genomic DNA and ddH2O made up to 25 !l. Polymerase chain reactions (PCRs) were run on an Eppendorf Mastercycler gradient thermocycler for 35 cycles under the following reaction conditions: Lid set to 94˚C, 94˚C 1 min, and 5057˚C 1 min (temperature range was dependant on primer annealing temperature), with an initial 3 min 94˚C denaturation step and a final 1.5 min 72˚C polymerization step. Amplification products were visually scored for the presence or absence of bands on 1% agarose gels stained with GelRed™ Nucleic Acid Gel Stain, diluted in water (Biotium, Hayward, CA 94545). Amplified PCR products were then prepared for sequencing by Macrogen, in Maryland USA.  62  3.3.4  Sequence and polymorphism data analysis SNPs in the gene sequences amplified for all individuals were identified using  phred/phrap/consed/polyphred (Ewing and Green, 1998; Ewing et al., 1998; Nickerson et al., 1997). This program was also used to edit and trim the DNA sequences before forming contiguous sequences (contigs) that corresponded with each individual that was sequenced. ClustalX (Larkin et al., 2007) was used to align the DNA sequences obtained for each gene, and then these alignments were edited using BioEdit version 7.0.5.3, (10/28/05) (Hall, 1999), before using DNA Sequence Polymorphism (DNAsp) (Librado and Rozas, 2009) to run a phase analysis. DNAsp uses statistical methods to infer haplotype phase, which allows for haplotype reconstruction from genotypic information (Librado and Rozas, 2009). I used DNAsp to perform calculations of the number of haplotypes represented by the individuals for a given gene, to estimate the amount of recombination between adjacent sites, the % average pairwise nucleotide diversity/site, the % of synonymous nucleotide diversity/site, the % of nonsynonymous nucleotide diversity/site, and the % nucleotide diversity per sequence for all the gene sequences obtained in the P. trichocarpa males and females were sampled. Nucleotide diversity values (! values) were calculated according to Nei (1987) and not Jukes and Cantor (1969) so that the values reported by this research would be comparable to those found in the literature for Populus species (Nei and Miller, 1990). Network (4.6.0.0, fluxusengineering.com) was used to construct the haplotype diagrams as visual representations of the haplotype data (Polzin and Daneschmand, 2003).  63  3.4 3.4.1  Results Investigation of gene content of scaffold 117 and the telomeric end of  chromosome 19 The first gene model of interest was annotated as a putative chloroplast terpene synthase that had three copies located on scaffold 117, and only one copy apparently located anywhere else in the poplar genome, on chromosome 19 (Table 3.1). The second group of genes consists of 10 gene models that are only found on scaffold 117 (Table 3.2). This research investigated the group of 10 genes unique to scaffold 117, as the putative chloroplast terpene synthase genes existed as multiple copies in the genome, making it difficult to develop specific primers that would differentiate between the different copies of these genes. With the release of the second version of the poplar genome on January 8th, 2010, it was important to update the information on the gene models, and find out if the gene models found on scaffold 117 were still associated with the telomeric region of chromosome 19, or if the content of scaffold 117 had been broken up and included in multiple other places in the genome. On the second version of the poplar genome, it appears that scaffold 117 has been incorporated into the telomeric end of chromosome 19. When the gene models located on scaffold 117 (version 1.1) were BLASTed against version 2.0 of the P. trichocarpa genome, the majority of the top hits were on a 1.7MB region of chromosome 19, and the number of gene models dropped from approximately 80 to 43.  This is because on version 1.1 of the genome, gene models sometimes  indicated that separate exons of the same gene were separate genes. The sequences of the putative chloroplast terpene synthase genes all BLASTed to chromosome 19 of version  64  2.2 of the poplar genome, in a region between 1075484 and 7014599bp, overlapping with six gene models. More reassuringly, the ten genes that were unique to scaffold 117 were all still located on chromosome 19, though some of them also produced blast hits with lesser amounts of sequence similarity on scaffolds 1, 3, 73, 121, 182 and 879. However, because scaffold 117 was incorporated into the telomeric region of chromosome 19 in the new assembly, it appears that scaffold 117 does not represent a separate gamete of chromosome 19, but is more probably a fragment of chromosome 19 that was not aligned correctly in version 1.1 of the genome. Primers were designed for 24 genes, just over half of the approximate 40 gene models in the region of chromosome 19 that corresponds to scaffold 117.  PCR  amplification of the 10 gene models unique to scaffold 117 was done (numbered one though ten, Figure 3.2). These genes were clustered in a 200kb region of scaffold 117, so genes were then chosen at 100kb intervals along the length of the scaffold to get a representative sample of SNP variation in genes on scaffold 117. Eight genes at 100 000bp intervals along the length of scaffold 117 were selected (labeled A though H, Figure 3.2). Given the differences in gene model annotation between version 1.1 and version 2.0 of the poplar genome, it is not surprising that many of the primers that were designed based on gene model sequence from version 1.1 of the genome did not successfully amplify the target sequences, or failed to produce PCR products at all. A list of the putative gene functions and location of all the genes worked with for this project is included in Appendix B.  65  3.4.2  Analysis of sequence and polymorphism data obtained for eleven genes  located on the telomeric region of chromosome 19 After performing a preliminary analysis of the haplotype data based on sequence obtained for using the gene model notation from version 1.1 of the poplar genome (http://genome.jgi-psf.org/Poptr1_1/Poptr1_1.home.html), information on the genes of interest in this study was updated to that available on version 2.0 of the genome, and thereafter version 2.0 of the poplar genome (http://www.phytozome.net) was used as the source of genomic information for designing primers for genes positioned on the telomeric end of chromosome 19. After looking at the SNP recombination rates for genes that sequence data had been collected for, gene SCA_117_10 showed the most interesting pattern of SNPs with regards to gender (Figure 3.3). This is a pattern of SNPs that would be expected in the region of a sex locus where recombination between alleles is suppressed. If these SNPS were sex-linked it would be expected that all the male haplotypes would group together, separate from the female haplotypes and vice versa. Given that this was the only gene that I sampled that showed this haplotype distribution, I decided to sequence genes on either side of this one to determine if they displayed similar patterns in SNP variation and recombination rates between adjacent sites (labeled SCA_19_1 through 5, Table 3.3). Genes that were evenly distributed in a 55 000 bp region on either side of the gene SCA_117_10 were chosen to see if the low recombination rate and distribution of haplotypes specific to males or females occurred in other genes in this region. After testing the primers for the 24 genes sampled from scaffold 117 for amplification in the sample of 22 individuals (13 females and 9 males), good quality  66  sequence was obtained for eleven genes (Table 3.3). Then a haplotype analysis was performed to look at SNP variation and see how it relates to sequence differences between gender (Table 3.3). When the estimated recombination between adjacent sites (R-values) was investigated for the genes located in the region of suppressed recombination on the telomeric end of chromosome 19 quite a lot of variability was found.  Table 3.1 Gene models located on scaffold 117 and chromosome 19 annotated as putative chloroplast terpene synthase in P. trichocarpa. Gene name  Location on P. trichocarpa genome  fgenesh4_pg.C_scaffold_117000025  Poptr1_1/scaffold_117:421459-422086  eugene3.01170028  Poptr1_1/scaffold_117:428369-431855  fgenesh4_pm.C_scaffold_117000002  Poptr1_1/scaffold_117:539441-542289  grail3.0085006801  Poptr1_1/LG_XIX:2752362-2753129  Table 3.2 Gene models unique to scaffold 117, updated with names and positions on version 2.0 of the P. trichocarpa genome. V 1.1 gene name and assigned code  V 2.0 best match gene name and location  eugene3.01170047(SCA_117_1)  POPTR_0019s01520 sca_19:1339019-1340779  fgenesh4_pg.C_sca_117000045(SC_117_2)  POPTR_0019s01530 sca_19:1347076-1348225  grail3.0117003001(SCA_117_3)  POPTR_0019s01540 sca_19:1348343-1349893  gw1.117.122.1 (SCA_117_4)  POPTR_0019s01560 sca_19:1379318-1382368  fgenesh4_pg.C_sca_117000051(SCA_117_5)  POPTR_0019s01570 sca_19:1391060-1395729  fgenesh4_pg.C_sca_117000053(SCA_117_6)  POPTR_0019s01560 and POPTR_0019s01570  fgenesh4_pg.C_sca_117000054(SCA_117_7)  POPTR_0019s01560 and POPTR_0019s01570  e_gw1.117.150.1(SCA_117_8)  POPTR_0019s01630 sca_19:1438061-1439700  gw1.117.169.1(SCA_117_9)  Between POPTR_0019s01660 sca_19:14608251462203 and 1670 sca_19:1463832-1466771  eugene3.01170072(SCA_117_10)  POPTR_0019s01790 sca_19: 1546894-1550419  67  Figure 3.2 Genes sampled along the length of scaffold 117 (version 1.1) and their corresponding positions on chromosome 19 (version 2.0). Bars indicate 100 000bp increments on scaffolds. Orange indicates no sequence data was obtained for the gene. Green indicates only 200bp of gene was sequenced, and no SNPs of interest were found/no sequence was produced so work with them was discontinued. Red indicates that good sequence for at least 800bp of the ORF of the gene was obtained. The ‘X’ by gene SCA_117_B indicates that this gene model did not exist in version 2.0 of the poplar genome.  68  Figure 3.3 Haplotype diagram for gene SCA_117_10 (gene model POPTR_0019s01790) based on 772bp sequenced in ten females and four males. In this diagram the circles represent nodes where pink indicates the proportion of female haplotypes, blue for male, and size of node is proportional to number of individuals with that haplotype. The length of the lines in not proportional to distance, but the red numbers indicate base pair position of SNPs. The red node indicates a haplotype that must exist for other nodes to be present, but was not observed in the sequences sampled.  69  Table 3.3 Overview of polymorphism data for genes located on the telomeric end of chromosome 19. n = number of chromosomes, S = number of variable sites, H = number of haplotypes, R = estimate of recombination between adjacent sites, !/site indicates % average pairwise nucleotide diversity/site, !sil indicates % of synonymous nucleotide diversity /site, !syn indicates % of synonymous nucleotide diversity /site, !nonsyn indicates % of nonsynonymous nucleotide diversity /site, and " is the % nucleotide diversity per sequence. Gene Gene ID on chromosome 19 n Length*(bp) S H R (%) !/site (%) !sil (%) !syn(%) !nonsyn(%) "/site (%) SCA_117_A POPTR_0019s01120 39 847 23 35 13.24 0.713 NA 0.659 0.736 0.643 SCA_117_2 POPTR_0019s01530 31 673 10 11 1.49 0.287 0.292 0.000 0.000 0.394 SCA_117_5 POPTR_0019s01570 31 815 9 2 0.00 0.071 0.061 0.233 0.082 0.276 SCA_117_F POPTR_0019s01670 45 853 13 15 0.23 0.338 NA 0.643 0.263 0.349 SCA_19_1 POPTR_0019s01700 37 528 29 21 1.31 1.804 3.031 3.248 1.266 1.406 SCA_19_2 POPTR_0019s01740 35 842 22 31 12.49 0.764 1.618 2.059 0.335 0.750 SCA_19_3 POPTR_0019s01780 43 731 12 19 3.03 0.412 0.258 0.190 0.568 0.413 SCA_117_10 POPTR_0019s01790 29 772 17 12 0.22 0.724 0.594 0.000 0.000 0.594 SCA_19_4 POPTR_0019s01830 31 838 14 15 1.98 0.452 0.724 0.623 0.284 0.448 SCA_19_5 POPTR_0019s01850 17 804 17 17 13.63 0.757 1.321 0.392 0.364 0.625 SCA_117_G POPTR_0019s01880 37 832 23 25 3.16 1.176 1.411 1.743 0.875 0.749 . Average: 34.1 775.9 17.2 18.5 4.62 0.682 1.034 0.890 0.434 0.604 *Length: excluding sites with gaps/missing data *n = this number of chromosomes is calculated by multiplying the number of diploid individuals sequenced for each gene, and adding the single haplotype of the reference sequence “Nisqually 1” from version 2.0 of the poplar genome. Example: For SCA_117_2 15 individuals were sequenced.  70  3.5 3.5.1  Discussion Identifying if the homogametic gender of Populus trichocarpa Identifying which gender is the homogametic one with respect to gender in P.  trichocarpa based on sequence data from chromosome 19 was complicated by the discrepancy in assembly data between version 1.1 and version 2.0 of the poplar genome. This discrepancy in alignment between the two assemblies of the poplar genome could be due to the fact that different software was used to create the two assemblies. Version 2.0 of the poplar genome, accessible at http://www.phytozome.net, was assembled using ARACHNE version 20071016HA, and the assembly covers 403 million base pairs of sequence with an average read depth of 7.45x assembled. ARACHNE is a wholegenome shotgun assembler that analyzes paired forward and reverse sequence reads that are obtained from both ends of plasmid clones, and assembles them with a reported 99% accuracy due to its ability to detect alignment errors caused by sequencing errors or false sequence overlaps caused by repeat DNA sequences in complex genomes (Batzoglou et al., 2002). Version  1.1  of  the  poplar  genome,  accessible  at  http://genome.jgi-  psf.org/Poptr1_1/Poptr1_1.home.html, was assembled using JAZZ, a group of programs developed by the Joint Genome Institute (JGI) designed specifically for working with large sequencing projects (Taylor and Semple, 2002). The first version of the poplar genome was estimated to consist of approximately 485 million base pairs and was sequenced with a read depth of approximately 7.5x (Tuskan et al., 2006). JAZZ uses a multistep process similar to that of ARACHNE to assemble whole genome shotgun  71  sequences, but has stricter assembly parameters that make it more likely that haplotypes will assemble separately (Shapiro, 2005). 3.5.2  Version 1.1 versus version 2.0 of the poplar genome The main difference between these two genome assemblers appears to be the way  in which they assemble supercontigs (ARACHNE) or scaffolds (JAZZ) which is how the genome assemblers take the shotgun sequences and group them into large contiguous assemblies that match up with the number of chromosomes in the genome. ARACHNE creates supercontigs by using the forward and reverse links from plasmid reads to orient and order unique contigs into longer sequences (Batzoglou et al., 2002). The JAZZ assembler attempts to build a scaffold by iteratively building and breaking sequence contigs, progressively including lower-quality sequence data (Taylor and Semple, 2002). Because of the discrepancy between the way in which the two versions of the genomes were assembled, it is difficult to know if scaffold 117 really represents an alternate haplotype of chromosome 19, that was detected by the JAZZ assembler (version 1.1) but not the ARACHNE assembler (version 2.0), or if the different way in which ARACHNE bridges gaps in the poplar genome assembly correctly incorporated the smaller scaffold 117 from the first version of the genome into the telomeric end of chromosome 19 in version 2.0 of the poplar genome. The sequence difference between the two gender-determining haplotypes of chromosome 19 may be too small to be detected at this point, even if both copies of chromosome 19 were sequenced since Nisqually-1, the genotype selected to be sequenced for the Populus trichocarpa whole genome assembly, was a female (Tuskan et al., 2006), the reported heterogametic gender in this species.  72  3.5.3  Developing a genetic marker on chromosome 19 that segregates with gender  in Populus trichocarpa I investigated 24 genes located on the 5’ telomeric end of chromosome 19 that was indicated to have suppressed recombination with scaffold 117 on version 1.1 of the poplar genome, looking for SNPs located in genes in this region that segregate with gender. There was a large amount of variability in the number of SNPs, or variable sites (S), detected in the sequences obtained for the genes that were studied (Table 3.3), however, none of the SNPs that were detected in any of the genes segregated with gender so I was unable to discover a genetic marker that could be used to sex P. trichocarpa individuals of unknown gender. Although the number of individuals that were sampled for each gene varied from eight to twenty-two, it appears that neither the length of sequence obtained for each gene, not the number of variable sites or the number of haplotypes observed, were correlated with the number of individuals sampled (Table 3.3). 3.5.4  Investigation of recombination rates in genes located in the telomeric region  of chromosome 19 in P. trichocarpa Based on the results of the investigation into the recombination rates of genes located on the 5’ telomeric end of chromosome 19, no overall trend of reduced recombination between adjacent SNPs was found in the gene sequences examined. In fact there was considerable variation in R-values even in genes that were adjacent to each other on the genome. An example of this is illustrated by the genes SCA_19_4 and SCA_19_5, which are only 21 134bp apart on chromosome 19, but differ in % R-value by a factor of 10. Three of the 11 genes I looked at, SCA_117_5 (POPTR_0019s01570), SCA_117_F (POPTR_0019s01670), and SCA_117_10 (POPTR_0019s01790), seemed  73  to show much lower recombination (Table 3.3). A pattern of low recombination between SNPs would be expected in the region of a sex locus where recombination between alleles is suppressed. Suppression of recombination would be expected in the region of a sex locus in order to maintain separate genders, but my examination of genes located on either side of these genes shows that these low R-values are not maintained across this region of chromosome 19. 3.5.5  Sex-linked markers on chromosome 19 of P. trichocarpa It appears that it is possible to develop genetic sex-linked markers that map to  chromosome 19 of P. trichocarpa, as three SSR markers that were identified in an interspecific cross between Populus tremula L. and Populus tremuloides Michx. were mapped to a central location on chromosome 19 (Pakull et al., 2011). It is interesting to note that none of the markers developed in the P. tremula x P. tremuloides cross were mapped to the region of chromosome 19 in P. trichocarpa that contained the genes that were investigated in this study. Also, all the fully sex-linked SSR markers that were mapped to a central position on chromosome 19 in P. trichocarpa appeared to be inherited from the P. tremuloides male parent (Pakull et al., 2011). These results seem to indicate that the male is the heterogametic gender in P. trichocarpa, which contradicts the finding of Yin et al. (2008) that provided evidence that in this species the female is the heterogametic gender, and also that a sex locus is located at the telomeric position on chromosome 19. It is possible that the evidence that indicates that different species of Populus show differences between which gender is heterogametic and where the sex locus is located on chromosome 19 can be explained by the idea that the evolution of sex paralleled speciation in this genus (Paolucci et al., 2010). However, I think it is more  74  likely that the emerging genetic evidence supports the theory that gender in Populus, and also in its sister genus Salix, is controlled by several sex-determining loci (AlströmRapaport et al., 1998), that in Populus appear to be located on chromosome 19. 3.5.6  Nucleotide diversity of genes located on the telomeric end of chromosomes  19 in P. trichocarpa Genetic variation in a species can be measured by looking at nucleotide diversity (!), and measuring genetic variation is important because it is related to most phenotypic variation observed and also can give clues to the evolutionary history of a species (Gilchrist et al., 2006). When I compared the average value of !/site (0.682%) observed for the coding regions of the eleven genes I sampled (Table 3.3) to the average level of nucleotide diversity (0.184%) observed in P. trichocarpa according to Gilchrist et al. (2006), I find that my average observed level of nucleotide diversity is higher, though my sample sizes were much smaller than the ones reported by Gilchrist et al. (2006). However, at least one of the genes I investigated, SCA_117_5, showed a !/site value of 0.071% (Table 3.3), which is in the same range of the !/site values for the genes investigated by Gilchrist et al. (2006), and this study also found that nucleotide diversity varies a great deal between genes, which is consistent with my observations. However, a more recent study of nucleotide diversity found an average !/site value of 3.38% for 3 separate genes, and this study had a sample size of 15 individuals, with similar lengths of sequence analyzed per gene to what I reported (Breen et al., 2009). It would appear that the nucleotide diversity I observed for the genes falls within an acceptable range for P. trichocarpa, with !/site values being gene dependant.  75  There are two likely explanations for my observation of much higher average amount of nucleotide diversity than the values reported by Gilchrist et al. (2006). First, I chose the genes based on their position on chromosome 19 to investigate recombination rates in this region of the poplar genome thought to contain a sex locus, and the genes in this study have functions annotated based only on sequence homology to genes characterized in other species, as yet unconfirmed by research in P. trichocarpa. The genes investigated by Gilchrist et al. (2006) were chosen based on data from evolutionary studies from other organisms, or because of their known functions in pathogen defense or wood quality, and therefore are probably under stricter genetic selection to avoid the accumulation of sequence mutations that would affect gene function (Gilchrist et al., 2006).  Choosing genes without prior knowledge of their function increases the  likelihood that the data from the loci will provide a more accurate and less biased view of genome wide patterns of polymorphism (Ingvarsson, 2008), so it may be that the genes I chose to work with represent a wider pattern of nucleotide diversity than those sampled by Gilchrist et al. (2006). Secondly, in this study between 8 and 22 individuals were sampled from one stand of trees, while Gilchrist et al. (2006) looked at between 37 and 40 trees, sampled from a wide geographic area, depending on the gene being investigated.  Those  researchers also sequenced gene fragments that were slightly longer than the sequences I investigated. The smaller tree sample size from a small geographic area, and shorter gene sequences used in this study may mean that the data reported here is not a good estimate for the average nucleotide diversity in the genes studied.  76  3.5.7  Nucleotide diversity observed in other species of Populus. Nucleotide diversity has been investigated in several other species of Populus,  notably P. balsamifera, which is the sister species to P. trichocarpa (Olson et al., 2010) and P. tremula, which is a more distant relative. The estimates for nucleotide diversity reported for three genes studied in P. balsamifera were similar in range to the !/site values found for the genes in this study, and Breen et al. (2009) had a comparable sample size (between 5 and 18 individuals) with an average sequence length of 609bp, about 100bp shorter than the average length of the gene sequences in this study (775.9bp, Table 3.3). !/site values reported for P. deltoides for the same three genes, also from the study by Breen et al. (2009), were also similar in range, and provide comparable values to support this. Several studies of P. tremula (Ingvarsson, 2005, 2008) have shown that this species has a high level of genetic variation relative to other Populus species. Nucleotide diversity (!/site) is reported at 1.1%, which is higher than my reported !/site of 0.682%, and five times greater that the average !/site values reported for P. trichocarpa (Gilchrist et al., 2006) and P. balsamifera (Breen et al., 2009). North American poplar species exhibit approximately 50% less population differentiation and lower nucleotide diversity compared to the values observed in P. tremula, but this difference may be explained by the fact that several of the studies of the North American species were conducted over small geographic scales compared to the studies of P. tremula, and may not have captured a species-wide estimate of nucleotide diversity (Breen et al., 2009). The lower nucleotide diversity observed in P. balsamifera compared to P. tremula may also indicate a historically lower effective population size in P. balsamifera, caused by population wide bottle necks experienced in response to the  77  expansion and retraction of ice sheets during the Quaternary period (Hewitt, 2004), followed by a recent population expansion (Keller et al., 2010). The range of !/site values found in my data is comparable to that reported in the literature for P. trichocarpa and other Populus species, so the !/site values reported in this study for the eleven genes that were sequenced adds to the information available on nucleotide diversity in P. trichocarpa. 3.6  Conclusions I showed that scaffold 117 reported in the version 1.1 genome sequence of P.  trichocarpa appears to have been a fragment of chromosome 19 in the poplar genome. There was little re-arrangement of the gene order between scaffold 117 of version 1.1, and the telomeric end of chromosome 19 on version 2.0 of the poplar genome. The gene model annotations in this region changed substantially as exons that were considered separate genes on version 1.1 of the genome were annotated as exons of the same gene on version 2.0 of the genome in many cases. I was ultimately unable to identify a way to distinguish the homogametic genotype from the heterogametic genotype in P. trichocarpa by using genomic markers in this region. I investigated 24 genes in the telomeric region of chromosome 19 looking for SNP variation in the genomic sequence, or reduced recombination rates in SNPs between males and females in these genes, which could be associated with genetic markers for gender and a sex locus. I observed large variability in the number of SNPs detected in the gene sequences studied, but was unable to discover a genetic marker that could be used to sex P. trichocarpa individuals of unknown gender. I found no overall trend of reduced recombination between adjacent SNPs in the gene sequences that were worked  78  with, and it appears from the data I collected that low recombination rates between SNPs are not maintained across the telomeric region on chromosome 19. If there is reduced recombination in the region of a sex locus on the telomeric end of chromosome 19, it would appear that it is very localized, as I looked at genes located less than 100kb from each other. Alternatively, it is possible that the sex locus is not in the ~1 000 000bp region I investigated. My investigation of nucleotide diversity in the telomeric region of chromosome 19 showed that nucleotide diversity varies a great deal between genes, even in the small region on the genome investigated. The ! values I observed for the genes investigated fall within the range previously reported for P. trichocarpa (Breen et al., 2009; Gilchrist et al., 2006), with !/site values being gene-dependant. It appears that the genetic mechanism for gender-determination in P. trichocarpa is more complicated than previously thought. However, research is now being conducted on gender-determination in multiple Populus species. These studies, along with maturing genetic resources such as the re-annotated version 2.0 of the P. trichocarpa genome make understanding the genetics of gender-determination in this species only a matter of time.  79  Chapter 4: Exploration of genome wide gender-biased gene-expression patterns to identify genes involved in gender-determination in Populus trichocarpa 4.1  Introduction In the maintenance of separate genders, a sex locus functions to control  differential gene-expression in males and females, as well as gender-biased geneexpression patterns. One approach to investigating gender-determination is to look at differential gene-expression patterns early in the development of male and female inflorescences or in vegetative tissues. Research into gene-expression and its role in gender-differentiation has provided information about the genetic basis for genderdetermination in three ways. The genes involved in controlling the development of male and female reproductive organs can be identified by relating specific gene-expression patterns in the floral meristems to organ position and development (Zik and Irish, 2003). Secondly, investigation of gene-expression differences between males and females in vegetative tissues may reveal genes that are responsible for secondary sexual characteristics. Many dioecious plant species exhibit gender-specific morphological differences in addition to their unisexual flowers, as well as small differences in ecological requirements (Lloyd and Webb, 1977). And thirdly, mapping genes that are differentially expressed between male and female individuals on to the physical map of the P. trichocarpa genome may indicate a region containing a sex locus. Clusters of gender-segregating genes may indicate areas of limited recombination in the genome, which are thought to be indicative of the initiation of sex chromosomes (Liu et al., 2004). Sex chromosomes start to evolve 80  when selection, acting through local suppression of recombination, tightly links all loci involved in gender-determination in a region that does not recombine, thereby ensuring that separate genders are maintained (Scotti and Delph, 2006). 4.1.1  The biology of gender-specific differences in dioecious species One technique used for identifying genes involved in gender-determination uses  homologous genes that are known to function in floral development in hermaphroditic model plants such as Antirrhinum and Arabidopsis to identify floral development genes that may control organ suppression in unisexual flowers (Ainsworth, 2000). There are many similarities between the genomes of P. trichocarpa and A. thaliana because they belong to the Eurorosid I clade of angiosperms (Jansson and Douglas, 2007) so it is plausible that extensive gene function is conserved between the two species. An example of this is the Populus PTD (Populus trichocarpa DEFICIENS) promoter gene, a B-class floral homeotic transcription factor, which is homologous to the MADS box genes DEFICIENS and APETALA3, that are found in Antirrhinum and Arabidopsis, respectively (Skinner et al., 2003). However, while there may be an evolutionary advantage for the genes to be inherited together without much recombination, genderdifferentiation genes involved in floral development do not need to be to be located at a sex locus as their expression could be trans-regulated by genes located at a sex locus. In fact, an in silico survey of the P. trichocarpa genome found no poplar MADS box genes located on chromosome 19 (Leseberg et al., 2006), which is the chromosome thought to contain the sex locus in this species (Yin et al., 2008).  81  4.1.2  Secondary sexual characteristics in plants Genes that show a gender-biased expression pattern in vegetative tissues may not  be directly involved in gender functions in the plant but may produce secondary sexual characteristics that identify male or female phenotypes outside of the floral tissues. There are many examples of secondary sexual characteristics being expressed in plants. Sexual dimorphism between the genders can result in differences in the life histories of males and females in a given species, as seen in Chamaelirium luteum, where female plants begin flowering at a greater age than males, flower less frequently and show a higher mortality rate than male plants in the same population (Meagher, 1984). The differences between males and females can also be morphological, as is seen in Asparagus officinalis where males usually produce more vigorous growth early in the growing season (Bracale et al., 1991), and in Cannabis sativa females of this species do not grow as tall as the males, and have larger root systems and leaf blades (Lloyd and Webb, 1977). In the genus Populus, numerous studies have indicated that there are vegetative morphological differences between males and females, as well as small gender related ecological preferences. Elevation seems to have an influence on the distribution of male and female individuals within populations of Populus tremuloides (Einspahr, 1960). Female P. tremuloides are more abundant than males at lower elevations, and female growth rates decreased with increasing elevation at more than twice the rate of males (Grant and Mitton, 1979). Though females of this species consistently showed a greater annual radial growth rate at all elevations than males, which contradicts a theory put forth by Lloyd and Webb (1977), sexual reproduction is more costly for females than for males (Grant and Mitton, 1979).  82  In P. augustifolia it was found that females are more flood tolerant than males, and a possible explanation for this is that seed-producing female trees located near streams would facilitate seed dispersal into recruitment areas along river beds (Nielsen et al., 2010). In P. trichocarpa it has been shown that male clones have higher dry weight fiber yields on average than females (Tschaplinski et al., 1994), though to date no reliable way has been developed to sex trees prior to their sexual maturity between the ages of five and ten years (Brunner, 2010), so investigating genes that are differentially expressed between males and females in vegetative tissues may yield a genetic marker for gender. 4.1.3  Clustering of differentially expressed genes near the sex locus Once genes with statistically significant differential expression in males and  females have been identified, their locations and linkage groups position can be determined using the sequenced P. trichocarpa genome. In P. trichocarpa it is thought that chromosome 19 may contain the sex locus (Yin et al., 2008), so investigating genderbiased gene-expression on this chromosome may yield insight into how gender is determined genetically in this species. Genes that are differentially expressed between the genders could be clustered around the sex locus because as suppression of recombination spreads along a chromosome from an initial mutation for female or male sterility (Nicolas et al., 2005), genes not directly involved in gender-determination could also begin to segregate with gender and cause slight phenotypic differences between the genders. Linkage of genes located at a sex locus is necessary in order to maintain separate genders (Charlesworth and Guttman, 1999), as without the suppression of  83  recombination of the female- or male-sterile mutations, the plant could revert to hermaphroditism (Ming et al., 2007). The evolution of recombination suppression at the sex locus is thought to be driven by sexually antagonistic genes, whose expression is advantageous in male and deleterious in females, or vice versa (Charlesworth and Mank, 2010). Genetic sexual antagonism has probably had a role in the evolution of dioecy because without it there would be no selective pressure to convert hermaphrodites into males (Bergero and Charlesworth, 2009). 4.1.4  Investigating gender-biased gene-expression using genome wide survey  approaches Gender-determining genes could be identified by investigating genes that are preferentially expressed in males or females using microarrays, or whole transcriptome expression data from next-generation sequencing. The advent of global gene-expression profiling approaches, and the availability of whole genome microarrays for Populus, made it possible to use microarrays to look at differential gene-expression between male and female individuals. This approach could identify genes controlling the expression of floral development genes produce separate male and female plants. Of more importance is the identification of genes whose expression is gender-biased in all tissues of the plant as these will provide potential RNA sex-markers and be possible candidates for sex linkage. The two main types of microarrays used in gene-expression experiments are cDNA arrays and oligonucleotide arrays. cDNA arrays are constructed using spotted cDNA clones that correspond to specific genes based on EST sequence information.  84  Oligonucleotide arrays are made using 16 to 20 pre-fabricated sequences 25-60 nucleotides long for each gene (Kafadar and Phang, 2003). Oligonucleotide microarrays, which allow examination of the expression of  >35000 genes, are available for P.  trichocarpa through NimbleGen Systems Inc. (USA), and Affymetrix microarrays for P. trichocarpa have been available since the fall of 2005. This study used NimbleGen oligonucleotide microarrays, which consisted of 24mer-70mer long nucleotide sequences, with a coverage of 6 to 20 probe sequences/gene, to determine the feasibility of microarray expression profiling to detect gender-specific gene-expression. In the five years since the microarray experiments were conducted in this study there have been many advancements in the field of collecting genome wide geneexpression and transcriptome data.  These include hybridization techniques such as the  microarray experiments outlined above, as well as sequence-based approaches, which I will describe briefly here. Initially, Sanger sequencing of cDNA or EST libraries were used to identify genes that were being expressed in given different tissues of P. trichocarpa (Sterky et al., 2004), and other organisms. But Sanger sequencing of cDNA libraries is slow, expensive, and not usually quantitative. Tag-based methods such as SAGE, CAGE and MPSS were developed as high-throughput processes to overcome some of the limitations of Sanger sequencing and produce precise information about gene-expression levels (Wang et al., 2009), but these methods also result in only portions of transcripts being analyzed and paralogues of genes are generally indistinguishable from each other. The development of high throughput, next-generation sequencing techniques such as those available from Illumina IG, Applied Biosystems SOLiD and Roche 454 Life  85  Science Systems, has allowed for the development of new methods for quantifying and mapping transcriptomes, which are the complete set of DNA transcripts contained in a given cell at a specific physiological or developmental stage (Wang et al., 2009). Currently this approach is being used to investigate SNP variation in P. trichocarpa to detect alternative splicing of genes and adaptive evolution in this species (Geraldes et al., 2011). 4.2  Objectives The goal of this project was to investigate gender-biased gene-expression, and to  identify genes that show an expression pattern in vegetative and floral tissues that segregates with gender. I hypothesize that the number of genes that are differentitally expressed between male and female reproductive structures in floral buds to be numerous due to the separate male (stamens) and female (ovules) organs present in the unisexual flowers. For this reason I focused on gene-expression in the leaf tissues to see if there are consistent gene-expression differences between male and female individuals in vegetative tissue. These leaf expressed genes may be the ones responsible for initiating genderrelated gene-expression, or they may be linked to such genes. The first objective was to look at genome wide gene-expression differences between male and female P. trichocarpa individuals, and see if any genes showed a statistically significant gene-expression difference between males and females.  The  second objective was to investigate any genes that showed gender-biased expression patterns, even if they were not statistically significant, with a larger sample size of female and male P. trichocarpa individuals using reverse-transcription (rt) PCR, to attempt to identify potential genomic sex-linked markers.  86  4.3 4.3.1  Methods Collection of and preparation of biological materials The plant tissue from floral buds and leaves of male and female P. trichocarpa  trees used for the first microarray experiment was collected for the lab in May of 2004 from natural stands located near Chilliwack, B.C. Plant tissue from leaf buds of male and female P. trichocarpa trees was also collected in March of 2006 from natural stands located on the UBC campus and these samples were used for the second microarray experiment (Figure 4.1). Collected tissues were stored at –80ºC until RNA was extracted from male and female leaf and floral buds following an RNA extraction protocol used for tree tissues (Kolosova et al., 2004), and samples were cleaned for use in cDNA preparation and microarray experiments with the QIAgen RNeasy MinElute Cleanup Kit (50) (Qiagen Inc., Toronto, Ont., Canada) or the Turbo DNA-free kit (Applied Biosystems/Ambion, Streetsville, Ont., Canada), and then stored at –80ºC to prevent degradation. 4.3.2  cDNA preparation and PCR conditions cDNA was prepared using the RevertAid" H minus First Strand cDNA Synthesis  Kit (Fermentas Life Sciences, CA, USA). PCR primers for this project were designed using Primer3 (v. 0.4.0, http://frodo.wi.mit.edu/primer3/), and tested in silico to determine the probability of primer dimers or unspecific primer-binding with Amplify 3X (v.3.1.4, http://engels.genetics.wisc.edu/amplify). Primers (Appendix A) were ordered from Integrated DNA Technologies Inc (Coralville, Iowa). DNA amplification for this project was performed as described in chapter two, section 2.3.2 of this thesis. Amplification products were visually scored for the presence or absence of bands on 1%  87  agarose gels stained with GelRed™ Nucleic Acid Gel Stain, 10,000X in water (Biotium, Hayward, CA 94545). Amplified PCR products were then prepared for sequencing by Macrogen, in Maryland USA.  Figure 4.1 Collecting leaf and floral buds from P. trichocarpa individuals. Trees were located on the University of British Columbia, Vancouver campus near the Triumf parking lot. Gregg Doughty, UBC arborist with plant operations, assisted in collecting. 4.3.3  Microarray gene-expression profiling experiment and statistical analysis of  the microarray data P. trichocarpa cDNA prepared from RNA extracted from male and female floral and leaf tissue was used to probe NimbleGen P. trichocarpa Affymetrix oligonucleotide microarray chips. The quality of the mRNA samples used in the microarray experiments was tested for degradation using the Agilent Bioanalyzer at the Centre for Molecular Medicine and Therapeutics (CMMT) facility, Vancouver, BC.  Two microarray  experiments were designed and these were sent to NimbleGen Systems Inc. (USA) to 88  perform the microarray hybridization experiments (Figure 4.2).  After receiving the  microarray expression data file from NimbleGen Systems, a simple Student’s tdetermined that the expression data from the floral tissues should be studied separately as the unisexual flowers contributed to almost 7000 genes being differentially expressed between the male and the female inflorescences, and these expression value differences in flowers reduced the signal from gender-biased gene-expression in leaf tissues samples to undetectable levels. The data for the leaf mRNA from the two microarrays were normalized together and the male and females leaf expression were compared to see if there was any difference in gene-expression patterns due to gender. When the leaf samples form both microarrays were added together this resulted in leaf mRNA samples from a total of five female and five male individuals (n = 5 for each gender). Normalization of the data from the arrays was necessary due to experimental variations in sample treatment, dye labeling, efficiency and detection. As a result, the fluorescence intensities of microarrays cannot be compared directly, but must be calibrated or “normalized” (Huber et al., 2002). In this study the microarray expression data was normalized using both Variance Stabilization Normalization (VSN) (Huber et al., 2002) and Quantile Normalization (Bolstad et al., 2003).  Two statistical methods were considered for analyzing the  microarray data once it had been normalized. A mixed effects model was fit to the data which adjusted for variation within (two probes per gene on the same array) and between arrays along with fixed effects for gender and experiment. Also, a LIMMA software package (www.bioconductor.org), designed to analyze experiments involving  89  Figure 4.2 Arrangement of P. trichocarpa mRNA samples on the NimbleGen Affymatrix oligonucleotide microarray chips. Numbers in brackets indicate the collection number of the individual tree the MRNA was prepared from.  90  comparisons of expression data from two colour microarrays, or log-intensity values from one channel Affymetrix arrays (Smyth, 2005) were used to analyze the normalized microarray data. The LIMMA analysis was conducted with an empirical Bayes modified error to compute estimates of differential expression for gender and experiment. After looking at the results from both normalization and stasistical analysis techniques, the Quantile Normalization, followed by the LIMMA analysis was chosen as this analysis identified the most likely candidate genes as having differentially expressed values between male and female individuals. 4.3.4  Investigation of gender-biased gene-expression patterns on chromosome 19  Chromosome 19 is thought to be involved in gender-determination in P. trichocarpa (Yin et al., 2008), and given that clustering of genes that are differentially expressed between the genders could occur around the sex locus due to recombination suppression spreading along a chromosome from an initial female or male sterility mutation (Nicolas et al., 2005), gender-biased gene-expression in male and female leaf tissues was investigated by looking at the microarray expression data for genes located on chromosome 19. The log of the ratio of male to female gene-expression values was plotted against the position of genes associated with microarray probe expression values along chromosome 19. Negative log expression values indicated a female-biased geneexpression, and positive log expression values indicated male-biased gene-expression. Genes showing either male- or female-biased expression values were identified and investigated in a larger sample of nine male and thirteen female individuals using reverse-transcription PCR. With the release of version 2.0 of the poplar genome, position and annotation data for the gene content of chromosome 19 was updated from version 1.1  91  to version 2.0 by performing BLAST searches of gene DNA sequences from version 1.1 against the version 2.0 genome sequence.  Reverse-transcription PCR primers were  designed based on version 2.0 of the poplar genome. 4.4 4.4.1  Results Results from the statistical analysis of the P. trichocarpa leaf tissue data  from both microarray experiments When the data from both microarray experiments was pooled the LIMMA statistical analysis of the data indicated that 1.17% of genes included on the microarrays were differentially expressed with respect to gender (Figure 4.3). This is a surprisingly small number as some 9.88% of genes are differentially expressed with respect to the different experiments (Figure 4.4). These figures are derived from the distribution of the parametric p-values, calculated by the LIMMA analysis. Given an experiment of this size, the biological noise clearly masks the effects (if any) of gender. There were two genes that showed a statistically significant difference in geneexpression between male and female P. trichocarpa individuals from the microarray experiments (Table 4.1). Reverse-transcription PCR was going to be used to verify if these two genes showed a gender-bias in their expression in a larger number of individuals. The resulting sequences obtained for these two genes were then going to be investigated to see if the sequences from difference individuals were exactly the same, or if they contained SNPs that differed between individuals or genders. By investigating SNP differentiation it may have been possible to determine why there is an expression difference between males and female using the sequence data collected.  92  Figure 4.3 Histogram of the parametric p-values plotting gene-expression in male leaf tissue (x axis) against the frequency of gene-expression in female leaf tissue (y-axis). The blue line indicates the threshold above which are the percentage of genes showing differential gene-expression between the genders, N = 5 for both genders. However, with the release of the second version of the poplar genome, the sequences of these two genes were BLASTed against the new genome and it was discovered that both genes have multiple hits. eugene3.00660277 BLASTs to two genes on scaffold 3 (POPTR_0003s03450 and POPTR_0003s03440), and to intergenetic regions on scaffold 43, 1184 and 19. fgenesh4_pg.C_scaffold_277000004 blasts to four genes on scaffold 17 (POPTR_0017s13140, POPTR_0017s13180, POPTR_0017s13220, 93  POPTR_0017s13260) (Appendix B). Given that these two genes did not appear to be single copy genes on version 2.0 of the genome, investigating their expression pattern in a greater number of male and female individuals would be problematic as it would be difficult to design primers to amplify specific copies of the gene, and it is unlikely that such a gene would be involved in gender-determination. Therefore work with these two genes was discontinued in favor of focusing research (reported in chapter three of this thesis) on genes located on chromosome 19, the chromosome indicated to contain the sex locus (Yin et al., 2008).  94  Figure 4.4 Histogram of the parametric p-values comparing gene-expression differences in leaf tissue between the two microarray experiments. The blue line indicates the threshold above which are the percentage of genes showing differential gene-expression between the two microarray experiments, N = 5 for both male and female mRNA samples. Table 4.1 Genes showing a statistically significant difference in the gene-expression between males and females in P. trichocarpa, on version 1.1 of the genome. Gene Name eugene3.00660277  Q-value 0.2460120  fgenesh4_pg.C_scaffold_277000004 0.2460120  Location on poplar genome, V. 1.0 Poptr1_1/scaffold_66:18973281898141 Poptr1_1/scaffold_277:17700-18283  95  4.4.2  Investigation of gender-biased gene-expression patterns on chromosome 19 Four genes that showed a gender-biased expression pattern in my microarray  results from working with version 1.1 of the poplar genome translated to genes that were also located on scaffold 19 on version 2.0 of the genome when a BLAST search was performed to update the information from version 1.1 to version 2.0 of the genome (Figure  4.5).  Two  of  the  genes,  POPTR_0019s03090  (SCA_19_9)  and  POPTR_0019s03100 (SCA_19_10), highlighted in orange (Figure 4.4), are right next to each other, and fairly close to the region of chromosome 19 where scaffold 117 was incorporated. These two genes show female biased expression, as well as a third gene, located at the other end of the chromosome, POPTR_0019s12360 (SCA_19_11), highlighted in red (Figure 4.4). POPTR_0019s13010 (SCA_19_7) was the only gene located on chromosome 19 that the statistical analysis of the microarray experiment indicated showed a nearly significant difference in expression values between males and females. It showed a malebiased expression pattern, and is highlighted in blue on Figure 4.5. The gene model from version 1.1 of the genome that showed the gender-biased expression blasted to two gene models on version 2.0 of the genome: POPTR_0019s13010 and POPTR_0019s13020. Primers were designed for both genes, but gene specific primers were only successfully designed for POPTR_0019s13010, so this gene was chosen to verify the microarray results in a larger sample of male and female leaf samples. Information on the position and putative gene functions for all the genes that were worked with is listed in Appendix B.  96  Figure 4.5 Comparison of the gene-expression values from the microarray data for chromosome 19, based on version 1.1 of the poplar genome, to the physical map chromosome 19 from version 2.0 of the poplar genome. On the y-axis negative values indicate female-biased gene-expression, and positive values indicate male-biased expression. The x-axis indicates the distance along chromosome 19 on both versions of the genome. The green box indicates the region containing the genes sampled on telomeric region of chromosome 19 during the investigation of SNP differences in the research outlined in chapter three of this thesis. Coloured boxes (orange, red and blue) indicate genes showing the most gender-biased expression that were located on chromosome 19, with arrows in corresponding colours indicating their approximate positions on version 2.0 of the chromosome. After designing specific primers for all four genes located in chromosome 19 that showed gender-biased expression pattern from the microarray experiments, reversetranscription PCR was performed to confirm this expression pattern in a larger sample of nine male and thirteen female P. trichocarpa. All successful PCR products obtained 97  were sent for sequencing to confirm that expression data for the target genes had been obtained. For  genes  POPTR_0019s03090  (SCA_19_9)  and  POPTR_0019s03100  (SCA_19_10) PCR amplification of bands was successful in a few individuals, both males and females for both genes, and it was confirmed from the good quality sequence obtained from the samples that the target sequence for each gene was being amplified. So it appears that both these genes are expressed in leaf tissue, as indicated by the microarray data, but with no discernable gender-bias. For the gene POPTR_0019s13010 (SCA_19_7) amplified bands were obtained, but it appears that the primers were not specific enough as the sequence obtained was not good quality, and didn’t match up well with the reference sequence from version 2.0 of the poplar genome. Also, when the best quality sequence produced for one female sample was BLASTed against version 2.0 of the poplar genome, the best hit was to gene POPTR_00190309, not the target sequence. Amplification  products  were  not  obtained  using  the  primers  designed  for  POPTR_0019s12360 (SCA_19_11). Redesigning primers for SCA_19_7 and SCA_19_11, and testing the primers under various PCR conditions also failed to yield better PCR results for these two genes. The cDNA used in these experiments was tested with a constituently expressed gene, POPTR_0006s19870, an elongation factor the Cronk laboratory uses as a positive control for PCR work with P. trichocarpa, and consistent amplification was observed in all samples with this gene, so I am confident that the lack of amplification for the target genes was not due to poor cDNA quality, but due to either ineffective primer binding or that the target gene was not expressed in the tissue sampled.  98  4.5  Discussion  4.5.1  Sample size on statistical power of microarray experiments Initially, when flower tissues were included in the microarray analysis a strong  gender-bias in gene-expression was detected between males and females, that corresponded to almost 7000 genes showing a positive gender-bias in their expression pattern.  This result was expected because given that flowers are unisexual in the  dioecious P. trichocarpa (Cronk, 2005), and the organs specific to male flowers, and therefore the genes involved in their development, would be more highly expressed in the male flowers than the females flowers, and vice versa. Of more interest from the point of view of developing a genetic sex-linked marker and investigating how gender is controlled genetically in this species would be genes that showed a gender-biased geneexpression pattern in vegetative tissue such as leaves, because ostensibly both males and females would have to express genes involved in determining or regulating gender, though they may be expressed at different levels in males or females. Therefore this investigation was focused on leaf tissue, and the leaf mRNA samples from the two microarray experiments were normalized together to increase the statistical power of this experiment. When the LIMMA statistical analysis was performed on the expression data for leaf tissue from the two microarrays, it was to shown that 1.17% of the genes included on the microarrays were differentially expressed with respect to gender (Figure 4.3). Given that the microarray experiment conducted consisted of only two arrays, with 5 female and 5 male P. trichocarpa leaf samples, it appears that biological variation masked any differential expression between females and males.  If there is gender-biased gene-  99  expression, the effect is very small and a much larger sample size would be necessary to detect this effect. By investigating genes that showed even non-statistically significant gender-biased expression patterns in a larger sample size (13 females and 9 males), if there was differential gene-expression between the genders it should have been detectible. Genes located on chromosome 19 were focused on (Figure 4.5), as research indicates that this chromosome may contain the sex locus in the genus Populus (Pakull et al., 2011). Unfortunately it proved difficult to amplify the four genes indicated as having a gender-bias by the microarray experiment analysis. While this research was able to confirm  that  two  of  the  genes  (POPTR_0019s3090/SCA_19_9  and  POPTR_0019s3100/SCA_19_10) were expressed in leaf tissue in a larger sample size, these results did not indicate a gender-bias in that expression pattern. When the DNA sequence obtained for these genes was investigated there were no gender-biased SNPs present. Gender effects may be present but a larger experiment will be needed to detect this with any statistical confidence, as the gender effects are clearly subtle in vegetative tissues in P. trichocarpa. 4.5.2  Effect of biological variation on interpreting microarray results One of the reasons I was unable to detect gene-expression differences between the  genders was due to the high amounts of biological variation. There is a ~10% difference in gene-expression levels between the two microarray experiments conducted, and the LIMMA statistical analysis indicates that these are real differences, not due to chance (Figure 4.4).  The difference in gene-expression levels between the two microarray  experiments may have been due to a number of factors. The P. trichocarpa individuals  100  that were sampled in the two experiments came from two different populations, located approximately 100km apart. The male and female individuals sampled for the first microarray experiment came from groves of P. trichocarpa growing in Chilliwack, B.C., and the individuals from both genders sampled for the second microarray experiment were located on the University of British Columbia endowment lands, in Vancouver B.C. While species in the genus Populus are all wind pollinated and one would expect a large effective population size (Hamrick et al., 1992), studies of SNP variation in P. balsamifera (Keller et al., 2010), and P. trichocarpa (Gilchrist et al., 2006), have shown that populations within species in this genus are adapted to local environmental conditions. This ability to adapt could result in variations in gene-expression levels between different stands of trees, even those within 100km of each other. Population substructure arises in P. trichocarpa because of dispersal patterns of seed and pollen as well. In fact, using microsatellite markers it has been shown that population substructure can be detected even when sampling populations of P. trichocarpa separated by a few hundred meters (Slavov et al., 2010). Another reason why there may have been a lot of biological variation between the two microarray experiments is that the samples collected from the different populations were collected at different times of year, and in different years. While the samples collected from the Chilliwack individuals were collected from young leaves in May of 2004, the samples UBC individuals were collected in March of 2006 from leaf buds just beginning to open. This means that the samples used for the microarray experiments were at different developmental stages and therefore gene-expression profiles could be  101  different in the leaf tissue used in the two experiments. Also, clonal field trials in Populus species have shown that the timing of spring bud flush is a highly heritable trait, with up to 98% of the variability observed in this trait being under genetic control, with environment influencing only 2% of the variability observed (Frewen et al., 2000). It appears from the results presented in this study that sampling from two different stands of trees, at different times of the year, and in different years, introduced an amount of biological variation into this experiment that was too large to effectively measure any gene-expression differences between leaf tissue in male and female individuals. With the amount of variation already introduced by using microarrays to look at gene-expression (variations in dye efficiency and detection, or fluorescence intensities, to name a few), it is important to make sure that samples are closely related biologically and environmentally so as to ensure that the gene-expression levels does not vary between samples greatly (Huber et al., 2002). This is much more possible to do when working with model species like Arabidopsis thaliana, where inbred lines can be quickly produced and maintained to reduce the amount of biological variation present in the sample individuals prior to conducting a microarray experiment. One such experiment very effectively compared varying amounts of gene-expression variation across seven Arabidopsis genotypes by making reciprocal crosses among the inbred lines (Vuylsteke et al., 2005). Unfortunately, controlling for biological variation in this manner is more difficult in P. trichocarpa due to its perennial life history, and the fact that sexing individuals is not possible until the individuals flower at between five and ten years of age (Brunner, 2010).  102  4.5.3  Interpreting gender-biased gene-expression using genome wide survey  approaches Unfortunately, with the release of version 2.0 of the poplar genome, it appears that some of the genes of interest were annotated differently between the two versions, making it problematic to investigate gene function or expression patterns of the genes indicated by the microarray experiment to have had a gender-biased expression in leaf tissues. This is a common problem when working with model organisms that have rapidly developing genomic resources, as gene structural and functional annotations can change as new data sets and more accurate information becomes available (van den Berg et al., 2010). Both version 1.1 and version 2.0 of the poplar genome used Fgenesh+ (http://www.softberry.com/berry.phtml) to predict gene models on the sequenced P. trichocarpa genome, but version 1.1 also used three other gene model prediction methods (Fgenesh, EuGene and GrailExp6) (Tuskan et al., 2006), whereas version 2.0 of the genome only used one other gene model predicting program, GenomeScan, if Fgenesh+ did not predict a model at a given locus (http://www.phytozome.net/poplar.php). Given the differences in how gene models were predicted between the two versions of the genome, it is not surprising that when I searched for the sequences of the two gene models indicated as having a statistically significant difference in geneexpression  between  males  and  females,  eugene3.00660277  and  fgenesh4_pg.C_scafford_277000004, on version 2.0 of the genome, these two gene models were now incorporated into different positions, and also had hits to multiple new gene models, making it challenging to further investigate their expression pattern in leaf tissue, gender-biased or otherwise. Genes involved in gender-differentiation are likely to  103  be located at the sex locus, and therefore genes with paralogues in multiple places in the genome are unlikely to be involved in maintaining separate genders. This is because current theories on the evolution of sex chromosomes seem to indicate that the evolution of a sex locus begins with a male or female sterile mutation around which genetic recombination becomes suppressed (Ming and Moore, 2007), and it is unlikely that a mutation of this kind would occur in multiple copies of the same gene, spread out on many chromosomes of the genome. 4.5.4  Considerations when working with the developing model system of Populus  trichocarpa When this study into gender-determination in P. trichocarpa was begun, version 1.1 of the poplar genome had just been released (Tuskan et al., 2006), so the microarray experiments were performed using gene annotation based on the first genome assembly for P. trichocarpa.  While this first assembly provided a very important tool for  developing poplar as a model species for genetic research, once work began with the genomic data, it became apparent just how much data needed to be added to the sequenced genome before it became as comprehensive a source of information as the annotated genomes for other model species such as Arabidopsis thaliana and Drosophila melanogaster. Most of the functional genomic research done in P. trichocarpa, and other Populus species, has focused on genes involved in wood development, and plant antipathogen biochemical pathways, as these characteristics are important to the development of poplars as sources of bio-fuel and pulp and paper (Jansson and Douglas, 2007). With version 1.1 of the poplar genome approximately 63.5% of the gene models had at least  104  some functional data or structural domain annotation (Tuskan et al., 2006), based on either functional studies in P. trichocarpa, or sequence homology with genes of known functions in Arabidopsis thaliana, or other model plant genomes.  Because I was  investigating gene-expression based on any detectible gender-biased expression pattern, many of the genes that were of interest in this study were either on genetic scaffolds that were not incorporated into the 19 chromosomes of P. trichocarpa (Table 4.1), or had no known function. This is not surprising, as though P. trichocarpa has great potential as a model species for plant genome evolution, genetic variation in response to ecological changes, and gender-determination studies (Cronk, 2005), until recently functional studies in P. trichocarpa have been few. The choice to work with the NimbleGen P. trichocarpa Affymetrix oligonucleotide microarray chips was based on these being a good option that was available to work on at the time, given that the resources to construct custom microarray chips were not available “in house”.  However, one draw back to working with  commercial microarrays is that they are often annotated prior to being made available for public use, so the structural and functional annotations for the genes they contain may not be the most up to date (van den Berg et al., 2010). The release of version 2.0 of the poplar genome in 2009 made it necessary for the functional and structural annotations of the genes of interest in this study to be updated, as it was important to work with the most current information available on the P. trichocarpa genome in order to be able to draw any valid conclusions from this study. This is a common occurrence for researchers working with newly available genome sequences as updates to the data set happen at a rapid pace and it is a challenge to  105  correctly interpret an experiment-based data set (such as the one reported here) with developing gene functional annotations (van den Berg et al., 2010). The rapid pace of development of genetic tools for genome sequencing frequently means that interpreting experimental results is challenging, and requires a lot of data management to ensure that the most current functional and structural gene data is being referenced. Gene re-annotation is becoming an important step in analyzing functional genomic data to improve its quality and quantity (van den Berg et al., 2010). With the availability and cost effectiveness of high throughput, next generation sequencing technologies, the ability to update gene annotations ensures that researchers are deriving conclusions about gene function based on the most up-to-date information, and are developing computational technologies to greatly streamline this process. 4.6  Conclusions I found that sample size is critical to conducting this kind of genome wide gene-  expression experiment as the results reported here showed that there was a greater geneexpression difference between the two microarray experiments performed than between genders.  Gender-biased gene-expression may be present in vegetative tissues of P.  trichocarpa, but these effects are subtle, and therefore a much larger sample size would be needed for a microarray experiment to be able to detect it with any statistical confidence. I was able to identify differential gene-expression patterns between male and female leaf tissues with the microarray experiments, however the investigation of the genes indicated by the microarray data to have differential expression between males and females using reverse-transcription PCR was unable to confirm the microarray results.  106  As a result of this I was unable to identify potential genomic sex-linked markers in P. trichocarpa. I detected a large amount of biological variation with the two microarray experiments. This result indicates that there is quite a lot of genetic variation even between two populations that are less than 100km apart. Although the samples used in the microarray experiments were all from young leaves, they were collected in different years, and it was possible to detect differences in gene-expression due to the slight differences in the developmental stage at which the samples are taken. Work with microsatellite markers has shown that population substructure can be detected in populations of P. trichocarpa even when sampling from stands of trees within a few hundred meters of each other (Slavov et al., 2010), and this work seems to support this as well. Another conclusion I can draw from this study is that working with a rapidly developing model system such as P.trichocarpa requires that one revisit data and update it frequently to reflect the latest and most accurate gene annotations available. The release of version 2.0 of the poplar genome made much more information available on the genes I was working with, which improved my project greatly, as I was able to more accurately place genes showing gender-biased expression on chromosome 19, the putative sex chromosome in P. trichocarpa. The advances in cost effective, high throughput next generation sequencing technologies that are currently available from Illumina IG, Applied Biosystems SoLiD and Roche 454 Life Sciences Systems (Wang et al., 2009) will make it much easier to generate and manage larger genetic data sets, and to keep the gene annotations up to date.  107  Data sets of the kind would make it possible to detect with greater accuracy the subtle gender-biased gene-expression differences in vegetative tissues in P. trichocarpa that my experiments may indicated exist.  108  Chapter 5: MADS-box genes in the genus Populus and their role in floral development 5.1  Introduction One of the major discoveries in the field of developmental evolution was that  certain gene families control the development of whole organs (Meagher, 2007). In the case of flowers, the MADS-box gene family has been found to be involved in the designation of floral organ identity and development. MADS-box genes that have been identified as having a role in plant organ development encode proteins that have a typical four domain MIKC structure (Alvarez-Buylla et al., 2000).  The gene family derives its  name from the highly conserved 55 amino acid DNA binding domain called the MADS domain after the genes that it was first discovered in MINICHROMOSOME MAINTENANCE 1 (MCM1), AGAMOUS (AG), DEFICIENS (DEF), and SERUM RESPONSE FACTOR (SRF) (Gramzow and Theißen, 2010). The second most conserved protein domain in this gene family is the keratin-like (K) domain, which is generally 70 amino acids in length with regularly spaced hydrophobic amino acids so it takes on a coiled-coil structure (Alvarez-Buylla et al., 2000). The Intervening (I) domain links the MADS and K domains, and along with the K domain is critical for mediating protein dimerization and specificity of interactions between MIKC-type proteins (Stellari et al., 2004). The fourth protein domain that is part of the MIKC structure is the poorly conserved carboxyl-terminal or C domain which functions as a trans-activation domain (Alvarez-Buylla et al., 2000).  109  5.1.1  Evolution of MADS-box genes in plants Given that MADS box genes play a central role in floral development they can  provide insight into how flowers have evolved (Meagher, 2007). From phylogenetic analyses it appears that an ancestral MADS-box protein domain duplicated in the common ancestor of the main eukaryotic kingdoms approximately one billion years ago, giving rise to the Type I and Type II MADS-box gene lineages found in animals, fungi and plants today (Alvarez-Buylla et al., 2000). MADS-box genes in plants that have characterized functions are predominantly Type II, and include a K domain, indicating that this plant-specific domain evolved early in the lineage (Alvarez-Buylla et al., 2000). One MADS-box gene has been found in an extant green alga (Gramzow and Theißen, 2010), and research in the moss Physcomitrella patens (Henschel et al., 2002) and the fern Ceratopteris sp. (Münster et al., 1997) indicate that this gene family is present in all land plant groups, and that the MIKC protein domain structure evolved and has been diversifying in function for over 650 million years (Nam et al., 2003). 5.1.2  MADS-box genes and their role in floral development The MADS-box gene family has generally been studied in hermaphroditic flower  development, by creating homeotic mutants that produce floral organs in inappropriate whorls of the flowers (Lebel-Hardenack and Grant, 1997). Analysis of floral mutants in Antirrhinum and Arabidopsis led to the formation of the ABC model for floral organ development (Coen and Meyerowitz, 1991; Zik and Irish, 2003). This model defines three regions of the floral meristem, each of which is controlled by a class of genes – A, B, or C (Zik and Irish, 2003). Region A consists of the first and second whorls of the floral meristem, region B comprises the second and third whorls and region C contains  110  the third and fourth whorls (Coen and Meyerowitz, 1991). Expression of A-class genes such as APETALA1 (AP1) and APETALA2 (AP2) alone specify sepal identity, A + Bclass genes specify petal development in the second whorl, B + C-class gene expression results in stamen formation in the third whorl and C-class genes alone specify carpel development in the fourth whorl (Zik and Irish, 2003). B-class genes include APETALA3 (AP3) and PISTILLATA (PI) in Arabidopsis, and the primary C-class gene is AGAMOUS (AG) (Kramer et al., 2004). Further research has indicated that two other classes of genes are also involved in the floral development pathway. D-class genes such as floral binding protein 71 (FBP71) have been found to be crucial for the determination of ovule identity in Petunia (Colombo et al., 1995), and Eclass genes, known as SEPALLATA1, 2, and 3 (SEP/1/2/3) which provide further levels of developmental control, are also required for specifying stamen, petal and carpel identity (Pelaz et al., 2000). Further research has indicated that D-function genes are not important in floral development outside of Petunia, so the current model for floral development that is generally applicable is the ABCE model (Theißen, 2001). Research done in Arabidopsis thaliana has been instrumental in identifying the role of MADS-box genes in floral development, and the DNA sequences from MADSbox genes in this model system have been used to identify genes with homologous functions and expression patterns in other species of flowering plants (Meagher, 2007), including Populus species. 5.1.3  MADS-box gene research in the genus Populus. The molecular genetics of floral development in Populus has been studied in a  few species, namely P. deltoides (Zhang et al., 2009), P. tremuloides and P. tremula  111  (Cseke et al., 2005), and P. trichocarpa (Brunner and Nilsson, 2004b). Populus spp. flowers are unisexual in all but a few cases, and consist of a very reduced perianth cup (Fisher, 1928), that appears as a single ridge encircling the male or female floral organs in some species like P. deltoides, or as a true lobate perianth in other Populus species (Kaul, 1995). Trees in the genus Populus have a life span of 100 to 200 years, and a long juvenile period, lasting seven to ten years, before the trees mature and begin flowering (Hsu et al., 2006). The transition from this juvenile phase to reproductive competency appears to have a genetic component given the strong developmental pattern of reproductive shoot formation (Yuceer et al., 2003).  Studies in P. trichocarpa, P.  deltoides and a P. tremula x P. tremuloides hybrid have indicated that the cycles of reproductive and vegetative growth in these tree species is coordinated by two genes, FLOWERING LOCUS T1 (FT1) and FLOWERING LOCUS T2 (FT2), which are homologous to the FT gene in Arabidopsis (Hsu et al., 2011). Over expression of both these genes seems to be associated with rapid shoot to flower conversion in P. deltoides (Hsu et al., 2006), and in P. tremula x P. tremuloides (Böhlenius et al., 2006). When it comes to floral development, research in A. thaliana has shown that the B-class gene AP1 is a direct target of FT, and AP3 is down-stream from both FT and AP1 in the development of petal and stamen identity (Hsu et al., 2006). Just as two homologs for the Arabidopsis gene FT have been found in Populus species. Two homologs for the Arabidopsis floral development genes AP1, PI, AP3, AG and SEEDSTICK (STK) have been identified in P. trichocarpa (Leseberg et al., 2006). The Populus AP1 homologs PTAP1-1 and PTAP1-2 are both expressed in the initial stages of developing floral  112  meristems (Brunner, 2010), but to date the gene-expression patterns of the P. trichocarpa PI homologs have not been investigated in detail. The P. trichocarpa MADS-box gene PTD shows homology with AP3, and shows an expression pattern similar to that found for AP3 and DEF, with no detectible expression in vegetative tissue, but spatial and temporal sex-specific expression in the inner whorl of male and female floral meristems (Sheppard et al., 2000). The two AG homologs found in P. trichocarpa, PTAG1 and PTAG2, show an AG-like floral tissue expression pattern, and phylogenetic analysis strongly supports their evolutionary orthology to C-class MADS-box genes (Brunner et al., 2000). Finally, homologs of E-class genes, which are involved in the developmental control of the inner whorls of flowers, have been shown to have similar expression to the A. thaliana gene SEPALLATA (SEP), in P. tremuloides and P. deltoides.  In P.  tremuloides, PTM3/4 and PTM6 genes are expressed at all stages of male and female flower development (Cseke et al., 2005), and PdMADS2 is expressed in the perianth cup of male inflorescences, and in the ovaries of female inflorescences in P. deltoides (Zhang et al., 2009). 5.2  Objectives I expected to find gender-biased expression patterns in genes involved in floral  development in P. trichocarpa, as flowers in this species are unisexual. Therefore, it was essential to collect the functional information available for genes involved in floral development in the Populus genus, and compare it to the expression data I generated from the microarray experiment (chapter four). I was particularly interested in B and Cclass MADS-box genes as these genes play a role in androecium and gynoecium  113  development (Meagher, 2007), and therefore in dioecious species such as P. trichocarpa these genes are good candidates for the separation of male and female functions. The first objective for this project was to create a comprehensive list of all Populus MADS-box genes that included the current annotation data from version 2.0 of the poplar genome, and the functional characterization data from expression studies. Secondly, I wanted to identify which of the poplar MADS-box genes would show gender-biased expression using male and female floral tissue on my microarray experiment. Thirdly, I wanted to relate my data on gender-biased expression pattern of floral MADS-box genes in poplar to the known function and expression patterns of homologous A. thaliana floral MADS-box genes. The fourth objective was to confirm the gene-expression bias observed in the microarray experiment in a larger sample size of male and female flower tissue using reverse-transcription PCR (rtPCR), and to add to the expression data available for MADS box genes in P. trichocarpa by focusing my work on genes that had little or no expression data available for them. 5.3 5.3.1  Methods Collection of and preparation of biological materials The plant tissue from floral buds and leaves of male and female P. trichocarpa  trees used for the first microarray experiment was collected in May of 2004 from natural stands located near Chilliwack, B.C. Additional flower buds were collected from three male and four female P. trichocarpa trees in March of 2006 from natural stands located on the UBC campus previously identified as males or females by myself and Dr. Cronk for cDNA preparation. These tissues were stored at –80ºC for future use. RNA was extracted from male and female floral buds following either an RNA extraction protocol  114  developed for tree tissues (Kolosova et al., 2004), or a modified protocol using Plant RNA Purification Reagent from Invitrogen (Catalog # 12322-012).  Samples were  cleaned for use in cDNA preparation and microarray experiments with the QIAgen RNeasy MinElute Cleanup Kit (50) (Qiagen Inc., Toronto, Ont., Canada) or the Turbo DNA-free kit (Applied Biosystems/Ambion, Streetsville, Ont., Canada), and then stored at –80ºC to prevent degradation. 5.3.2  cDNA preparation and PCR conditions cDNA preparation was done using the RevertAid" H minus First Strand cDNA  Synthesis Kit (Fermentas Life Sciences, CA, USA). PCR primers for this project were designed using Primer3 (v. 0.4.0, http://frodo.wi.mit.edu/primer3/), and tested in silico to determine the probability of primer dimers or unspecific primer binding with Amplify 3X (v.3.1.4, http://engels.genetics.wisc.edu/amplify).  Primers used in this project were  ordered from Integrated DNA Technologies Inc (Coralville, Iowa) (Appendix A). DNA amplification for this project was performed as described in chapter two, section 2.3.2 of this thesis, except that for this project 50ng of DNA was used/reaction. Amplification products were visually scored for the presence or absence of bands on 1% agarose gels stained with GelRed™ Nucleic Acid Gel Stain, 10,000X in water (Biotium, Hayward, CA 94545).  Amplified PCR products were sent for sequencing by Macrogen, in  Maryland USA. 5.3.3  Microarray gene-expression profiling experiment In the fall of 2005 P. trichocarpa cDNA prepared from RNA extracted from male  and female floral and leaf tissue was used to probe NimbleGen P. trichocarpa Affymetrix oligonucleotide microarray chips. The extracted mRNA samples were verified to be of  115  suitable quality for working with microarrays by testing them for degradation using the Agilent Bioanalyzer at the CMMT facility, Vancouver, BC. Two male and two female leaf mRNA samples were sent, as well as one male and one female flower sample (6 samples total) to NimbleGen Systems Inc. (USA) to perform the microarray hybridization. From the data file the floral tissue hybridization data was separated and it was observed that many genes showed a gender-bias, particularly in the male-biased direction, most probably because these genes are involved in pollen production. Genderbiased expression was calculated by taking the log intensity value of the female floral expression for a given gene, and subtracting it from the log intensity value of the male floral expression for the same gene. To narrow the study to genes likely to be directly involved in floral development, the top 15 genes were identified that 1) were annotated as having MADS-box functions and that 2) showed the greatest differences in gender-biased expression values between the male and female flower tissue samples, with log intensity values above a threshold of 0.05 for male inflorescences, and below -0.05 for female inflorescences. 5.3.4  Updating information on MADS-box genes in P. trichocarpa from version  1.1 to version 2.0 of the poplar genome The first comprehensive study of MADS-box genes in P. trichocarpa was that of Leseberg et al. (2006), who used an in silico approach to identify 105 putatively functional MADS-box genes in version 1.1 of the poplar genome. With the release of version 2.0 of the poplar genome, in order to correctly identify the functional annotations for the MADS-box genes that showed gender-biased expression patterns in the data, the annotation of the MADS-box gene family presented by Leseberg et al. (2006) needed to  116  be updated to version 2.0 of the poplar genome. To create a dataset of all the putative MADS-box genes on version 2.0 of the poplar genome that corresponded to the PtMADS-box gene model sequences identified by Leseberg et al. (2006), the version 1.1 MADS-box gene sequences were BLASTed against version 2.0 of the poplar genome. 5.3.5  Phylogenetic analysis of MADS-box genes in P. trichocarpa In order to correctly identify sequence homology between the genes showing a  gender-biased expression pattern in my experiments and MADS-box genes with characterized functions in A. thaliana and other Populus species, I performed a phylogenetic analysis by aligning the M, I, and K protein domains from the A. thaliana genes PI, AP1, AP3, AG, SHP1, SHP2 and SEP 1/2/3/4 with the genes that showed novel expression data from the microarray and cDNA experiments (M1, M2, M4, M6, F2, SCA19_12, MADS_1 and MADS_2). The MIK protein domains from all of the Populus MADS-box genes with characterized function (PTAG1, PTAG2, PTAP1-1, PTAP1-2, PTD, PTM/3/4/5 and PtMADS31) were also included, as well as all the PtMADS genes from Brunner and Nilsson (2004) for a total of 35 genes. The Populus trichocarpa MIK protein  sequences  were  taken  from  version  2.0  of  the  poplar  genome  (http://www.phytozome.net), and the A. thaliana MADS-box genes were from the TAIR data base (http://www.arabidopsis.org/). Protein alignments were initially compiled using the T-COFFEE webserver (available at http://www.tcoffee.org/) (Notredame et al., 2000), and then manually edited using BioEdit version 7.0.5.3, (10/28/05) (Hall, 1999). The alignment included the MADS-box domain of about 60 amino acids, I domain, and the K domain, for a total protein sequence of between 163 and 187 amino acids (Appendix E). The C domain was  117  excluded from the alignment, as it did not align well across the majority of the MADSbox protein sequences studied. Only MADS-box genes from version 2.0 of the poplar genome that corresponded to a single MADS-box gene annotated on version 1.1 of the poplar genome were included in the alignment. The phylogenetic programs RAxML 7.2.8 (Stamatakis, 2006) with the LG +G model for amino acid substitution, and GARLI 1.0 (Zwickl, 2006), with the WAG amino acid substitution model were used to construct the phylogenetic trees. These amino acid substitution models were the ones that had the lowest Akaike Information Criterion (AIC) score for each program, respectively. These two programs produced similar unrooted phylogenetic trees and bootstrap values, but the RAxML analysis yielded a phylogenetic tree with shorter branch lengths, so this tree was used to represent the MIK protein sequence homology in this study. 5.4 5.4.1  Results Updating available MADS-box gene research from version 1.1 to version 2.0  of the poplar genomes To identify the most current MADS-box gene annotations and identify which of these genes have been characterized with expression studies, I updated the entire list of P. trichocarpa MADS-box genes (PtMADS genes) identified by Leseberg et al. (2006) to the most current annotation data available for version 2.0 of the poplar genome. Because of the discrepancy between the way in which the two versions of the genome were assembled, a number of genes were identified that had changed position in the genome. Scaffolds that were previously not included in the sequence of chromosomes had been added. Several genes that were single genes on version 1.1 but BLASTed to two separate gene models on version 2.0 were observed (indicated in gray on Figure 5.1). In addition  118  some genes that were two or more gene models from version 1.1 of the poplar genome were annotated as single gene models on version 2.0 (indicated in yellow on Figure 5.1). Some of the genes previously annotated as MADS-box genes on version 1.1 of the poplar genome now have either no known annotated function, had a best BLAST hit in an intergenic region, or in the case of PtMADS90, did not have a single hit on version 2.0 of the genome. Finally, it had been previously reported that there were no floral MADS-box genes located on chromosome 19 (Leseberg et al., 2006), and I was able to identify two PtMADS-box genes located on that chromosome. A number of floral MADS-box genes have been studied to confirm their functions and expression patterns in the genus Populus, and I was able to ascertain that all of these corresponded to PtMADS-box genes identified by Leseberg et al. (2006) either by direct comparison with the sequences of characterized P. trichocarpa, or by identifying the homologous P. trichocarpa gene if the MADS-box gene had been studied in another Populus species. Of the twelve MADS-box genes that had been previously characterized by expression studies in Populus, all twelve were located on the second version of the poplar genome (Figure 5.1).  After updating the list of putative MADS-box genes  reported in Leseberg et al. (2006) to version 2.0 of the poplar genome the total number of poplar MADS-box genes indicated by the updated data set is now 88, down from 105. The methods used may not have identified all the MADS-box genes in the poplar genome, and given the dynamic nature of genome annotation and the complexity of the poplar genome, it is conceivable that additional MADS-box genes could be detected in Populus. However, my data can provide a jumping off point for further studies of this gene family in Populus. 119  Figure 5.1 Distribution of 88 MADS-box genes on P.trichocarpa chromosomes. The locations of the genes are approximate and not directly proportional to their real physical distances. Genes indicated in black have no expression studies associated with them to date. Genes in gray BLASTed to multiple genes on version 2.0 of the genome. Genes in yellow indicate two or more gene models from version 1.1 of the poplar genome that are now annotated as single gene models on version 2.0. Genes in green have been investigated in other expression studies, but showed no gender-bias expression on the microarray. Genes in light pink show a female-biased expression pattern on the microarray, and genes in blue show a male-biased expression pattern. One gene (purple) was equally expressed in male and female floral tissue. The two genes indicated in orange show male-biased expression, while the one gene indicated in dark pink on chromosome 19 showed female-biased expression from the microarray and cDNA data. Further genomic information on the genes indicated in this figure can be found in Appendix D.  120  5.4.2  Identifying P. trichocarpa MADS-box genes showing gender-biased  expression using a microarray approach The initial microarray experiment indicated that certain MADS-box genes showed a gender-bias in their expression in floral tissue.  This result is not unexpected,  considering the lack of female organs in male flowers, and vice versa. When male and female floral tissue were hybridized on the microarray, many genes showed male-biased expression, probably because these genes are involved in pollen production. To narrow this study to genes likely to be directly involved in floral development, I identified the top 14 genes that showed gender-biased expression, and were also annotated as MADS-box genes in the microarray file. One of these 14 genes is no longer annotated as a MADSbox gene, but 13 remain identified as such. The sole gene that showed a male genderbiased expression pattern but is not a MADS-box gene was labeled as MALE (M)5 (Table 5.1). It is annotated as a kinesin motor family protein, similar to the AtNACK1 kinesin-like protein in A. thaliana on version 2.0 of the poplar genome. As the protein sequence for M5 did not align with the protein sequences for the MADS-box genes in this study, it appears that the current annotation for this gene is correct. Comparatively little work has been done to characterize MADS-box gene-expression patterns in P. trichocarpa, but by comparing these results to those available in the literature, I was able to identify which of the genes indicated by the microarray as having gender-biased expression corresponded to Populus MADS-box genes that had previously been characterized (Table 5.1). The genes that showed the most female-biased expression on the microarray were SCA19_12 and FEMALE (F) 2.  Neither of these genes had been previously 121  characterized with expression data, but both of which group with the A. thaliana STK gene (Figure 5.2).  The remaining four genes that showed female-biased gender  expression on the microarray are floral MADS-box genes that have been previously studied in Populus – PTAG1 and PTAG2 (Brunner et al., 2000), PTD (Sheppard et al., 2000), PTAP1-1 (Brunner and Nilsson, 2004b) and PTM2 (Cseke et al., 2005), with PTAP1-1 from P. trichocarpa homologous to PTM2 in P. tremuloides.  One gene,  characterized as PTAP1-2 in P. trichocarpa and PTM1 in P.tremuloides, was expressed in both male and female floral tissues on the microarray, indicating that this MADS-box gene plays a role in both male and female floral development. The two genes that showed the most male-biased expression were MADS_1 and MADS_2, and like SCA19_12 and F2, these genes have not been previously studied in detail in P.trichocarpa either, so I am reporting novel gender-biased expression for these four genes. The remaining four MADS-box genes showing male-biased expression, M1, M2, M4, M6, were identified as MADS-box genes in P. trichocarpa by Leseberg et al. (2006). However, no previous Populus expression data appears to be available for any of these six genes, so my research reports novel gender-biased expression for these genes. Five other MADS-box genes have been studied in Populus, PTM3, PTM4, PTM5, PTM6 (Cseke et al., 2005), and PtMADS31 (Zhang et al., 2009), but none of these E-class genes showed gender-biased expression in floral tissues in this study, and the expression data from the previous studies indicates that these five genes are expressed in both male and female floral tissues (Cseke et al., 2005; Zhang et al., 2009).  122  Table 5.1 Relating available information on the expression of MADS-box genes in Populus species to MADS-box genes showing gender-biased expression patterns. All data shown is from this study unless otherwise indicated by superscript. Characterized Corresponding Gender-bias Gender-bias from Position on version Species of Gene Gene Name Gene Model in this study other studies 2.0 poplar genome Characterization SCA19_12 POPTR_0019s10580 F N/A scaffold_19: 12132258-12133799 P. trichocarpa F2  .  POPTR_0013s09980  F  N/A  scaffold_13: 9955912-9966253  P. trichocarpa  POPTR_0011s03140  F  No bias  scaffold_11: 3200188-3207704  P. trichocarpa  PTAP1-1 [PTM2 ] POPTR_0008s09800  F  N/A  scaffold_8: 6076113-6078596  P. trichocarpa  N/A  scaffold_10: 15345650-15351172  P.trich/P. tremuloides  PTAG21 2 2  3  3  PTAP1-2 [PTM1 ]  POPTR_0010s16380  No bias  PTD4  POPTR_0007s13660  F  No bias  scaffold_7: 13686094-13687851  P. trichocarpa  PTAG11  POPTR_0004s06300  F  No bias  scaffold_4: 4997208-5005680  P. trichocarpa  M1  POPTR_0004s11420  M  N/A  scaffold_4: 10038841-10040605  P. trichocarpa  M2  POPTR_0012s05960  M  N/A  scaffold_12: 5936482-5942087  P. trichocarpa  M4  POPTR_0005s12000  M  N/A  scaffold_5: 8770246-8772014  P. trichocarpa  M5  POPTR_0002s02910  M  N/A  scaffold_2: 1789707-1795019  P. trichocarpa  M6  POPTR_0017s13410  M  N/A  scaffold_17: 13578276-13585496  P. trichocarpa  MADS_1  POPTR_0002s07920  M  N/A  scaffold_2: 5450756 – 5453302  P. trichocarpa  MADS_2  POPTR_0005s20480  M  N/A  scaffold_5: 19532615 – 19535235  P. trichocarpa  POPTR_0008s09790  N/A  No bias  scaffold_8: 6063858 – 6070604  P. deltoides  PTM3 and PTM4  POPTR_0004s11440  N/A  No bias  scaffold_4: 10100959 – 10108181  P. tremuloides  PTM56  POPTR_0014s07010  N/A  N/A  scaffold_4: 10100959 – 10108181  P. tremuloides  PTM63  POPTR_0001s13650  N/A  No bias  scaffold_1: 10729860 – 10732842  P. tremuloides  PtMADS315 3  1  3  .  (Brunner et al., 2000) 2(Brunner and Nilsson, 2004b) 3(Cseke et al., 2005) 4(Sheppard et al., 2000) 5(Zhang et al., 2009) 6(Cseke et al.,  2007) 123  5.4.3  Identification of putative gene function based on protein sequence homology  for P. trichocarpa MADS-box genes The Arabidopsis thaliana genome only contains one copy of the majority of the floral MADS-box genes that are involved in floral development, while studies in Populus species have shown many MADS-box genes in this species have two homologs to every one A. thaliana MADS-box gene. An example of this is seen with the discovery of PTAG1 and PTAG2 in P. trichocarpa, which are homologous to AG in A. thaliana (Brunner et al., 2000), which is likely due to the fact that there has been a whole genome duplication event since the divergence of Populus and Arabidopsis (Tuskan et al., 2006). It is similarly logical to assume that the Populus MADS-box gene homologues have also probably differentiated in function following their origins in genome duplication events in evolutionary history of the genus. With this in mind, I conducted a phylogenetic analysis that included the eleven of the MADS-box genes involved in floral development in A. thaliana, the MADS-box genes that showed gender-biased expression from the microarray experiment (13), and seven PtMADS-box genes from the phylogenetic analysis performed by Brunner and Nilsson (2004), which were included because these genes had been previously identified as being the P. trichocarpa homologs of A. thaliana MIKC group proteins known to be involved in floral development (Brunner and Nilsson, 2004b). From the phylogenetic analysis it appears that the female-biased PTD gene and the male-biased M4 group closest to the A. thaliana gene AP3, while MADS_1 and MADS_2 group with PI.  As expected, PTAG1 and PTAG2 group with AG, and  PtMADS31, an E-class gene first characterized in P. deltoides, groups with the SEP genes 124  from A. thaliana. SCA19_12 and F2 group most closely with STK. M1, M2 and M6 group with PTAP1-2 and PTAP1-1/PTM1 with the A. thaliana gene AP1. The overall trend in my data seems to be that most floral MADS-box genes in A. thaliana, have two P. trichocarpa homologs (Table 5.2). Also, it appears that the P. trichocarpa homologs for C-class genes such as AG and STK show a female bias in their expression pattern, whereas the putative PI P. trichocarpa homologs, which are B-class genes, show a malebiased expression pattern.  125  Table 5.2 Populus MADS-box floral genes that have been characterized with expression patterns in floral tissues and their A. thaliana homologs. Homologs were identified based on protein sequence homology of the MADS-box protein domain. All genes listed here showed gender-biased gene-expression in the microarray experiment, and some of these genes have been characterized in other studies, as indicated by superscript. Gene class refers to the ABC model of floral development. Characterized Gene Name  Corresponding Gene Model  A. thaliana MADS-box homolog  A/B/C/E-class gene .  SCA19_12  POPTR_0019s10580  SEEDSTICK (STK)  C-class  F2  POPTR_0013s09980  SEEDSTICK (STK)  C-class  PTAG11  POPTR_0004s06300  AGAMOUS (AG)  C-class  PTAG21  POPTR_0011s03140  AGAMOUS (AG)  C-class  PTAP1-12  POPTR_0008s09800  APETALA1 (AP1)  A-class  PTAP1-22  POPTR_0010s16380  APETALA1 (AP1)  A-class  4  PTD  POPTR_0007s13660  APETALA3 (AP3)  B-class  M1  POPTR_0004s11420  homology uncertain  N/A  M2  POPTR_0012s05960  homology uncertain  N/A  M4  POPTR_0005s12000  APETALA3 (AP3)  B-class  M6  POPTR_0017s13410  homology uncertain  N/A  MADS_1  POPTR_0002s07920  PISTILLATA (PI)  B-class  MADS_2  POPTR_0005s20480  PISTILLATA (PI)  B-class  1  .  (Brunner et al., 2000) 2(Brunner and Nilsson, 2004b) 3(Sheppard et al., 2000)  126  Figure 5.2 Un-rooted phylogenetic tree produced using a Maxmimum Likelihood (ML) analysis to show the homology in protein sequence data between the MADS-box domain from the poplar floral MADS-box genes and A.thaliana floral MADS-box genes. Colour shading indicates the class of floral MADS-box genes. Blue = A-class, purple = B-class, pink = C-class, and green = E-class. 127  5.4.4  Confirming the MADS-box gene microarray results using cDNA expression  experiments I confirmed the gender-biased expression pattern observed on the microarray for one gene showing female expression bias (SCA19_12) and two genes exhibiting male expression bias (MADS_1 and MADS_2) by using reverse-transcription PCR with a sample of floral tissue from four female individuals and three males. I chose to further investigate SCA19_12 because it showed the most female-biased expression value, and it is located on chromosome 19, which is the putative sex chromosome in P. trichocarpa. I investigated MADS_1 and MADS_2 because they showed the most male-biased expression values and appear to be the P. trichocarpa PI homologs, and PI homologs have not previously been investigated in P. trichocarpa. When testing the primers designed to amplify cDNA of SCA19_12, I discovered that this gene is expressed in female, but not male, floral tissues (Figure 5.3). Also, this gene does not appear to be expressed in the leaf tissue of male or female individuals (Figure 5.4). The bands resulting from the cDNA amplification of SCA19_12 were sequenced, and by comparing the PCR product sequence to the sequence for this gene that is annotated on the poplar genome, it was confirmed that these bands represent the target sequence. The two genes MADS_1 and MADS_2 showed amplification products in male, but not female, floral tissues (Figure 5.3), and like SCA19_12, these genes do not appear to be expressed in the leaf tissues of either gender. The bands observed when amplifying these two genes were confirmed to be the target sequence by sequencing the PCR products, and comparing the sequence to the poplar genome.  128  Figure 5.3 Electrophoresis gel photo showing differential expression patterns measured by PCR of cDNA from male and female P. trichocarpa flower tissues. Four female and three male individuals were tested. MADS_1 and MADS_2 are genes expressed in male but not female floral tissues. SCA19_12 is expressed is female floral tissues but not male floral tissues. The positive control for this experiment was EF, a constitutively expressed transcription factor. ! DNA of known concentration was used to give an estimate of cDNA concentration in the bands on the gel, and a 1kb ladder was used to estimate the fragment size in the bands.  129  Figure 5.4 Electrophoresis gel photo showing no detectible expression in male and female P. trichocarpa leaf tissue of the MADS box genes that had differential expression patterns in male and female floral tissues. Leaf tissue from thirteen female and nine male individuals was tested with MADS_1, MADS_2 and SCA19_12. The positive control for this experiment was EF, a constitutively expressed transcription factor. ! DNA of known concentration was used to give an estimate of cDNA concentration in the bands on the gel, and a 1kb ladder was used to estimate the fragment size in the bands.  130  5.5 5.5.1  Discussion MADS-box genes in the second version of the P. trichocarpa genome. The release of the second version of the P. trichocarpa genome in January 2010  provided an opportunity to conduct a survey of the current knowledge on MADS-box genes involved in floral development in Populus, and to provide a comprehensive overview of the functional and expression studies available for this gene family in P. trichocarpa. Using the genome wide analysis of the MADS-box gene family conducted by Leseberg et al. (2006) as a starting point, and including studies done on floral MADS-box genes from P. tremula, P. tremuloides, and P. deltoides as well as the expression data generated using a microarray approach in P. trichocarpa in this study, I created a body of MADS-box gene data that was based on version 2.0 of the poplar genome. To date, all of the published studies of MADSbox genes in Populus have relied on version 1.1 of the poplar genome. My work updates the knowledge of these genes to the latest gene position and annotation data available for version 2.0 of the poplar genome. One of the most interesting findings made during this process is that two floral P. trichocarpa MADS-box (PtMADS) genes were located on chromosome 19, when it had been previously reported that there were no floral MADS-box genes located on this chromosome (Leseberg et al., 2006).  This is particularly intriguing because chromosome 19 is thought to  be the emerging sex chromosome in P. trichocarpa and other Populus species that have been investigated (Paolucci et al., 2010). In Populus, depending on the species, the putative sex locus appears to map either to a telomeric position, or an internal position (Paolucci et al., 2010). In P. trichocarpa the sex locus is thought to occur in a telomeric position (Yin et al., 2008).  SCA19_12 and  131  PtMADS42 both exist in an internal position, so it is unlikely that they are directly involved in gender-determination at the putative sex locus. But if a multi-locus system controls gender-determination in Populus as has been proposed (Alström-Rapaport et al., 1998), it is possible that one or both of these genes could be involved in gender-determination. Alternatively, they could be targeted by gender-determining genes located at the site of the putative sex locus at the telomeric end of chromosome 19. 5.5.2  Gender development in poplar flowers The model species that have been used to study floral development include  Arabidopsis thaliana, Antirrhinum majus, and Petunia hybrida, all of which are hermaphroditic species that develop bisexual flowers (Zluvova et al., 2006). The genetics of floral development have also been studied in the dioecious species Silene latifolia and Rumex acetosa, which both develop unisexual flowers, but appear to abort the male organs in female flowers, and vice versa, early in floral development. In S. latifolia, floral bud development is classified into twelve stages, and prior to stage five of development, flowers of both genders develop as bisexual flower primordia. However, once male flowers reach stage six the gynoecium primordia stops differentiating, and in female flowers after stage seven the development of stamen primordia is arrested, so anthers do not develop (Grant et al., 1994) In Rumex acetosa four whorls of floral organs are initiated in both male and female flowers, but in each gender development of the non-gender appropriate set of organs (ie: stamens in female flowers and carpels in male flowers) is stopped soon after initiation (Ainsworth et al., 2005). In P. trichocarpa however, there is no evidence of the initiation or abortion of inappropriate sets of organs in either the male of female flowers, and the evidence of a reduced perianth suggests that the unisexual flowers in Populus are a result of evolution  132  favoring fewer floral whorls (Boes and Strauss, 1994). This evolution of fewer floral whorls is probably a result of the evolution of dioecy in Populus being via the monoecious pathway (chapter one). The monoecious pathway would allow these flowers the long evolutionary time required for the extensive diversification of gene function necessary for the development of unisexual flowers.  Therefore, studying the genes involved in floral  differentiation in P. trichocarpa may provide insight into how the ABC genetic model of floral development works in flowers that are strictly unisexual, and may indicate which floral MADS-box genes have differentiated to have only male or female expression. Given that B and C-class genes are respectively involved in the development of male and female floral organs, these classes of genes seem to be likely candidates to have a role in the gender asymmetry that is found in unisexual flowers (Meagher, 2007). 5.5.3  Comparing the MADS-box genes involved in floral development in A. thaliana  and P. trichocarpa In Populus the study of genes involved in floral development is still a developing field. Using a microarray approach to do a genome wide survey of floral tissue gene expression proved useful in identifying genes that are involved in floral development, and allowed a gender-bias in the expression of some of these genes to be observed. Comparing the MADS-box genes expressed in floral tissues in P. trichocarpa to floral MADS-box genes already well characterized in A. thaliana provides a valuable tool for inferring floral gene functions in P. trichocarpa.  This is possible because the P. trichocarpa genome has been  shown to contain orthologs of the major genes that regulate floral development in A. thaliana (Brunner, 2010). P. trichocarpa and A. thaliana both belong to the rosid clade of the Core  133  Figure 5.5 Comparing differences in MADS-box gene-expression in floral tissues between A. thaliana and P. trichocarpa. The lines on the right next to the labels A, B, C and D indicate the whorls in which these classes of genes are expressed. Eudicots, which allows for the comparison of gene functions between the two species (Brunner et al., 2004a). Based on the ABC model of MADS-box gene-expression in A. thaliana (Coen and Meyerowitz, 1991), and the gene-expression studies done in Populus by Brunner et al. (2000), Skinner et al., (2003), Cseke et al. (2005) and Zhang et al. (2010), as well as the data reported in this study, a comparison has been made here between MADS-box gene-expression patterns in the perfect flowers of A. thaliana, and the unisexual flowers found in P. trichocarpa (Figure 5.5). Comparing the floral MADS-box gene-expression  134  patterns found in P. trichocarpa to those that have been studied in A. thaliana provides information about how the ABC model has been altered to result in the development of unisexual flowers. 5.5.4  A-class gene-expression in P. trichocarpa inflorescences In the ABC genetic model of floral development, genes APETALA1 (AP1) and  APETALA2 (AP2) are A-class genes, that, when expressed alone specify sepal development, and when expressed with B-class genes form petals (Theißen, 2001). In A. thaliana, AP1 is an A-class gene that is initially expressed in the entire floral meristems, but later its expression is limited to specifying sepal and petal identity (Zik and Irish, 2003).  The  microarray data presented in this study indicates that PTAP1-1 has a female-biased expression pattern, which is interesting because if this gene is an A-class gene like AP1, it should be expressed in the first whorl of floral organs (Theißen, 2001), which corresponds to the sepals, or perianth cup in Populus flowers, which is found in both male and female flowers (Boes and Strauss, 1994). The PTAP1-2 gene was expressed in both male and female floral tissues according to the microarray data, indicating that this MADS-box gene plays a role in both male and female floral development. AP2, the second A-class gene, was not included in the phylogenetic analysis in this study because while it plays a role in floral meristem identification and is initially expressed throughout the floral meristems in A. thaliana (Zik and Irish, 2003), it is not a MADS-box gene but rather a ethylene responsive element binding protein (EREBP), a family of at least 12 proteins in A. thaliana that are characterized by a specific AP2 protein domain (Okamuro et al., 1997).  135  Three genes indicated as having male-biased expression from the microarray results, M1, M2 and M6, were identified as being MADS-box genes in P. trichocarpa by Leseberg et al. (2006), however, no previous Populus expression data appears to be available for these three genes, so this research introduces novel gender-biased gene-expression data for M1, M2 and M6. These three genes grouped most closely with PTAP1-1 and PTAP1-2 in the phylogenetic analysis (Figure 5.2). It is possible that PTAP1-1, PTAP1-2, M1, M2 and M6 have similar roles in floral development in P. trichocarpa, with PTAP1-1 and PTAP1-2 having specific female functions, and with M1, M2 and M6 having specific male functions. Further investigation of the putative A-class genes in P. trichocarpa is required to determine their function in floral development, and confirm the gender-biased expression that was observed in this study. 5.5.5  B-class gene-expression in P. trichocarpa inflorescences B-class function in the ABC model of floral development is contributed to by two  genes, APETALA3 (AP3) and PISTILLATA (PI), which together with A-class genes specify petal development, or when paired with C-class genes are involved in stamen development (Theißen, 2001). The two MADS-box genes that showed the highest male-biased expression pattern in the microarray data were MADS_1 and MADS_2. When the sequence of the MADS-box protein domain for these two genes was aligned with MADS-box genes from P. trichocarpa and A. thaliana, both of these genes were positioned most closely in the tree with the A. thaliana gene PI (Figure 5.2). cDNA expression patterns confirmed that MADS_1 and MADS_2 are expressed in male, but not in female floral tissues, and that there was no detectible expression of these two genes in leaf tissues (Figures 5.3 and 5.4). In A. thaliana, PI is necessary for the specification of organ identity in the second and third whorls of  136  developing perfect flowers, and is therefore involved in designating petal and stamen identity (Goto and Meyerowitz, 1994). As flowers in P. trichocarpa do not have petals, it is most likely that these two genes are involved in stamen development, which would also explain why they have a male-biased expression pattern in floral tissue. In A. thaliana, AP3 plays a role in determining the development of the second and third whorls of organs in perfect flowers, the petal and stamens (Krizek and Meyerowitz, 1996). Two genes grouped most closely with AP3 according to the phylogenetic analysis performed in this study, M4 and PTD (Figure 5.2). PTD is already identified as the P. trichocarpa homolog of AP3 (Sheppard et al., 2000) and showed a female-biased expression according to the microarray data in this study. This is an interesting result because AP3 is a B-class gene that is required to specify petal and stamen identity (Krizek and Meyerowitz, 1996), and female P. trichocarpa flowers have neither petals, nor stamens. Sheppard et al. (2000) reported that PTD is expressed initially in the inner whorls of both male and female flowers, but once reproductive primordia began to form the spatial expression pattern became gender-specific, with PTD expression continuing in the stamen primordia, but excluded from the carpel primordia. M4 exhibited male-biased expression according to the microarray data, and this study reports novel expression data for this gene. Flowers of P. trichocarpa do not have petals (Boes and Strauss, 1994), therefore it may be plausible that the male-biased expression pattern that was observed for the gene M4, along with its protein sequence similarities to AP3, indicate that this gene has a role in stamen development in the male flowers. If PTD and M4 are the result of a genome duplication that has occurred in Populus since the genus diverged from Arabidopsis between 100 and 120 million years ago (Tuskan 137  et al., 2006), it could be that these duplicate genes have diverged in function as unisexual flowers evolved in Populus. There is evidence from whole genome studies that genes that are retained as duplicates in the genome frequently diversify in function, or develop subfunctionalization (Adams and Wendel, 2005). Further work will be required to test if this expression pattern is an indication of a differentiation of function of AP3 in a unisexual flower. 5.5.6  C-class gene-expression in P. trichocarpa inflorescences In the ABC model of floral development, C function in the third and fourth whorls of  flowers is contributed to by the gene AGAMOUS (AG) (Theißen, 2001), and to a lesser extent AGAMOUS-like genes such as SHATTERPROOF1 (SHP1) and SHATTERPROOF2 (SHP2) which are involved in seed dispersal (Liljegren et al., 2000). In A. thaliana, both SHP1 and SHP2 are C-class genes (Table 5.2), positively regulated by AG, with AG directly regulating SHP2 by binding to sequences in the SHP2 promoter (Zik and Irish, 2003). SHP1 and SHP2 are required for fruit dehiscence in A. thaliana, and are functionally redundant in this species, as mutations in either of these genes independently cause no phenotypic changes, but the double mutant shp1 shp2 result in fruits that will not break open (Liljegren et al., 2000). SEEDSTICK (STK) is another C-class gene that is necessary for the development of the funiculus, which connects the fruit to the seed, and plays a role in seed dispersal (Pinyopich et al., 2003). The two genes that showed the most female-biased expression on the microarray were SCA19_12 and F2, and neither of these genes had been previously characterized with expression data, but both of these genes group most closely with the A. thaliana SEEDSTICK (STK) gene (Figure 5.2). While SCA19_12 and F2 show different expression levels on the  138  microarray, both are expressed in female flowers, and not in males. SCA12_19 may be involved in at least carpel development, if not the development of the fruit, as STK is involved in this process in A. thaliana (Pinyopich et al., 2003). cDNA expression patterns confirmed that SCA19_12 is expressed in female floral tissues, but not in male inflorecences, and that there was no detectible expression of this gene in leaf tissues (Figures 5.3 and 5.4). Now that SCA12_19 and F2 have been identified by this work, and the expression of SCA19_12 has been confirmed to be strongly female flower specific, future research could take a more detailed look at specific expression patterns of SCA19_12 and F2 in female flowers using an in situ approach or perhaps by creating transgenic A. thaliana mutants. The remaining two genes that showed female-biased gender expression in this microarray study are floral MADS-box genes that have been previously studied in Populus. PTAG1 and PTAG2 have been shown to have an AG-like expression pattern in flowers. While this data shows that they have a female-biased expression pattern in floral tissues, previous work has indicated that they are also consistently expressed in vegetative tissues (Brunner et al., 2000). The phylogenetic analysis performed in this study confirms that PTAG1 and PTAG2 are positioned most closely with A. thaliana AG, based on sequence similarity of the MIK protein domains. 5.5.7  E-class gene-expression in P. trichocarpa inflorescences After the inception of the ABC model of floral development it was found that a fourth  class of genes was important in specifying the different whorls in floral meristems, and this class of genes was called the E-class (Theißen, 2001).  This group of genes, named  SEPALLATA (SEP1, SEP2, SEP3 and SEP4), are important in the quartet model of floral organ identity in Arabidopsis because they code for proteins that form quartets with the other  139  three classes of MADS-box genes to result in the development of floral organs (Theißen, 2001).  E-class genes are expressed throughout the developing floral meristems in  Arabidopsis (Figure 5.5). E-class genes in Populus have been characterized in the species P. tremuloides and P. deltoides, but not in P. trichocarpa. PtMADS31, the P. trichocarpa homolog of PdMADS2, an E-class gene first characterized in P. deltoides (Zhang et al., 2009), grouped with the A. thaliana SEP genes, as does PTM5 (Cseke et al., 2007), an E-class gene first characterized in P. tremuloides (Table 5.1).  These genes showed no gender-biased gene-expression in this  microarray study (Table 5.1), and both genes have been previously shown to be expressed in both male and female floral tissues, as well as some vegetative tissues. In P. deltoides, PdMADS2 is highly expressed in the perianth cups of male inflorescences, and the ovaries of female flowers, and also shows abundant expression in the peduncles of both genders (Zhang et al., 2009). In P. trichocarpa, PtMADS31 has not been the subject of expression studies to date, so further research would be required to establish if PtMADS31 has a similar expression pattern to PdMADS2 in P. trichocarpa flowers. In P. tremuloides, PTM5 has been shown to be a member of the SOC1/TM3 class of MADS-box genes and is involved in protein-protein interactions with itself and other MADS-box proteins (Cseke et al., 2007), but the P. trichocarpa homolog has not yet been studied. Three other MADS-box genes have been studied in Populus, PTM3, PTM4, and PTM6 (Cseke et al., 2005), and while previous work has shown that these genes are expressed in both male and female tissues, none of these E-class genes showed gender-biased expression in floral tissues according to this microarray study (Table 5.1). PTM3 and its duplicate PTM4 are related to the SEP1 and SEP2 genes found in A. thaliana, and PTM6 is  140  related to SEP3, and the concentration of expression of these genes in the inner whorls that contain the sexual organs of both male and female flowers suggests that in P. deltoides these genes are important in ensuring reproductive viability (Cseke et al., 2005). Given that SEPALLATA homologs in Populus appear to be expressed in all floral tissues studied to date in this genus, it is plausible that they play a similar role to the SEPALLATA genes that form protein quartets in A. thaliana to specify floral organ development. 5.6  Conclusions This research updates the known floral MADS-box genes to the most current genomic  annotations and locations, and places 88 floral MADS-box genes on version 2.0 of the poplar genome. My microarray experiment identified 14 putative MADS-box genes that showed gender-biased expression patterns in male and female inflorescences, and 13 of these genes were confirmed as correctly annotated as MADS-box genes. Novel expression patterns for nine floral MADS-box genes were identified with this microarray data, and the expression patterns of three of these genes was investigated in further detail using reverse-transcription PCR. I was able to show that SCA19_12 was expressed in female but not male floral tissues, and based on protein sequence homology is the P. trichocarpa homolog of the A. thaliana STK gene. I also discovered that MADS_1 and MADS_2 were expressed in male but not female floral tissues, and are homologs to the A. thaliana gene PI. None of these three genes were expressed in leaf tissues of male or female individuals at a detectible level.  141  Chapter 6: Conclusions The research presented in this thesis adds to the current knowledge available on the model organism Populus trichocarpa with respect to the use of the genetic resources available for this species, and investigated the development of sex-linked markers in P. trichocarpa that could be used to identify the gender of trees prior to flowering. This thesis is comprised of four projects that took various approaches to discover how gender in P. trichocarpa might be regulated at a genetic level. There were two principal ideas underlying the four projects. The first idea was to develop a genetic marker that could be used to sex P. trichocarpa individuals of unknown gender using vegetative tissue. Having a marker like this would be useful to Populus breeding programs as it would allow the selection of either all male or all female progeny for clonal propagation, and it would allow the gender of experimental crosses to be identified prior to these individuals reaching the age of first flowering, which can take as long as 15 years in some Populus species. The second idea was to investigate the genes involved in floral development in P. trichocarpa and compare geneexpression patterns in this dioecious, unisexual flower species with those of the homologous genes in the model organism A. thaliana, which produces hermaphroditic flowers. In this thesis, chapters two, three and four report the results of projects that investigated the development of a genetic marker for gender in vegetative tissues, and chapter five details the results of a study of MADS-box genes involved in floral development in P. trichocarpa.  142  6.1  Developing a genetic marker to identify the gender of an individual using  vegetative tissues SCAR-markers have been discovered that segregate with gender in pedigreed families of Salix viminalis, a finding that led to the development of the hypothesis that gender in this species may be determined genetically via several loci, perhaps functioning in an epistatic way (Alström-Rapaport et al., 1998). Because Salix is the sister genus to Populus in the Salicaceae, and it is probable that both these genera evolved dioecy, and therefore genetic sex-determination mechanisms via the monoecious pathway, I wanted to see if the sex-linked markers discovered in S. viminalis could be used to develop a genetic maker for gender in vegetative tissues in other Salix species and in P. trichocarpa. I identified the homologous sequence in P. trichocarpa for the Salix viminalis sexlinked marker SCAR-354 on chromosome 15 on the poplar genome. I was also able to amplify the Salix SCAR 354 marker sequence as well as the adjacent gene sequence for a Ssu72-like protein.  I characterized the SCAR marker sequence, and identified distinct  regions of the marker that varied in the degree to which they were conserved between the species sampled. When I investigated the gene-anchored sequence obtained in S. arctica and S. reticulata male and female individuals I found some evidence that gender-biased SNPs do exist in this sequence, which may explain why the SCAR 354 marker segregates with gender in pedigreed families of S. viminalis (Alström-Rapaport et al., 1998). However, the gender bias observed was not statistically significant. I was unable to confirm that the SCAR 354 marker that segregated with sex in S. viminalis was also a sex-linked marker in other Salix species or in P. trichocarpa,  143  Research into the genetics of gender-determination in Populus indicates that, as in Salix, one or two chromosomal regions or sex loci are involved in gender-determination. The development of the sequenced P. trichocarpa genome has allowed other researchers to associate the putative sex loci in this species to chromosome 19. For the project reported on in chapter three of this thesis, I investigated 24 genes in the telomeric region of chromosome 19 looking for SNP variation in the genomic sequence, or reduced recombination rates in SNPs between males and females in these genes, which could be associated with genetic markers for gender and a sex locus. The reasoning behind this was that it has been shown that areas of reduced recombination between the genders in a genome can indicate the initiation of a sex chromosome (Liu et al., 2004). I observed large variability in the number of SNPs detected in the gene sequences studied, but discovered no genetic marker that could be used to sex P. trichocarpa individuals of unknown gender. There was no overall trend of reduced recombination between adjacent SNPs in the gene sequences investigated, and the data collected shows that low recombination rates between SNPs are not maintained across the telomeric region on chromosome 19. If there is reduced recombination in the region of a sex locus on the telomeric end of chromosome 19, it would appear that it is very localized, as I looked at genes located less than 100kb from each other. Alternatively, it is possible that the sex locus is not in the ~1Mbp region I investigated. Gender-determination mechanisms have been investigated in at least six species of Populus (P. alba, P. detoides, P. nigra, P. tremula, P. tremuloides and P. trichocarpa), using pedigreed families and mapping maternal and paternal genetic markers in progeny of known gender (Paolucci et al., 2010). These studies, along with maturing genetic resources such as the re-annotated version 2.0 of the P.  144  trichocarpa genome makes it likely that developing a full understanding the genetics of gender-determination in this species will only be a matter of time. Shortly after the release of version 1.1 of the poplar genome I conducted two microarray experiments to look for gene-expression patterns that differed between male and female leaf tissues. I was able to identify a number of genes that showed small differences in expression levels between males and females, however the majority of these differences were not statistically significant, and my subsequent investigation using reverse transcription PCR of the genes was unable to confirm the microarray results. As a result I was unable to identify potential genomic sex-linked markers in P. trichocarpa using this method. Sample size and replication are both critical when conducting this kind of gene-expression experiment, as illustrated by the observation that there was a greater gene-expression difference between the two microarray experiments performed than between genders. Gender-biased gene-expression may be present in vegetative tissues of P. trichocarpa, but if so, these effects are subtle, and therefore a much larger sample size and better environmental control would be needed for a microarray experiment to be able to detect it with any statistical confidence. Another conclusion I came to during the course of this project was that revisiting data and updating it frequently to reflect the latest and most accurate gene annotations available is essential when working with a rapidly developing model system like P. trichocarpa. The release of version 2.0 of the poplar genome provided much additional information on the genes I was working with, which greatly improved the quality of my analysis, as I was able to more accurately place genes showing gender-biased expression on chromosome 19, the putative sex chromosome in P. trichocarpa.  145  6.2  Investigation of the genetics of floral development in P. trichocarpa P. trichocarpa produces flowers that appear to be unisexual throughout their  development, as they only initiate either male or female floral organs. There is no evidence that organs of the opposite gender are initiated and then later aborted during floral development. Studying the genetics of floral development in P. trichocarpa provides an opportunity to investigate how the genes involved in this process have differentiated to have either male- or female-specific expression.  In chapter five of this thesis a microarray  experiment using male and female inflorescence tissues showed that a number of floral MADS-box genes were differentially expressed between males and females. Since this gene family has never been systematically investigated in Populus, a phylogenetic analysis of P. trichocarpa MADS-box genes was conducted to compare the MIK protein domains of these genes to floral MADS-box genes that have had their functions thoroughly characterized in A. thaliana. I discovered that among the MADS-box genes that had a gender-biased expression pattern in Populus floral tissues there were often two P. trichocarpa homologs for each of the A. thaliana genes I looked at. There appears to have been some divergence in function of each paralogue, as they do not always share the same gender-bias in their expression, or the same levels of expression in a given tissue. My microarray experiment identified 14 putative MADS-box genes that showed putative gender-biased expression patterns in male and female floral tissue, and 13 of these genes were confirmed as MADS-box genes. I identified which of these genes had been previously characterized in Populus as having functions in flower development in this species, and which of these genes were identified as having floral expression patterns for the first time by my research. Novel expression patterns for nine floral MADS-box genes were  146  identified with this microarray data, and the expression patterns of three of these genes were investigated in further detail using reverse transcription PCR. I was able to show that SCA19_12 was expressed in female but not male floral tissues, and determined that SCA19_12 is the P. trichocarpa homolog of the A. thaliana STK gene based on protein sequence homology. I also discovered that Populus MADS_1 and MADS_2 were expressed in male but not female floral tissues, and are homologs to the A. thaliana gene PI. None of these three genes were expressed in leaf tissues of male or female individuals at a detectable level. With the release of the second version of the poplar genome I was able to update the known floral MADS-box genes to the most current genomic annotations and locations, and the total number of floral MADS-box genes according to my survey is now 88, down from the 105 first reported in Leseberg et al. (2006). Interestingly, I confirmed that there are two floral MADS-box genes located on chromosome 19, whereas it was previously reported that MADS-box genes were distributed across all chromosomes in P. trichocarpa except chromosome 19. One of these genes, SCA19_12, is expressed only in female floral tissues, and not in male floral tissues, or leaf tissues of either gender. Chromosome 19 is the location of the putative sex locus in P. trichocarpa, and the female of this species is thought to be the heterogametic gender (Yin et al., 2008), so it is interesting that a gene that is only expressed in female flowers is found on this chromosome. 6.3  Future directions and applications of this research Given that the conserved Ssu72-like gene associated region consistently amplified in  the Salix species I sampled, this marker may be useful for phylogenetic and population studies of willows. Current molecular phylogenetic studies in Salix have used chloroplast  147  genetic markers to resolve the phylogenetic relationships in this taxonomically difficult genus (Chen et al., 2010). Using the gene-anchored primers developed in this work it may be possible to discover species-specific single feature polymorphisms (SFPs), and these SFPs could prove to be useful phylogenetic markers for identifying Salix species, which can be difficult to classify based on morphological traits alone. Advances in cost effective, high throughput next-generation sequencing technologies that are currently available from Illumina IG, Applied Biosystems SoLiD and Roche 454 Life Sciences Systems (Wang et al., 2009) will make it much easier to generate and manage larger genetic data sets, and to keep the gene annotations up to date. Currently, 20 P. trichocarpa accessions are included in a population transcriptome resequencing project which promises to provide much more detailed information about gene annotation of the P. trichocarpa genome (Geraldes et al., 2011). Data sets of this kind would make it possible to detect with greater accuracy the subtle gender-biased gene-expression differences in vegetative tissues in P. trichocarpa that my experiments indicated exist. Future research into the genetics of gender-determination in poplar needs to address why genetic markers for some species of Populus indicate that the male is the heterogametic gender, and in other species they indicate that the female is heterogametic. Genetic data from resequencing the transcriptomes of 20 P. trichocarpa individuals could be used to identify genetic markers linked to gender once these trees have reached sexual maturity and can be sexed by observing floral gender. This is because this resource allows genotypic variability, which includes genetic difference between the genders, to be linked to geneexpression (Geraldes et al., 2011). Also, investigating chromosome 19 in the transcriptomes  148  of multiple P. trichocarpa individuals could explain why at least two regions on chromosome 19 in Populus species have been identified as being involved in sex-determination. This research constitutes the first survey of Populus floral MADS-box genes conducted using the updated genome annotations available for version 2.0 of the poplar genome and therefore provides a jumping off point for further research into the genetics of floral development in P. trichocarpa. A logical next step for continuing work on MADS-box genes involved in floral development in P. trichocarpa would be to confirm the gender bias I observed in the six other genes that were identified as having novel gender-biased expression with the microarray experiment using the cDNA synthesis technique used to investigate SCA19_12, MADS_1 and MADS_2 in a larger sample size of floral tissues of male and female P. trichocarpa individuals. 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In The University of Texas at Austin (Austin, The University of Texas), pp. 125.  165  Appendices Appendix A List of PCR primers designed to amplify genes studied in chapters three, four and five  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  eugene3.01170047  P.trichocarpa V 1.1 Genome  SCA117_1_1F  20  60.14  TGAAGAGCTCGAGACCCAGT  211  eugene3.01170047  P.trichocarpa V 1.1 Genome  SCA117_1_1R  20  60.04  TGCCTTGTCGATCCCTATTC  211  fgenesh4_pg.C_scaffold_117000045  P.trichocarpa V 1.1 Genome  SCA117_2_1F  20  59.84  TGTGCCGAAGAAAATGACAG  243  fgenesh4_pg.C_scaffold_117000045  P.trichocarpa V 1.1 Genome  SCA117_2_1R  20  59.73  GAGCGTTGCATGTGAATGTT  243  grail3.0117003001  P.trichocarpa V 1.1 Genome  SCA117_3_1F  20  60.22  AAGCATCAAATGGCGAAGAC  238  grail3.0117003001  P.trichocarpa V 1.1 Genome  SCA117_3_1R  20  59.94  ACCCTTGCCTCAATTCCTTT  238  gw1.117.122.1  P.trichocarpa V 1.1 Genome  SCA117_4_1F  20  60.09  TGGAGGCAACTTGTTTTTCC  155  gw1.117.122.1  P.trichocarpa V 1.1 Genome  SCA117_4_1R  20  59.86  TCCTCAAATCCCATCCAGAC  189  166  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  gw1.117.122.1  P.trichocarpa V 1.1 Genome  SCA117_4_2R  20  60.33  CTTACCCATCCGCAACATTC  208  fgenesh4_pg.C_scaffold_117000051  P.trichocarpa V 1.1 Genome  SCA117_5_1F  20  59.62  ACAGTGGGCATCACAGTACG  150  fgenesh4_pg.C_scaffold_117000051  P.trichocarpa V 1.1 Genome  SCA117_5_1R  20  59.98  GCTATGGTGGCGAGAGAAAG  150  fgenesh4_pg.C_scaffold_117000051  P.trichocarpa V 1.1 Genome  SCA117_5_2R  20  59.98  AGCTATGGTGGCGAGAGAAA  151  fgenesh4_pg.C_scaffold_117000053  P.trichocarpa V 1.1 Genome  SCA117_6_1F  20  60.01  TTTGATGGCCTGAATGATGA  153  fgenesh4_pg.C_scaffold_117000053  P.trichocarpa V 1.1 Genome  SCA117_6_1R  20  60.17  CTCCCCATATCTCGAAGCAA  212  fgenesh4_pg.C_scaffold_117000053  P.trichocarpa V 1.1 Genome  SCA117_6_2F  20  59.62  GAAGATTGCCGCTAGCACTT  151  fgenesh4_pg.C_scaffold_117000053  P.trichocarpa V 1.1 Genome  SCA117_6_2R  20  60.01  TCATCATTCAGGCCATCAAA  151  fgenesh4_pg.C_scaffold_117000054  P.trichocarpa V 1.1 Genome  SCA117_7_1F  20  60.07  ATCTCGGACTCCCGACTTTT  211  fgenesh4_pg.C_scaffold_117000054  P.trichocarpa V 1.1 Genome  SCA117_7_1R  20  60.13  TTTTGAGCAGCCTGAGAGGT  211  167  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  e_gw1.117.150.1  P.trichocarpa V 1.1 Genome  SCA117_8_1F  20  60.14  AACATCAGGCGTGTGCATTA  177  e_gw1.117.150.1  P.trichocarpa V 1.1 Genome  SCA117_8_1R  20  60.12  ACCTTTGCCGAGTCAACATC  177  e_gw1.117.150.1  P.trichocarpa V 1.1 Genome  SCA117_8_2R  20  59.81  TTTGAAGGCATGAGATGTGC  247  gw1.117.169.1  P.trichocarpa V 1.1 Genome  SCA117_9_1F  20  59.99  TGAGGCTTTTGTGGAGCTTT  150  gw1.117.169.1  P.trichocarpa V 1.1 Genome  SCA117_9_1R  20  59.67  CAAAACAGGGCATCTCTTCC  150  gw1.117.169.1  P.trichocarpa V 1.1 Genome  SCA117_9_2F  20  60.30  TGGAGCGGAAACACTACACA  214  gw1.117.169.1  P.trichocarpa V 1.1 Genome  SCA117_9_2R  20  60.07  CAGGGCATCTCTTCCACAAT  214  eugene3.01170072  P.trichocarpa V 1.1 Genome  SCA117_10_1F  20  60.30  AAAGAATGGAGGGGGACTTG  242  eugene3.01170072  P.trichocarpa V 1.1 Genome  SCA117_10_1R  20  60.49  AGCCTTGGGGACTAAGGTTG  242  fgenesh4_pg.C_scaffold_117000045  P.trichocarpa V 1.1 Genome  SCA117_2WG_1F  20  60.07  AGGCCTTCCCTCATGAAAGT  853  168  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  fgenesh4_pg.C_scaffold_117000045  P.trichocarpa V 1.1 Genome  SCA117_2WG_1R  20  59.94  CGAGATTTTGCCAACCATTT  853  fgenesh4_pg.C_scaffold_117000045  P.trichocarpa V 1.1 Genome  SCA117_2WG_2F  20  59.85  CATGGCTGCATGATAGCACT  922  fgenesh4_pg.C_scaffold_117000045  P.trichocarpa V 1.1 Genome  SCA117_2WG_2R  20  59.94  CGAGATTTTGCCAACCATTT  922  gw1.117.122.1  P.trichocarpa V 1.1 Genome  SCA117_4WG_1F  20  60.07  CATGGAGGATGGCTTTGACT  980  gw1.117.122.1  P.trichocarpa V 1.1 Genome  SCA117_4WG_1R  20  60.17  CTCCCCATATCTCGAAGCAA  980  fgenesh4_pg.C_scaffold_117000051  P.trichocarpa V 1.1 Genome  SCA117_5WG_1F  21  59.92  TGCTATCAACAAGCACAGTGG  852  fgenesh4_pg.C_scaffold_117000051  P.trichocarpa V 1.1 Genome  SCA117_5WG_1R  22  57.81  AATAGCTTTGTCGTACGCTTTC  852  gw1.117.169.1  P.trichocarpa V 1.1 Genome  SCA117_9WG_1F  20  60.15  GGGTAGGGATCACGGAGATT  559  gw1.117.169.1  P.trichocarpa V 1.1 Genome  SCA117_9WG_1R  20  59.67  AAAACAGGGCATCTCTTCCA  559  eugene3.01170072  P.trichocarpa V 1.1 Genome  SCA117_10WG_1 F  20  59.80  TTCTCATCAAGGGAGGCACT  853  169  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  eugene3.01170072  P.trichocarpa V 1.1 Genome  SCA117_10WG_1 R  20  60.49  AGCCTTGGGGACTAAGGTTG  853  eugene3.01170072  P.trichocarpa V 1.1 Genome  SCA117_10WG_2 F  20  59.80  TTCTCATCAAGGGAGGCACT  897  eugene3.01170072  P.trichocarpa V 1.1 Genome  SCA117_10WG_2 R  21  60.26  TCAGTCTTGGGCAACTTGTGC  897  gw1.117.255.1  P.trichocarpa V 1.1 Genome  SCA117_A_1F  20  60.07  ATGGGCTCCTCCTTCTTGTT  935  gw1.117.255.1  P.trichocarpa V 1.1 Genome  SCA117_A_1R  20  59.99  GGATCACCATGGAAATTTGG  935  eugene3.01170002  P.trichocarpa V 1.1 Genome  SCA117_B_1F  27  57.93  TCAGGATATGATGTATTATATG GACAA  705  eugene3.01170002  P.trichocarpa V 1.1 Genome  SCA117_B_1R  22  60.12  TTCATTGGTGACATATGGTCGT  705  fgenesh4_pg.C_scaffold_117000010  P.trichocarpa V 1.1 Genome  SCA117_C_1F  21  59.64  TTATACAGGCAATGTGCAACG  619  fgenesh4_pg.C_scaffold_117000010  P.trichocarpa V 1.1 Genome  SCA117_C_1R  24  59.67  TTCTCACCATCAAGTCTTAAAT CG  619  fgenesh4_pg.C_scaffold_117000025  P.trichocarpa V 1.1 Genome  SCA117_D_1F  20  59.52  CTAAAGGTCGCTGCAAGGTT  634  170  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  fgenesh4_pg.C_scaffold_117000025  P.trichocarpa V 1.1 Genome  SCA117_D_1R  20  59.59  AAAAGGTGTTTGGGGGTTTT  634  eugene3.01170041  P.trichocarpa V 1.1 Genome  SCA117_E_1F  24  59.43  CTTTTGGAAACTATGGATCAAA CA  617  eugene3.01170041  P.trichocarpa V 1.1 Genome  SCA117_E_1R  25  60.25  GAATGAAAATACATGCTTCTCC AAC  617  eugene3.01170064  P.trichocarpa V 1.1 Genome  SCA117_F_1F  20  60.03  TTCTTCCGGAGGCCTTAAAT  925  eugene3.01170064  P.trichocarpa V 1.1 Genome  SCA117_F_1R  20  60.03  TTGCCTCCTCCCTATTCCTT  925  gw1.117.220.1  P.trichocarpa V 1.1 Genome  SCA117_G_1F  20  59.56  CCTTGGAGAAGGTGGTTTTG  867  gw1.117.220.1  P.trichocarpa V 1.1 Genome  SCA117_G_1R  20  60.12  ATTGGTCCACAGGAGCAGTC  867  gw1.117.235.1  P.trichocarpa V 1.1 Genome  SCA117_H_1F  20  59.85  GACAACACCCACATCGACAC  853  gw1.117.235.1  P.trichocarpa V 1.1 Genome  SCA117_H_1R  20  59.99  TGCTGGAGACTGATTTGTGC  853  e_gw1.117.150.1  P.trichocarpa V 1.1 Genome  SCA117_8WG_1F  20  59.88  AACCCAGTACACCCAAGTCG  957  171  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  e_gw1.117.150.1  P.trichocarpa V 1.1 Genome  SCA117_8WG_1R  20  59.83  CATGAGCGAGCCATTGTAAA  957  POPTR_0019s01700  P.trichocarpa V 2.0 Genome  SCA19_1_1F  20  59.97  AAATATTGGTGCCAGCGTTC  633  POPTR_0019s01700  P.trichocarpa V 2.0 Genome  SCA19_1_1R  20  60.30  GCTGGCTGGTGTTGTTTCTT  633  POPTR_0019s01740  P.trichocarpa V 2.0 Genome  SCA19_2_1F  20  60.05  CACCGCAAAGCACTGTTAGA  970  POPTR_0019s01740  P.trichocarpa V 2.0 Genome  SCA19_2_1R  20  60.05  GTCCAGAAATCCTGCCAAAA  970  POPTR_0019s01780  P.trichocarpa V 2.0 Genome  SCA19_3_1F  20  59.96  AGGGGTTCGAGCTTTGGTAT  863  POPTR_0019s01780  P.trichocarpa V 2.0 Genome  SCA19_3_1R  20  60.04  CCTGACATCTTTTCCCCTGA  863  POPTR_0019s01830  P.trichocarpa V 2.0 Genome  SCA19_4_1F  20  59.98  CAGATCAGAGGCTGGTGTCA  957  172  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  POPTR_0019s01830  P.trichocarpa V 2.0 Genome  SCA19_4_1R  20  59.91  AACCTGGCTGTGCAGAAAGT  957  POPTR_0019s01850  P.trichocarpa V 2.0 Genome  SCA19_5_1F  20  59.99  AGTCAAAGCTCCGAAGACCA  942  POPTR_0019s01850  P.trichocarpa V 2.0 Genome  SCA19_5_1R  20  59.85  CTGGACTCATAGCACGACCA  942  POPTR_0019s01870  P.trichocarpa V 2.0 Genome  SCA19_6_1F  20  59.55  GTTCAGTTGGATGTGCCAAA  979  POPTR_0019s01870  P.trichocarpa V 2.0 Genome  SCA19_6_1R  20  59.78  CCCAGATAACCTTTCCACCA  979  POPTR_0019s13010  P.trichocarpa V 2.0 Genome  SCA19_7_1F  20  60.42  AAGCCAAACGAAGCAACAAG  939  POPTR_0019s13010  P.trichocarpa V 2.0 Genome  SCA19_7_1R  20  59.43  ATGACGGAACCTCGGTTAAA  939  173  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  POPTR_0019s03090  P.trichocarpa V 2.0 Genome  SCA_19_9_1F  20  60.05  GGAATGGAAGTTCGAGTGGA  912  POPTR_0019s03090  P.trichocarpa V 2.0 Genome  SCA_19_9_1R  20  59.70  CATACCTGCCACTCTGGTCA  912  POPTR_0019s03100  P.trichocarpa V 2.0 Genome  SCA_19_10_1F  20  59.97  CAAGGAATGGGTGTTCGAGT  899  POPTR_0019s03100  P.trichocarpa V 2.0 Genome  SCA_19_10_1R  20  60.04  TCTTAAGATGCCCCATTTGC  899  POPTR_0019s12360  P.trichocarpa V 2.0 Genome  SCA_19_11_1F  20  60.05  GGCCCAAACCAAACCTTATT  992  POPTR_0019s12360  P.trichocarpa V 2.0 Genome  SCA_19_11_1R  20  60.49  ACCACTGGGTATCCCACCTT  992  POPTR_0019s10580  P.trichocarpa V 2.0 Genome  SCA_19_12_1F  20  60.20  GCAGACATGGGAAGAGGAAA  722  174  Gene name  Organism and genome  Primer name  Length  Tm  Primer sequence  Product size  POPTR_0019s10580  P.trichocarpa V 2.0 Genome  SCA_19_12_1R  20  59.50  CAAAATACCACTTGCACACCA  722  POPTR_0002s07920  P.trichocarpa V 2.0 Genome  MADS_1_1F  23  60.03  CCTCACACCTTCACTTTGAAAT C  879  POPTR_0002s07920  P.trichocarpa V 2.0 Genome  MADS_1_1R  20  60.11  TTTCACGGGCTAAGTGGTTC  879  POPTR_0005s20480  P.trichocarpa V 2.0 Genome  MADS_2_1F  21  58.63  CCTCCTCACACCTTCAATTTC  874  POPTR_0005s20480  P.trichocarpa V 2.0 Genome  MADS_2_1R  20  59.58  TAGGGTTTTGAATGCACACG  874  175  Appendix B List of gene names, genomic locations, and functional annotations for genes studied in chapters two, three and four  Working name for gene  Ssu72-like  N/A  Poplar genome version 1.1 gene name  Poplar genome version 2.0 gene name  Location on version 2.0 poplar genome  Function annotation for gene  scaffold_15: 4923219 - 4926558  DEAD/DEAH box helicase  Overlaps with 2 genes on scaffold_3:POPTR_0003s03450 and POPTR_0003s03440  POPTR_0003s03450: scaffold_3: 3897472 - 3898473 and POPTR_0003s03440:scaffold_3: 3882517 - 3883355  There are no functional annotations for these two loci  POPTR_0017s13140:scaffold_17: 13300328 - 13301186 POPTR_0017s13180: scaffold_17: 13358880 - 13359781 POPTR_0017s13220: scaffold_17: 13422659 - 13423406 POPTR_0017s13260: scaffold_17: 13439842 - 13440015  POPTR_0017s13140: Rapid ALkalinization Factor (RALF) POPTR_0017s13180: There are no functional annotations for this locus POPTR_0017s13220: Rapid ALkalinization Factor (RALF) POPTR_0017s13260: There are no functional annotations for this locus  fgenesh1_pg.C_LG_XV 000380 POPTR_0015s04700  eugene3.00660277  N/A  fgenesh4_pg.C_scaffold_277 000004  Overlaps with 4 genes on scaffold_17: POPTR_0017s13140, POPTR_0017s13180, POPTR_0017s13220, POPTR_0017s13260  SCA 117 A  gw1.117.255.1  POPTR_0019s01120  scaffold_19: 849164 - 851761  Protein tyrosine kinase  SCA 117 B  eugene3.01170002  Sequence not included in V 2.0  N/A  N/A  176  Working name for gene  Poplar genome version 1.1 gene name  Poplar genome version 2.0 gene name  Location on version 2.0 poplar genome  Function annotation for gene POPTR_0019s01190 and POPTR_0019s02960: There are no functional annotations for these gene  SCA 117 C  fgenesh4_pg.C_scaffold_117 000010  POPTR_0019s01190 POPTR_0019s02960  POPTR_0019s01190 sca_19:914104917075 POPTR_0019s2960 scaffold_19: 2875642 - 2876509  SCA 117 D  fgenesh4_pg.C_scaffold_117 000025  POPTR_0019s01270  scaffold_19: 1075484 - 1078988  Terpene synthase family, metal binding domain  SCA 117 E  eugene3.01170041  POPTR_0019s01340  scaffold_19: 1177994 - 1181330  terpene synthase/cyclase family protein; similar to myrcene/ocimene synthase  SCA 117 1  eugene3.01170047  POPTR_0019s01520  POPTR_0019s01520 = scaffold_19: 1339019 - 1340779  Transferase family  SCA 117 2  fgenesh4_pg.C_scaffold_117 000045  POPTR_0019s01530  POPTR_0019s01530 = scaffold_19: 1347076 - 1348225  there are no functional annotations for this gene  SCA 117 3  grail3.0117003001  POPTR_0019s01540  POPTR_0019s01540 = scaffold_19: 1348343 - 1349893  Transferase family  SCA 117 4  gw1.117.122.1  POPTR_0019s01560  scaffold_19: 1379318 - 1382368  Leucine Rich Repeat  SCA 117 5  fgenesh4_pg.C_scaffold_117 000051  POPTR_0019s01570  Apoptotic ATPase, protein binding  SCA 117 6  fgenesh4_pg.C_scaffold_117 000053  Overlaps with 2 genes on scaffold_19: POPTR_0019s01560 POPTR_0019s01570  POPTR_0019s01570 = scaffold_19: 1391060 - 1395729 POPTR_0019s01560 = scaffold_19: 1379318 - 1382368 POPTR_0019s01570 = scaffold_19: 1391060 - 1395729  fgenesh4_pg.C_scaffold_117 000054  Overlaps with 2 genes on scaffold_19: POPTR_0019s01560 POPTR_0019s01570  POPTR_0019s01560 = scaffold_19: 1379318 - 1382368 POPTR_0019s01570 = scaffold_19: 1391060 - 1395729  SCA 117 7  Apoptotic ATPase  Apoptotic ATPase  177  Working name for gene SCA 117 8  SCA 117 F  Poplar genome version 1.1 gene name  Poplar genome version 2.0 gene name  e_gw1.117.150.1  POPTR_0019s01630  eugene3.01170064  Overlaps with 2 genes on scaffold_19: POPTR_0019s01660 and POPTR_0019s01670  Location on version 2.0 poplar genome POPTR_0019s01630 = scaffold_19: 1438061 - 1439700 POPTR_0019s01660 = scaffold_19: 1460825 - 1462203 POPTR_0019s01670 = scaffold_19:scaffold_19: 1463832 1466771 POPTR_0019s01660 = scaffold_19: 1460825 - 1462203 POPTR_0019s01670 = scaffold_19:scaffold_19: 1463832 1466771  Apoptotic ATPase  Function annotation for gene Alginate lyase  Apoptotic ATPase  SCA 117 9  gw1.117.169.1  Between 2 genes on scaffold_19:POPTR_0019s01660 and POPTR_0019s01670  SCA 117 10  eugene3.01170072  POPTR_0019s01790  scaffold_19: 1546894 - 1550419  Transcription elongation factor  SCA 117 G  gw1.117.220.1  scaffold_19: 1630220 - 1633020  Serine/threonine protein kinase  SCA 117 H  gw1.117.235.1  POPTR_0019s01880 Oversaps with 2 genes on scaffold_19: POPTR_0019s01940 and POPTR_0019s01950  scaffold_19: 1728118 - 1733133  Serine/threonine protein kinase  SCA 19 11  eugene3.00190854  POPTR_0019s12360  scaffold_19: 13401257 - 13403527  Chitin recognition protein  SCA 19 12  eugene3.00190689  POPTR_0019s10580  scaffold_19: 12132258 - 12133799  K-box region, regulation of transcription, DNA-dependent  178  Appendix C List of gene names, genomic locations, and functional annotations for genes studied in chapter five  Working name for gene  Poplar genome version 2.0 gene name  Location on version 2.0 poplar genome  Function annotation for gene  SCA19_12  POPTR_0019s10580  scaffold_19: 12132258 12133799  K-box region, regulation of transcription, DNA-dependent  F2  POPTR_0013s09980  scaffold_13: 9955912 9966253  MADS box transcription factor  F3  POPTR_0011s03140  scaffold_11: 3200188 3207704  MADS-box transcription factor  F4  POPTR_0008s09800  scaffold_8: 6076113 6078596  MADS-box protein regulation of transcription, DNA-dependent  PTAP2 and PTM1  POPTR_0010s16380  scaffold_10: 15345650 15351172  MADS-box transcription factor  PTAP2 and PTM1  POPTR_0010s16380  PTD  POPTR_0007s13660  scaffold_10: 15345650 15351172 scaffold_7: 13686094 13687851  PTAG1  POPTR_0004s06300  scaffold_4: 4997208 5005680  SRF-type transcription factor (DNA-binding and dimerisation domain)  M1  POPTR_0004s11420  scaffold_4: 10038841 10040605  MADS-box protein  M2  POPTR_0012s05960  scaffold_12: 5936482 5942087  MADS box transcription factor  M4  POPTR_0005s12000  M5  POPTR_0002s02910  scaffold_5: 8770246 8772014 scaffold_2: 1789707 1795019  M6  POPTR_0017s13410  scaffold_17: 13578276 13585496  MADS-box transcription factor SRF-type transcription factor (DNA-binding and dimerisation domain)  MADS-box protein involved in regulation of transcription, DNAdependent Kinesin-like protein involved in microtubule motor activity MADS box transcription factor  179  Working name for gene  Poplar genome version 2.0 gene name  Location on version 2.0 poplar genome  Function annotation for gene  MADS_1  POPTR_0002s07920  scaffold_2: 5450756 5453302  MADS box transcription factor  MADS_2  POPTR_0005s20480  PtMADS31  POPTR_0008s09790  PTM3 and PTM4  POPTR_0004s11440  scaffold_4: 10100959 10108181  MADS box transcription factor  PTM5  POPTR_0014s07010  scaffold_14: 5281611 5290965  MADS box transcription factor  PTM6  POPTR_0001s13650  scaffold_1: 10729860 10732842  Regulation of transcription, DNA-dependent  scaffold_5: 19532615 19535235 scaffold_8: 6063858 6070604  MADS box transcription factor MADS box transcription factor  180  Appendix D List of all MADS-box genes reported in Leseberg et al. (2006) updated to version 2.0 of the poplar genome Leseberg name  PtMADS1  PtMADS2  PtMADS3 + 4  PtMADS5  Locus Name  POPTR_0001s08500 and POPTR_0001s08510  POPTR_0001s33600  POPTR_0001s29100  POPTR_0001s13660  Location  POPTR_0001s08500=scaffold_1: 6475276 - 6476293 and POPTR_0001s08510=scaffold_1: 6478981 - 6480482  Research on this gene  Functional annotations for this locus  BLAST hit to 2 genes on version 2.0 of the poplar genome.  POPTR_0001s08500=Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN POPTR_0001s08510=Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN  scaffold_1: 31831353 - 31831577  scaffold_1: 27887506 - 27887724  These two genes are annotated as a single gene on version 2.0 of the genome.  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN  scaffold_1: 10747910 - 10748155 P. trichocarpa homolog for PTM6 SEP-class gene in Populus tremuloides, Cseke et al 2005  PtMADS6  POPTR_0001s13650  scaffold_1: 10729860 - 10732842  PtMADS7  POPTR_0002s10580  scaffold_2: 7672322 - 7674041  PtMADS8  POPTR_0002s11030  scaffold_2: 8048500 - 8050136  Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN  PtMADS9  POPTR_0002s15300  scaffold_2: 11431129 - 11433605  Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN  181  Leseberg name  Locus Name  Location  PtMADS10  POPTR_0002s02990  scaffold_2: 1842523 - 1843856  PtMADS11  POPTR_0002s07920  scaffold_2: 5450756 - 5453302  PtMADS12  POPTR_0003s11960  scaffold_3: 12666516 - 12672466  PtMADS13  POPTR_0003s16800 and POPTR_0003s16810  POPTR_0003s16800 = scaffold_3: 16329000 - 16330057 POPTR_0003s16810 = scaffold_3: 16335820 - 16337536  PtMADS14 and 15  PtMADS16  POPTR_0003s16850  POPTR_0004s11430  scaffold_3: 16367370 - 16368102  scaffold_4: 10049087 - 10049726  Research on this gene  MADS_1 MADS-box gene that this research has shown to be only expressed in male floral tissue, not in female flowers tissue, or leaf tissue from either gender.  BLAST hit to 2 genes on version 2.0 of the poplar genome. These two genes are annotated as a single gene on version 2.0 of the genome.  Functional annotations for this locus Pfam:01486 K-box regionPfam:00319 SRFtype transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor KEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor POPTR_0003s16800 = Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN POPTR_0003s16810 = Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN  182  Leseberg name  Locus Name  Location  PtMADS17  POPTR_0004s11440  scaffold_4: 10100959 - 10108181  PtMADS18  POPTR_0004s06300  scaffold_4: 4997208 - 5005680  PtMADS19  POPTR_0009s08270  scaffold_9: 7613164 - 7616576  PtMADS20  POPTR_0009s06060  scaffold_9: 5973865 - 5977200  Functional annotations for this locus Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor KEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN KEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  PtMADS21  POPTR_0005s17950  scaffold_5: 15803268 - 15804499  Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN  PtMADS22  POPTR_0005s20480  scaffold_5: 19532615 - 19535235  PtMADS23  POPTR_0005s19420  scaffold_5: 18264598 - 18266213  Research on this gene  PTAG1 (Brunner et al., 2000)  MADS_2 MADS-box gene that this research has shown to be only expressed in male floral tissue, not in female flowers tissue, or leaf tissue from either gender.  Pfam:01486 K-box regionPfam:00319 SRFtype transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factorKEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN  183  Leseberg name  Locus Name  Location  PtMADS24  POPTR_0006s04730  scaffold_6: 3266443 - 3269027  PtMADS25  POPTR_0007s13660  scaffold_7: 13686094 - 13687851  PtMADS26  POPTR_0007s14310  scaffold_7: 14130991 - 14137078  PtMADS27 and 28 and 29  POPTR_0007s03280  scaffold_7: 1888163 - 1889185  PtMADS30  POPTR_0007s07620  scaffold_7: 5972870 - 5975044  PtMADS31  POPTR_0008s09790  scaffold_8: 6063858 - 6070604  Research on this gene  F6  These three genes are annotated as a single gene on version 2.0 of the genome.  Zhang et al, 2009., in P.deltoides, found that homolog to this gene is involved in foral development in males and females - SEP type expression, E-class gene.  Functional annotations for this locus Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  Panther:11945 MADS BOX PROTEIN Pfam:01486 K-box regionPfam:00319 SRFtype transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor  Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  184  Leseberg name  PtMADS32  Locus Name  POPTR_0008s09800  Location  Research on this gene  Functional annotations for this locus  scaffold_8: 6073641 - 6078620  PTM2 and PTAP1 sequence blasts to this sequence  Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor KEGGORTH:09264 MADS-box transcription factor, plant GO:0006355 regulation of transcription, DNAdependent  PtMADS33  POPTR_0010s16380  scaffold_10: 15345650 15351172  PTAP2 (Brunner et al., 2004) and PTM1 (Cseke et al, 2005) are same gene, and BLAST to this gene so it shows both male and femlae biased expression.  PtMADS34  POPTR_0011s03140  scaffold_11: 3200188 - 3207704  PTAG2 from Brunner et al., 2000 These two genes are annotated as a single gene on version 2.0 of the genome.  PtMADS35 and 36  POPTR_0012s10190  scaffold_12: 11018129 11024115  PtMADS37  POPTR_0012s14020  scaffold_12: 13998081 14005758  Pfam:01486 K-box regionPfam:00319 SRFtype transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factorKEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor KEGGORTH:09264 MADS-box transcription factor, plant Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  185  Leseberg name  Locus Name  Location  PtMADS38  POPTR_0012s14770  scaffold_12: 14522968 14525400  PtMADS39  POPTR_0012s05960  scaffold_12: 5936482 - 5942087  PtMADS40  POPTR_0014s07000  scaffold_14: 5271849 - 5273587  PtMADS41  POPTR_0014s07010  scaffold_14: 5281611 - 5290965  PtMADS42  POPTR_0019s10540  scaffold_19: 12079388 12082794  PtMADS43  POPTR_0019s10580  scaffold_19: 12126290 12134031  Research on this gene  Male biased expression on the microarray study, no expression data other than this research.  Functional annotations for this locus Pfam:01486 K-box regionPfam:00319 SRFtype transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN  P. trichocarpa homolog for PTM5 SEP-class gene in Populus tremuloides, Cseke et al., 2005  SCA19_12 - MADSbox gene on Chromosome 19 that this research has shown to be only expressed in female floral tissue, not in male flowers tissue, or leaf tissue from either gender.  Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  Pfam:01486 K-box regionPfam:00319 SRFtype transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor  186  Leseberg name  Locus Name  Location  PtMADS44  POPTR_0015s11040  scaffold_15: 11727470 11733274  PtMADS45  POPTR_0015s14950  scaffold_15: 14342335 14344327  PtMADS46  POPTR_0015s14010  scaffold_15: 13758181 13760371  Research on this gene  Functional annotations for this locus Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  PtMADS47 and 48  POPTR_0017s07170  scaffold_17: 5557162 - 5558254  These two genes are annotated as a single gene on version 2.0 of the genome.  PtMADS49  POPTR_0017s13390 and POPTR_0017s13410  POPTR_0017s13390 = scaffold_17: 13572863 13573949 POPTR_0017s13410 =  Used POPTR_0017s13410 in the phylogeny= M6 Male  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEIN POPTR_0017s13390 = there are no functional annotations for this locus POPTR_0017s13410 = Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  POPTR_0017s13390  POPTR_0017s13390 = scaffold_17: 13572863 13573949  BLAST hit to 2 genes on version 2.0 of the poplar genome.  POPTR_0017s13390 = there are no functional annotations for this locus  scaffold_13: 9955912 - 9966253  Female biased expression on the microarray study, no expression data other than this research.  Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  PtMADS50  PtMADS51  POPTR_0013s09980  187  Leseberg name  Locus Name  PtMADS52  POPTR_0013s10200  Location scaffold_13: 10214970 10216137  PtMADS53  POPTR_0003s16840  scaffold_3: 16351961 - 16352345  PtMADS54 and 55  POPTR_0003s16830  scaffold_3: 16347023 - 16350147  PtMADS56  POPTR_0001s25850  scaffold_1: 24929361 - 24929969  PtMADS57 and 75  PtMADS58  POPTR_0005s14660 Hits between POPTR_0002s06270 and POPTR_0002s06280  scaffold_5: 11665785 - 11666699  Research on this gene  Functional annotations for this locus Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN  These two genes are annotated as a single gene on version 2.0 of the genome.  These two genes are annotated as a single gene on version 2.0 of the genome.  Pfam:01486 K-box region Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  Does not align with reading frame of a gene.  PtMADS59  POPTR_0004s13590  scaffold_4: 13919101 - 13919616  PtMADS60  POPTR_0004s13600  scaffold_4: 13937678 - 13938214  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  PtMADS61  POPTR_0009s05070  scaffold_9: 5245134 - 5245484  Panther:11945 MADS BOX PROTEIN  188  Leseberg name  Locus Name  Location  Research on this gene  PtMADS62 and 63  POPTR_0009s08740 and POPTR_0009s08750  POPTR_0009s08740=scaffold_9: 7896599 - 7897252 POPTR_0009s08750=scaffold_9: 7900899 - 7901868  These two genes are annotated as to the same two genes on version 2.0 of the genome.  PtMADS64  POPTR_0006s27460 and POPTR_0006s27470  POPTR_0006s27460=scaffold_6: 25357649 - 25358757 POPTR_0006s27470=scaffold_6: 25361513 - 25362412  PtMADS65  POPTR_0007s03410  scaffold_7: 1958989 - 1959761  PtMADS66  POPTR_0008s04110  scaffold_8: 2294059 - 2294718  PtMADS67 and PtMADS73  POPTR_0012s11190  scaffold_12: 11727312 11728355  PtMADS68 and PtMADS70  POPTR_0016s12640  scaffold_16: 12022576 12023214  BLAST hit to 2 genes on version 2.0 of the poplar genome.  Functional annotations for this locus POPTR_0009s08740=Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor POPTR_0009s08750=Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor POPTR_0006s27460= Panther:11945:SF19 MADS BOX PROTEIN POPTR_0006s27470=Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  189  Leseberg name  Locus Name  Location  PtMADS69  POPTR_0017s07460  scaffold_17: 5801036 - 5801934  PtMADS71  POPTR_0004s00400  scaffold_4: 124160 - 124946  PtMADS72  POPTR_0004s01080  Research on this gene  scaffold_4: 542625 - 543137  PtMADS74 and PtMADS77  POPTR_0017s06710  scaffold_17: 5128975 - 5129586 ::  These two genes are annotated as a single gene on version 2.0 of the genome.  PtMADS76  POPTR_0017s06800 and POPTR_0017s06810  POPTR_0017s06800=scaffold_17: 5188482 - 5189081 POPTR_0017s06810=scaffold_17: 5192102 - 5192713  BLAST hit to 2 genes on version 2.0 of the poplar genome.  PtMADS78  POPTR_0016s12650  scaffold_16: 12031661 12032191  Functional annotations for this locus Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor POPTR_0017s06800=Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor POPTR_0017s06810=Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN GO:0003700 transcription factor activity GO:0043565 sequence-specific DNA binding GO:0006355 regulation of transcription, DNAdependent GO:0005634 nucleus  190  Leseberg name  Locus Name  Location  PtMADS79  POPTR_0001s22220  scaffold_1: 20864473 - 20865483  Functional annotations for this locus Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEIN  scaffold_1: 31907047 - 31908123  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  PtMADS80  POPTR_0001s33740  PtMADS81  POPTR_0002s25700  scaffold_2: 22772493 - 22773710  PtMADS82  POPTR_0007s03260  scaffold_7: 1869051 - 1877508  PtMADS83  POPTR_0008s02120  scaffold_8: 1011737 - 1012369  PtMADS84  POPTR_0010s18860  scaffold_10: 16975875 16976760  PtMADS85  POPTR_0010s24540  scaffold_10: 20709023 20709655  PtMADS86  POPTR_0012s11170  scaffold_12: 11722595 11723767  Research on this gene  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:01486 K-box region Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor  191  Leseberg name  PtMADS87 PtMADS88 and PtMADS89  Locus Name  Location  POPTR_0015s08890  scaffold_15: 10253929 10254633  POPTR_0004s23090  PtMADS90  scaffold_4: 21882354 - 21883063 No BLAST hit on version 2.0 of the poplar genome for this gene.  PtMADS91  POPTR_0002s09290  scaffold_2: 6583977 - 6587230  PtMADS92  POPTR_0005s16500  scaffold_5: 13345268 - 13345565  PtMADS93  POPTR_0007s02770  scaffold_7: 1612601 - 1612946  PtMADS94  POPTR_0007s05980  scaffold_7: 4126423 - 4128527  PtMADS95  POPTR_0008s08780  scaffold_8: 5431595 - 5435799  PtMADS96  POPTR_0010s17450  scaffold_10: 16038303 16041326  Research on this gene  Functional annotations for this locus  These two genes are annotated as a single gene on version 2.0 of the genome.  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  192  Leseberg name  Locus Name  Location  PtMADS97  POPTR_0018s08400  scaffold_18: 9348208 - 9348865  PtMADS98 and PtMADS99  PtMADS100  POPTR_0007s10345  POPTR_0006s21630  scaffold_7: 10218463 - 10218950  scaffold_6: 20654652 - 20656061  Research on this gene  These two genes are annotated as a single gene on version 2.0 of the genome.  Functional annotations for this locus Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  PtMADS101  POPTR_0013s01880  scaffold_13: 1089954 - 1090806  PtMADS102  POPTR_0013s00350  scaffold_13: 120118 - 120792  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor  scaffold_16: 4631275 - 4632726  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain)Panther:11945 MADS BOX PROTEINKOG:0014 MADS box transcription factor  PtMADS103  POPTR_0016s06850  PtMADS104  POPTR_0005s00420  scaffold_5: 134895 - 135674  PtMADS105  POPTR_0013s11210  scaffold_13: 11884123 11884934  Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN KOG:0014 MADS box transcription factor Pfam:00319 SRF-type transcription factor (DNA-binding and dimerisation domain) Panther:11945 MADS BOX PROTEIN  193  Appendix E Protein alignment of MADS-box protein domain for Populus trichocarpa MADS-box genes used to construct the phylogenetic tree in chapter five Gene name >AP1 >PTAP1_1 >PTAP1_2 >M1 >M2 >M6 >PI >MADS_1 >MADS_2 >AP3 >PTD >PMADS30 >M4 >AG >PTAG1 >PTAG2 >SHP1 >SHP2 >STK >SCA19_12 >F2 >PTM3_PTM4 >SEP1 >SEP2 >SEP3 >SEP4 >PTM5 >PtMADS31 >PtMADS7 >PtMADS12 >PtMADS17 >PtMADS21 >PMADS27 >PtMADS82  10 MSRGRVQLKR MGRGRVQLKR MGRGRVQLKR ---------MGRGRIQLKR MGRGRVQLKR MGRGKIEIKR MGRGKIEIKR MGRGKIEIKR MARGKIQIKR MGRGKIEIKK MGRGKIEIKK MGRGKIEIKK SGRGKIEIKR LGRGKVEIKR LGRGKVEIKR LGRGKIEIKR IGRGKIEIKR MGRGKIEIKR MGRGKIEIKR MGRGKIEIKR MGRGRVELKR MGRGRVELKR MGRGRVELKR MGRGRVELKR MGRGKVELKR MVRGKTQMRR MGRGRVELKR MAREKIKIKK MVRGKTQMKR MGRGRVELKR MAREKIKIKK MTRKKIQIKK MTRKKIQIKK  20 IENKINRQVT IENKINRQVT IENKINRQVT ---------IENKINRQVT IENNISRQVT IENANNRVVT IENSSNRQVT IENASNRQVT IENQTNRQVT IENPTNRQVT IENPTNRQVT IENPTNRQVT IENTTNRQVT IENTTNRQVT IENTTNRQVT IENTTNRQVT IENTTNRQVT IENSTNRQVT IENTTNRQVT IENTTNRQVT IENKINRQVT IENKINRQVT IENKINRQVT IENKINRQVT IENKINRQVT IENATSRQVT IENKINRQVT IDNVTARQVT IENATSRQVT IENKINRQVT IDNVAARQVT IDNTAARQVT IDNTAARQVT  30 FSKRRAGLLK FSKRRTGLLK FSKRRTGLLK ---------FSKRRSGLLK FSKRRTGLLK FSKRRNGLVK YSKRRSGIIK YSKRKNGIIK YSKRRNGLFK YSKRRNGIFK YSKRRNGIFK YSKRRNGIFK FCKRRNGLLK FCKRRSGLLK FCKRRNGLLK FCKRRNGLLK FCKRRNGLLK FCKRRNGLLK FCKRRNGLLK FCKRRNGLLK FAKRRNGLLK FAKRRNGLLK FAKRRNGLLK FAKRRNGLLK FAKRRNGLLK FSKRRNGLLK FAKRRNGLLK FSKRRRGLFK FSKRRNGLLK FAKRRNGLLK FSKRRRGLLK FSKRRRGLFK FSKRRRGLFK  40 KAHEISVLCD KAHEISVLCD KANEISVLCD ---------KAHEISVLCD KAHEISVLCD KAKEITVLCD KAKEITVLCD KAKEITVLCD KAHELTVLCD KAQELTVLCD KAQELTVLCD KAQELTVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAYELSVLCD KAFELSVLCD KAYELSVLCD KAEELSVLCD KAFELSVLCD KAYELSVLCD KAEELSVLCD KAYELSTLCD KAYELSTLCD  50 AEVALVVFSH AEVALIVFSH AEVALIVFSH -----NFVEH AEVALIVFST ADVAVIVFST AKVALIIFAS AQVSLVIFAS AQVSLVIFAS ARVSIIMFSS AKVSLIMFSN AKVSLIMFSN AKVSLIMFSN AEVALIVFSS AEVALIVFSS AEVALIVFSS AEVALVIFST AEVALVIFST AEVALIVFST AEVALIVFSS AEVSLIVFSS AEVALIIFSN AEVALIIFSN AEVSLIVFSN AEVALIIFSN AEIALLIFSN AEVALIVFSP AEVALIIFSN AEVAVIIFSA AEVALIVFSS AEVALIIFSN VEVAVIIFSA AEIALTVFSA AEIALMVFSA  60 KGKLFEYSTD KGKLFEYSTN KGKLFEYSTD SSEIFHFF-G KGKLFEYATD KGKLFEYSTD NGKMIDYCCP SGRMHEYCSP SGRMHEYCSP SNKLHEYISP TNKLNEYISP TNKFHEYISP TNKFHEYISP RGRLYEYSNN RGRLYEYSND RGRLYEYSNN RGRLYEYANN RGRLYEYANN RGRLYEYANN RGRLYEYANN RGRLYEYANN RGKLYEFCST RGKLYEFCSS RGKLYEFCST RGKLYEFCSS RGKLYEFCSS RGKLYEFAST SGKLFEFCSS TGKLFEYSSS RGKLYEFSSS RGKLYEFCST TGKLFEYSSS TGKLFEYSNT TGKLFEYSNS  70 -S-CM-----A-CM-----D-SM-------SM-----S-CM-----S-SM-----SMDL-----STTV-----STTV-----NTTT-----STST-----STTT-----STTT-----S--V-----S--V-----S--V----–S--FIYLLL -S--V-----N--I-----N--I-----N--I-----S-NM-----S-NM-----S-NM-----S-SM----PS-GM-----S--M-----S-NM-----S--M-----S--I-----S-NM-----S--M-----R--T-----S--M-----  80 ------------------------------------------------------------------------------------------------------------------------------------------------EKKKKKKKKK ----------------------------------------------------------------------------------------------------------------------------------------------------------  194  Gene name >AP1 >PTAP1_1 >PTAP1_2 >M1 >M2 >M6 >PI >MADS_1 >MADS_2 >AP3 >PTD >PMADS30 >M4 >AG >PTAG1 >PTAG2 >SHP1 >SHP2 >STK >SCA19_12 >F2 >PTM3_PTM4 >SEP1 >SEP2 >SEP3 >SEP4 >PTM5 >PtMADS31 >PtMADS7 >PtMADS12 >PtMADS17 >PtMADS21 >PMADS27 >PtMADS82  90 ------------------------------------------------------------------------------------------------------------------------------------------------NLWIYSSHVV ----------------------------------------------------------------------------------------------------------------------------------------------------------  100 110 EKILERYERY SYAERQLIAP EKILERHERY SYAERQLVAT EKILERYERY SYAERQLVAT ESILERYERC SYLEQQLVPN ERILERYERY SYAERQLLAN ESILERYERC SYAEQQFVPH GAMLDQYQKL SG--KK-WDVDLLDKYHKQ SG--KRLWDVDLLDKYHKQ SG--KRLWDKEIVDLYQTI SDVDVWA--KKIYDQYQNA LGIDLWG--KKIYDQYQKA LGIDLWS--KKIYDQYQKA LGIDLWS--KGTIERYKK AISDNSNTGSV KSTIERYKK ASADSSNTGSV KSTIERYKK ACADSSNNGSV RGTIERYKK ACSDAVNPPSV RGTIERYKK ACSDAVNPPTI RSTIERYKK ACSDSTNTSTV RSTIDRYKK ASSDSSNASSI RSTIDRYKK VSSDSSNTASI LKTLERYQK CSYGAEEVN-K LKTLDRYQK CSYGSIEVNNK LKTLERYQK CSYGSIEVNNK LRTLERYQK CNYGAPEPNVP ARTVDKYRK HSYATMDPN-Q QETIERYRR HVKENNTNKQP ATTIEKYQR FSYGALEGG-Q KDVLARYNL HSNNLDKINPP NRTIERYQK RAKDVGISSKM LKTLERYQK CSYGAEEVN-K KDVLARYNL HSNNLDKLNQP ID------- ----------GQVIERRNL HPKNINTLDQP  120 130 –ESDVNTNWS MEYNRLKAKI -DLDSQG-NW TLEYNRLKAK -DLDSQG-NW TLEYNRLKAK -GSEHQE-SW SLEHPKLMAR DDPENHG-SW TLEYAKLKAR -GPEHQG-SW FLEHPKLRAR AK---HE-NL SNEIDRIKKE AK---HE-NL SNEIDRIKKE AK---HE-NL SKEIDRIKKE TQ---YE-RM QETKRKLLET TQ---YE-KM QEHLRKLNDI AQ---YE--- ------IS-I AQ---YE--- ------IS-I AEIN-AQ-YYQ QESAKLRQQ SEAN-AQ-YYQ QEAAKLRSQ SEAN-AQ-FYQ QEAAKLRSQ TEAN-TQ-YYQ QEASKLRRQ TEAN-TQ-YYQ QEASKLRRQ QEIN-AA-YYQ QESAKLRQQ TEIN-AQ-YYQ QESAKLRQQ TEIN-AQ-YYQ QESAKMRQQ PAKE-LE-SSY REYLKVKAR PAKE-LE-NSY REYLKLKGR PAKE-LE-NSY REYLKLKGR SRELAVELSSQ QEYLKLKER SAKD-LQ-DKY QDYLKLKSR VEQN-ML-QLK EEAASMIKK SEKETQQ-NNY QEYLKLKTR SLE--LQL-EN SNHMRLSKE VQDN-IQ-PVK EDTFTLAKK PAKE-LE-SSY REYLKVKAR SLE--LQL-EN SNHMRLRKE ----------- --------SLE--KQL-DG GVHAMLIKE  140 ELLERNQRHY VELLQRNHRH VELLQRNHRN VEILQRNLRN VDVLQRNQRH VELLQRNLRN NDSLQLELRH NESMQIELRH NDSMQIELRH NRNLRTQIKQ NHKLRQEIRK NHKLKKEIRQ NHKLKKEIRQ IISIQNSNRQ IGNLQNSNRH IGNLQNSNRN IRDIQNSNRH IRDIQNLNRH IQTIQNSNRN IQMLQNSNRH IQLLQNSNRH FEALQRTQRN YENLQRQQRN YENLQRQQRN YDALQRTQRN VEILQHSQRH IEHLEVSKRK VDVLQRSQRN VSEKSHQLRR IELLEVSKRK FEALQRTQRN VSEKSHQLRR ---------IAKKNRELRH  150 LGEDLQAMSP YLGEDLDSVS YLGEDLDSMS YAGQELDPLS FMGEDLDSLN YTGQDLDPLS LKGEDIQSLN LKGQDISSLP LKGEDISSLH RLGECLDELD YHVIKTQNET RIGEDLNELS RIGEDLNELS LMGETIGSMS MLGEALSSLS MLGESLSALS IVGESLGSLN ILGESLGSLN LMGDSLSSLS LMGDAVSNLS LMGEAVSNLS LLGEDLGPLN LLGEDLGPLN LLGEDLGPLN LLGEDLGPLS LLGEELSEMD LLGECLGSCT LLGEDLGNLG MRGEDLHGLN LLGEGLETCS LLGEDLGPLN MRGEELQGLN ---------MRGEDLQGLD  160 KELQNLEQQL LKELQNLEQQ LKELQNLEQQ LKELQYLEQQ IKELQNLEHQ YKELQHLEQK LKNLMAVEHA HKELMAIEEA HTELMAIEEA IQELRRLEDE YRKKVS-D-IDHLRVLEQN IDHLRVLEQN PKELRNLEGR VKELKSLEIR VKELKSLEIK FKELKNLEGR FKELKNLESR VKELKQVENR VKELKQLENR VKELKQLENR TKELEQLERQ SKELEQLERQ SKELEQLERQ TKELESLERQ VNELEHLERQ VEELQQIEQQ TMELDQLENQ IEELQQLEKA TDDLQQLENQ TKELEQLERQ IEELQQLEKV ---------LEELQKLEKI  195  Gene name >AP1 >PTAP1_1 >PTAP1_2 >M1 >M2 >M6 >PI >MADS_1 >MADS_2 >AP3 >PTD >PMADS30 >M4 >AG >PTAG1 >PTAG2 >SHP1 >SHP2 >STK >SCA19_12 >F2 >PTM3_PTM4 >SEP1 >SEP2 >SEP3 >SEP4 >PTM5 >PtMADS31 >PtMADS7 >PtMADS12 >PtMADS17 >PtMADS21 >PMADS27 >PtMADS82  170 DTALKHIRTR IDTALKLIRE IDTALKHIRA IDTALKRIRS IDSALKHVRS IDTALKSVRS IEHGLDKVRD LDTGLAAVRK LDAGLAAVRK MENTFKLVRE ---------MTEALNGVRG MTEALNGVRG LERSITRIRS LEKGISRIRS LEKGIGRIRS LEKGISRVRS LEKGISRVRS LEKAISRIRS LERGITRIRS LERGMTRIRS LESSLNQVRS LDGSLKQVRS LDGSLKQVRC LDSSLKQIRA VDASLRQIRS LERSVSTIRA LDSSLKQIRS LEVGLSRVLE LGRSLTRIRA LESSLNQVRS LEVGLCCVLE --TSICEILMEGSLRRLVE  180 KNQLMYESIN RKNHLMYQSI RKNHLMSQSI RKNQLIHESL RKNQL----RKNQLVHESL HQMEILISKKQMEFHSMLKQMEYHSMLRKFKSLGNQI KKN-ILPLQF RKYHVIKTQT RKYHVIKTQT KKNELLFSEI KKNELLFAEI KKNELLFAEI KKNELLVAEI KKHEMLVAEI KKHELLLVEI KKHELLLAEI KKHELLLAEI TKTQYMLDQL IKTQYMLDQL IKTQYMLDQL LRTQFMLDQL TKARSMLDQL RKNQVFKEQI RKGQFVLDEL TKGERIMNEI RKNQLFRERI TKTQYMLDQL TKGERIMNEI ---------EKGGKIINEI  190 ELQKKE---SELQIKE--SELQRKE--NELRKKE-----------AEMQKKE--R---RNE--E---QNE--E---QNE--ETTKKKN--HASK-----ETYKKKV--ETYKKKV--DYMQKRE--EYMQKRE--EYMQKRE--EYMQKRE--EYMQKRV--ENAQKRE--EYLQKRE--EYMQKRE--ADLQNKE--SDLQNKE--SDLQGKE--NDLQSKE--SDLKTKE--ELLRQKE--SELQRKE--STLERKG--EKLKGEE--ADLQNKE--STLERKG--LMLPMYE--DALKTKG---  196  

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