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Building the pollen wall : the role of tapetum transport proteins and sporopollenin translocation in… Quilichini, Teagen Danielle 2014

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BUILDING THE POLLEN WALL: THE ROLE OF TAPETUM TRANSPORT PROTEINS AND SPOROPOLLENIN TRANSLOCATION IN THE FORMATION OF THE SPECIALIZED CELL WALL ENCASING ARABIDOPSIS THALIANA POLLEN  by Teagen Danielle Quilichini  B.Sc., The University of Calgary, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)   March 2014  © Teagen Danielle Quilichini, 2014 ii  Abstract   A durable framework of sporopollenin and a lipid-rich pollen coat form the outer (exine) wall encasing pollen grains. The sporopollenin and pollen coat constituents of the exine are contributed by surrounding sporophytic tapetal cells by pre- and post-mortem events, respectively. The objective of this study was to investigate the transport and assembly of exine components from tapetal cells to developing pollen in the anthers of Arabidopsis thaliana. High-pressure freezing/freeze substitution and transmission electron microscopy revealed the sequence of developmental events in the anther that lead to sporopollenin deposition to form the exine and the dramatic differentiation and death of the tapetum, which produces the pollen coat. Through its tight co-expression with genes required for sporopollenin biosynthesis, a member of the ATP-binding cassette transporter superfamily, ABCG26, was found to be expressed preferentially in tapetal cells during early exine formation, and identified as a candidate gene encoding a sporopollenin export protein. Phenotypic abnormalities in the abcg26 mutant were first apparent in early uninucleate microspores as a lack of exine formation and sporopollenin deposition, and indicated that ABCG26 is required for normal male fertility, exine formation and pollen maturation. Further, two-photon microscopy of abcg26 anthers revealed large fluorescent vacuoles in tapetal cells with corresponding loss of fluorescence on microspores, consistent with abnormal ABCG26-mediated export activity in the mutant. These tapetum inclusions were not observed in double mutants of abcg26 and genes encoding the proposed sporopollenin polyketide biosynthetic metabolon (ACOS5, PKSA PKSB and TKPR1), providing a genetic link between transport by ABCG26 and polyketide biosynthesis. Genetic analysis also showed that hydroxycinnamoyl spermidines, known components of the pollen coat, were exported from tapeta prior to programmed cell death in the absence of polyketides, raising the possibility that they are incorporated into sporopollenin prior to pollen coat deposition. I propose a model where ABCG26-exported polyketides traffic from tapetal cells to form the sporopollenin backbone, in coordination with trafficking of additional constituents prior to tapetum programmed cell death.  iii  Preface   A version of Chapter 3 has been published as: Quilichini, T.D., Friedmann, M.C, Samuels, A.L., and Douglas, C.J. (2010). ATP-binding cassette transporter G26 is required for male fertility and pollen exine formation in Arabidopsis. Plant Physiol. 154: 678-690 © Copyright American Society of Plant Biologists, 2010 ( Text, figures and tables were reproduced with permission. Dr. Michael Friedmann and Prof. Carl Douglas identified the research question, and Dr. Friedmann performed in situ hybridization experiments (Figure 3.6) and initial abcg26 mutant analyses, including light microscopic analyses of chemically fixed bud samples. With Prof. Lacey Samuels’ assistance in designing optimized cryo-fixation methods for pollen wall analysis, Teagen Quilichini performed all other experiments, analyzed the data and prepared the manuscript. All authors provided valuable input towards the final manuscript.   The bulk of Chapters 1, 2 and 4 will be submitted for publication as a literature review and as two primary research articles, respectively, and are first-authored by Teagen Quilichini, with Prof. Carl Douglas and Prof. Lacey Samuels as co-authors. Figures in Chapter 1 are reproduced with permission (where noted), or were produced by Teagen Quilichini. Research questions and experiments in Chapters 2 and 4 were designed by the three co-authors and were performed by Teagen Quilichini. Undergraduate laboratory assistants Samantha Kang, Ada Roman and Hans Liu, under the supervision of Teagen Quilichini, performed genetic crosses and the extensive genotyping required for analyses presented in Chapter 4. Derrick Horne, Bradford Ross, and Dr. Kim Rensing provided technical assistance for the cryo-SEM images presented in Chapters 1, 2 and 4. Teagen Quilichini wrote these articles with the assistance of the co-authors.   For Chapter 5, Prof. Carl Douglas, Prof. Lacey Samuels and Teagen Quilichini identified the research question and experimental design. Under the supervision of Teagen Quilichini, Sheila Wang and Samantha Kang performed genotyping and SEM analyses. Lexcy Li performed complementary genetic and phenotypic analyses not included here. Teagen Quilichini performed all other experiments and analyzed the data. iv  Table of Contents  Abstract.......................................................................................................................................... ii	  Preface........................................................................................................................................... iii	  Table of Contents ......................................................................................................................... iv	  List of Tables ..................................................................................................................................x	  List of Figures............................................................................................................................... xi	  List of Symbols and Abbreviations .......................................................................................... xiv	  Acknowledgements .................................................................................................................... xix	  Chapter 1: Introduction ................................................................................................................1	  1.1	   Evolution of spores and pollen in the plant life cycle........................................................ 1	  1.2	   Morphological events in angiosperm pollen development ................................................ 1	  1.2.1	   Stages of pollen and pollen wall development in the model plant Arabidopsis thaliana ................................................................................................................................... 1	  1.2.2	   Structure and functions of the stratified pollen wall ................................................... 5	  1.3	   Composition and biosynthesis of the pollen exine ............................................................ 7	  1.3.1	   Priming for exine formation........................................................................................ 7	  1.3.2	   Sporopollenin composition: insights provided by biochemical analyses ................... 9	  1.3.3	   Sporopollenin composition: insights provided by genetic analyses ......................... 10	  1.3.4	   Model for sporopollenin biosynthesis and the sporopollenin metabolon ................. 16	  1.3.5	   Pollen coat (tryphine)................................................................................................ 17	  1.3.6	   The sporophytic contribution to the pollen wall ....................................................... 19	   The origin of the primexine ............................................................................... 20	  v	   The origin of the exine ....................................................................................... 21	  1.4	   Exine translocation from the tapetum .............................................................................. 23	  1.4.1	   Sporopollenin translocation and assembly................................................................ 24	  1.4.2	   Orbicules ................................................................................................................... 26	  1.5	   Candidate proteins for sporopollenin trafficking from tapetal cells ................................ 27	  1.5.1	   ATP-binding cassette transport proteins and the G subfamily ................................. 27	  1.5.2	   Lipid transfer proteins............................................................................................... 29	  1.6	   Research questions, objectives and significance of findings ........................................... 30	  Chapter 2: New views of tapetal cell structure and pollen exine development in Arabidopsis thaliana..........................................................................................................................................32	  2.1	   Introduction...................................................................................................................... 32	  2.2	   Results.............................................................................................................................. 34	  2.3	   Discussion ........................................................................................................................ 54	  2.4	   Materials and methods ..................................................................................................... 58	  2.4.1	   Plant growth .............................................................................................................. 58	  2.4.2	   cryo-scanning electron microscopy analysis ............................................................ 58	  2.4.3	   High-pressure freezing and freeze substitution......................................................... 59	  2.4.4	   Light microscopy and transmission electron microscopy......................................... 59	  Chapter 3: ATP-binding cassette transporter G26 is required for male fertility and pollen exine formation in Arabidopsis ..................................................................................................61	  3.1	   Introduction...................................................................................................................... 61	  3.2	   Results.............................................................................................................................. 64	  vi  3.2.1	   ABCG26 is required for male fertility....................................................................... 64	  3.2.2	   ABCG26 expression pattern ...................................................................................... 70	  3.2.3	   Microspore development is impaired in abcg26-1 ................................................... 72	  3.2.4	   Pollen wall and tapetum development in abcg26-1 .................................................. 75	  3.2.5	   YFP:ABCG26 is localized at the plasma membrane ................................................ 82	  3.3	   Discussion ........................................................................................................................ 84	  3.3.1	   The abcg26-1 mutant has partial Arabidopsis male fertility .................................... 85	  3.3.2	   Function of ABCG26 in the tapetum ........................................................................ 85	  3.3.3	   Locular inclusions in abcg26-1................................................................................. 87	  3.3.4	   Potential ABCG26 substrates ................................................................................... 88	  3.4	   Materials and methods ..................................................................................................... 89	  3.4.1	   Mutant isolation ........................................................................................................ 89	  3.4.2	   Plant growth .............................................................................................................. 90	  3.4.3	   Quantitative real-time PCR analysis ......................................................................... 91	  3.4.4	   In situ hybridization .................................................................................................. 92	  3.4.5	   Genetic complementation of the abcg26-1 mutant ................................................... 93	  3.4.6	   Scanning electron microscopy .................................................................................. 93	  3.4.7	   Transmission electron microscopy ........................................................................... 94	  3.4.8	   Subcellular localization of ABCG26 in planta ......................................................... 94	  Chapter 4: ABCG26-mediated polyketide trafficking and hydroxycinnamoyl spermidines contribute to pollen wall exine formation prior to tapetum programmed cell death in Arabidopsis thaliana .....................................................................................................................96	  vii  4.1	   Introduction...................................................................................................................... 96	  4.2	   Results.............................................................................................................................. 99	  4.2.1	   Tapetal cells of abcg26 anthers accumulate intrinsically fluorescent compounds in vacuole-like bodies ............................................................................................................... 99	  4.2.2	   The polyketide synthesis pathway is required for the accumulation of vacuolar inclusions in the tapetal cells of abcg26 mutants................................................................ 107	  4.2.3	   Discrete, extracellular orbicule-like bodies surrounding tapetal cells and microspores accumulate in key sporopollenin biosynthetic mutants ...................................................... 110	  4.2.4	   Role of phenylpropanoid and flavonoid biosynthetic enzymes in sporopollenin biosynthesis......................................................................................................................... 118	  4.2.5	   Polyketide product-dependent extracellular fluorescent bodies (ORBs) are hydroxycinnamoyl spermidines .......................................................................................... 122	  4.2.6	   HC spermidines do not contribute to autofluorescence of vacuolar inclusions in tapetal cells of the abcg26 mutant ...................................................................................... 124	  4.2.7	   The incorporation of HC spermidines in the pollen wall........................................ 126	  4.3	   Discussion ...................................................................................................................... 127	  4.3.1	   In planta characterization of the putative substrate of ABCG26............................ 128	  4.3.2	   The polyketide biosynthetic pathway and transport by ABCG26 are linked ......... 128	  4.3.3	   Orbicule-like bodies in polyketide mutants provide insight into trafficking of novel exine components................................................................................................................ 130	  4.3.4	   HC spermidines are exported from tapetal cells prior to programmed cell death and are incorporated into exine.................................................................................................. 130	  4.4	   Materials and methods ................................................................................................... 133	  viii  4.4.1	   Plant growth ............................................................................................................ 133	  4.4.2	   Two-photon laser scanning microscopy analysis ................................................... 133	  4.4.3	   Spectral analysis (lambda scans) ............................................................................ 134	  4.4.4	   Transmission electron microscopy analysis............................................................ 134	  4.4.5	   Acetolysis................................................................................................................ 135	  Chapter 5: The putative role of lipid transfer proteins in sporopollenin transport ............136	  5.1	   Introduction.................................................................................................................... 136	  5.2	   Results............................................................................................................................ 137	  5.2.1	   Lipid transfer protein candidate selection............................................................... 137	  5.2.2	   SALK_038995 heterozygous plants exhibit reduced fertility and pollen defects .. 139	  5.3	   Discussion ...................................................................................................................... 147	  5.4	   Materials and methods ................................................................................................... 149	   Critical point drying and SEM analysis ........................................................... 149	   Pollen stains ..................................................................................................... 150	  Chapter 6: Conclusions .............................................................................................................151	  6.1	   Major findings of this dissertation ................................................................................. 151	  6.2	   Questions arising from this research and future directions............................................ 153	  6.2.1	   What is the substrate of ABCG26? ......................................................................... 154	  6.2.2	   What is the dimerization partner(s) of ABCG26 exhibit in planta? ....................... 156	  6.2.3	   ABCG26 subcellular localization: tapetum polarized secretion? ........................... 157	  6.2.4	   Are hydroxycinnamoyl spermidines present in the sporopollenin biopolymer? .... 157	  ix  6.2.5	   How does sporopollenin traffic through the locule and assemble on the primexine of developing microspores? .................................................................................................... 158	  References ...................................................................................................................................160	   x  List of Tables  Table 3.1: Primer sequences used for PCR and RNA probe generation ...................................... 90	  Table 5.1: Arabidopsis thaliana LIPID TRANSFER PROTEIN (LTP) genes co-expressed with ACOS5......................................................................................................................................... 138	  Table 5.2: Sequences of primers used for PCR. ......................................................................... 138 xi  List of Figures  Figure 1.1: Stages of anther development in Arabidopsis thaliana................................................ 2	  Figure 1.2: Morphology and surface structure of mature pollen from Arabidopsis anthers........... 4	  Figure 1.3: Diagrammatic representation of the Arabidopsis pollen wall in cross-section ............ 7	  Figure 1.4: A model for the synthesis of sporopollenin in Arabidopsis ....................................... 17	  Figure 1.5: An immature Arabidopsis anther with all cell types present...................................... 20	  Figure 2.1: Overview of pollen development in the Arabidopsis anther ...................................... 37	  Figure 2.2: Pollen morphology through microsporogenesis and microgametogenesis ................ 43 Figure 2.5: The binucleate tapetum in Arabidopsis ...................................................................... 46	  Figure 2.6:  Tapetum development during exine formation in cryo-fixed Arabidopsis anthers... 48	  Figure 2.7: Tapetum development during pollen coat formation in cryo-fixed Arabidopsis anthers....................................................................................................................................................... 49	  Figure 2.8:  Late tapetum organelle ultrastructure........................................................................ 51	  Figure 2.9: The mature Arabidopsis pollen wall .......................................................................... 53	  Figure 3.1: Identification of Arabidopsis ABCG26 T-DNA insertion alleles .............................. 65	  Figure 3.2: Phenotypic comparison of Arabidopsis T-DNA insertion alleles .............................. 69 Figure 3.5: Accumulation of ABCG26.1 and ABCG26.2 transcripts in abcg26-1 mutants expressing the ABCG26.2 transgene............................................................................................. 70	  Figure 3.6: ABCG26 tissue-specific expression pattern during microspore development........... 71	  Figure 3.7: Key stages in Arabidopsis pollen wall development relative to ABCG26 expression........................................................................................................................................................ 73	  Figure 3.8: Microspore development in wild-type (Col-0) and abcg26-1 mutant anthers ........... 74	  xii  Figure 3.9: Pollen wall structure in wild-type (Col-0) and abcg26-1 mutant plants .................... 76	  Figure 3.10: Transmission electron micrographs of microspore primexine and exine formation in wild-type (Col-0) and abcg26-1 anthers ....................................................................................... 80	  Figure 3.11: Transmission electron micrographs of tapetum ultrastructure in wild-type and abcg26-1 anthers ........................................................................................................................... 81	  Figure 3.12: Transmission electron micrographs of locular inclusions in abcg26-1 mutant anthers at the uninucleate stage of development ....................................................................................... 82	  Figure 3.13: Subcellular localization of an EYFP:ABCG26 fusion protein in Arabidopsis mesophyll protoplasts ................................................................................................................... 83	  Figure 4.1: Vacuolar inclusions are present in abcg26 mutant tapetal cells............................... 101	  Figure 4.2: Variation in the severity of the abcg26 anther phenotype........................................ 103	  Figure 4.3: Controls for the timed Nile Red staining of anthers................................................. 105	  Figure 4.4: abcg26 tapetum vacuole cores stain with Nile Red ................................................. 110 Figure 4.7: acos5 mutant exhibits tapetum-associated autofluorescent bodies, not observed in wild-type ..................................................................................................................................... 112	  Figure 4.8: cyp703a2, pksa pksb, and tkpr1 mutants exhibit tapetum-associated autofluorescent bodies, not observed in wild-type ............................................................................................... 113	  Figure 4.9: Ultrastructure of extracellular orbicule-like bodies in acos5 and abcg26 acos5 mutant anther locules .............................................................................................................................. 116	  Figure 4.10: Extracellular bodies are observed in pksa pksb and tkpr1 anther locules .............. 116	  Figure 4.11: Extracellular autofluorescent bodies in acos5 anther locules do not stain with Nile Red .............................................................................................................................................. 117	  Figure 4.12: Microspore emission from traced regions of interest in developing anthers ......... 120	  xiii  Figure 4.13: Spectral emission profiles for wild-type, 4cl3, tt4, sht, and fah1 mutant microspores within developing anthers and prior to tapetal cell death ........................................................... 121	  Figure 4.14: Hydroxycinnamoyl spermidines constitute orbicule-like bodies in acos5............. 122	  Figure 4.15: The acos5 4cl3 and acos5 tt4 double mutants exhibit the same phenotype as the acos5 single mutant..................................................................................................................... 124	  Figure 4.16: abcg26 and sht abcg26 tapetum vacuolar inclusions appear indistinguishable by two-photon microscopy and TEM .............................................................................................. 126	  Figure 4.17: The chemical recalcitrance of the sporopollenin polymer in wild-type and sht mutant pollen is indistinguishable .............................................................................................. 127	  Figure 5.1: Identification of Arabidopsis LTPH1 T-DNA insertion alleles ............................... 140	  Figure 5.2: Pollen and pollen exine structure in wild-type and SALK 038995 heterozygous mutant plants ............................................................................................................................... 142	  Figure 5.3: Microspore development in wild-type and SALK 038995 heterozygous mutant anthers ......................................................................................................................................... 145	  Figure 5.4: Anther and pollen morphology at the bicellular pollen stage in SALK 038995 heterozygous mutant anthers....................................................................................................... 146	  Figure 5.5: Autophagic bodies in pollen from SALK 038995 heterozygous mutant plants....... 147	   xiv  List of Symbols and Abbreviations  ∆   Delta 1°P   Primary parietal cell layer  1°Sp    Primary sporogenous cell layer 2°P    Secondary parietal cell layer 2-P   Two-photon 4CL3   4-Coumarate:CoA ligase 5H   5-hydroxyguaiacyl rich 7D   7-day-old seedling α   Alpha ABC   ATP-binding cassette ABCG   ATP-binding cassette transporter, subfamily G ACOS   Acyl-CoA Synthase ACP   Acyl carrier protein AMS   aborted microspores Ar   Archesporial cell ATA   Arabidopsis thaliana anther ATP   Adenosine triphosphate bp   Base pair °C   Degrees Celsius C   Callose CALS   Callose synthase CAMV 35S  Cauliflower mosaic virus promoter CCR   Cinnamoyl-CoA reductase cDNA   complementary DNA CER   Eceriferum Co   Connective cell CoA   Coenzyme A Col-0   Columbia-0 xv  CT   Threshold cycles CYP   Cytochrome P450 DET   De-etiolated DEX   Defective exine DNA   Deoxyribonucleic acid DRL   Dihydroflavonol Reductase-Like e.g.   exempli gratia EDTA   Ethylenediaminetetraacetic acid E. coli   Escherichia coli El   Elaioplast En   Endothecium Ep   Epidermis ER   Endoplasmic reticulum et al.    et alii Ex   Exine EYFP   Enhanced yellow fluorescent protein F5H   Ferulate 5-hydroxylase Fb   Fibrous bands FLWR   Flower FM4-64  N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino) phenyl)     hexatrienyl) pyridinium dibromide  g   gram GFP   Green fluorescent protein GPI   Glycosylphosphatidylinositol GRP   Glycine-rich protein HC   Hydroxycinnamoyl HCl   Hydrochloric acid HPF   High-pressure freeze i.e.   id est In   Intine xvi  k   Kile (prefix) - 10001 L    Litre L1-L3   Stamen primordia cell layers LAP   Less adherent pollen Lo   Locule LTP   Lipid transfer protein LTPG    Glycosylphosphatidylinositol-anchored lipid transfer protein μ    Micro (prefix) - 1000−2 m    Meters min   Minute M    Molar  MALDI  matrix-assisted laser desorption ionization MC   Meiotic cell MgCl2   Magnesium chloride ML   Middle layer MMC   Microspore mother cell MS   Male sterile MSp   Microspore Mt   Mitochondrion n   Nano (prefix) - 1000- N   Nucleus NAD(P)H  Reduced nicotinamide adenine dinucleotide (phosphate) NEF   No exine formation ORB   Orbicule-like body OX   Over-expression P   Pollen PAP   Plastid-lipid associated protein PC   Pollen coat PG   Pollen grain PM   Plasma membrane xvii  pro   Promoter PKS   Polyketide synthase PW   Primary wall REF   Reduced epidermal fluorescence rER   Rough endoplasmic reticulum RNA   Ribonucleic acid rPE   Remnant primexine rPW   Remnant primary wall rT   Remnant tapetum RT-PCR  Reverse transcription polymerase chain reaction   s   Seconds SD   Standard deviation SE   Standard error SEM   Scanning electron microscopy SHT   Spermidine hydroxycinnamoyl transferase SIMS   secondary ionization mass spectrometry Sm   Septum Sp   Sporogenous cells St   Stomium StR   Stomium region T   Tapetum T-DNA  Transfer-deoxyribonucleic acid TAG   Triacylglycerol Td(s)   Tetrad(s) TDE   Transient defective exine TEM   Transmission electron microscopy TKPR   Tetraketide α-pyrone synthase  TMD   Transmembrane domain ToF   time-of-flight Tris    Tris(hydroxymethyl)aminomethane  xviii  Ts   Tapetosome TT4   Transparent testa4 UPEX    Uneven pattern of exine v   Volume V   Vacuole VR   Vascular region w   Weight WBC   White brown complex YFP   yellow fluorescent protein YL   Young leaf YS   Young stem xix  Acknowledgements   I owe my deepest gratitude to Prof. Carl Douglas and Prof. Lacey Samuels. It has been an honour to be supervised and mentored by such a talented pair of scientists. To Carl, the great adventurer, I have learned so much under your kind and patient supervision and I feel privileged to have had the opportunity to work with you. Thank you for your extraordinary leadership and encouragement every step of the way. To Lacey, the master mentor, your energy, creativity and enthusiasm for science are inspirational. Thank you for keeping me challenged, positive and ambitious throughout this degree.   I owe a great deal of gratitude to many people who have supported my graduate research over the last five years. To my committee members, Prof. Ljerka Kunst and Prof. Reinhard Jetter, thank you for your support, constructive feedback, guidance throughout my program, and for your feedback on this thesis. I am grateful to Dr. Michael Friedmann for initiating a project on ABCG26, and for graciously allowing me to pursue research related to this gene for my graduate work. I’d also like to extend my heartfelt thanks to all members of the Friedmann family for making Vancouver feel more like home. I had the great opportunity to work with a number of outstanding students during my degree, including Sheila Wang, Samantha Kang, Ada Roman, Hans Liu, Joanna Majarreis, Joanna Urban, and Edric Chou. Thank you for the hard work that you contributed to my research projects and for giving me positive mentorship experiences. I’d like to thank our collaborator, Prof. Clint Chapple, who kindly hosted me at Purdue University, Lafayette Indiana. Also, a big thank you to Yi Li and Nick Anderson for being tremendous hosts. I greatly appreciate the data analysis assistance provided by Cuong Li, a PhD candidate at the University of Victoria, British Columbia and the Bindley Biosciences Centre staff, Purdue University. I am especially grateful for the hours of training, technical assistance, support and expertise provided by Derrick Horne, Brad Ross, Kevin Hodgson and Garnet Martens of the UBC BioImaging Facility.   I have been fortunate to have worked in two friendly, encouraging lab environments and will always be thankful for my fellow lab mates in the Douglas and Samuels labs. In particular, I want to thank previous members of the Douglas lab, Dr. Clarice de Azevedo Souza, Dr. Sung Soo Kim, and Dr. Etienne Grienenberger, whose work on sporopollenin biosynthesis made my xx  investigations into transport processes possible. I’d also like to thank Heather McFarlane, Etienne Grienenberger, Ryan Eng, Gabriel Levesque-Tremblay, Rebecca Smith, Mathias Schuetz, Albert Cairó, Yoshi Watanabe, Kim Carruthers, Caitlin Donnelly, Anika Benske, Miranda Meents, Grant McNair, Tegan Haslam, Eryang Li, and Yuanyuan Liu for their supportive friendships, trouble-shooting advice, and for giving me many happy graduate school memories. Finally, I want to extend gratitude to all members of the UBC Botany community, who have been amazing colleagues and have made my graduate student experience memorable.  Funding for this research and related travel opportunities was provided by an NSERC Postgraduate Scholarships (Masters and Doctoral), a UBC Four-Year Fellowship, a UBC Department of Botany Entrance Scholarship, and an NSERC CREATE “Working on Walls” training grant. Thank you to all members of the Working on Walls community for enhancing my graduate student experience.   Finally, I am forever thankful for the support and love of my family and friends. Tracy, thank you for years of laughter and friendship, no matter the distance. Shea, thank you for seeing my potential and for being a loyal brother and friend. Mom, you are my inspiration. I couldn’t have asked for a stronger, more talented and hard-working female role model and I am endlessly grateful to have you as my mother. Thank you for believing in me unconditionally. Graeme, every day I thank my lucky stars (and UBC Botany) that we met. Thank you for your enduring support and encouragement over the last five years. I’m looking forward to sharing many more adventures with you.   1 Chapter 1: Introduction  1.1 Evolution of spores and pollen in the plant life cycle    For the evolution of plant life on land, a number of adaptations were required to support terrestrial life and its inherent stresses (Cronk, 2009; Kenrick and Crane, 1997). Among these adaptations, external barriers encasing the dominant plant body and its reproductive offshoots were forefront in preventing desiccation and ensuring the transmission of genetic information (Wallace et al., 2011). The cycling between the sporophytic and gametophytic generations present in land plants is made possible by the reproductive cells they produce, namely the haploid spores produced by the sporophyte (through meiosis) and the haploid gametes produced by the gametophytes (through mitosis). However, these reproductive cells, such as the male spores of seed plants, often require prolonged survival as independent entities, necessitating the fortification of their cell walls with polymers capable of withstanding physical, chemical and biological stresses. Sporopollenin, a structurally robust biopolymer, served this critical function in the evolution of land plants by enveloping and protecting the spores of early branches of seedless plants and the pollen grains (microgametophytes) formed by the male reproductive organ, the anther, of seed producing plants (Wallace et al., 2011). Sporopollenin appears to be conserved in its properties and to have been critical in the evolution of land plants.  1.2 Morphological events in angiosperm pollen development   1.2.1 Stages of pollen and pollen wall development in the model plant Arabidopsis thaliana   Investigation of anther and microspore development by light and transmission electron microscopy (TEM) is the basis for much of our knowledge of pollen development and pollen wall formation, and provided starting points for understanding the biosynthetic mechanism of sporopollenin formation. Sanders et al. (1999) divided Arabidopsis thaliana (Arabidopsis) anther development into 14 stages based on distinct cellular events that begin with stamen primordia formation and conclude with anther dehiscence (Figure 1.1). In the first four stages of anther   2 primordia differentiation, periclinal divisions in archesporial cells produce primary sporogenous and parietal cells, which give rise to all anther cell types: the epidermis, endothecium, middle layer, tapetum, and microspore mother cells (MMCs, also known as microsporocytes). In stages 6 and 7, MMCs undergo two meiotic divisions, transforming from diploid mother cells into haploid uninucleate microspores. Throughout the meiotic divisions, callose (β-1,3-glucan) forms a transient wall between the plasma membrane and primary cell wall of microspores, facilitating meiotic synchrony and the formation of a new, specialized primexine wall (that is not visible by light microscopy, see section 1.3.1 for further information). By stage 8, callose dissolution, presumably due to the secretion of tapetum-derived callase, releases free uninucleate microspores from tetrads, marking the beginning of exine proper formation (Stieglitz, 1977).   Figure 1.1: Stages of anther development in Arabidopsis thaliana. Light microscopy of transverse sections through chemically-fixed and embedded anthers. Anther stages are numbered (1 to 14) and tissues present are labeled. Ar, archesporial cell; Co, connective tissue; Ep, epidermis; En, endothecium; Fb, fibrous bands; L1-L3, stamen primordia cell layers; MC, meiotic cell; ML, middle layer; MMC, microspore mother cells; MSp, microspores; PG, pollen grains; 1°P, primary parietal layer; 2°P, secondary parietal cell layers; 1°Sp, primary sporogenous layer; Sm, septum; Sp, sporogenous cells; St, stomium; StR, stomium region; T, tapetum; Tds, tetrads; VR, vascular region. Bar over stage 1 = 25 μm (applies to stages 1-4). Bar over stage 6 = 25 μm (applies to stages 5-8). Bar over stage 9 = 50 μm (applies to stages 9-14). Reproduced with permission from Sanders et al., 1999; Copyright © 1999, Springer-Verlag.       3  After the release and maturation of uninucleate microspores in the anther locule, two microspore mitotic divisions transform the microspores into tricellular pollen, while the tapetal cells degenerate (anther stage 11). The first mitotic division occurs asymmetrically, yielding bicellular pollen grains consisting of a peripheral generative cell surrounded by a vegetative cell. The second division occurs in the generative cell and forms the male germ unit (or sperm cells) (McCormick, 1993). In many angiosperms, the second mitotic division occurs after pollen tube germination, however, in Arabidopsis, the tricellular pollen forms prior to anther anthesis (Edlund, 2004). Also during anther stage 11, the tapetal cells degenerate, and fibrous bands of thickenings in the endothecium and connective cells become visible. In insect-pollinated species, tapetal cell death releases pollen coat components that are deposited on and within the crevices of the exine, completing the formation of the pollen wall (Blackmore et al., 2007). The intine, pecto-cellulosic in composition, forms late in pollen development between the exine and the pollen plasma membrane as the pollen grains become engorged with starch granules and oil bodies (Ariizumi and Toriyama, 2011).   Anther dehiscence releases pollen, the mature male gametophyte, into the environment (Figure 1.2A,B). Pollen that successfully reaches a compatible stigma will hydrate, and the vegetative cell will extend a tube that grows through the style and toward the ovule, by intrusive tip growth. After degeneration of the synergid, the pollen tube ruptures to release two sperm cells into the embryo sac for double fertilization and the lifespan of the pollen grain is complete.     4  Figure 1.2: Morphology and surface structure of mature pollen from Arabidopsis anthers. Scanning electron micrographs showing the reproductive organs of a bisexual Arabidopsis flower, after sepal and petal removal (A), with increasing magnification on the pollen in panels B, C and D, respectively. A) Anther dehiscence has released mature pollen grains, seen in abundance on the anther surfaces, stigma and outer carpel wall. B) Magnification from panel A, showing pollen grains on the epidermal cells of the carpel.  C) Critical point dried pollen grain with reticulate exine and one visible aperture (of three). D) Surface structure of mature pollen grain, showing reticulate pattern of sporopollenin created by tecta. Bars = 100 μm (A), 20 μm (B), 2 μm (C), 1 μm (D).     5 1.2.2 Structure and functions of the stratified pollen wall    In considering the central importance of the male spore in plant sexual reproduction, together with the inherent stresses of a terrestrial environment, it is not surprising that the evolution of a tough wall surrounding these vulnerable structures would be an adaptive advantage for plants. Indeed, the outer wall of spores/pollen are assembled into architectural masterpieces of extraordinary strength and chemical inertness (Scott, 1994). It is the sporopollenin component of the exine that gives spores/pollen their characteristic resistance to terrestrial stresses. However, the spore and pollen wall functions reach beyond protective armour, as suggested by the diversity of surface features they exhibit.   The surfaces of pollen grains often appear intricately decorated due to species-specific sculpturing of their outer walls (Figure 1.2C,D). Such features have presumably adapted to hold additional wall materials such as tapetum-derived pollen coat constituents that are important for the physiology of the pollen in its interaction with the stigma. Apertures or colpi intercept the decorated outer wall of pollen in species-specific locations and frequencies, and serve as the typical sites of pollen tube emergence (Figure 1.2C). Spore/pollen surface features have also facilitated plant identification in the fossil record, aiding our understanding of the evolution and distribution of land plants over geological time.   Despite the great diversity of pollen surface structure, spore/pollen walls exhibit common features in cross-section, typically consisting of an inner intine composed of pectin, cellulose, and hemicellulose, and an outer exine composed of sporopollenin (Heslop-Harrison, 1968a). The intine appears as a light band by TEM, directly external to the pollen plasma membrane and is produced by the vegetative cell (Hess, 1993). The intine maintains the structural integrity of pollen grains, as Arabidopsis plants with mutations in primary cell wall cellulose synthases produce collapsed or malformed pollen grains with aberrant pollen walls that lack or have uneven intine cellulose (Persson et al., 2007). After pollen tube emergence from the exine shell, the intine serves as the only cell wall encasing the growing pollen tube, and is rapidly remodeled to assist growth while preventing premature rupture (Chebli et al., 2012). Mutations affecting the integrity of the intine have been shown to produce pollen grains unable to grow pollen tubes (Persson et al., 2007). There is also evidence for the presence of hydrolytic enzymes and antigens   6 required for compatible (or incompatible) stigmatic interactions, held within the intine (Knox and Heslop-Harrison, 1970; 1971).  Sporopollenin provides the rigid and sculptured framework of the exine, which serves to encapsulate and protect the contents of spores/pollen, and to assist in stigmatic capture (Figure 1.3). For many species, including Arabidopsis, this structured backbone is additionally covered by a pollen coat (or tryphine), which primarily serves in pollen stigmatic adhesion, recognition, and hydration and can be extracted from the underlying sporopollenin with organic solvents (Edlund, 2004; Murphy, 2006; Piffanelli et al., 1998). The Arabidopsis pollen coat contains a mixture of lipids, proteins, carotenoids, flavonoids, and hydroxycinnamoyl spermidines, providing diverse functions in support of pollen survival and germination (Preuss et al., 1993; Hülskamp et al., 1995; Mayfield, 2001; Mayfield and Preuss, 2000; Piffanelli et al., 1998; Grienenberger et al., 2009). In Arabidopsis, proteinaceous coat constituents are necessary for recognition by the stigma and, together with lipids, contribute to stigmatic adhesion after pollen capture (Edlund, 2004; Zinkl et al., 1999). Pollen coat lipids prevent water loss, facilitate pollen hydration on the stigma and, when applicable, lipid-derived volatiles and pigmentation primarily from carotenoids and flavonoids provide olfactory and visual cues, respectively, to pollinators and protect against oxidative damage and UV radiation (Edlund, 2004). However, pollen hydration mechanisms show dramatic variation among species and can be categorized broadly by stigmatic surface features. In dry stigma species, such as Arabidopsis, the stigma either lacks or is covered by a thin layer of exudate (or pellicle) on a continuous cuticle that requires penetration for pollen tube growth (Elleman et al., 1992; Zinkl et al., 1999). In these species, the pollen coat carries many of the necessary lipids and proteins that mediate the highly specific pollen recognition and hydration processes. However, pollen hydration on wet stigma plants, such as tobacco, petunia and lily, is largely unregulated and facilitated by the large volume of stigmatic exudates rather than pollen coat constituents.     7  Figure 1.3: Diagrammatic representation of the Arabidopsis pollen wall in cross-section. Layers of the pollen wall exist outside the pollen cytoplasm and are directly appressed to the plasma membrane. Layers of the pollen wall are broadly categorized as intine and exine. Within the outer exine wall, nexine is a thin layer of wall neighbouring the intine, and the sexine forms the outer sculptured component. Baculae and tecta form the sculptured components of the sexine, which are encased by the pollen coat. Based on Suzuki et al., 2008 and reproduced with permission; Copyright the author © 2008 Oxford Journals.   1.3 Composition and biosynthesis of the pollen exine  1.3.1 Priming for exine formation    The formation of the sculptured pollen wall requires the establishment of a primexine wall early on in microspore development. Based primarily on staining experiments, the primexine is thought to be predominantly cellulosic in composition and forms on the surface of microspores in the tetrad stage (Heslop-Harrison, 1968c; Paxson-Sowders et al., 1997). Within the primexine matrix, structured components known as probaculae and protecta emerge at regular intervals along an undulating microspore plasma membrane and are required for the patterning, deposition and structure of sporopollenin constituents of the exine. In the subsequent anther stage of development, early free microspores rapidly form a sporopollenin-based outer exine wall, guided by probaculae and protecta and taking the shape of structured pillars (baculae) and caps (tecta), respectively (Paxson-Sowders et al., 1997)   8  Prior to the formation of primexine, a transient callose wall forms between the plasma membrane and primary cell wall of each microspore. Several functions for the callose surrounding early microspores have been postulated, including aiding meiotic synchrony, functioning as a molecular filter between the tapetum and microspores, creating a barrier for the prevention of microspore fusion, and providing glucose to free microspores upon its digestion by callase (Ariizumi and Toriyama, 2011; Dong et al., 2005). Callose also serves to separate the primary cell wall from the MMCs to make room for the formation of a new, specialized wall, the primexine (Heslop-Harrison, 1968c). CALLOSE SYNTHASE5 (CALS5) plays an important role in the formation of callose around early microspores in Arabidopsis (Nishikawa et al., 2005; Dong et al., 2005). In mutants with decreased CALS5 expression, such as cals5 and ruptured pollen grain1 (rpg1), the callose around MMCs and tetrads is reduced, and microspores lacking sculptured sporopollenin baculae and tecta tend to abort prematurely (Nishikawa et al., 2005; Dong et al., 2005; Guan et al., 2008; Sun et al., 2013). Callose is also required for protecta formation on probaculae, but not for the positioning of probaculae, suggesting that the boundary formed by callose serves as a platform for protectum formation (Worrall et al., 1992). These data support a crucial role for callose in early stages of exine formation and subsequent pollen development in Arabidopsis.   The primexine is composed primarily of polysaccharides and is believed to provide the anchoring site for sporopollenin (Heslop-Harrison, 1968c; Ariizumi and Toriyama, 2011). In the 1980s, centrifugation experiments with early-meiosis phase lily MMCs demonstrated the importance of the microspore plasma membrane and positioning of the microspore’s cytoskeleton, endoplasmic reticulum (ER) and vesicles in the formation of primexine and apertures, as microspores with disrupted cytoskeleton and ER had abnormal patterning, disruptions in baculae positioning and abnormal apertures (Sheldon and Dickinson, 1983; Dickinson and Sheldon, 1986). In support of these findings, genetic analyses have identified genes required for normal primexine formation and plasma membrane undulation in Arabidopsis, including DEFECTIVE EXINE1 (DEX1) (Paxson-Sowders et al., 2001), NO EXINE FORMATION1 (NEF1) (Ariizumi et al., 2004), NO PRIMEXINE AND PLASMA MEMBRANE UNDULATION (NPU) (Chang et al., 2012) and RPG1 (Guan et al., 2008; Sun et al., 2013), and mutations in these genes result in microspores with defective probaculae formation and   9 positioning. These data indicate that together, the primexine, plasma membrane and callose wall contribute to exine wall deposition, sculpting and patterning (Suzuki et al., 2008; Dong et al., 2005). As the primexine function appears to be intimately linked with sporopollenin anchoring and exine patterning, primexine mutants are further reviewed in the discussion of sporopollenin translocation and assembly (see section 1.4.1).   1.3.2 Sporopollenin composition: insights provided by biochemical analyses   Despite the importance of the sporopollenin biopolymer in plant life, our understanding of its composition, structure, biosynthesis, transport and polymerization has remained largely incomplete. In the late 1960s, the predominant model for sporopollenin proposed a monomeric backbone of polymerized carotenoids, based primarily on their known accumulation in anthers correlated with pollen formation, and their ability to polymerize in vitro into insoluble polymers resembling sporopollenin (Brooks and Shaw, 1968). One of the first studies to indicate that the carotenoid model for sporopollenin composition required revision found no substantial decrease in sporopollenin accumulation after the application of different carotenoid synthesis inhibitors, norflurazon and Sandoz 9789, to developing Curcubita pepo anthers (Prahl et al., 1986; 1985). Additionally, labeling studies, primarily in developing Tulipa sp. anthers, found a substantial portion of radioactive phenylalanine incorporated in sporopollenin, indicating a role for phenolics in building this biopolymer (Prahl et al., 1986; Gubatz et al., 1993; 1992; Herminghaus et al., 1988). Since then, lipids have also been recognized for their role in sporopollenin formation, as treatment of Zea mays with a thiocarbamate herbicide inhibiting chain elongation during long-chain (C > 18) fatty acid biosynthesis altered sporopollenin chemistry, as determined by Fourier transform infrared spectroscopy (FTIR) (Wilmesmeier and Wiermann, 1995). Since these and other exploratory studies revealed the uncertainty in sporopollenin composition, new analytical techniques emerged and are continuously being developed to study sporopollenin, and models for the composition of this fascinating organic compound continue to be revised.  In an attempt to elucidate the composition and structure of sporopollenin, a number of biochemical approaches have been applied in the field of pollen biology to augment our knowledge of sporopollenin constituents. The challenge in elucidating the composition of   10 sporopollenin lies primarily in its recalcitrance to solubilization, which requires analysis of sporopollenin components remaining after acetic anhydride treatment, extended ozonolysis, nitrobenzene oxidation, potash fusion or aluminum iodide treatment (Southworth, 1974; Schulze Osthoff and Wiermann, 1987; Kawase and Takahashi, 1995). Gas-liquid chromatography coupled to mass spectrometry (MS) found fatty acids and oxygenated cinnamic acid derivatives to be the main components of sporopollenin, however, some argue that impurities have falsely been identified as components of sporopollenin (Van Bergen et al., 1995; 2004). Using pyrolysis-MS, ferulic acid and 4-coumaric acid constituents have been identified in sporopollenin (Rozema et al., 2001; Wehling et al., 1989). Inhibitor treatments, radiolabeling, and chemical breakdown studies have yielded complementary information on the constituents present in sporopollenin, which is typically accepted to contain phenolics and polyhydroxylated aliphatics, covalently coupled by ether and ester bonds (Guilford et al., 1988; Scott, 1994; Wiermann et al., 2005; Ahlers et al., 2000). Because the limited solubility of sporopollenin poses challenges to many chemical analyses, the application and optimization of polymer analyses could prove to be informative alternatives. Through the application of solid-state nuclear magnetic resonance (NMR), aromatic and aliphatic moieties as well as oxygen functionalities have been identified in the sporopollenin biopolymer (Guilford et al., 1988; Hemsley et al., 1992; Ahlers et al., 1999; 2000; 2003). The application of matrix-assisted laser desorption ionization time-of-flight MS for sporopollenin structure elucidation achieved limited success and will require matrix optimization (Moore et al., 2006). Altogether, data from chemical analyses have provided insight into the composition of sporopollenin and the cross-linking that exists within this biopolymer. However, as analyses employing harsh treatments to sporopollenin increase the likelihood of modified fragment release from the original biopolymer, our understanding of sporopollenin biochemistry requires further investigation.   1.3.3 Sporopollenin composition: insights provided by genetic analyses    The clearly established progression of cellular events in Arabidopsis anther and pollen development and the crucial role of pollen in sexual plant reproduction allow mutants with phenotypic changes in pollen wall development and fertility to be identified. Specifically,   11 numerous gene products have been implicated in sporopollenin biosynthesis and deposition, as mutations in their genes produce plants with defects in pollen wall formation immediately following tetrad release, at the time of high sporopollenin flux from the tapetum, often impairing male fertility.  Among these genes, a subset of transcriptional regulators with tapetum-preferential expression have clarified the importance of the sporophyte in sporopollenin formation and have assisted in uncovering the genetic networks required for exine development. MALE STERILITY 1 (MS1/HKM), encoding a plant homeodomain (PHD) motif-containing putative transcription factor, is required for tapetum development and exine formation through the apparent direct or indirect regulation of over 260 genes, primarily thought to function in exine component synthesis (Ito and Shinozaki, 2002; Ito et al., 2007; Yang et al., 2007; Wilson et al., 2001). The ABORTED MICROSPORES (AMS) gene encodes a basic helix-loop-helix transcription factor that plays a role similar to MS1 in tapetum and post-meiotic microspore development, with ams mutants displaying alterations in over 500 genes associated with pollen wall development and tapetum function (Xu et al., 2010). Additionally, AtMYB80/MYB103/MS188, an R2R3-type transcription factor, is required for the callose wall and exine formation as well as tapetum maturation in Arabidopsis (Zhang et al., 2007; Phan et al., 2011). MS1, AMS and MYB80 are expressed in tapetal cells at a time surrounding sporopollenin synthesis and deposition, and appear to be important regulatory points in the formation of the pollen wall. These regulators have been and will continue to serve as important tools in the identification of genes involved in processes related to sporopollenin synthesis, traffic and assembly.   Reverse genetic screens of candidate genes involved in anther and pollen development have identified a number of Arabidopsis genes required for sporopollenin synthesis and deposition whose protein products have been biochemically characterized. One of the first such studies identified ACYL-COA SYNTHETASE5 (ACOS5) as being critical for pollen development and sporopollenin biosynthesis. ACOS5 belongs to a novel class of enzymes related to, but functionally distinct from the phenylpropanoid enzyme 4-coumarate:CoA ligase (4CL). It was originally identified as a 4CL-like gene encoding an acyl-CoA synthetase of unknown function specifically expressed in developing anthers of Arabidopsis and with putative orthologs in other species (Souza et al., 2008). In vitro assays revealed the surprising preference   12 of recombinant ACOS5 enzyme for non-phenolic substrates, delineating a distinct function for this enzyme from true 4CLs involved in phenylpropanoid metabolism (Kienow et al., 2008; Souza et al., 2009). Recombinant ACOS5 has broad activity toward medium and long-chain fatty acids, including hydroxylated forms, and is thought to produce an acyl-CoA ester that is critical for the synthesis of sporopollenin. In support of this hypothesis, the acos5 mutant is male sterile and developing microspores show defects upon release from tetrads, failing to form exine and aborting shortly after the start of the free microspore stage (stage 9 of anther development). Furthermore, ACOS5 is specifically expressed in tapetal cells at a time corresponding to the appearance of sporopollenin on free microspores, consistent with a function in sporopollenin biosynthesis (Souza et al., 2009).   The identification of ACOS5 as a key enzyme in exine formation facilitated the identification of additional genes encoding enzymes with putative functions in sporopollenin biosynthesis, primarily through in silico co-expression analyses. Two such co-expressed chalcone synthase-like genes were found to function in exine formation, as mutations in these genes resulted in “less adherent pollen” (LAP) with abnormal exine patterning (Dobritsa et al., 2010). These enzymes, named LAP6 and LAP5, exhibited broad in vitro activity, performing malonyl-CoA condensations with 4-coumaroyl-CoA and fatty acyl-CoAs of varying chain lengths (Mizuuchi et al., 2008; Dobritsa et al., 2010). In vitro, these enzymes exhibited preference for medium length and hydroxylated fatty acyl-CoA esters (products of acyl-CoA synthetase and fatty acid hydroxylase enzymes), and through successive condensations with malonyl-CoA, yielded tri- and tetraketide α-pyrones. These data suggested that tri- and tetraketide α-pyrones were likely in planta products of the two enzymes, earning these enzymes the names POLYKETIDE SYNTHASE A (PKSA) and PKSB (Kim et al., 2010). In support of their putative role in sporopollenin precursor synthesis, PKSA and PKSB are expressed specifically in the tapetum at a time surrounding sporopollenin synthesis and deposition, and reverse genetic analysis showed that pksa pksb double mutants are male sterile and completely lack the formation of exine after tetrad release (Dobritsa et al., 2010; Kim et al., 2010).   Also co-expressed with ACOS5 were two oxidoreductase genes, DIHYROFLAVONOL REDUCTATASE LIKE1 (DRL1) and CINNAMOYL-COA REDUCTASE LIKE6 (CCRL Tang et al., 2009; Hamberger et al., 2007). The ability of the corresponding recombinant proteins to   13 reduce the ketone function of tetraketide α-pyrones (produced by PKSA and PKSB) in the presence of NADPH suggested that these enzymes work downstream of PKSA and PKSB, and they were thus renamed TETRAKETIDE α-PYRONE REDUCTASE1 (TKPR1) and TKPR2 (Grienenberger et al., 2010). Consistent with their hypothesized roles in sporopollenin biosynthesis, a strong male-sterile phenotype was observed for mutant alleles of TKPR1, (Tang et al., 2009; Grienenberger et al., 2010), which lacked or had severely perturbed exine. However, loss-of-function TKPR2 mutants had only minor perturbations in their exine walls and produced fertile plants (Grienenberger et al., 2010). Thus, despite the nearly identical substrate preferences observed for recombinant TKPR1 and TKPR2, their in vivo functions likely differ, possibly due to different timing of expression in tapetal cells over the course of microspore development (Grienenberger et al., 2010), or differences in subcellular localization (Grienenberger et al., 2010).  The numerous oxygen atoms in sporopollenin, which are commonly present in ether (and to a lesser extent ester) groups, support the prediction that high levels of covalent coupling among subunits results in the characteristic recalcitrance of sporopollenin (Ahlers et al., 2000). Molecular-genetic studies support this prediction, as two cytochrome P450 enzymes capable of hydroxylating the fatty acid constituents of predicted sporopollenin monomer precursors have been implicated in sporopollenin biosynthesis, thereby creating functional groups with polymerization and/or cross-linking potential.  These cytochrome P450 enzymes, CYP703A2/DEX2 and CYP704B1 catalyze the in-chain and ω-hydroxylation of fatty acids, with substrate chain lengths of C10-C14 and C16-18, respectively (Morant et al., 2007; Dobritsa et al., 2009b). In support of the predicted roles of CYP703A2 and CYP704B1 in sporopollenin formation, cyp703a2 mutants exhibit strong reductions in male fertility with pollen grains that lack exine (Morant et al., 2007), and cyp704b1 mutant pollen lacks normal exine (Dobritsa et al., 2009b), though is not sterile. Both genes are specifically expressed in developing anthers. Altogether, based on the above data, ACOS5, PKSA, PKSB, TKPR1, CYP703A2/DEX2, and CYP704B1 are proposed to sequentially synthesize polyhydroxylated long-chain α-pyrones that form critical sporopollenin constituents or precursors (Grienenberger et al., 2010).   Along with polyhydroxylated long-chain α-pyrones, fatty alcohols may also form a key subset of the aliphatic constituents of sporopollenin. MALE STERILITY2 was first identified as a   14 gene required for exine formation in Arabidopsis, based on the severe reduction in male fertility and loss of exine formation observed in ms2 mutants (Aarts et al., 1997). Despite contrasting reports on the fertility of ms2 mutants with different insertion alleles, all ms2 mutants characterized to date exhibit severe exine defects (Aarts et al., 1997; Dobritsa et al., 2009b). Based on sequence homology with characterized fatty acyl reductases (FARs), MS2 was predicted to function in the production of sporopollenin fatty alcohols by reducing fatty acyl-CoA substrates. Escherichia coli (E. coli) strains expressing MS2 formed C14:0, C16:0, and C18:1 alcohols from endogenous bacterial fatty acids (Doan et al., 2009). In a subsequent study on MS2, Chen et al. (2011) found the enzyme localized to the plastid. With this information, in vitro assays were repeated using purified recombinant MS2 and a range of substrates, including long-chain fatty acids esterified to acyl carrier protein (ACP). These assays found that in the presence of NAD(P)H, MS2 has strong preference for C16:0-ACP (or palmitoyl-ACP), producing C16:0 alcohol (Chen et al., 2011). Interestingly, these assays found no MS2 activity against acyl-CoA substrates, including C16 acyl-CoA, which serve as the preferred substrates for other known fatty acid reductases. These data suggest that MS2 functions as a plastid-localized fatty acyl-ACP reductase and its fatty alcohol products form sporopollenin constituents or precursors in Arabidopsis.   ECERIFERUM 3 (CER3/FLP1/WAX2/YRE), commonly called FACELESS POLLEN1 in the pollen literature, encodes a protein involved in alkane biosynthesis, although its specific biochemical function is not known (Bernard and Joubès, 2012). CER3 functions in alkane biosynthesis in stem and silique epidermal cells, as well as in the synthesis of pollen wall lipids, as cer3 mutant pollen exhibit increased coat covering and acetolysis-sensitive sporopollenin (Ariizumi et al., 2003; Chen et al., 2003; Kurata et al., 2003; Rowland et al., 2007). Although some cer mutants exhibit male fertility defects, this phenotype has typically been associated with alterations to pollen coat constituents, and fertility can be recovered if plants are grown under high humidity.  CER3 appears to have multiple functions in pollen wall lipid production as, in addition to acetolysis-sensitive pollen (suggesting altered sporopollenin), cer3 mutants exhibit altered lipid body morphology and abundance (suggesting altered pollen coat). Thus, alkanes may form additional aliphatic components of sporopollenin in Arabidopsis.   Beyond the aliphatic constituents of sporopollenin, one or more phenolic components in   15 this polymer have been long documented. These phenolics appear to be derived from hydroxycinnamic acids of the phenylpropanoid pathway, as mutants (such as reduced epidermal fluorescence 3-2) affecting C4H expression fail to develop pollen, and wall fluorescence in such mutants is abolished (Schilmiller et al., 2009). The importance of the phenylpropanoids in building the pollen exine was further demonstrated in the male sterility of a transgenic Arabidopsis line over-expressing FERULATE 5-HYDROXYLASE in a CAFFEIC ACID O-METHYLTRANSFERASE mutant (comt) background that produces high levels of an unusual 5-hydroxyguaiacyl (5H)-rich lignin (Weng et al., 2010). These 5H-rich plants produced pollen that appeared devoid of exine, suggesting that manipulations of the monolignol pathway could alter the flux of select phenolics into sporopollenin synthesis. Based on these data, the authors suggest that reductions in available p-coumaroyl-CoA and feruloyl-CoA, substrates for a spermidine hydroxycinnamoyl transferase (SHT) involved in the synthesis of pollen coat constituents, could cause the observed sporopollenin deficiencies (Weng et al., 2010). However, reductions of hydroxycinnamoyl spermidines, as in sht mutants, do not have fertility defects or major exine deficiencies (Grienenberger et al., 2009).  In contrast to the characterized gene products described above, a number of genes encoding proteins of unknown function are also required for normal exine formation in Arabidopsis, and the proteins encoded by these genes may participate in sporopollenin biosynthesis or deposition in Arabidopsis. These include DEFECTIVE IN EXINE PATTERNING 1 (DEX1), NO EXINE FORMATION1 (NEF1), LESS ADHERENT/ADHESIVE POLLEN 3 (LAP3), and TRANSIENT DEFECTIVE EXINE (TDE1/DET2) (Paxson-Sowders et al., 2001; Ariizumi et al., 2004; Dobritsa et al., 2009a; Rowland et al., 2007; Ariizumi et al., 2008). For extensive reviews on the putative roles of these genes in sporopollenin synthesis and/or deposition, the reader is referred to recent reviews (Blackmore et al., 2007; Ariizumi and Toriyama, 2011). Additionally, two genetic screens employed in Arabidopsis with the goal of finding novel genes required for pollen wall formation identified 12 KAONASHI genes (meaning “faceless” in Japanese), and 14 genes required for exine formation (Suzuki et al., 2008; Dobritsa et al., 2011). The application of forward and reverse genetics, molecular biology and biochemical analyses has enabled the identification and characterization of numerous genes, encoding proteins and enzymes required for exine formation. Together, these studies have   16 provided clues regarding the mechanism of sporopollenin biosynthesis, particularly pertaining to its aliphatic constituents, and have defined a key role for the tapetum in pollen wall formation.   Despite our limited understanding of sporopollenin composition, structure and biosynthesis, this extracellular matrix is thought to be conserved in its composition among all land plants whether it is present in the spores of early diverging land plants, or in the pollen of seed-bearing trees and flowering species. Comparative phylogenetic and genomic analyses support this hypothesis, as putative orthologs of ACOS5, PKSA, PKSB, CYP703A2 and CYP704B1 are broadly distributed in flowering plants, and early diverging Physcomitrella patens.  They are absent from Chlamydomonas reinhardtii and thus support a key role for these genes in the progression of plant life onto land (Souza et al., 2009; Colpitts et al., 2011; Morant et al., 2007; Dobritsa et al., 2009b; Koduri et al., 2010; Ríos et al., 2013).  1.3.4 Model for sporopollenin biosynthesis and the sporopollenin metabolon   With the insights provided by molecular genetic and biochemical studies, sporopollenin has most recently been suggested to contain hydroxylated tetraketide α-pyrones, based on the synthesis of these components by the successive actions of recombinant sporopollenin biosynthetic enzymes ACOS5, PKSA, PKSB, and TKPR1 (Grienenberger et al., 2010). In the proposed model, the ACOS5-derived fatty acyl-CoA is a central precursor required for the synthesis of sporopollenin monomers in the tapetum, and serves as a substrate for PKSA and PKSB, which generate a tetraketide that is subsequently reduced by TKPR1. The fatty acid precursors (or possibly the tetraketides produced downstream of PKSA and PKSB) are additionally hydroxylated by CYP703A2 and CYP704B1, and the polyhydroxylated tetraketide sporopollenin precursor serves as a critical aliphatic component or monomer of sporopollenin (Figure 1.4). This hydroxytetraketide sporopollenin monomer is then exported into the anther locule by an unknown mechanism for polymerization into the pollen exine or further modified by unknown enzymes, prior to its export (Figure 1.4). In support of this model, ACOS5, PKSA, PKSB and TKPR1 interact in pairwise combinations and co-localize at the ER, and may form the core of a sporopollenin metabolon, facilitating efficient sporopollenin synthesis in Arabidopsis tapetal cells (Lallemand et al., 2013).   17   Figure 1.4: A model for the synthesis of sporopollenin in Arabidopsis. Two putative pathways for the synthesis of tetraketide α-pyrones are presented. The synthesis of fatty acids in tapetum plastids produces substrates for CoA esterification by ACOS5. In pathway 1, the variable-length CoA esters serve as substrates for condensation with two or three malonyl-CoAs producing triketide or tetraketide α-pyrones, respectively, in a reaction catalyzed by polyketide synthase enzymes (PKSA and PKSB). These products may be hydroxylated by cytochrome P450 enzymes (CYP703A2 and CYP704B1) prior to the specific reduction of the tetraketide α-pyrone at the carbonyl function by tetraketide α-pyrone reductases (TKPR1 and TKPR2). In pathway 2, polyhydroxylation of the fatty acyl-CoA esters occur prior to PKS-catalyzed condensations with malonyl-CoAs and TKPR-catalyzed reduction of the tetraketide α-pyrone carbonyl function. The polyhydroxylated tetraketide α-pyrone product is putatively exported from the tapetum by an ATP-binding cassette transport protein (such as ABCG26) or by lipid transfer proteins (LTPs), to directly become a sporopollenin building block, or after further biochemical modification.  Reproduced with permission from Grienenberger et al., 2010; Copyright American Society of Plant Biologists © 2010 (  1.3.5 Pollen coat (tryphine)    The pollen coat is a generic term given to the waxy or oily covering on the scaffold created by sporopollenin, and which typically constitutes a mixture of lipids, proteins and other metabolites. This coating can be minimal, as in anemophilous (wind-pollinated) species, or substantial and adhesive as in entomophilous (insect-pollinated) species such as Arabidopsis. In angiosperms, two types of pollen coats are recognized: the pollenkitt, which is nearly universally   18 present in entomophilous species, and tryphine, which to date is known only in Brassicaceae and represents more heterogeneous mixtures contributed by tapetal cells (Pacini and Hesse, 2005). Tryphine, referred to herein as the pollen coat, will be the focus of this review, as it has been extensively studied in Arabidopsis.   In contrast to the poorly understood composition of sporopollenin, the pollen coat layer of many pollen exines has been more thoroughly characterized, facilitated by its ease of extraction with non-polar organic solvents (Doughty et al., 1993). Lipid analyses on Brassicaceae pollen coats have identified a mixture of primarily neutral-ester lipids, free fatty acids, very-long-chain wax esters, volatile lipid derivatives, flavonoids, alkanes and hydroxycinnamoyl spermidines (Piffanelli et al., 1998; Hsieh and Huang, 2007; Grienenberger et al., 2009). From genetic studies, a number of mutants with altered pollen coat lipids, such as the eceriferum (cer) mutants with altered very long-chain acyl lipid synthesis, have provided the opportunity to investigate the function of the coat in plant reproduction (Preuss et al., 1993; Hülskamp et al., 1995; Koornneef et al., 1989). The cer mutants are characterized by a glossy, wax-deficient stem, but in the case of mutants of CER3/WAX2/YRE/FLP1 (Rowland et al., 2007; Chen et al., 2003; Ariizumi et al., 2003; Kurata et al., 2003), CER6/CUT1/POP1/KCS6 (Fiebig et al., 2000; Millar et al., 1999; Preuss et al., 1993) and ABCG11/WBC11/DESPERADO (Bird et al., 2007; Panikashvili et al., 2007), defects also extend to the pollen coat. Mutations altering pollen coat lipid constituents typically exhibit conditional male-sterile phenotypes, which can be recovered under high humidity and suggest a function for certain pollen coat lipids in pollen hydration (Preuss et al., 1993).    The pollen coat also contains a number of proteins, the majority of which are classified as either processed oleosin proteins or self-incompatibility proteins (Piffanelli and Murphy, 1998; Kim, 2002; Kim and Huang, 2003). Although the precise role of the oleosin proteins in the pollen coat of Brassicaceae species remains to be determined, their highly basic nature and divergence from seed oleosins have suggested that they play structural roles similar to other cell wall-associated proteins, and may assist in channeling water between the stigma and pollen grain (Dickinson, 1995; Ross and Murphy, 1996; Huang et al., 2013a).     19 1.3.6 The sporophytic contribution to the pollen wall   The different layers of the pollen wall are deposited at different stages of anther development and are contributed by different cells within the anther. Generally, components of the exine are synthesized by the surrounding sporophytic tapetal cells (see section 1.4), while components of the intine are generated by the microspores. Although both sporopollenin and pollen coat constituents of the exine arise in tapetal cells, the timing of their deposition and release differ; sporopollenin is exported from intact tapeta (Figure 1.5) and the pollen coat is deposited as tapetal cells disintegrate (see section 1.4.3 for details on pollen coat constituents as they exist in tapetal cells). Thus, many exine-related studies require methods of exposing immature anther sporophytic tissues, often by embedding and sectioning anthers, or by cryo-fracturing anthers as shown below, to allow the ultrastructural features of tapetal cells to be examined (Figure 1.5).    20  Figure 1.5: An immature Arabidopsis anther with all cell types present. Cryo-SEM of cryo-fractured anther with all cell layers of the sporophytic anther wall present: epidermis (outermost), endothecium, middle layer, and tapetum (innermost). The sporophytic tissues surround a locule filled with developing microspores at the late uninucleate stage of development (anther stage 9). Ultrastructural features visible in these early microspores include an enlarged vacuole, nucleus with nucleolus, and exine radiating from each microspore plasma membrane. En, endothecium; Ep, epidermis; Ex, exine; Lo, locule; M, microspore; ML, middle layer; N, nucleolus; T, tapetum; V, microspore vacuole. Bar = 5 μm. The origin of the primexine   It has long been assumed that the barrier formed by callose prevents cell wall component transfer between the tapetum and developing microspores, and thus, primexine components must   21 be provided solely by the microspores. This view has been challenged by genetic studies, as homozygous mutants with primexine abnormalities show no defects in their respective heterozygous states, indicating that the sporophytic generation functions in primexine formation (reviewed by Ariizumi and Toriyama, 2011). Additionally, there is uncertainty regarding the first appearance of sporopollenin on the surface of microspores, and whether this sporopollenin is supplied by both microspores and tapetal cells, or uniquely by the tapetum. According to Heslop-Harrison (1968), early primexine is acetolysis-sensitive, while late primexine bearing developed probaculae in Lilium longiflorum becomes acetolysis-resistant, suggesting the presence of sporopollenin in the latter (Heslop-Harrison, 1968c). However, as the material fortifying the late-primexine walls of tetrads of microspores did not stain in a manner similar to mature sporopollenin, it was considered to differ from sporopollenin, and was termed protosporopollenin. Recently, a method to remove callose from Brassica rapa tetrads was developed to allow the early wall around microspores to be examined by SEM (Kirkpatrick and Owen, 2013). After callose digestion by cellulase, pectolyase and cytohelicase, the persistence of patterned microspore walls with probaculae and protecta suggested that sporopollenin may already be present on microspores in tetrads (Kirkpatrick and Owen, 2013). Interestingly, in Arabidopsis, tapetum-expressed genes encoding enzymes or proteins required for sporopollenin synthesis or accumulation, such as ACOS5, PKSA, PKSB, TKPR1 and ABCG26, show no apparent primexine defects, suggesting that tapetum-supplied sporopollenin is not present around callose-encased tetrads. Thus, further studies on the composition and origin of primexine constituents are of great importance to our understanding of pollen wall patterning and assembly. Future studies on the formation and composition of the primexine, particularly regarding the location of enzymes and proteins involved in its synthesis and anchoring to the microspore, will provide valuable insight into the processes governing pattern formation and sporopollenin trafficking for exine assembly. The origin of the exine    Most genes encoding enzymes or proteins implicated in outer pollen wall formation have expression patterns that appear specific to the sporophytic generation (Reviewed by Wallace et   22 al., 2011). In most cases, the expression of Arabidopsis sporopollenin-related genes has been localized to tapetal cells in coordination with sporopollenin biosynthesis and deposition, although some genes required for sporopollenin formation appear to have dual expression in the tapetum and microspore. For example, ACOS5, PKSA, PKSB, TKPR1, TKPR2, and MS2 transcripts are specifically detected in tapetal cells by in situ hybridization, and PKSA and PKSB proteins are present in the tapetum at comparable developmental stages as demonstrated by immunolocalization (Souza et al., 2009; Kim et al., 2010; Grienenberger et al., 2010; Aarts et al., 1997). Interestingly, the expression of these genes appears highly restricted to the stages surrounding sporopollenin deposition on microspores, suggesting that mechanisms for high efflux from intact tapetal cells are in place at these early stages of uninucleate microspore development. Similarly, characterized mutations in KNS4-10, CER3/FLP1/YRE/WAX2, CYP704B1, and NEF1 are recessive, and plants heterozygous for mutations in these genes produce 100% wild-type appearing microspores, suggesting that these sporopollenin-related genes function in sporophytic cells (Suzuki et al., 2008; Ariizumi et al., 2004). These data support the requirement for active translocation of sporopollenin precursors from sporophytic tapetal cells to microspores, prior to tapetum programmed cell death.   In addition to the tapetum-specific contribution to sporopollenin, the pollen coat represents a second major component of the outer pollen wall in Arabidopsis dependent on tapetal cells. These cells accumulate high levels of metabolites and proteins, primarily within specialized organelles called tapetosomes and elaioplasts, which are the sources of many pollen coat components. Tapetosomes are storage organelles that lack a visible outer membrane enclosure and are specific to the plant tapetum (Wu et al., 1997). Tapetosomes are ER-derived and produce a plethora of metabolites, including alkanes and triacylglycerol (TAG)-rich oil droplets, which are structurally maintained by oleosins and surrounded by vesicles associated with flavonoids (Hsieh and Huang, 2005; 2007). Elaioplasts are specialized plastids bound by an outer membrane and filled with numerous steryl ester-rich globuli (Wu et al., 1999). Like many other specialized plastids, the neutral lipids of elaioplasts are held within the globuli, maintained in part by plastid lipid-associated proteins (PAPs) (Kim et al., 2001). In the final stage of tapetum development, programmed cell death releases tapetum-stored materials into the locule. Interestingly, not all lipids or proteins detected in the tapetum are present in the pollen coat after   23 expulsion from tapetum elaioplasts and tapetosomes (Ting et al., 1998).  For example, tapetosome-derived oleosins, although present in the final pollen coat, are cleaved such that the C-terminus is present in the coat, while the abundant tapetosome TAGs appear absent, or minimally present in the pollen coat (Murphy and Ross, 1998; Wu et al., 1997). Thus, even amongst tapetum constituents destined for the pollen coat, different processes governing their modification, degradation, and transfer to pollen grains exist (Ting et al., 1998). In summary, the sporopollenin and pollen coat layers of the exine both move from their site of synthesis in tapetal cells to the surface of microspores/pollen grains by poorly understood mechanisms of active secretion and release after cell rupture and modification, respectively.    1.4 Exine translocation from the tapetum   Tapetal cells are derived from the L2 and L3 anther cell layers, forming the innermost cell layer of the sporophytic anther wall, and their direct proximity to developing microspores facilitates their nutritive role in pollen development (Goldberg et al., 1993). Tapeta in Spermatophytes are broadly grouped into the secretory (or parietal or glandular) type or the amoeboid (or periplasmodial or invasive) type, differing primarily in the extent of their intrusion into the locule during microspore development (Pacini, 2010). Amoeboid tapeta intrude into the locule, encasing microspores and providing direct nutrition to microspores, while secretory tapeta retain their shape at the locule periphery, requiring the transport of materials to microspores through the locule fluid (Pacini, 2010). In Arabidopsis, in which tapetal cells are of the secretory type, the sporopollenin-based wall around microspores forms prior to tapetum programmed cell death. After programmed cell death, tapetum lysis releases their cytoplasmic contents into the locule, and many of these components fill the exine crevices as pollen coat constituents. Different components of the pollen exine contributed by tapetal cells are incorporated into the pollen wall at different times in development and have different modes of transport from their sites of synthesis to their sites of accumulation or assembly in the pollen wall.     24 1.4.1 Sporopollenin translocation and assembly    As tapetal cells represent the major site of sporopollenin synthesis for most plant species, there is a need for rapid and efficient sporopollenin export precisely upon the release of free microspores, however the underlying molecular mechanisms supporting this high and rapid efflux are poorly understood. A number of proteins required for exine formation have been hypothesized to function in sporopollenin export from the tapetum, or in the translocation of sporopollenin constituents from secretory tapeta through the locule to the microspore surface. ATP-Binding Cassette (ABC) transport proteins are candidates for exporting sporopollenin from tapetal cells in Arabidopsis, and testing the hypothesis that ABCG26 is required for pollen wall development is the basis of Chapter 3. A number of lipid transfer protein (LTP) genes are expressed in the tapetum and have proposed roles as sporopollenin couriers, particularly through the locule fluid (see section 1.5.2). One rice LTP, OsC6, may function in sporopollenin trafficking, as mutants silencing OsC6 are impaired in exine development, resulting in reduced fertility (Zhang et al., 2010). Structures associated with the tapetum in some species, such as viscin threads (or strands bridging tapetal cells and microspores) and orbicules (see Section 1.4.2 below), also have proposed roles in the trafficking of sporopollenin or other microspore components, although their transport capabilities have yet to be demonstrated. Also, their absence in many species and persistence after exine formation is complete suggest they may have other functions (Rowley and Morbelli, 2009; Huysmans et al., 1998; 2000). In summary, the export and translocation of sporopollenin components from tapetal cells is poorly understood, intensified by our lack of full understanding of the biochemical nature of sporopollenin components.   With little information on the mechanisms governing sporopollenin traffic out of tapetal cells and through the locule, we arguably know even less about the processes and proteins governing sporopollenin assembly into the intricately patterned and sculptured pollen exine. Over the last decade, a number of Arabidopsis exine patterning or deposition mutants have been characterized, shedding some light on the proteins involved in pollen outer wall assembly. DEFECTIVE IN EXINE FORMATION1 (DEX1) encodes a protein of unknown function that is required for Arabidopsis exine patterning (Paxson-Sowders et al., 2001). The dex1 mutant lacks   25 the hallmark plasma membrane invaginations of callose-encased tetrads, and upon their release, dex1 microspores exhibit random sporopollenin-like deposits and abort prematurely. A plant protein of unknown function, NO EXINE FORMATION1 (NEF1), has a proposed role in the deposition of sporopollenin, as nef1 mutants exhibit sporopollenin-like aggregates in the locule and primexine abnormalities (Ariizumi et al., 2004). NEF1 encodes an integral membrane protein of plastids, and disruption of this gene alters lipid accumulation in the tapetum plastids, although the link between this predicted plastid membrane protein’s function in tapetum lipid metabolism and its function in sporopollenin anchoring requires further investigation. The tomato tapetum-produced GLYCINE-RICH PROTEIN92 (LeGRP92) also has proposed sporopollenin deposition capabilities, as its down-regulation results in uneven exine deposition and reduced pollen viability (McNeil and Smith, 2009). The broad localization of LeGRP92 in the callose wall of MMCs, tetrads, microspore exine and orbicules suggests that this protein may function as an intermediary between the primexine and sporopollenin precursors, guiding the deposition of the exine (McNeil and Smith, 2009). Two putative glycosyl transferases encoded by At1g27600/SPG2/IRX9-like and At1g33430/ UPEX1/UPEX2 were found to function in Arabidopsis exine patterning, possibly in the polymerization or anchoring of sporopollenin to the microspore primexine (Dobritsa et al., 2011).   Although the evidence for protein-assisted sporopollenin assembly is growing, the rapid form assumed by sporopollenin on the microspore surface suggests that properties enabling self-assembly of the biopolymer may contribute to exine wall formation (Hemsley et al., 1996). Despite the broad range of species-specific patterning, an underlying reticulate, hexagonal arrangement of bacula is a uniting feature of pollen exine and spore walls in most species, suggesting that sporopollenin self-assembly may in part account for the complex form of the exine (Scott, 1994; Hemsley et al., 1996). To test the contribution of self-assembly in exine patterning, spore wall construction has been examined in vitro by removing the leading role of the genome (i.e. in specifying primexine or glycocalyx formation, prior to sporopollenin formation) and applying sporopollenin-mimicking substances (fatty acids) under different conditions (Gabarayeva and Grigorjeva, 2013). Some exine-like patterns were observed in these and other wall-forming simulations, and provide preliminary evidence for the contribution of self-assembly in the elaborate outer wall formed by pollen and spores.   26  1.4.2 Orbicules     Orbicules (also called Ubisch bodies) are small, typically spherical granules (most often with a diameter of less than 5 µm), which are of particular interest in exine translocation studies, as they appear to arise in tapetal cells, and have been proposed to carry sporopollenin or components of the exine from the tapetum to developing microspores (Wang et al., 2003). These particles, when treated with various chemicals and stains, have the same properties as the exine, and have long been thought to contain sporopollenin in a number of species (Ubisch, 1927). Orbicules can also exhibit different layers, with an inner core positively staining for polysaccharides, unsaturated lipids and proteins, and an outer encasing resembling sporopollenin (Suarez-Cervera et al., 1995). One rice protein, RAFTIN1, synthesized by tapetal cells, appears to move to microspore exines in orbicules, suggesting that orbicules may also function in the traffic of sporophyte-derived proteins that are required for pollen development (Wang et al., 2003). Similarly, the glycine-rich protein required for exine formation in tomato (LeGRP92) discussed above was localized to orbicules and the early exine in this species (McNeil and Smith, 2009). In support of their proposed sporopollenin-trafficking function, orbicules are abundant outside the locule-facing edge of secretory-type tapeta, typically appear in coordination with sporopollenin synthesis and accumulation on microspores, and contain acetolysis-resistant sporopollenin (Vinckier et al., 2005; Huysmans et al., 2010).   Although there are a number of observations that appear to link sporopollenin translocation with orbicules, the function of these small bodies remains a topic of great uncertainty. Evidence against a role for orbicules in sporopollenin trafficking comes primarily from their absence in many species. For example, while secretory tapeta are associated with orbicules in rice and wheat, Arabidopsis orbicules have not been found. Orbicules are also typically absent in species with amoeboid tapeta, although exceptions such as species of Malvaceae exist (Galati et al., 2006). Although the role of orbicules in pollen development is unknown, a long list of functions beyond sporopollenin transport processes has been proposed for orbicules (Huysmans et al., 1998). Among these, orbicules may simply be by-products of tapetal cell metabolism, or may function to assist pollen dispersal, or programmed cell death by   27 the tapetum (Huysmans et al., 2000). The persistence of orbicules after pollen wall formation and their absence in select taxa with secretory-type tapeta have made their proposed function in sporopollenin trafficking controversial.  The mechanism by which orbicules exit tapetal cells is also largely unknown. Mature orbicules are typically locule-localized, external but often in close association with the tapetum prior to its programmed cell death. Tapetum ER is the site of orbicule progenitor (or pro-orbicule) formation, prior to their extrusion from tapetal cells (Echlin and Godwin, 1968; Suarez-Cervera et al., 1995; Vinckier et al., 2005). In Triticum, the pro-orbicule is encased in a second membrane as it exits the tapetum, and this body is delivered by exocytosis from the tapetum into the locule (El-Ghazaly and Jensen, 1986). Because Arabidopsis is the primary model for genetic and molecular biology studies of pollen and anther development to date, our understanding of orbicules, which are absent in Arabidopsis, remains limited.  1.5 Candidate proteins for sporopollenin trafficking from tapetal cells    As discussed in section, genes encoding sporopollenin synthetic machinery appear to be primarily expressed within the tapetum. This information, together with the observation that the sporopollenin wall around microspores forms prior to tapetum rupture, suggests that sporopollenin precursors are likely exported by tapetal cells. Two classes of proteins, ABC transport proteins and LTPs, are strong candidates for the translocation of sporopollenin precursors and are the focus of the research described in this thesis.  1.5.1 ATP-binding cassette transport proteins and the G subfamily   The ABCG half transporter encoding gene, ABCG26, which is tightly co-expressed with ACOS5, is of particular interest in pollen exine assembly, due to its potential role in the transport of sporopollenin components from their site of synthesis in tapetal cells to the anther locule, similar to the function of ABCG11 and ABCG12 in wax secretion from epidermal cells (Pighin et al., 2004; Bird et al., 2007; Quilichini et al., 2010). This protein belongs to a large and ubiquitous family of ABC proteins with 129 members in Arabidopsis, 51 in humans, 55 in   28 Drosophila melanogaster, and 58 in Caenorhabditis elegans (Sanchez-Fernandez, 2001; Sanchez-Fernandez et al., 2001). The large number and diversity of ABC proteins present in Arabidopsis could assist in plant survival and defense, when considering their sessile nature and metabolic diversity (Sanchez-Fernandez, 2001). Given that ABC transporters are ubiquitous across the three domains of life, it is likely that the ABC gene subfamilies in Arabidopsis are ancient, with ancestral homologs of each subfamily present in the earliest land plants and algae. Additionally, considering the prevalence of gene duplication events in the course of land plant evolution, ABC subfamilies may have unique expansion patterns within lineages, as seen for the monosaccharide transporter gene family (Taylor and Raes, 2004; Johnson and Thomas, 2007; Johnson et al., 2006). As sporopollenin appears to have evolved in coordination with the transition of plant life to a terrestrial setting, processes supporting sporopollenin synthesis and translocation may have similarly been selected for. Based on the early bud expression pattern and co-expression of ABCG26 with genes required for sporopollenin synthesis, this transport protein is a strong candidate for a translocator of sporopollenin precursors in Arabidopsis.   ABC transporters are commonly characterized by the presence of two highly conserved ATP-binding cassette (ABC) domains and two transmembrane domains (TMDs), each composed of multiple transmembrane alpha-helices (Verrier et al., 2008). ABCG26 and other half-size transporters of the same subfamily [composed of 28 members, formerly called the White-Brown Complex (WBC) subfamily] represent an exception to this general domain structure, as they contain one ABC domain fused to one TMD and require dimerization to form a functional transport protein (Higgins and Linton, 2001). In Arabidopsis, the importance of a number of half-size ABCG transport proteins in plant development has been demonstrated, including ABCG11 and ABCG12 in cuticular lipid export, ABCG9, ABCG11 and ABCG14 in vascular development, ABCG13 in flower cuticle deposition, and ABCG22 and ABCG25 in ABA transport (Pighin et al., 2004; Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007; Panikashvili et al., 2010; Quilichini et al., 2010; Choi et al., 2011; Dou et al., 2011; Kuromori et al., 2011a; Le Hir et al., 2013; Panikashvili et al., 2011; Kuromori et al., 2011b; 2010). Among these characterized half-size ABCGs, ABCG11 has the ability to homodimerize or heterodimerize with ABCG12, ABCG14 and ABCG9 to generate apparently different functionalities, while other transporters appear to strictly form heterodimers, as reported for   29 ABCG12 and ABCG14 (Bird et al., 2007; McFarlane et al., 2010; Le Hir et al., 2013). These data suggest that the expression patterns among ABCGs, together with their interaction promiscuity/specificity may account, in part, for the broad metabolite transport capabilities required by plants.   1.5.2 Lipid transfer proteins   In Arabidopsis, there are a number of lipid transfer proteins (LTPs) co-expressed with ACOS5, indicating that one or more of these proteins may function in the export of sporopollenin precursors from tapetal cells. LTPs were initially defined as small extracellular proteins capable of phospholipid transfer between membranes in vitro and possess a hydrophobic cavity shown to bind lipids in vitro (Kader, 1996). LTPs are expressed with high developmental and tissue specificity, commonly in defined cell layers such as the epidermis in maize seedlings and elongating Arabidopsis stem cells (Suh, 2005; DeBono et al., 2009), in cotton fibers (Ma et al., 1995), the protoderm in carrot (Sterk et al., 1991), in tobacco embryos (Sossountzov et al., 1991), in the cell walls of stigma and pollen in Arabidopsis (Thoma et al., 1994), as well as in the tapetum of tobacco (Koltunow et al., 1990), Brassica napus (Foster et al., 1992), maize (Lauga et al., 2000), and Arabidopsis (Rubinelli et al., 1998; Ariizumi et al., 2002). Although the in vivo function of most plant LTPs remains unclear, a glycosylphosphatidylinositol (GPI)-anchored LTP (LTPG) required for cuticular wax export was recently characterized (DeBono et al., 2009). Facilitated co-expression analyses have revealed genes encoding LTPs with similar expression patterns to ACOS5, suggesting a role for these proteins in sporopollenin transport, polymerization and/or deposition. The identification of LTPs expressed specifically in the tapetum of tobacco (Koltunow et al., 1990), Brassica napus (Foster et al., 1992), maize (Lauga et al., 2000), and Arabidopsis (Rubinelli et al., 1998; Ariizumi et al., 2002) suggests that one or more LTPs may function in pollen wall transport and/or assembly, but this has yet to be demonstrated.       30 1.6 Research questions, objectives and significance of findings   Based on the literature surrounding sporopollenin formation, it has been established that components of sporopollenin are synthesized in the tapetum and that these components are found polymerized as sporopollenin on the surface of pollen grains. Our expanding knowledge of sporopollenin composition and the processes governing its synthesis in the tapetum are in stark contrast to our modest understanding of the mechanisms involved in its export from tapetal cells and its traffic through the locule to early microspores. Therefore, this thesis addresses the major question of how constituents of the exine wall (sporopollenin) exit tapetal cells.  The broad goal of my research is to elucidate the export pathway of sporopollenin components from tapetal cells to developing pollen grains. More specifically, I tested the potential role of transport proteins in the traffic of sporopollenin components from the tapetal cells to developing microspores. Arabidopsis thaliana (Arabidopsis) was selected as the model organism to study sporopollenin trafficking because of the well-established developmental progression of anthers during pollen wall formation, the knowledge of the genes required for sporopollenin synthesis, its ease of genetic manipulation, and the small size of male reproductive organs that are conducive to imaging and microscopic analysis. The developmental co-expression of a gene encoding a membrane transport protein from the ATP-binding cassette (ABC) transporter superfamily with genes required for sporopollenin biosynthesis in Arabidopsis suggested a possible mechanism for the export of tapetum-synthesized sporopollenin monomers. Initial work performed by Dr. M. Friedmann in the Douglas lab revealed that ABCG26 was expressed in the tapetum of anthers during the developmental phase coinciding with sporopollenin synthesis and deposition in Arabidopsis. This preliminary work led to the hypothesis that ABCG26 functions to export sporopollenin constituents from tapetal cells. The identification of multiple genes encoding putative LTPs through co-expression analyses with ACOS5, suggested additional export components in the traffic of sporopollenin. Based on these data, a second hypothesis was developed that multiple proteins, including lipid transfer proteins aid in the transport of constituents of sporopollenin during pollen grain formation.   To address these hypotheses, my thesis had four specific objectives: 1) To obtain detailed morphological and ultrastructural information on the pollen wall and tapetum during anther   31 development in Arabidopsis using the most up-to-date methods of sample preparation an imaging (Chapter 2), 2) To characterize the biological function of ABCG26 in Arabidopsis pollen wall formation (Chapter 3), 3) To characterize the biochemical nature of sporopollenin constituent(s) exported from the tapetum by ABCG26 (Chapter 4), and 4) To investigate the role of lipid transfer proteins in sporopollenin traffic from tapetal cells (Chapter 5). In the 6th and final chapter of this thesis, I summarize the key findings and conclusions of Chapters 2 through 5, explore the implications of these findings, and discuss how my results will direct future studies in the field.    32 Chapter 2: New views of tapetal ultrastructure and pollen exine development in Arabidopsis thaliana  2.1 Introduction  The highly reduced gametophyte of flowering plant species forms within the sporophytic tissues of the anther. These microgametophytes, or pollen grains, play a central role in plant reproduction, carrying the sperm cells to the female reproductive structures for double fertilization. The critical role fulfilled by pollen grains in the life cycle of flowering plants has required these structures be fortified against the harsh elements of terrestrial life. The wall encasing pollen is highly specialized, consisting of an inner (intine) cellulosic wall, and an outer (exine) wall containing a highly stable and recalcitrant biopolymer called sporopollenin (Southworth 1974; Scott 1994). In many species including Arabidopsis thaliana, the exine additionally contains a pollen coat, which covers the patterned sporopollenin framework and aids in hydration and species-specific recognition by the stigma. Pollen development relies heavily on the surrounding tissues of the anther wall, especially the tapetum, but also the middle layer, endothecium, and epidermal cell layers. The epidermis prevents anther water loss and aids gas exchange and, together with endothecial cells, provides structural support to the anther wall (Goldberg et al., 1993). Secondary wall thickenings in the endothecial cells are required for pollen dispersal by anther dehiscence along specialized cells of the anther epidermis (Keijzer, 1987; Mitsuda et al., 2005). In contrast, cells of the middle layer and tapetum undergo cell death and are absent from mature anthers prior to dehiscence. Although the precise role of the middle layer is not known, the middle layer becomes tapetum-like if its degeneration is postponed, as in the Arabidopsis fat tapetum mutant (Sanders et al., 1999). The middle layer also has secretory activity and in dioecious plants, such as Actinidia deliciosa (kiwifruit), may regulate the production of male-sterile and male-fertile flowers by altering the timing of their own cell death (Falasca et al., 2013). The gametophytic reliance on sporophytic tissues is particularly clear in the case of tapetal cells. Tapetum-specific cell ablation studies demonstrated the specific and critical role served by tapetal cells in the development of pollen grains, as microspores aborted in their absence, while other anther cell types appeared   33 unaffected (Goldberg et al., 1995; 1993; Mariani et al., 1990; Koltunow et al., 1990). The delay of tapetum degeneration that occurs in mutants of Arabidopsis MALE STERILE1 (MS1) and rice TAPETUM DEGENERATION RETARDATION (TDR), respectively, results in collapsed microspores (Li et al., 2006; Yang et al., 2007).  Both signaling between microspore and tapetal cells, as well as complex transcriptional regulation of tapetum proliferation and cell death, are postulated to be necessary for the critical timing of development in the tapetum (reviewed by Parish and Li, 2010). Among the many essential roles of tapetal cells, this sporophytic cell layer is a key player in forming the highly sculptured sporopollenin wall of the exine. The role of the tapetum in microspore wall formation is evident early in anther development, when microspore tetrads are freed from their callose encasement by tapetum-secreted callase (Dong et al., 2005). In the absence of normal callose dissolution in Arabidopsis, as in the myb103 mutant, defective exine patterning is observed (Zhang et al., 2007). The tapetum-specific expression of genes required for sporopollenin synthesis indicates that an abundance of sporopollenin is exported from intact tapetal cells during the free uninucleate microspore stage of pollen development (see reviews by Ariizumi and Toriyama, 2011; and Liu and Fan, 2013). Recently, immunolocalization and protein-protein interaction studies have supported a model where the enzymes producing sporopollenin precursors are organized in a metabolon within the tapetum (Lallemand et al., 2013). However, as our knowledge of these functional components has progressed, our understanding of the dynamic cellular structure of the tapetum and the ultrastructure of the developing pollen wall over the course of anther development has lagged.  The tapetum is a transitory tissue and, together with the many structures associated with early microspores, appears to be sensitive to chemical fixation protocols employed for light and electron microscopy (Sanders et al., 1999; Ariizumi et al., 2005; de Azevedo Souza et al., 2009).  A study in Brassica campestris used cryo-fixation to document early microspore wall development in the tetrad stage, but examination of tapetum ultrastructure was not within the scope of the work (Fitzgerald and Knox, 1995). In a study characterizing DEFECTIVE EXINE1 (DEX1) protein function in pollen wall formation, the formation of structured components within the primexine matrix, known as probaculae and protecta, and the importance of microspore undulations in the formation of these structures were demonstrated by comparing high-pressure frozen and freeze   34 substituted Arabidopsis wild-type and dex1 mutant anthers in the tetrad stage (Paxson-Sowders et al., 2001). These studies validated cryo-fixation as a useful approach to preserve the delicate and transitory structures of early microspore walls in the tetrad stage, which serve as a scaffold for the subsequent polymerization of tapetum-derived sporopollenin into baculae and tecta.  In contrast to sporopollenin, which appears to be actively exported from the intact tapetum early in microspore development, components of the lipidic pollen coat are produced and stored in the tapetum and deposited onto pollen upon tapetum programmed cell death (Heslop-Harrison, 1968b; Edlund, 2004). The pollen coat components are liberated upon the breakdown of two specialized organelles in the tapetum, elaioplasts and tapetosomes, which accumulate at late stages of pollen development (Piffanelli et al., 1998; Hsieh et al., 2003). Elaioplasts are specialized plastids that are rich in steryl esters, free polar lipids, and plastid lipid-associated proteins and are filled with globuli (Kim et al., 2001; Ting et al., 1998; Piffanelli and Murphy, 1998). Tapetosomes are unique densely packed organelles that are intimately associated with the endoplasmic reticulum (ER), and consist of a fibrous meshwork of vesicles, fibrils, and oil bodies containing oleosin proteins, alkanes and flavonoids (Hsieh and Huang, 2005; 2007; Huang et al., 2013b).  The biochemical data on the components of tapetosomes and elaioplasts have been correlated with their ultrastructure in chemically fixed specimens, however these structures are rich in membranes and lipids, so structural features are sometimes ambiguous.  The objective of this study was to define the ultrastructure of the developing anther of Arabidopsis using cryo-fixation techniques in order to preserve the delicate tapetum tissue and developing pollen grains during production of the complex pollen cell wall.  The goal is to provide a detailed analysis of wild-type cell structure in Arabidopsis anthers, which will serve as a baseline for studies examining cellular and cell wall architecture in mutants affected in microspore and tapetum development.    2.2 Results   The classic study by Sanders et al., (1999) defined the stages of pollen and anther development in Arabidopsis using light microscopy. Here, we re-examined these stages using   35 high-pressure freezing and freeze-substitution combined with transmission electron microscopy (TEM). We first undertook an overview of pollen development using light microscopy to follow the development of all cell layers of the anther through the meiotic events of microsporogenesis and mitotic events of microgametogenesis (Figure 2.1).  In addition, corresponding pollen morphology and nuclei associated with these development stages are shown (Figure 2.2). Following cryopreservation, freeze substitution and resin embedding, the sectioned anthers captured near the first microspore meiotic division were cytoplasmically dense and the cell layers were not separated (Figure 2.1A,B), as seen in earlier studies employing chemical fixation. Tapetal cells at the tetrad stage (Figure 2.1C) contained enlarged vacuoles and were easily distinguished from surrounding sporophytic layers by their dense cytoplasmic contents and binucleate nature. The tapetum had a tendency to separate from the surrounding middle layer cells as part of the embedding process, but unlike images from chemically fixed material, our fixation method showed that the tapetum and locule were tightly associated.  At more advanced stages of microsporogenesis, the tapetum became less vacuolated and began to accumulate dense organelles (Figure 2.1D,E). Throughout these stages, the tapetum formed a continuous ring of tightly packed cells (Figure 2.1A-E), unlike some chemically fixed samples, in which tapetal cells were separated by gaps (Sanders et al., 1999). The preservation method employed here additionally revealed an abundance of locule fluid around microspores through to the early tricellular stage of pollen development (Figure 2.1D-G), in contrast to previous studies where the medium between microspores and tapetal cells appeared as an empty space (Sanders et al., 1999).     36    37 Figure 2.1: Overview of pollen development in the Arabidopsis anther. Development is divided into meiotic divisions (left panel) and mitotic divisions (right panel). A) Diploid microspore mother cells. B) Meiotic microspore mother cells surrounded by a callose wall. Two polar nuclei (arrows) with lack of internal cell walls suggest that meiosis is not complete. C) After meiosis, tetrads of haploid microspores encased in thick callose are present. D) Early uninucleate microspores with darkly staining exine walls. E) Uninucleate microspores in the ring-vacuolate stage. F) Following the first mitotic division, bicellular pollen grains contain one small generative cell and one large vegetative cell (arrows indicate generative cell). Tapetosomes and elaioplasts are visible in the tapetum (F), and become more abundant as bicellular pollen mature (G). H) After the second mitotic division, tricellular pollen is surrounded by tapetum. Thickenings in the endothecial cell walls are visible in cells marked by asterisks. I) Tapetum cytoplasmic content in the locule following cell death. The intine wall surrounds each pollen (arrows). J) Mature pollen grains following tapetum break-down. En, endothecium; Ep, epidermis; M, microspore; ML, middle layer; MMC, meiotic microspore mother cell; P, pollen; T, tapetum; Td, tetrad. Bars = 10 μm.     38  Figure 2.2: Pollen morphology through microsporogenesis and microgametogenesis. Light microscope images of pollen in key stages of development are shown, beginning at the top, centre panel with microspore mother cells (MMC), and moving clockwise from meiotic cells with polar nuclei, to tetrads of microspores, uninucleate microspores, bicellular pollen, and concluding with tricellular pollen. Images with white or black text indicate stage of development within microsporogenesis or microgametogenesis, respectively. Purple arrows indicate a meiotic or mitotic division that occurs within mother cells/microspores/pollen grains between the images they connect. Unlabeled arrows indicate nuclei. The four asterisks denote four microspores of one tetrad. Arrows labeled ‘v’ or ‘g’ designate the vegetative nucleus and generative cell(s) of pollen, respectively. The double headed arrow in late bicellular pollen indicates the mitotic division underway in the generative cell. Bars = 5 μm.    39  After the two meiotic divisions and cytokinesis transform callose-encased MMCs to tetrads of haploid microspores (Figure 2.1A-C), microsporogenesis concludes with the release of free haploid microspores from their callose encasement (Figure 2.1D). At this stage of development, a dramatic change in the free microspores was observed, as a structured exine wall of sporopollenin became visible (Figure 2.1D,E).  At early stages, free uninucleate microspores appeared cytoplasmically dense and contained small but numerous vacuoles around a centrally located nucleus (Figure 2.1D, Figure 2.2). In more mature uninucleate microspores, the cytoplasm was in a peripheral location as the vacuoles coalesced into one enlarged vacuole (Figure 2.1E, Figure 2.2). In our samples, free microspores appeared round, and the cytoplasmic contents filled the area formed by the microspore wall, in contrast to previous studies in which microspores frequently appeared shrunken or detached from their walls. With their densely staining exine wall, polarized uninucleate microspores, often called ring-vacuolate microspores (Blackmore et al., 2007), represent the final stage of microsporogenesis before the divisions of microgametogenesis that will form mature pollen grains (Fitzgerald and Knox, 1995).  In stages of bicellular pollen grain development, a small generative cell was visible within a large vegetative cell (Figure 2.1F,G, arrows indicate generative cell; see Figure 2.2 for enlargement), consistent with the asymmetry of the first mitotic division. Each pollen grain contained multiple tiny vacuoles, unlike the enlarged vacuole of the previous stage, and the large vegetative cells appeared rich in subcellular organelles (Figure 2.2). After the second mitotic division during which the generative cell had produced two male gametes (or sperm cells), tricellular pollen grains were larger in volume and filled with numerous lipid bodies and starch granules (Figure 2.1H, Figure 2.2).  Numerous darkly-staining organelles were prominent in tapetal cells surrounding pollen in bicellular and early tricellular stages of development (Figure 2.1F-H). These organelles, although not clearly discernable in the light microscope, were consistent in size and shape with elaioplasts and tapetosomes (Figure 2.1F-H). The locule fluid, which stained with toluidine blue in early bicellular pollen-containing anthers (Figure 2.1F), was weakly stained in the late bicellular pollen stage (Figure 2.1G), and appeared absent in locules containing tricellular pollen (Figure 2.1H). Simultaneously, tapetal cells gradually appeared thinner and became almost entirely occupied by densely staining organelles (Figure 2.1H). The surrounding endothecial cells became highly vacuolated, and the cell corners stained light blue   40 with toluidine blue stain, which is typical for lignified cell walls (see cells marked by asterisks in Figure 2.1H). Also in the tricellular pollen stage, the intine, or inner pollen wall, was clearly visible by light microscopy as a lightly-staining band between the darkly-staining exine and pollen cytoplasm (arrows in Figure 2.1I). In the late tricellular pollen stage, the presence of tapetal cell cytoplasmic constituents in the locule indicated that cell disintegration after programmed cell death had occurred (Figure 2.1I). Finally, mature, slightly oblong tricellular pollen grains, ready for release by anther dehiscence, filled the locule space formed by the endothecium and epidermis (Figure 2.1J).   From this analysis of anther development, most anther cell types over the course of microsporogenesis and microgametogenesis were clearly visible and preserved in high quality by the cryo-fixation and embedding techniques employed. However, this analysis provided limited information on the middle layer, as these cells appear to be easily damaged in the embedding process and were poorly resolved by light microscopy. To view the middle layer of the anther wall and microspores with associated locule fluid in a near-native and hydrated state, immature anthers were cryo-fractured longitudinally and visualized by cryo-scanning electron microscopy (cryo-SEM, Figure 2.3). In two of the four microspore-filled locules, all anther wall cell layers were visible (Figure 2.3A). Each locule revealed uninucleate microspores in the ring-vacuolate stage of development (Figure 2.3B and equivalent to the stage depicted in Figure 2.1E).  Surrounding microspores, the fluid of the locule appeared semi-ordered with a lattice-like substructure. While this may reflect some ice crystal formation after plunge freezing, with separation of solutes into the eutectic phase between ice crystals, the regularity of the pattern is intriguing and its underlying cause is worthy of further investigation. Furthermore, this analysis showed that the tapetum bulges into the locule in close association with nearby microspores, rather than forming the typical rigid shape of cell wall-encased plant cells. Interestingly, in contrast to the current models that describe loss of the middle layer early in development, the middle layer cells, directly external to the tapetum, clearly contained cytosolic contents, in agreement with findings by Owen and Makaroff, 1995 (Figure 2.3B). A wide band of endothecial cells was visible, external to the thin band of middle layer cells and below a surface layer of epidermal cells. A waxy cuticle was observed on the outermost face of epidermal cells.  The cryo-SEM method employed here revealed two poorly characterized features of immature   41 anthers: the lattice-like substructure of the locule fluid and persistence of the middle layer into the latest stage of uninucleate microspore development.      42 Figure 2.3: Cryo-SEM of hydrated Arabidopsis anther. A) Two of the four locules filled with developing pollen grains (microspores) exposed by fracture of the anther. B) A magnified view of one locule and surrounding sporophytic cell layers from panel A. Microspores are in the ring-vacuolate uninucleate stage of development (anther stage 9). The microspores and surrounding locule fluid are encased in four cell layers of sporophytic tissues: the innermost tapetum, T; followed by a thin middle layer, ML; the endothecium, En; and outermost epidermis, Ep. Ex, exine; Lo, locule; M, microspore; N, nucleolus; T, tapetum, V, vacuole. Bars = 5 μm (B,C).  The light microscopy analysis suggested that the cryo-fixed anthers retained structural elements that were not preserved in earlier studies using chemical fixation, but the sub-micron details were beyond the limit of light microscopy resolution. In order to trace the development of the multilayered pollen wall, cryo-fixation combined with TEM was used to examine its origins, beginning with the primary cell wall surrounding meiotic microspore mother cells (MMCs), through primexine formation around microspores in tetrads, and sporopollenin-based exine wall formation around free uninucleate microspores.  MMCs demonstrated a high degree of symplastic continuity, with abundant cytomictic channels interconnecting the cytoplasm of MMCs within each anther locule (Figure 2.4A, arrowheads indicate cytomictic channels). The plasma membrane and primary cell wall of meiotic MMCs (Figure 2.4B,C) were separated by the previously characterized callose wall (Dong et al., 2005), and no primexine wall was visible. In the tetrads, the primexine wall first became visible on the surface of individual microspores (Figure 2.4D-F). With the exception of aperture sites, the microspore plasma membrane appeared invaginated at regular intervals and acquired “spacers” in each of the pockets formed by its undulations, which appeared electron-dense, consistent with previous reports (Figure 2.3E; Fitzgerald and Knox, 1995; Paxson-Sowders et al., 1997). Separated by spacers, the peaks along the microspore plasma membrane correlated with sites of probaculae assembly (arrowheads in Figure 2.4E,F). In a late tetrad stage, probaculae and protecta were distinguished as electron dense cones topped by caps, respectively and extended from the microspore plasma membrane at regular intervals within the primexine matrix (arrowheads in Figure 2.4F). Although little is known about the composition of probaculae and protecta, these structures appear to contain an early form of sporopollenin (Heslop-Harrison, 1968c).     43  Figure 2.4: Microspore exine wall development in cryo-fixed Arabidopsis. High-pressure frozen/freeze substituted anthers examined using TEM reveal stages of exine deposition. A) Microspore mother cells (MMCs) with cytomictic channels (marked by arrowheads). B) Early MMCs primary cell wall displaced by an electron lucent layer of callose, between the primary wall and the MMC plasma membrane. C) Thick callose wall on each side of primary cell wall of MMCs. D) Tetrads fill the locule, representing a key developmental stage for primexine formation (early and late primexine stages, as in E and F, respectively). E) Within the primexine matrix, spacers form in the pits created by the undulating plasma membrane. The raised portions of plasma membrane along the microspore surface (arrowheads in E) mark the sites of   44 probaculae and protecta formation (arrowheads in F). G) Early free uninucleate microspores accumulate in the locule.  H) Sporopollenin forms at the microspore’s baculae and tecta, initially appearing to be lamellar (arrows). I) As exine matures, it assumes a homogenous appearance in baculae and tecta. Remnant primexine matrix surrounds baculae shortly after microspore release (H and I), but is not observed as uninucleate microspores mature (K and L). J) Mid-stage uninucleate microspores in the locule. K) Uninucleate microspores with sculptured, electron-dense baculae (marked by asterisks) and tecta (arrowheads), as well as a thin layer of early intine. L) Tangential section of uninucleate microspore with baculae (marked by asterisks) and tecta (arrowheads). C, callose; En, endothecium; Ep, epidermis; Ex, exine; In, intine; Lo, locule; ML, middle layer; MMC, microspore mother cell; Msp, microspore; N, nucleus; PM, plasma membrane; PW, primary wall; rPE, remnant primary exine; rPW, remnant primary wall; Td, tetrad; T, tapetum. Bars = 5 μm (left panel) and 500 nm (middle and right panels).  Previous studies using cryo-fixation in Arabidopsis and Brassica sp. have focused specifically on wall formation within the tetrad stage of pollen development. Although a number of studies have examined the microspore wall after tetrad release, the chemical fixation methods employed typically resulted in shrunken microspores and the separation of the microspore plasma membrane and exine wall. Therefore, we extended our investigation of microspore wall ultrastructure in cryo-fixed anthers through the uninucleate stages of microspore development. At this stage, in contrast to clusters of microspores in the tetrad stage, individual microspores were surrounded by a homogenous locule of medium electron-density (Figure 2.4G,J).  Microspores at this early uninucleate stage typically exhibited an exine of baculae and tecta that was homogenously electron-dense (Figure 2.4I). Upon close examination, the exine wall of a subset of free microspores contained lamellae, visible as electron-translucent lines within an electron-dense wall, and situated roughly parallel to the microspore plasma membrane (see arrows in Figure 2.4H). Remnants of the primexine matrix were observed surrounding the sculptured exine wall, appearing as a loose mesh between the locule and the sporopollenin (Figure 2.4I).  In the absence of cytoplasmic shrinkage and separation from the cell wall, the intine wall, a thin band of wall material between the exine and the microspore plasma membrane, was identified in early free microspores (Figure 2.4I), and became clearly distinguished by the mid-free microspore stage (Figure 2.4K).  Surrounding the developing intine, the exine wall around mid-uninucleate microspores consisted of highly electron-dense baculae and tecta (Figure 2.4K, see asterisks and arrowheads, respectively). In sections cut tangentially to the mid-uninucleate   45 microspore through the exine, baculae appeared round and discrete, connected by tecta, where captured (Figure 2.4L). In summary, the first evidence of the patterned outer wall was identified on the plasma membrane of microspores in tetrads. Deposition of the interior layer of the pollen wall, the intine, was observed by the end of the free uninucleate microspore stage.  No gaps or locule fluid were observed between layers of the pollen wall and the callose wall or microspore plasma membrane, in contrast to many previous reports. To obtain high quality information on tapetum ultrastructure over the course of microspore development, we began by examining anthers in developmental stages near the time of microspore meiosis. Tapetal cells in Arabidopsis are binucleate from the MMC stage (Figure 2.5A) to the late uninucleate microspore stage of anther development (Figure 2.5B). Although the binucleate state of tapetal cells presumably persists until the tapetal cell death program is initiated, this state is difficult to capture as elaioplasts and tapetosomes rapidly consume the majority of the cellular space. As early as the MMC stage of pollen development, the tapetal cells were distinguishable from the middle layer, endothecial and epidermal cell layers of the anther wall by their dense cytoplasm and binucleate nature (Figure 2.6A).  These tapetal cells were cytoplasmically dense, with ER, ribosomes, mitochondria and vacuoles (Figure 2.6B). In subsequent stages, the tapetum appeared metabolically active based on the high abundance of ER, Golgi apparatus, mitochondria and proplastids in the tetrad and free uninucleate stages of microspore development (Figure 2.6C-H). Where microspores were not closely appressed to the tapetum, the locule-facing edge of tapetal cells often appeared wavy with occasional invaginations (see asterisks in Figure 2.6D,F,H). Thus, during the period of peak sporopollenin synthesis, from the tetrad stage through the uninucleate microspore stage, the tapetum is rich in ER and the plasma membrane, lacking a cell wall, is often in contact with microspores.     46  Figure 2.5: The binucleate tapetum in Arabidopsis. Two nuclei with associated nucleoli (asterisks) are present in tapetal cells. Both nuclei are observed throughout microspore development from the microspore mother cell stage (anther stage 5) (A) to the ring-vacuolate uninucleate microspore stage (anther stage 9) (B), after which elaioplasts and tapetosomes become abundant and fill the majority of tapetal cells. C, callose; El, elaioplast; Lo, locule; Msp, microspore; Mt, mitochondrion; N, nucleus; T, tapetum; V, vacuole. Bars = 2 μm.    47    48 Figure 2.6:  Tapetum development during exine formation in cryo-fixed Arabidopsis anthers. High-pressure frozen/freeze substituted anthers examined using TEM reveal tapetum ultrastructure in stages associated with microsporogenesis. A) Tapetal cells form the innermost layer of the anther wall, adjacent to microspore mother cells. B) Higher magnification view of tapetum adjacent to MMC (anther stage 5), with tapetal cell wall present. C) Tapetum around callose-encased tetrads. D) Higher magnification view of tapetum lacking visible cell wall on locule-facing plasma membrane. E) In the early uninucleate microspore stage (anther stage 9), developing microspore in close proximity to plasma membrane (PM) of tapetum during exine deposition. F) Higher magnification view of tapetum. Tapetum PM exhibits occasional invaginations (asterisks in D, F and H). G) Late uninucleate microspore stage of development, with well-developed exine on microspores, which are adjacent to tapetum.  H) Higher magnification view of tapetal developmental stage shown in G, with abundant ER, and organelles beginning next stage of differentiation.  C, callose; En, endothecium; Ep, epidermis; ER, endoplasmic reticulum; Ex, exine; G, Golgi body; Lo, locule; ML, middle layer; MMC, microspore mother cell; Mt, mitochondrion; Msp, microspore; N, nucleus; Pp, proplastid; PM, plasma membrane; rER, rough endoplasmic reticulum; rPW, remnant primary wall; Td, tetrad; T, tapetum; V, vacuole. Bars = 5 μm (left panel) and 1 μm (right panel).  In addition to sporopollenin biosynthesis, tapetal cells are known to synthesize numerous lipid-rich pollen coat constituents during the pollen mitotic divisions.  We investigated tapetum ultrastructure in detail at these stages in cryo-fixed anthers (Figure 2.7). Proplastids and abundant ER observed at earlier stages were replaced by elaioplasts, tapetosomes and morphologically distinct rough and smooth ER (Figure 2.7A). At the tricellular pollen stage, elaioplasts and tapetosomes filled the majority of each tapetal cell, pushing lamellar smooth ER (asterisks in Figure 2.7B) and rough ER (arrows in Figure 2.7B) to the cortical cytoplasm. With the preservation methods we employed, tapetal cells remained intact through the bicellular and early tricellular pollen stages (Figure 2.7A-B). Consistent with tapetum rupture due to programmed cell death, the contents of tapetal cells were scattered in the locule and appeared closely associated with the crevices of tricellular pollen exine (Figure 2.7C), or were absent from locules, in which the tricellular pollen grains were engorged and exhibited pollen coats (Figure 2.7D).    49  Figure 2.7: Tapetum development during pollen coat formation in cryo-fixed Arabidopsis anthers. High-pressure frozen/freeze substituted anthers examined using TEM reveal tapetum ultrastructure in stages associated with microgametogenesis. A) During bicellular pollen stage (early anther stage 11), the tapetal cells contained differentiating elaioplasts and tapetosomes, as well as dilations of rough ER with fibrillar material in their lumen (arrows). B) In the tricellular pollen stage, most tapetal cell space is consumed by elaioplasts and tapetosomes, with smooth ER (asterisk) and rough ER (arrows) in a peripheral position. Immediately following tapetum programmed cell death, tapetum contents is scattered in the locule and within the spaces created by the sculptured exine (C). (D) The tapetum is absent from the locule, but tapetum-derived pollen coat constituents fill the cavities of the sporopollenin-based exine framework on tricellular pollen, completing pollen wall formation (anther stage 12). El, elaioplast; En, endothecium; Ex, exine; Lo, locule; Mt, mitochondrion; P, pollen grain; PC, pollen coat; rT, remnant tapetum; T, tapetum; Ts, tapetosome. Bars = 1 μm.    50 Close inspection of tapetal cells at the late stages of pollen development revealed organelle substructure not previously reported. Although cells were primarily filled by elaioplasts and tapetosomes in late bicellular and tricellular stages of pollen development, distinct forms of ER were also observed. Dilations of rough ER with fibrillar material in the lumen were observed throughout the tapetal cytoplasm (arrows in Figure 2.8A), alongside developing tapetosomes, mitochondria, Golgi bodies, and lamellar ER (see asterisks in Figure 2.7B, 6B). Large sheets of lamellar smooth ER were also observed specifically in very late stages of tapetal cell development, just prior to their rupture (asterisks in Figures 2.7B and 2.8B). In the final stages of development, the tapetum exhibited fragmented cytoplasm and lacked free ribosomes (Figure 2.8B-D).      51  Figure 2.8:  Late tapetum organelle ultrastructure. High-pressure frozen/freeze substituted anthers examined using TEM. A) Rough ER (arrows) contains electron-dense fibrous material. B) Smooth ER (asterisk) is extensive and in abundance in the periphery of late-stage tapetal cells. C) Elaioplast ultrastructure. These specialized plastids contain circular globuli in their stroma, magnified in E and F. The elaioplast globuli contain electron-translucent oil bodies (E), an electron-dense meshwork (F), and are contained in a membrane. D) Tapetosome ultrastructure, showing bundles of tightly associated linear tubes (magnified in G) and an intricate matrix of coiled fibrils (H) that make up the majority of each tapetosome. Arrows indicate circular electron-translucent regions. El, elaioplast; Ex, exine; Lo, locule; P, pollen grain; T, tapetum; Ts, tapetosome. Bars = 500 nm.    52 Close inspection of elaioplasts (Figure 2.8C) and tapetosomes (Figure 2.8D) revealed clear substructure within these organelles. Elaioplasts contained numerous round globuli within their stroma. Within each elaioplast globule, two main components of varying texture and electron density were visible (specified by boxed regions in Figure 2.8C, and magnified in Figure 2.8E,F). The electron-translucent component, presumably containing steryl esters, formed small and often circular inclusions within an electron-dense meshwork. The preservation methods used here revealed a large proportion of electron-dense constituents associated with elaioplast globuli. In addition to the globuli of elaioplasts, occasional membrane-like protrusions were observed between the globuli (Figure 2.7A).  In contrast to elaioplasts, tapetosomes transitioned from a near circular appearance in bicellular pollen locules to amorphous multi-lobed shapes that occupied much of the cellular space and lacked an encasing membrane (Figure 2.8A,C,D). Within tapetosomes, distinct components were differentiated, including clusters of rod-like tightly packed membrane tubes (Figure 2.8G) that occurred within a network of branching tubules in a dense matrix (Figure 2.8H). The sample preservation methods employed here revealed a high level of substructure within tapetosomes that was difficult to correlate with previously reported tapetosome structures. The dense matrix with associated tubules may be ER-derived membranes housing flavonoids, as have been identified in Brassica sp. tapetosomes (Hsieh and Huang, 2007; 2005). The nature of the densely packed rod-like structures of tapetosomes (Figure 2.8G) has not been described previously, and their chemical composition is unknown.   Finally, we employed TEM to view the subcellular details of the cryo-fixed mature pollen cell wall at high resolution. The ultrastructure of the mature pollen wall was striking: the detailed structures of the intine and exine, tightly appressed to the plasma membrane of the microspore, were revealed (Figure 2.9, labeled blue in B). The intine appeared as a thick electron-lucent band with microfibrils running parallel to the vegetative cell plasma membrane  (Figure 2.9, pink band in B). The exine exhibited two main constituents, a homogenous and sculpted component consistent with the sporopollenin-based wall, and a heterogeneous component consistent with the pollen coat (Figure 2.9, false-coloured green and orange, respectively). The sporopollenin component of the exine wall appeared uniformly electron-dense, with baculae that extended radially from the intine and tecta that formed at the external end of baculae, roughly parallel to   53 the pollen plasma membrane. A thin band of comparable electron-density was also observed at the interface between the intine and pollen coat in the crevices between baculae. In contrast to the homogenous appearance of the sculptured sporopollenin, the pollen coat component of the exine appeared highly heterogeneous. Within the pollen coat, two electron-translucent components were observed, one appearing as long and rectangular, the other appearing roughly star-shaped (see arrowheads in Figure 2.9B). As the images were taken in cross-section, it is possible that these two electron-translucent constituents represent the same component of the coat, captured in different view planes. While the remainder of the pollen coat was electron-dense, it exhibited previously undescribed regions of regularly patterned lamellae (see white asterisk in Figure 2.9B).   Figure 2.9: The mature Arabidopsis pollen wall. A) Transmission electron micrograph of a wild-type pollen grain in cross section, revealing the stratified pollen wall outside the pollen grain plasma membrane and underlying cytoplasmic contents. B) Coloured overlay of the micrograph from panel A, highlighting and differentiating the sporopollenin (green) and pollen coat (orange) constituents of the pollen exine, from the intine (pink), plasma membrane (black) and pollen cytoplasm (blue). Two structural features of the sporopollenin wall, known as the bacula and tectum, are labeled. Bars = 500 nm.        54 2.3 Discussion   In this study, high quality cryo-preservation methods were employed to clarify the major events and morphological markers in Arabidopsis anther development, while additionally defining novel features within tapetum organelles and microspore walls. High-pressure freezing and cryo-fixation methods enabled cellular ultrastructure to be rapidly preserved, preventing many of the artifacts and distortions observed when conventional methods of chemical fixation were used (Gilkey and Staehelin, 1986). In particular, cryo-fixation prevented the separation between cells of the tapetum, microspore shrinkage and wall detachment, the loss of locule fluid, and the deformation of subcellular membranes and organelles. The application of anther plunge-freezing and cryo-SEM additionally enabled the observation of all layers of the anther wall without the artifacts caused by tissue embedding. This approach revealed novel information about the poorly characterized middle layer, locule fluid and the morphology of tapetal cells. This study highlights the numerous cellular events required to assemble the multi-layered pollen wall, and the importance of optimized sample preparation when analyzing pollen and tapetum development, which will be particularly important in mutants with altered tapetum and/or microspore wall development.   The dependence of developing pollen grains on tapetal cells has been well established (Ariizumi and Toriyama, 2011; Liu and Fan, 2013). However, a detailed analysis of Arabidopsis tapetum ultrastructure through pollen development is lacking in the literature, likely due to the challenges associated with preservation.  The TEM analysis in this study revealed several important features of tapetal cells.  First, the tapetum lacks a cell wall as early as the tetrad stage (Figure 2.1 and Figure 2.3), as hypothesized by Owen and Makaroff (1995).   Second, the tapetum ultrastructure during exine deposition showed no signs of an active endomembrane secretion system, e.g. post-Golgi vesicles, consistent with a plasma membrane transporter mediated export of sporopollenin components.  These observations support the model that ATP binding cassette (ABC) transporters, such as Arabidopsis ABCG26 and rice ABCG15 (Quilichini et al., 2010; Qin et al., 2013), export sporopollenin components from the tapetum. Finally, these data provide evidence that tapetal cells were distinguishable into the tricellular pollen stage   55 (anther stage 12; Figure 2.1H), in contrast to previous reports indicating that tapetal cells were absent from locules with tricellular pollen (Sanders et al., 1999).    The most striking features of the tapetum revealed in this TEM analysis were tapetosomes and elaioplasts, two highly specialized organelles that distinguish late-stage tapetal cells from all other plant cells, and are the major contributors to pollen coat formation. Tapetosomes and elaioplasts are typically described as spherical and approximately 3 µm in diameter (Wu et al., 1997). In contrast, we found that the size, shape and number of tapetosomes varied dramatically based on the developmental stage sampled. They appeared numerous and round in bicellular pollen stages, and amorphous and branching through a large portion of the tapetum cytoplasm in tricellular pollen stages.   The biogenesis of tapetosomes has been proposed to resemble oil body biogenesis (Hsieh and Huang, 2004), but Arabidopsis tapetosomes are morphologically distinct, consisting of tightly bunched membrane tubules in a dense matrix rather than discrete spherical oil bodies as found in seeds (Figure 2.8).  While tapetosomes were contiguous with ER, their relationship to the dilated rough ER cisternae and the abundant smooth ER is presently unknown.  In Brassica napus (Brassica), cell fractionation, biochemical analysis and confocal microscopy have revealed that flavonoids co-localize with ER and tapetosomes, while alkanes were associated with tapetosome oil bodies (Hsieh and Huang, 2007). Tapetosome oil droplets additionally contain triacylglycerols (TAGs), and are believed to be coated by a layer of phospholipids and oleosin proteins (Hsieh and Huang, 2004).  The distribution of flavonoids and TAGs within the branching tubules and rod-like structures observed in Arabidopsis tapetosomes are not known. The two distinct regions of the tapetosomes observed in our study, i.e. branched tubules and rod-like clusters, are consistent with earlier work where Platt et al., (1998) observed that chemically-fixed Brassica tapetosomes (then called lipid bodies) had two distinct regions, as differentiated by their relative electron-densities, although in that study, the substructure was not clear.   As in tapetosomes, substructural features within Arabidopsis tapetum elaioplasts complement the previous biochemical and ultrastructural data. In Brassica napus, elaioplasts contain abundant steryl esters, which constitute the predominant neutral lipids in these specialized plastids and are believed to form the contents of globuli (Wu et al., 1999; Platt et al., 1998).  Based on their presence and abundance in elaioplasts, phospholipids, glycolipids,   56 monogalactosyldiacylglycerols (MGDG) and plastid-associated proteins (PAPs) are additionally predicted to associate with and/or maintain the surface of globuli (Wu et al., 1999; Kim et al., 2001). We identified two components of varying texture and electron-density within elaioplast globuli not previously described. Based on the chemical data available, we predict that the electron-dense meshwork we observed in globuli represents lipid and PAP-based protein scaffold that encases electron-lucent (or extracted) steryl esters.  Instead of rectangular crystals within globuli of elaioplasts (Platt et al., 1998), the electron-translucent areas of globuli exhibited a variety of shapes in our samples (Figure 2.8).   Although the cryo-fixation methods we employed preserved novel features of tapetum cellular content and membranes, the fragility of middle layer cells still presented challenges. Although the middle layer is often cited as a cell layer that ruptures in the tetrad stage of development (Sanders et al., 1999), there is evidence that the middle layer cells persist into the tricellular stage of pollen development in Arabiopsis (Owen and Makaroff, 1995).  With cryo-SEM, it was clear that middle layer cells persist after the release of free microspores and into the late stages of uninucleate microspore development, where they appear alive based on their subcellular contents. Although the function of middle layer cells in Arabidopsis pollen development is unknown, our data suggest that their cell walls, as in tapetal cells, may be reduced or altered. In our cryo-preserved samples and chemically fixed samples (Owen and Makaroff, 1995), there was spatial separation of tapetal cells from the anther wall.  Given that cryo-fractured samples examined by cryo-SEM did not show this separation, the previously observed separation may be an artifact of the embedding process, rather than a native state that aids secretion, as proposed previously (Owen and Makaroff, 1995).   While most cell layers of the sporophytic anther wall have defined roles in pollen development and pollen cell wall formation, the medium that fills the locule and is in direct contact with both the tapetal cells and developing pollen grains, is poorly understood. Our images clearly show that locule fluid was present from the tetrad stage through to the early bicellular pollen stage, after which it appeared to become less dense, and was minimal or absent from locules filled with tricellular pollen grains. Interestingly, this is also the time frame in which sporopollenin is likely to traffic from tapetal cells to microspores, and, during this time frame, lipid transfer proteins are abundant in the locule fluid (Huang et al., 2013). Although   57 some studies have suggested that traffic of pollen wall components occurs through the locule in vesicles, we found no evidence to support this hypothesis. While the locule fluid appears homogeneous when embedded anthers are sectioned for light microscopy (Figure 2.1) and TEM (Figure 2.4-2.8), it exhibited a lattice-like appearance in cryo-fractured anther locules at comparable stages (Figure 2.3). This is likely due to the formation of ice crystals upon freezing; however, the ordered nature of these ice crystals may point to structural organization within the locule, such as the viscin thread-like structures between microspores and tapetal cells described in Betula pendula anther locules (Rowley and Morbelli, 2009). If this interpretation is correct, the locule in Arabidopsis, or other flowering species with secretory tapeta, and its protein content, could aid traffic to the developing microspores.    The uninucleate microspore stage represents a critical transition when the sporopollenin framework assembles.  Intermediate stages of this process were observed, suggesting that visible transitions occur in the exine as the framework matures. Judging by the maturity of the patterned wall, the first step in sporopollenin polymerization is characterized by lamellae within the exine wall, followed by the transition of the exine into the homogenous baculae and tecta in all subsequent stages (Figure 2.4).  The lamellar appearance in exine formation in some species has been hypothesized to indicate early sporopollenin polymerization, much like the models for suberin and cutin assembly, but early lamellar exine has not been previously described in Arabidopsis or other dicot microspore walls previously (Scott, 1994; Bernards, 2002). In Poa annua and Lilium sp., the initial lamellar appearance of the exine is replaced by a homogenous exine and is hypothesized to indicate the progression from partially polymerized to polymerized acetolysis-resistant sporopollenin (Rowley, 1962; Dickinson and Heslop-Harrison, 1968). This hypothesis is supported by our data, as the lamellar appearance associated with exine was only observed in early free microspores (Figure 2.4H), while the homogenous sporopollenin wall was observed in all examined mid-late stage uninucleate microspore walls (Figure 2.4K). While the exact time frame over which sporopollenin export and assembly occurs is not known, the sporopollenin framework built in uninucleate microspore stages did not appear altered in subsequent stages, suggesting that this wall is complete by late-uninucleate pollen development stages. Recent analyses of the timing of gene expression and mutants phenotypes in the   58 polyketide synthase/sporopollenin biosynthetic pathway are consistent with this model (de Azevedo Souza et al., 2009; Kim et al., 2010; Grienenberger et al., 2010).    After tapetum programmed cell death, the deposition of the pollen coat marks the completion of wall formation on Arabidopsis tricellular pollen grains. Using our fixation methods, mature pollen grains exhibited extensive heterogeneity within the pollen coat. The structures of the coat observed here are likely to include neutral lipids, alkanes, and very long-chain wax esters (Preuss et al., 1993; Ariizumi et al., 2003; Fiebig et al., 2000), as well as flavonoids (Hsieh and Huang, 2007), hydroxycinnamoyl spermidine metabolites (Grienenberger et al., 2009), and proteins such as lipid transfer proteins, oleosins and self-incompatibility proteins (Huang et al., 2013b; Piffanelli et al., 1998).  In conclusion, we applied cryo-fixation and diverse imaging techniques to the analysis of anther, pollen wall, and tapetum development in wild-type Arabidopsis anthers. These data provide a baseline of the morphological features in developing Arabidopsis anthers, especially in the tapetum. Future application of these methods to functional studies of mutant plants with altered pollen wall formation will be particularly revealing.   2.4 Materials and methods  2.4.1 Plant growth   Wild-type Arabidopsis (Arabidopsis thaliana, Columbia-0) seeds were sterilized in ethanol and germinated on agar plates with Murashige and Skoog medium, pH 5.7. Seeds were grown under 24 hour light at 28ºC for seven days, then transferred to soil (Sunshine mix 4, Sungrow Horticulture), and grown to maturity at 20ºC under long day light conditions (20ºC under 16/8 hour light/dark cycles).  2.4.2  cryo-scanning electron microscopy analysis   Stamens were dissected from multiple immature buds (0.7 to 1.0 mm long) and submerged in sterile water within copper sample holders (type B hats, Ted Pella Inc.). The   59 stamen-filled chamber was sealed with a second hat and secured to a cryo-EM specimen block with Tissue-Tek optimal cutting temperature compound (Ted Pella, Inc.).  The block was submerged in liquid nitrogen, transferred under vacuum to a fracturing chamber and fractured by removing the upper hat from the sample. The sample block was moved under vacuum (using the VCT100 system, Leica Microsystems) to a pre-cooled cryo-SEM (-135ºC, S-4700 Field Emission SEM, Hitachi), sublimed to remove surface ice (-95ºC for 20 minutes), and imaged in an uncoated state at -118ºC with an accelerating voltage of 5.0kV.   2.4.3  High-pressure freezing and freeze substitution   For high-pressure freezing, buds representing a range of anther developmental stages were dissected from flowering wild-type plants (4-6 weeks old). Perianth was removed from each bud, and remaining stamens and gynoecium were submerged in extracellular cryoprotectant (0.2 M sucrose between two type B hats that formed a chamber, Ted Pella, Inc.). High-pressure freezing was performed immediately (EM HPM 100, Leica Microsystems), and samples were transferred under liquid nitrogen to cryovials of freeze substitution medium containing 2% w/v osmium tetroxide in anhydrous acetone with 8% v/v dimethoxypropane. Sample vials underwent cryosubstitution within a slurry of dry ice and acetone at -80ºC for three days, then the vials were warmed to -20ºC by the end of one week. Sample vials were warmed to room temperature, and the substitution medium was replaced with acetone. Samples were embedded in Spurr’s epoxy resin over four days, reaching 10% v/v resin in acetone on day 1, 60% on day 2, and 100% resin on day 4. Resin-embedded samples were transferred to capsules (BEEM embedding capsules #70020, Electron Microscopy Sciences) and polymerized at 60ºC for 2 days. Four sets of wild-type stamen samples were independently prepared by the methods described and used for light microscopy and TEM analyses.   2.4.4 Light microscopy and transmission electron microscopy    Resin-embedded samples were sectioned to a thickness of 0.3–0.5 µm for light microscopy and 50-70 nm for TEM, with a microtome (Ultracut E, Leica-Reichert) and diamond   60 knife (Ultra 45º, Diatome). For viewing and staging anther development by light microscopy (with a Leica DMR microscope), sections were stained on glass slides with Toluidine Blue in 1% sodium borate, and sealed beneath a permount-attached coverslip. TEM sections were collected on copper 100-hex grids with a 0.5% Formvar coating with silver-gray interference colour, and stained, first with uranyl acetate (2% in 70% (v/v) methanol, 15 minutes), then with Reynolds lead citrate (6-8 minutes). Grid-mounted samples were viewed on a Hitachi H7600 TEM at an accelerating voltage of 80.0kV. Images were captured with an AMT Advantage (1 megapixel) CCD camera (Advanced Microscope Technologies).    61 Chapter 3: ATP-binding cassette transporter G26 is required for male fertility and pollen exine formation in Arabidopsis  3.1  Introduction    The central importance of spores in plant reproduction and the inherent stresses of a terrestrial environment have led to the evolution of a toughened wall surrounding the male gametophyte in angiosperms and gymnosperms and haploid spores in nonflowering plants. Sporopollenin, a structurally robust biopolymer, gives the outer walls of spores and pollen their unparalleled strength and resistance to terrestrial stresses (Scott, 1994).   The composition of sporopollenin is thought to include phenolics and polyhydroxylated unbranched aliphatics, coupled by ester and ether linkages, which provide this biopolymer with its characteristic resistance to chemical degradation (Guilford et al., 1988; Wiermann et al., 2005; Scott et al., 2004). Through the application of solid-state NMR, aromatic, aliphatic, and oxygen functionalities have been identified in the sporopollenin biopolymer (Hemsley et al., 1994; Ahlers et al., 1999; 2000; 2003). However, despite the technologies available, our knowledge of the polymer structure of sporopollenin is far from complete.   Investigation of anther and microspore development by light microscopy and transmission electron microscopy (TEM) have improved our knowledge of pollen wall formation and provided a context for understanding the mechanism of sporopollenin formation. In Arabidopsis (Arabidopsis thaliana) anther development, 14 stages can be distinguished based on distinct cellular events (Sanders et al., 1999). In stages 1 through 4, archesporial cells of the anther primordia divide periclinally to produce inner sporogenous cells and outer parietal cells, from which all anther cell types differentiate: epidermis, endothecium, middle layer, tapetum, and microspore mother cells destined to become pollen grains. Microspore mother cell meiosis produces haploid microspore tetrads, which are encased in callose (Dong et al., 2005; Nishikawa et al., 2005). At the tetrad stage (stage 7), a microspore-derived matrix of polysaccharides known as the primexine forms between the microspore and surrounding callose, followed by the emergence of probaculae, which serve as a scaffold for sporopollenin deposition (Suzuki et al., 2008; Wilson and Zhang, 2009). After the release of microspores from callose-encased tetrads in   62 stage 8, the sporopollenin-based exine wall forms by the deposition of sporopollenin generated in the tapetum, producing the sculptured baculae and tecta of the exine. During the continued maturation of microspores into tricellular pollen grains, a cellulosic and pectin-rich intine forms between the exine and microspore plasma membrane (Blackmore et al., 2007). The final component of the pollen wall, the waxy pollen coat or tryphine, is deposited on and within the crevices of the mature pollen grain exine in the late stages of pollen development, coinciding with tapetum degeneration in stage 10 (Blackmore et al., 2007; Grienenberger et al., 2009).   The production of viable pollen requires precise spatial and temporal coordination of both gametophytic and sporophytic developmental and metabolic events (Blackmore et al., 2007). The sporophytic tapetum, a single cell layer encasing the anther locule, plays key functions in pollen development. It supplies nutrients, structural components, and enzymes essential to the survival and development of microspores (Blackmore et al., 2007). Specifically, the tapetum functions in the production and secretion of structural components of the pollen wall, both as the source of exine sporopollenin precursors to early uninucleate microspores and later as the source of waxy pollen coat components (Heslop-Harrison, 1968a; Bedinger, 1992; Wilson and Zhang, 2009). In Arabidopsis, rice (Oryza sativa), wheat (Triticum aestivum), and other plants with secretory tapeta, the spatial separation of the tapetum from developing microspores requires the movement of substances, including sporopollenin components, from the tapetum to the developing microspore.   Analyses of male-sterile mutants defective in pollen wall formation, primarily in the model plant Arabidopsis, have revealed genes required for normal sporopollenin biosynthesis or deposition, including MALE STERILE2 (MS2), DEFECTIVE IN EXINE PATTERNING1 (DEX1), DEFECTIVE IN EXINE PATTERNING2 (DEX2/CYP703A2), FACELESS POLLEN1 (FLP1/WAX2/YRE/CER3), NO EXINE FORMATION1 (NEF1), TRANSIENT DEFECTIVE EXINE (TDE1), ACYL-COA SYNTHETASE5 (ACOS5), DIHYDROFLAVONOL 4-REDUCTASE-LIKE1 (DRL1), CYP704B1, LESS ADHERENT POLLEN3 (LAP3), and CALLOSE SYNTHASE5 (CALS5/LAP1); (Aarts et al., 1997; Paxson-Sowders et al., 2001; Ariizumi et al., 2003; 2004; 2008; Dong et al., 2005; Nishikawa et al., 2005; Morant et al., 2007; Rowland et al., 2007; Souza et al., 2009; Dobritsa et al., 2009b; 2009a; Tang et al., 2009). The expression of MS2, ACOS5, CYP703A2, CYP704B1, and DRL1 in the tapetum at the time of exine formation, together with   63 acetolysis-sensitive exine or the apparent absence of exine on developing microspores, suggest that these genes are involved in sporopollenin biosynthesis (Aarts et al., 1997; Morant et al., 2007; Souza et al., 2009; Dobritsa et al., 2009b; Tang et al., 2009). These mutants have provided clues regarding the biosynthesis of sporopollenin and have defined a key role for the tapetum as the site of sporopollenin biosynthesis in pollen wall formation.   Based on their putative roles in the export of extracellular polymer components, such as cutin precursors (Bird, 2008), we hypothesized that ATP-binding cassette (ABC) transporters could be required for sporopollenin export from the tapetum to the developing microspore. ABC transporters represent one of the largest protein superfamilies, known primarily for their function in substrate translocation across membranes (Theodoulou, 2000; Higgins and Linton, 2001). A wide range of transport capabilities have been discovered for plant ABC transporters, including ion, lipid, hormone, secondary metabolite, xenobiotic, and peptide transport (Theodoulou, 2000; Rea, 2007). In Arabidopsis, functions have been reported for four of the 28 ABC transporters in the G subfamily of half-size transporters: AtABCG12/ WBC12/CER5 and AtABCG11/WBC11/COF1 are required for lipid export to the cuticle (Pighin et al., 2004; Bird et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007), AtABCG19/WBC19 over-expression increases antibiotic resistance (Mentewab and Stewart, 2005), and AtABCG25 is involved in abscisic acid transport (Kuromori et al., 2010).   In this study, we identified a gene encoding an ABC transporter, ABCG26, that is tightly coexpressed with genes required for sporopollenin biosynthesis and is highly expressed in the early flower bud, supporting a role for this gene in pollen wall formation. We examined the function of ABCG26 in pollen wall development using reverse genetics and found that ABCG26 is required for normal male fertility, exine formation, and pollen maturation and that ABCG26 is expressed in the tapetum at the time of exine formation.         64 3.2 Results  3.2.1 ABCG26 is required for male fertility   To test the hypothesis that ABC transporters are required for the export of sporopollenin precursors or other constituents of the exine from the tapetum to developing microspores, it was necessary to identify candidate ABC transporters out of the over 120 annotated ABC transporter genes in the Arabidopsis genome (Verrier et al., 2008). Since ACOS5 is preferentially expressed in the tapetum and encodes an acyl-CoA synthetase required for sporopollenin biosynthesis (Souza et al., 2009), this gene was used for coexpression analysis to identify potential Arabidopsis transporters on the PRIMe database ( The Arabidopsis ABCG26 (formerly WBC27) gene At3g13220 shows a high coefficient of coexpression in the tissue and development version 1 data set (r2 = 0.95), along with several other genes involved in sporopollenin biosynthesis (Souza et al., 2009). ABCG26 encodes a predicted half-size ABC transporter in the G subfamily, formerly called the White-Brown Complex subfamily. The highly conserved protein architecture of ABCG26 is typical of this subfamily, where dimerization of two half-transporters produces a functional transport unit with two transmembrane domains and two ABC domains (Higgins and Linton, 2001; Verrier et al., 2008).   To test a role for ABCG26 in male fertility and pollen development, three ABCG26 T-DNA insertion lines (SALK_062317, SAIL_318_B09, and SAIL_885_F06, named abcg26-1, abcg26-2, and abcg26-3, respectively) were obtained from the Arabidopsis Biological Resource Center (Alonso, 2003). PCR genotyping identified plants homozygous for each ABCG26 T-DNA insertion, and sequencing confirmed insertion in the fifth exon (abcg26-1) or the 5´ untranslated region (abcg26-2, abcg26-3) of ABCG26 (Figure 3.1A). Plants homozygous for each of the mutant alleles exhibited highly similar mutant phenotypes of severe impairment in male fertility (Figure 3.2), and the abcg26-1 mutant was used for further experiments.     65  Figure 3.1: Identification of Arabidopsis ABCG26 T-DNA insertion alleles. A) Diagrammatic representation of ABCG26 with T-DNA insertions in exon 5 (abcg26-1, SALK_062317) and the 5’ untranslated region (abcg26-2, SAIL_318_B09; abcg26-3, SAIL_885_F06). Black boxes represent exons, and the positions of T-DNA insertions are indicated by triangles. Arrowheads labeled P8 to P11 represent primer positions used in (B). B) RT-PCR detection of ABCG26 transcripts in wild-type (Col-0) and abcg26-1 flowers. ACTIN2 expression was monitored as a control. C) Mature wild-type (left) and abcg26-1 mutant (right) plant morphology. Terminal siliques are shown (insets). (D, E) Mature wild-type and abcg26-1 mutant flowers, respectively. Bars = 200 μm. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   Reverse transcription (RT)-PCR analysis revealed a strong reduction of ABCG26 mRNA in the abcg26-1 mutant relative to the wild-type, suggesting that abcg26-1 is a loss-of-function allele of ABCG26 (Figure 3.1B). In homozygous abcg26-1 mutant plants, most siliques failed to mature and produce seeds (Figure 3.1C), and most abcg26-1 anthers appeared brown, dry, and lacked any obvious pollen grains (Figure 3.1E). However, occasional abcg26-1 anthers with bulbous yellow surfaces consistent with pollen that was not shed were identified, and occasional fully fertile siliques were observed, suggesting a low rate of fertile pollen formation. Wild-type pollen applied to homozygous abcg26-1 stigmas on flowers with brown, dry anthers produced fertile progeny, demonstrating normal female fertility and strong male sterility of abcg26-1. No other morphological differences between mature wild-type and abcg26-1 plants were observed, except that abcg26-1 mutant plants flowered for a longer time and grew taller (50 +/- 7 cm at   66 maturity [mean +/- SD; n = 12 plants] versus 29 +/- 2 cm at maturity for the wild-type [n = 12 plants]; Figure 3.1C). Growth under elevated humidity did not recover fertility in abcg26-1 plants.    Figure 3.2: Phenotypic comparison of Arabidopsis T-DNA insertion alleles. A) From left to right: mature wild-type (Col-0), abcg26-1, abcg26-2, and abcg26-3 mutant plant morphologies. B) From left to right: mature wild-type (Col-0), abcg26-1, abcg26-2, and abcg26-3 mutants representative terminal siliques. Arrows show representative small, sterile siliques. C) Characterization of fertility in wild-type (Col-0), abcg26-1, abcg26-2, and abcg26-3 mutant   67 plants. Fertility was measured according to seed production per silique along the primary stem (n=12 plants for each genotype). Small siliques contained 1.3 +/- 0.4 seeds, medium siliques 13.5 +/- 0.8 seeds, and large siliques 48.3 +/- 2.3 seeds. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   To confirm that the observed male-sterile phenotype is caused by the abcg26-1 T-DNA insertion mutation, we cloned ABCG26 cDNA from developing flower buds in order to complement the abcg26-1 mutant with a wild-type copy of ABCG26. However, during the isolation of the expected annotated ABCG26 cDNA, a second transcript, ABCG26.2 (At3g13220.2), was found, and sequencing revealed that it corresponded to an alternatively spliced ABCG26 variant resulting from the retention of intron 7 in the transcript (Figure 3.3A). The retention of intron 7, which encodes multiple in-frame stop codons, is predicted to produce a truncated protein if it were translated.     Figure 3.3: Flower preferred expression of ABCG26 splice variants. A) Diagrammatic representation of ABCG26 splice variants ABCG26.1 (At3g13220.1) and ABCG26.2 (At3g13220.2). Boxes and lines represent exons and introns, respectively. B) Expression of ABCG26 splice variants in various organs, assayed by quantitative reverse transcription-PCR.   68 Expressed levels in the measured plant organs are shown as fold-change relative to ABC26.1 flower expression (set at 100), and were normalized using ACTIN2 as a reference gene. FLWR, flower; 7D, 7 day old seedling; YS, young stem; YL, young leaf; MS, mature stem; ML, mature leaf; MR, mature root. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   Constructs containing either the major or minor splice variant of ABCG26 cDNA driven by the cauliflower mosaic virus 35S promoter were generated and transformed into the homozygous abcg26-1 mutant line. The presence of each construct, as well as the T-DNA insertion in the T1 progeny of homozygous abcg26-1 plants, was confirmed by PCR. Figure 3.4 shows the fertility phenotypes of T1 progeny relative to those of wild-type ecotype Columbia (Col-0) and abcg26-1 plants, assayed by silique development and seed set. Although occasional fertile siliques were observed in abcg26-1 mutant plants, very few fully developed siliques were observed, and clear fertility differences were observed between multiple mutant and wild-type individuals (85% of siliques on wild-type plants were classified as large, indicative of full fertility, whereas 1% of siliques on abcg26-1 plants were large; Figure 3.4). When assayed in a similar manner, T1 progeny from multiple lines expressing cDNA for the major transcript (ABCG26.1) restored wild-type fertility levels to the abcg26-1 mutant line, with numerous fully developed siliques and normal seed set (Figure 3.4). Wild-type fertility levels were also restored in multiple lines expressing cDNA for the minor transcript (ABCG26.2), which contains the retained intron. This could suggest that a truncated protein was functional, surprising given the high conservation of ABC transporter protein architecture. Alternatively, when the transcript with the retained intron was expressed, functional protein could have been produced by splicing out of the retained intron with its premature stop codon. RT-PCR was performed to assess whether the lines complemented with ABC26.2 contained the full-length or alternative transcripts. ABCG26.1 transcript was detectable in such lines (Figure 3.5), suggesting that complementation by the ABCG26.2 construct was due to the accumulation of ABCG26.1 mRNA and ABCG26.1 protein rather than by truncated ABCG26.2 protein.     69  Figure 3.4: Genetic complementation of abcg26-1 mutation. A) Characterization of fertility in wild-type (Col-0), abcg26-1 mutants, and abcg26 mutants containing the full length ABCG26.1 or ABCG26.2 transgene under control of the CAMV-35S promoter. Fertility was measured according to seed production per silique along the primary stem. Small siliques contained 1.3 +/- 0.4 seeds (n=20), medium siliques 13.5 +/- 0.8 seeds, and large siliques 48.3 +/- 2.3 seeds. B) From left to right: representative terminal siliques from wild-type plants, abcg26-1 mutant plants, abcg26-1 mutant plants expression the 35S:ABCG26.1 transgene, and abcg26-1 mutant plants expression the 35S:ABCG26.2 transgene. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   70   Figure 3.5: Accumulation of ABCG26.1 and ABCG26.2 transcripts in abcg26-1 mutants expressing the ABCG26.2 transgene. Lane 1, 1kb DNA ladder; lanes 2 and 3, no template controls; lanes 4 and 5, cDNA template from abcg26-1 seedlings expressing the ABCG26.2 transgene. Primers P13 and P14 were used for ABCG26.2 transcript amplification (lanes 2 and 4), and primers P12 and P13 were used for ABCG26.1 transcript amplification (lanes 3 and 5), as described in Materials and Methods. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   3.2.2 ABCG26 expression pattern   To evaluate the expression pattern of ABCG26, we queried the Genevestigator Arabidopsis expression database (Zimmermann et al., 2005). Results suggested expression of ABCG26 specifically in immature flower buds, and ABCG26 expression was absent from apetala3 and agamous mutants that lack stamens, suggesting a stamen-specific expression pattern. To confirm the flower-specific expression pattern of ABCG26, the expression levels of both ABCG26 splice variants were tested in various Arabidopsis organs by quantitative RT-PCR (Figure 3.3B). These results revealed a highly flower-preferred expression pattern for both ABCG26 splice variants, with little or no transcripts detected in other organs. ABCG26.1 is the major transcript and accumulates approximately 3-fold higher relative to ABCG26.2.    71  To determine whether ABCG26 is expressed in sporophytic or gametophytic tissues, we used in situ hybridization. Figure 3.6 shows that ABCG26 transcripts were transiently localized to tapetal cells in developing anthers. Tapetum-localized expression was first observed at stage 6 (Sanders et al., 1999), when microspore mother cells became apparent in locules. Maximum expression was observed at stages 7 and 8, corresponding to tetrad formation and free microspore release, respectively, and expression declined at stages 9 and 10 and disappeared by stage 11 in conjunction with tapetum degeneration. Relative to the sense RNA hybridization controls (shown for stages 8 and 11; Figure 3.6), some ABCG26 signal also appeared in developing microspores. The timing of tapetum-localized ABCG26 expression is similar to that of ACOS5, which encodes a key enzyme in sporopollenin biosynthesis, and coincides with the onset of exine formation and sporopollenin biosynthesis (Souza et al., 2009), suggesting that ABCG26 could be involved in this process.    Figure 3.6: ABCG26 tissue-specific expression pattern during microspore development. In situ hybridization was used to localize ABCG26 mRNA in wild-type (Col-0) developing flower bud sections using an ABCG26 gene-specific antisense probe and a control ABCG26 sense probe. Dark staining demarks sites of probe hybridization. From left to right: stage 6 locules contained microspore mother cells and a hybridization signal in the tapetum was detectable; stage 7 and 8 locules with tetrads and free microspores, respectively exhibited maximum hybridization in the tapetum; tapetum hybridization weakened in stages 9 and 10, and hybridization signal disappeared by stage 11 with tapetum degeneration; hybridization with the control sense probe produced no signal in stage 8, but staining was detected in mature pollen grains by stage 11. Anther developmental stages are numbered according to Sanders et al. (1999). M, microspores; MMC, microspore mother cell; PG, pollen grain; T, tapetum; Td, tetrad. Bars = 20 μm. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (     72 3.2.3 Microspore development is impaired in abcg26-1   To identify the stage at which pollen development is affected in the abcg26-1 mutant, we used light microscopy to compare anther and microspore development in wild-type (Col-0) and abcg26-1 flower buds using the stages defined by Sanders et al. (1999; Figure 3.8). In all early stages, including the completion of meiosis in stage 7, both wild-type and abcg26-1 microspore tetrads were encased by callose and appeared normal (Figure 3.8, A and B). However, by stage 9, characterized by the release of free microspores and formation of the exine wall, abcg26-1 anthers differed significantly from the wild-type. Figure 3.8D shows that free microspores in abcg26-1 exhibited signs of incipient degeneration. In stages 10 to 11, the degeneration of the tapetum and formation of tricellular mature pollen proceeded in wild-type anthers (Figure 3.8E), while pollen grains in the abcg26-1 mutant generally lacked cytoplasm and exhibited misshapen morphologies (Figure 3.8F). By stage 12, all abcg26-1 mutant anthers observed in multiple sections were devoid of pollen grains, while abundant mature pollen was evident in wild-type anthers (Figure 3.8, G and H). Despite impaired microspore development in abcg26-1, no additional changes in anther morphology were observed in the tapetum or integuments, and growth and expansion proceeded as in wild-type plants. These data suggest that impaired microspore development occurs in the abcg26-1 mutant at the developmental stage, when sporopollenin deposition and exine formation normally occur.       73  Figure 3.7: Key stages in Arabidopsis pollen wall development relative to ABCG26 expression. Stages 5 through 13 represent anther development, as described by Sanders et al., 1999. Key stages of pollen development are listed immediately below the stages of anther development. Relevant pollen wall components and events in pollen development are listed on the left. A solid black line is used to illustrate the currently available knowledge or accepted model for the formation and appearance of pollen wall components and the timing of meiosis and mitosis, relative to the stage of anther development. Dashed black lines represent information that remains unconfirmed or highly variable. The timing of ABCG26 expression in the tapetum, based on in situ hybridization (Figure 3.6), is shown below in red; solid red line for maximal expression, dashed red line for weak expression. MMC, microspore mother cell; Early UC, early unicellular; Late UC, late unicellular; BCP, bicellular pollen; TCP, tricellular. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (    74  Figure 3.8: Microspore development in wild-type (Col-0) and abcg26-1 mutant anthers. Anther developmental stages are according to Sanders et al. (1999). Images in the left panels (A, C, E, and G) show wild-type anthers and those in the right panels (B, D, F, and H) show abcg26-1 anthers. A and B, Microspore development appeared identical in stage 7 wild-type and abcg26-1 anthers, when a callose wall encases tetrads of microspores. C and D, Following the release of free microspores from tetrads at stage 8, abcg26-1 microspores showed signs of degradation not observed in wild-type microspores. E and F, In stages 10 and 11, characterized by tapetum degeneration, severe abnormalities in abcg26-1 microspores were visible in comparison with wild-type microspores. G and H, At stage 12, pollen grains were not observed in abcg26-1 mutant anthers, whereas tricellular pollen grains were present in wild-type anthers. Bars = 50 μm. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   75  3.2.4 Pollen wall and tapetum development in abcg26-1    The observed degeneration of microspores at the time of pollen wall formation indicated that defects in pollen wall and exine formation could be the primary cause of pollen abortion and male infertility in abcg26-1 mutants. To examine pollen wall and tapetum morphology at higher resolution in the abcg26-1 mutant relative to wild-type plants, scanning electron microscopy (SEM) and TEM were used. The surface of wild-type pollen, viewed with SEM, exhibited a characteristic reticulate exine (Figure 3.9). In contrast, SEM revealed that abcg26-1 anthers occasionally released small amounts of pollen or pollen-like structures with smooth surfaces and collapsed morphology (Figure 3.9B). This loss of normal pollen cell wall morphology and collapse of the microspores suggests a severe pollen wall defect, but since occasional fully fertile siliques were observed in the mutant, low numbers of fertile pollen grains were likely formed.     76  Figure 3.9: Pollen wall structure in wild-type (Col-0) and abcg26-1 mutant plants. A and B, Scanning electron micrographs of pollen from wild-type (A) and abcg26-1 mutant (B) plants. C to F, Transmission electron micrographs of chemically fixed sections taken from wild-type (C and E) and abcg26-1 mutant (D and F) anthers between stages 9 and 10 of anther development (late unicellular stage). C and D, Low-magnification images showing tapetal cells, locules, and free microspores. E and F, High-magnification images showing pollen wall ultrastructure and locular inclusions (unlabeled arrows), which were only observed in the abcg26-1 mutant. Ba, Bacula; DEx, defective exine; Ex, exine; Lo, locule; M, microspore; Ne I, nexine; T, tapetum; Te, tectum. Bars = 5 μm (A–D) and 100 nm (E and F). Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   77   Since the sporopollenin-based exine of pollen grains is deposited by stage 10 of anther development, the cell structure of the developing anthers and tapetum was examined using TEM focusing on this stage. Uninucleate microspores of chemically fixed abcg26-1 anthers at stage 10 exhibited thin walls and apparently defective exine deposition, as compared with wild-type microspores that had pollen walls with a typical thick reticulate exine (Figure 3.9, C and D). At higher magnification, tectum and bacula structures of wild-type exine deposited on the underlying nexine were obvious in wild-type microspores (Figure 3.9E). In contrast, walls of abcg26-1 microspores were devoid of baculae and tectum, containing instead a thin, amorphous defective wall layer (Figure 3.9F). No differences in the tapetum of abcg26-1 anthers compared with wild-type tapetal cells at the equivalent developmental stages were observed in chemically fixed material. Interestingly, coil-shaped electron-dense inclusions were consistently observed in abcg26-1 locules but were never observed in the locules of wild-type anthers (Figure 3.9, E and F).   From light microscopy analyses, pollen development showed signs of disruption in abcg26-1 early free uninucleate microspores, in coordination with the onset of exine formation. In the images obtained after chemical fixation, however, a clearly delineated primexine was not observed in wild-type plants. To improve the ultrastructural preservation of microspores and facilitate high-resolution imaging of both primexine and exine analysis in developing wild-type and abcg26-1 microspores, we used high-pressure freezing (HPF) and freeze substitution to preserve stamens during anther development. At stage 5, both wild-type and abcg26-1 mutant microspore mother cells were surrounded by a uniform matrix between the microspore primary cell wall and the plasma membrane (Figure 3.10, A and B). By stage 7, tetrads encased in callose were present in both the wild-type and abcg26-1 and appeared identical, showing primexine and probacula deposition between the plasma membrane and callose wall (Figure 3.10, C and D). Shortly after their release from tetrads at stage 8, however, abcg26-1 early uninucleate microspores failed to develop well-defined baculae, which were obvious in wild-type microspores at the same stage (Figure 3.10, E and F). The nexine layer in wild-type and abcg26-1 mutant anthers was observed as a faint electron-translucent layer immediately outside the plasma membrane in these HPF-fixed specimens (Figure 3.10, E–H). At the late unicellular stage   78 (stage 9), wild-type microspores had a nearly complete pollen wall composed of nexine, bacula, and tectum (Figure 3.10G). However, abcg26-1 microspores at the same stage lacked any defined bacula and tectum structure (Figure 3.10H); instead, ill-defined electron-dense deposits were observed on the surface surrounding the nexine layer. The absence of defects in primexine but visible defects in exine formation in the abcg26-1 mutant demonstrate that contributions of primexine from the microspore were not affected by ABCG26 mutation, while the deposition of tapetum-derived sporopollenin was severely affected.     79    80 Figure 3.10: Transmission electron micrographs of microspore primexine and exine formation in wild-type (Col-0) and abcg26-1 anthers. Pollen wall development between stages 5 and 8 of anther development is shown in sections taken from wild-type (A, C, E, and G) and abcg26-1 (B, D, F, and H) flowers. A and B, Microspore mother cells in stage 5. C and D, Tetrads in stage 7. E and F, Early unicellular microspores. G and H, Middle unicellular stage microspores. Ba, Bacula; CW, callose wall; Lo, locule; M, microspore; MMC, microspore mother cell; Ne I, nexine, PM, plasma membrane; Te, tectum. Arrowheads indicate probacula. Bars = 500 nm (A–H) and 2 μm (I and J). Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   No differences in abcg26-1 tapetal cell ultrastructure, relative to wild-type tapetal cells, were observed in TEM samples (Figure 3.11). In stages 5 through 9, wild-type and abcg26-1 tapetal cells were binucleate, cytoplasmically dense with numerous vesicles, and contained no inclusions. However, as initially observed in TEM micrographs of chemically fixed anthers (Figure 3.9, D and F), HPF-fixed abcg26-1 anther locules after stage 7 (tetrad stage) consistently contained electron-dense inclusions (commonly between one and 10 inclusions per locule per section) that were never observed in wild-type anther locules (Figure 3.12). These structures were often in the shape of rings or flattened rings (Figure 3.12, A–C) and at higher magnification (Figure 3.12, D–F) appeared as coils with two laminar surfaces.     81  Figure 3.11: Transmission electron micrographs of tapetum ultrastructure in wild-type and abcg26-1 anthers. Anther development in wild-type (A, C, E, G) and abcg26-1 (B, D, F, H) plants between stages 5 and 8 is shown. (A, B) Microspore mother cells in stage 5. (C, D) Tetrads in stage 7. (E, F) Early unicellular microspores. (G, H) Middle unicellular microspores. M, microspore; MMC, microspore mother cell; T, tapetum; Td, tetrad. (A-H) While pollen wall development was severely compromised in abcg26-1 mutant anthers, no differences in abcg26-1 tapetal cell ultrastructure relative to wild-type cells were observed. Bars = 500 nm. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (    82  Figure 3.12: Transmission electron micrographs of locular inclusions in abcg26-1 mutant anthers at the uninucleate stage of development. A and D, Chemically fixed abcg26-1 mutant buds. B, C, E, and F, High-pressure frozen fixed abcg26-1 stamens. All samples were embedded in Spurr’s resin. Lo, Locule; M, microspore; T, tapetum. Arrows in A to C indicate locule inclusions. Bars = 500 nm (A–C) and 100 nm (D–F). Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (  3.2.5  YFP:ABCG26 is localized at the plasma membrane   To test the subcellular localization of ABCG26, we generated a yellow fluorescent protein (YFP):ABCG26.1 N-terminal fusion under the control of a double 35S promoter and used an Arabidopsis protoplast transient expression system to monitor YFP localization. Such N-terminal fusions to ABCG transporters have been shown to retain activity (Pighin et al., 2004; Bird et al., 2007). Observation of multiple protoplasts expressing the construct showed differing levels of transgene expression, as monitored by YFP fluorescence (Figure 3.13). In protoplasts expressing YFP at low levels, fluorescence was localized to the periphery of protoplasts (Figure 3.13B), where it colocalized with FM4-64, a styrl dye known to insert in the plasma membrane (Bolte et al., 2004). In transformed protoplasts that expressed the YFP at higher levels,   83 fluorescence was also observed at the nuclear envelope and in trans-vacuolar strands, in a pattern consistent with localization of some of the protein in the endoplasmic reticulum (Figure 3.13, E and F). When combined with the in situ hybridization data, the predominantly plasma membrane localization suggests that ABCG26 functions at the plasma membrane of tapetal cells.    Figure 3.13: Subcellular localization of an EYFP:ABCG26 fusion protein in Arabidopsis mesophyll protoplasts. A to D, Transmitted light (A) and confocal microscopy of protoplast transiently expressing EYFP-ABCG26 (B), stained with FM4-64 (C), and an overlay of image B and C (D). E and F, Transmitted light (E) and confocal microscopy of a protoplast transiently expressing EYFP-ABCG26 at high levels (F). Bars = 5 μm. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (     84 3.3  Discussion   In this study, we identified a gene encoding an ABC transporter, ABCG26, that is required for normal male fertility and pollen development in Arabidopsis. Recently, Xu et al. (2010) reported that the ABORTED MICROSPORES (AMS) basic helix-loop-helix transcription factor is required for microspore and tapetum development and used microarray analysis of an ams mutant to identify ABCG26 (formerly WBC27) as an AMS downstream target. In agreement with our findings, Xu et al. (2010) reported that the ABCG26 mutant (abcg26-1 allele) exhibits altered male fertility. Here, we explored the underlying mechanism of decreased male fertility in abcg26 loss-of-function mutants by following pollen development from the callose-encased tetrad stage to the onset of dramatic defective exine formation following microspore release. In concert with the absence of an exine on the surface of abcg26-1 free microspores, mature anthers were nearly devoid of mature pollen grains, and the reduced male fertility phenotype was confirmed quantitatively. The transient tapetum-localized expression of ABCG26 during postmeiotic stages of pollen development, together with the tight coexpression of ABCG26 with genes required for sporopollenin biosynthesis, demonstrate that ABCG26 is required for exine formation on the microspore surface and suggest that ABCG26 could be involved in sporopollenin precursor export from tapetal cells into anther locules. This work further highlights the critical role played by the exine in microspore and pollen function, since the profound exine defects observed in the abcg26-1 mutant correlate with severely compromised pollen grain development and male fertility.   A number of Arabidopsis mutants that exhibit varying degrees of fertility and impaired sporopollenin biosynthesis and defective exine formation have been described (Aarts et al., 1997; Ariizumi et al., 2003; Morant et al., 2007; Souza et al., 2009; Dobritsa et al., 2009b; Tang et al., 2009). ABCG26 is unique as the only transporter described thus far and differs from previously characterized defective exine mutants that affect sporopollenin biosynthetic enzymes or transcription factors required to coordinate tapetum developmental and biosynthetic pathways. Thus, our analysis of ABCG26 function provides new insights into the mechanisms underlying pollen wall deposition.     85 3.3.1 The abcg26-1 mutant has partial Arabidopsis male fertility    In the abcg26-1 mutant, fertility was strongly reduced: most anthers failed to produce mature pollen and appeared brown and dry (Figure 3.1, C and E) and most siliques failed to produce seeds (Figure 3.4A). However, partial fertility occasionally arose late in development, visible as seed-containing siliques (Figure 3.4A). Occasional abcg26-1 anthers were observed that did not produce visible pollen but exhibited bulbous surface protrusions, suggesting the production of pollen grains unable to disperse from the anther. When viewed by SEM, pollen-like structures with smooth surfaces were observed in some manually dissected abcg26-1 anthers, consistent with the hypothesis that some pollen grains with thin defective exine survive to maturity in some mature abcg26-1 anthers (Figure 3.9). It is possible that sufficient defective exine is present on the surface of such developing microspores to permit pollen grain maturation.   Similar low levels of fertility are reported for the ms2 and cyp703a2 mutants that exhibit defective exine and sporopollenin deposition (Aarts et al., 1997; Morant et al., 2007), while acos5 and drl1 mutants are completely male sterile (Souza et al., 2009; Tang et al., 2009). The reasons underlying this variation in fertility among mutants with apparently similar exine defects remain unclear, but it could result from the chemically distinct composition of the defective exine.   3.3.2 Function of ABCG26 in the tapetum    As microspores progress through defined stages of development, the formation of the pollen wall requires contributions from the gametophytic and sporophytic anther tissues (Blackmore et al., 2007). In situ hybridization revealed ABCG26 expression in the tapetum and microspore at stages of microspore development associated with the synthesis and deposition of the pollen exine, specifically at the tetrad and early uninucleate stages. Figure 3.11 illustrates this pattern of expression relative to other events in microspore and pollen development, which closely resembles the tapetum-specific expression of a number of genes encoding enzymes and proteins required for sporopollenin biosynthesis (Aarts et al., 1997; Morant et al., 2007; Souza et al., 2009; Dobritsa et al., 2009b). This pattern is consistent with a role for ABCG26 in the export   86 of sporopollenin precursors or other components required for exine formation, rather than of other pollen wall components, from the tapetum. The detection of ABCG26 transcript in tetrad and early uninucleate microspores indicates that an additional role for ABCG26 in the export of primexine components from developing microspores is possible. However, the lack of a detectable primexine phenotype in abcg26-1 mutants suggests that the primary function of AGBC26 is not in the gametophyte.   The dual expression of ABCG26 primarily in the tapetum at the time of sporopollenin biosynthesis as well as expression in the developing microspores at the time of primexine formation led us to examine the function of ABCG26 in these stages of development by thorough phenotypic analysis of the abcg26-1 mutant. Using light and electron microscopy, signs of microspore degeneration and reduced pollen wall staining were first visible in abcg26-1 free microspores (stage 8), in contrast to wild-type microspores at the equivalent stage. At later stages of anther development, more pronounced defects in microspore development in the mutant were obvious, and by stage 12, pollen grains were absent in the abcg26-1 mutant. These data suggest that microspore development failed in the abcg26-1 mutant as a result of defective pollen wall formation.   Previous studies have demonstrated the improved ultrastructural preservation and elimination of chemical fixation artifacts in tissues preserved by HPF followed by freeze substitution (Kiss and Staehelin, 1995; Paxson-Sowders et al., 2001). We used this technique to examine the ultrastructure of abcg26-1 pollen at multiple stages of pollen development in anthers so that the precise appearance of pollen wall defects could be determined. When HPF-fixed anthers were examined using TEM, it became clear that the abcg26-1 loss-of-function mutant had normal primexine but displayed defects in exine formation that were first apparent in early uninucleate microspores (Figure 3.10). The absence of defective primexine on the abcg26-1 mutant tetrads suggests that the function, if any, of ABCG26 in the gametophytic generation is not in primexine formation. The first pollen wall defects observed were in exine wall formation on early uninucleate microspores, suggesting that microspore development failed due to the lack of sporophytic contributions from the tapetum (Figure 3.10, F and H). The similarity of the abcg26-1 mutant phenotype to sporopollenin biosynthetic mutants, such as ms2, cyp703a2, acos5, and drl1 (Aarts et al., 1997; Morant et al., 2007; Souza et al., 2009; Tang et al., 2009), is   87 consistent with a function for ABCG26 in the export of sporopollenin or other exine components from the tapetum.   The preferential localization of YFP:ABCG26 to the plasma membrane (Figure 3.13) further supports a role for ABCG26 in the export of sporopollenin or other components required for exine formation from tapetal cells into the locule. Endoplasmic reticulum localization observed for YFP:ABCG26 in some protoplasts could be due to high levels of transgene expression, resulting in an overabundance of ABCG26 and leading to impaired trafficking to the plasma membrane. Alternatively, ABCG26 trafficking to the plasma membrane could require a partner ABCG, as demonstrated for the human ABCG5/ABCG8 heterodimer sterol transporter, where one half-transporter partner is retained in the endoplasmic reticulum when its partner is mutated (Graf, 2003).  3.3.3 Locular inclusions in abcg26-1   Since ABCG26 transporter activity appears to be required for the export of tapetum-derived sporopollenin or other components required for exine formation, we examined the tapetum of the abcg26-1 mutant for signs of accumulation due to such defective export. No differences in the structure of the tapetum in abcg26-1 anthers were observed when compared with wild-type tapetal cells at the equivalent developmental stages in chemically fixed and HPF-fixed specimens (Figure 3.7). This suggests that, if the absence of ABCG26 transport results in the accumulation of exine components in the tapetum, such components are either not visible by electron microscopy or were extracted in the fixation process. It is also possible that such components do not accumulate due to rapid catabolic breakdown in tapetal cells. Unexpectedly, however, abcg26-1 locules after the tetrad stage contained coiled inclusions observed in both chemically fixed (Figure 3.12, A and D) and HPF-fixed (Figure 3.12, B, C, E, and F) tissue.   Sporopollenin-like accumulations have been previously described in locules of Arabidopsis pollen development mutants, including dex1, nef1, tde1, and cals5 (Paxson-Sowders et al., 2001; Ariizumi et al., 2004; 2008; Dong et al., 2005; Nishikawa et al., 2005). The accumulations in all of these mutants except abcg26-1 appear as homogenous, electron-dense globular aggregations, interpreted as sporopollenin-like debris. The unique coiled structure of the   88 abcg26-1 locule accumulations suggests that they differ in composition from previously described material. One possibility is that abcg26-1 locule accumulations represent a component of the pollen exine that normally copolymerizes with sporopollenin constituents that may be exported by ABCG26 but fails to do so in the absence of ABCG26-exported constituents in abcg26-1 mutant locules. Although the composition of the coiled locular inclusions observed in abcg26-1 is unknown, they resemble the trilamellar lipidic inclusions observed in the cytoplasm of mutant stem epidermal cells of two previously characterized ABC transporter mutants, abcg11-3 and abcg12-1, defective in cuticular wax export (Pighin et al., 2004; Bird et al., 2007).   3.3.4 Potential ABCG26 substrates   Defects in abcg26-1 microspores are in the structural component of the exine, primarily composed of sporopollenin. Based on a number of chemical analyses, sporopollenin is thought to be a copolymer of simple phenolic compounds and polyhydroxylated unbranched aliphatics, coupled by ester and ether linkages, which provide this biopolymer with its characteristic resistance to chemical degradation (Guilford et al., 1988; Wiermann et al., 2005; Scott et al., 2004). The lack of knowledge regarding the specific constituents of sporopollenin poses challenges in hypothesizing the chemical nature of the cargo that could be transported by ABCG26 from the tapetum into the locule. Recently, genetic analyses have identified a small number of genes encoding enzymes required for the biosynthesis of sporopollenin. Among these, MS2, CYP703A2, DRL1, ACOS5, and CYP704B2 appear to be involved in generating and modifying fatty acid-derived components of sporopollenin and are specifically expressed in the tapetum with similar developmental timing to ABCG26. According to the recent model for sporopollenin biosynthesis proposed by de Azevedo Souza et al. (2009), the ACOS5 enzyme produces a fatty acyl-CoA that is a central intermediate required for the synthesis of sporopollenin constituents that is further modified by other tapetum-expressed enzymes. One attractive hypothesis is that ABCG26 substrate(s) include such ACOS5-derived aliphatic sporopollenin constituents and that after ABCG26-mediated secretion into the locule, they are further transported by an unknown mechanism to the surface of developing microspores for polymerization into the pollen exine. Transport of such molecules is consistent with the function   89 of related Arabidopsis ABCG11 and ABCG12 proteins in the secretion of lipid constituents of cuticular wax (Pighin et al., 2004; Bird et al., 2007) and human ABCG transporters in lipid transport of lipids and steroids (van Meer et al., 2006; Velamakanni et al., 2007). However, alternative hypotheses are possible. For example, ABCG26 could transport a tapetum-generated signaling molecule required for the coordination of exine formation or for transport of an unknown component of the exine required for sporopollenin precursor polymerization and/or exine assembly on the developing microspore wall.   Given the promiscuity in substrates transported by many ABC transporters (Higgins and Linton, 2001; Yazaki, 2006) and the potential for both ABCG26 homodimeric and heterodimeric transporters in the tapetum plasma membrane, ABCG26 could participate in the transport of a suite of related substrates generated by a tapetum-localized sporopollenin precursor biosynthetic pathway. The abcg26-1 mutant provides a potential tool for gaining novel insights into the composition of sporopollenin precursors that are exported to the locule, since it may be possible to use biochemical profiling approaches to identify unpolymerized sporopollenin precursors that accumulate in the anther tapetal cells in the absence of functional ABCG26.   3.4  Materials and methods  3.4.1 Mutant isolation   Arabidopsis (Arabidopsis thaliana) lines with a T-DNA insertion in ABCG26 were identified at The Arabidopsis Information Resource and obtained from the Arabidopsis Biological Resource Center (SALK_062317, SAIL_318_B09, and SAIL_885_F06). Genomic DNA was isolated from the plants, and homozygous individuals were identified using PCR and primers P1 to P3 for abcg26-1, P4 to P6 for abcg26-2, and P5 to P7 for abcg26-3 (Table 3.1). To test ABCG26 expression levels in the mutant (abcg26-1), intron-spanning primer sets upstream (P9 and P10) and downstream (P11 and P12) of the T-DNA insertion site were used to amplify cDNA by PCR (Table 3.1). PCR was performed with Taq polymerase (Finnzymes) in a 20-µL reaction under the following conditions: 95ºC for 2 min, 35 cycles of 94ºC for 30 s, 57ºC for 30 s, and 72ºC for 1 min, and finally 72ºC for 10 min. The amplified genomic DNA fragments from   90 the homozygous lines were gel purified and sequenced to confirm the site of T-DNA insertion. Since the homozygous abcg26-1 mutants retained some partial fertility, it was possible to collect small amounts of homozygous mutant seed.   Table 3.1: Primer sequences used for PCR and RNA probe generation. Reproduced with permission from Quilichini et al., 2010; Copyright American Society of Plant Biologists © 2010 (   3.4.2 Plant growth   Wild-type (Col-0) and abcg26 mutant Arabidopsis seeds were germinated on Murashige and Skoog medium plates at pH 5.7. Seeds were imbibed at 4ºC in the dark, grown at 28ºC under continuous light for 1 week, transplanted to soil (Sunshine Mix 4; Sungrow Horticulture), and raised to maturity at 20ºC in 18 h of light and 6 h of dark. Measurements of plant height were   91 taken at the onset of senescence after height growth had ceased, and variance was expressed as SD of 12 plants measured.   3.4.3 Quantitative real-time PCR analysis    Arabidopsis total RNA was isolated from the specified tissues frozen in liquid nitrogen and ground to a fine powder, and DNase-treated RNA was extracted using the Aurum Total RNA Fatty and Fibrous Tissue Kit (Bio-Rad) following the manufacturer’s instructions. The quality of the RNA samples was assessed by visual inspection of the rRNA bands on a 1% agarose gel. RNA was quantified spectrophotometrically, and 2.0 mg of RNA was used to generate cDNA by Omniscript Reverse Transcriptase (Qiagen) as described by the manufacturer. Standard curves were generated for each primer pair using flower cDNA dilutions to obtain the following primer efficiencies: 101.5% (r2 = 0.944) for primers P12 and P13, producing a 134-bp ABCG26.1 amplicon; 116.2% (r2 = 0.978) for primers P14 and P13, producing a 191-bp ABCG26.2 amplicon; and 91.1% (r2 = 0.979) for primers P15 and P16, producing the ACTIN2 amplicon (Table 3.1). For the quantitative RT-PCR analysis of Arabidopsis gene expression, 10 ng of cDNA was incubated with 10 µL of IQ5 SYBR Green Supermix (Bio-Rad) and 300 nM of each forward and reverse primer (Table 3.1) in a total volume of 20 µL. After an initial denaturation step at 95ºC for 3 min, 40 cycles at 95ºC for 10 s, 53ºC for 10 s, and 72ºC for 20 s were carried out, followed by a melting curve ranging from 94ºC to 53ºC. Quantitative RT-PCR products were run on a gel and sequenced to confirm that single correct amplicons were amplified. Two biological replicates were carried out and gave similar results. Representative data are shown for three technical replicates of one biological replicate in Figure 3.3B, with SD shown for technical replicates. A t-test comparing ABCG26.1 and ABCG26.2 expression levels indicated a significant difference (P < 0.01). Threshold cycles (CTs) were adjusted manually, and CT values were subtracted from those of the flower sample (used as the reference tissue) to generate a ∆CT value. Fold changes were calculated from the ∆CT values and normalized by dividing the corresponding ∆CT value for the ACTIN2 control gene.     92 3.4.4 In situ hybridization   Wild-type Arabidopsis (Col-0) unopened flower buds were fixed in 50% ethanol, 5% acetic acid, and 3.7% formaldehyde under vacuum, dehydrated in an ethanol series, and stained with 0.1% eosin. Samples were then passed through a xylene-ethanol series and embedded in Paraplast (Sigma) with four changes of Paraplast. The blocks were sectioned at 8 mm thickness using a microtome and mounted onto precharged slides (Probe-On; Fisher Scientific). Sections from developing flower buds at multiple stages of development were selected for hybridization to sense and antisense probes.   For sense and antisense ABCG26 probe synthesis, an 822-bp DNA template corresponding to the ABCG26 region spanning exons 3 and 4 was PCR amplified from plasmid DNA containing the cloned full-length cDNA using gene-specific forward and reverse primers (Table 3.1; P17–P20) and Taq polymerase (Finnzymes) at 49ºC annealing and 68ºC extension for 40 cycles. A T7 polymerase-binding site was incorporated into the forward primer for sense probe amplification and in the reverse primer for antisense probe amplification. Digoxigenin (Roche)-labeled probes were transcribed off the template using T7 polymerase (Roche) following the manufacturer’s instructions. Reactions were stopped using EDTA and precipitated in ethanol and LiCl. Probes were shortened to 200-bp fragments by limited carbonate hydrolysis at 60ºC and quantified by dot blot dilutions compared with known samples on Hybond XL membranes (GE Healthcare).   Samples were dehydrated through an ethanol series and subjected to hybridization at 55ºC for 16 h. Both full-length and partial sense and antisense probes were used at 20 ng per slide in 50% formamide, 10% dextran sulfate, 13 Denhardt’s solution, 0.5 mg of tRNA, 10 mM Tris, pH 7.5, 1 mM EDTA, and 300 mM NaCl. The slides were washed in 23 SSC at 55ºC followed by four washes in 0.23 SSC at 55ºC, then 0.23 SSC at 37ºC, 0.23 SSC at room temperature, and finally with phosphate-buffered saline at room temperature. The slides were covered with 1 mL of blocking buffer (1% Roche blocking agent in 100 mM malic acid and 150 mM NaCl, pH 7.5) for 45 min at room temperature, washed in BSA solution (1% bovine serum albumin, 100 mM Tris, pH 7.5, 150 mM NaCl, and 0.3% Triton X-100), and hybridized to anti- digoxigenin antibody (Roche) at 1:1,250 dilution for 90 min. The slides were then washed in   93 BSA solution followed by 100 mM Tris, pH 9.5, 100 mM NaCl, and 50 mM MgCl2. The color reaction was started by the addition of 5-bromo-4-chloro-3-indolyl phosphate/nitroblue tetrazolium chloride color reagent (Roche), and the color was allowed to develop in the dark for 16 h before visualizing on an Olympus Provis AX-70 microscope.   3.4.5  Genetic complementation of the abcg26-1 mutant   The 2,058-bp fragment encoding ABCG26 was amplified from flower cDNA using Platinum High Fidelity Taq Polymerase (Invitrogen) with primers spanning the full-length cDNA and including a stop codon. The amplified band was gel purified and cloned into a Gateway-compatible entry vector (pCR8/GW/TOPO; Invitrogen), and the gene was confirmed by sequencing. Additionally, a second unexpected transcript of 2,215 bp was amplified (approximately 10% of the time) that retained intron 7 and encoded premature stop codons (truncating at 2,182 bp downstream of ATG in the first stop codon). The genes were recombined into a Gateway-compatible destination binary vector (pDG2; modified pSM-3 vector based on pCAMBIA1390 vector) using LR Clonase II enzyme mix (Invitrogen) according to the manufacturer’s instructions. Agrobacterium tumefaciens LBA4404 was transformed with one of the described destination vectors and used to transform homozygous abcg26-1 plants by the floral dip method (Clough and Bent, 1998). T1 seeds were grown on Murashige and Skoog plates with 50 mg L-1 hygromycin, and surviving plants were grown to maturity and allowed to self-pollinate. T2 plant genotypes were confirmed at the ABCG26 locus. PCR confirmed the presence of the transgene.   3.4.6  Scanning electron microscopy    Anthers were removed from fully opened flowers and placed on aluminum SEM stubs (Ted Pella, Inc.). Samples were coated with 7 nm gold using a Cressington 208HR High Resolution Sputter Coater and examined with a Hitachi S-2600N Variable Pressure SEM device.     94 3.4.7  Transmission electron microscopy    For chemical fixation, wild-type (Col-0) and abcg26-1 mutant flower buds were dissected from plants for immediate fixation in 4% formaldehyde and 2.5% glutaraldehyde in 50 mM sodium cacodylate buffer (pH 6.9). Samples were postfixed in 1% osmium tetroxide, dehydrated in an ethanol series, and incubated in acetone followed by 100% propylene oxide for 30 min. Samples were gradually embedded in Spurr’s resin and polymerized at 60ºC for 3 d.   For HPF followed by freeze substitution, unopened flower buds were excised from Col-0 and abcg26-1 plants, stripped of sepals and petals, and submerged in 0.2 M Suc in uncoated copper type B sample holders (Ted Pella, Inc.). Samples were frozen at high pressure with a Leica EM HPM 100 high-pressure freezer followed by freeze substitution and embedding in Spurr’s resin (Spurr, 1969) as described by McFarlane et al. (2008). Freeze substitution medium contained 2% (w/v) osmium tetroxide in acetone with 8% (v/v) dimethoxypropane.   Toluidine blue-stained thick sections (0.3–0.7 µm) were viewed by light microscopy to determine the stage of anther development as described by Sanders et al. (1999). Samples for representative stages of anther development were thin sectioned (less than 70 nm) with a Reichert Ultracut E ultramicrotome and mounted on 100-mesh 0.5% Formvar-coated grids. Sections were stained with 2% uranyl acetate in 70% (v/v) methanol for 15 min followed by Reynold’s lead citrate for 5 to 8 min. A Hitachi H7600 TEM device was used to examine the stained samples, and images were taken with an AMT Advantage (1 megapixel) CCD camera (Advanced Microscope Technologies).   3.4.8 Subcellular localization of ABCG26 in planta   ABCG26 was PCR amplified in a 50 µL reaction from digested TOPO-A vector containing full-length ABCG26, and the resulting PCR product was separated on a 2% ethidium bromide agarose gel and purified using a Qiagen quick gel extraction kit. Primers used for amplification (Table 3.1; P21 and P22) added restriction cut sites for subsequent ligation into the pE3150 vector provided by Dr. S. Gelvin. This produced an N-terminal fusion with enhanced yellow fluorescent protein (EYFP) driven by a double 35S promoter. The vector was   95 maxiprepped using the Qiagen Endo-free Maxi kit, and 10 µg was used per protoplast transformation. Arabidopsis leaf protoplasts were isolated and transformed as described by Tiwari et al. (2006) with a 2 to 3 h leaf digestion. Protoplasts were imaged with a Perkin-Elmer Spinning Disc confocal microscope. YFP was detected with a 491-nm laser and a 528/38 nm emission filter. FM4-64 (10 mM)-stained protoplasts were imaged using a 561 nm laser with a 593/40 nm emission filter within 15 min of staining. Images were processed with Volocity 4.3.2 (Improvision).   96 Chapter 4: ABCG26-mediated polyketide trafficking and hydroxycinnamoyl spermidines contribute to pollen wall exine formation prior to tapetum programmed cell death in Arabidopsis thaliana  4.1 Introduction   Mature pollen grains are surrounded by a specialized and complex cell wall that serves critical functions in grain survival, stigmatic recognition and hydration. The outer pollen wall (exine) is composed of a sculptured and structurally rigid sporopollenin backbone, which in entomophilous species is additionally covered by a lipid-based pollen coat or tryphine (Piffanelli et al., 1998; Hsieh and Huang, 2007). The components of the outer pollen wall are contributed by surrounding secretory tapetal cells, which form the innermost cell layer of the sporophytic anther wall. The sporopollenin component of the exine assembles over a short window of development. It begins following microspore meiosis and the release of free microspores from callose-bound tetrads, nears completion by the first pollen mitotic division, and concludes prior to tapetum programmed cell death and the second mitotic division (Piffanelli et al., 1998). As sporopollenin deposition progresses, tapetal cells accumulate an array of lipids, proteins, flavonoids, and phenolic spermidine conjugates, primarily in storage organelles called tapetosomes and elaioplasts, destined for the pollen coat in Arabidopsis (Piffanelli et al., 1998; Grienenberger et al., 2009; Hsieh and Huang, 2007). After sporopollenin deposition into the backbone of the exine is complete, tapetum programmed cell death releases lipid-rich contents into the locule, filling the exine crevices to form the pollen coat.   Elucidating the chemical composition of sporopollenin has posed great challenges due to its extreme recalcitrance to degradation. Based on biochemical analyses, sporopollenin is thought to contain a mixture of phenolics and aliphatic derivatives (Guilford et al., 1988; Ahlers et al., 1999; Hemsley et al., 1992; Wehling et al., 1989). Genetic approaches, primarily in Arabidopsis and rice, have revealed a number of genes encoding enzymes and proteins required for sporopollenin biosynthesis and deposition, providing clues regarding the composition and mechanism of sporopollenin biosynthesis, and substantiating the key role of the tapetum as the   97 source of sporopollenin precursors (reviewed by Ariizumi and Toriyama, 2011). Among the Arabidopsis genes required for sporopollenin formation, a number encode enzymes with characterized biochemical activities, including MALE STERILITY 2 (MS2) (Aarts et al., 1997; Chen et al., 2011), ACYL-COA SYNTHETASE5 (ACOS5) (Souza et al., 2009), two POLYKETIDE SYNTHASES  PKSA/LAP6 and PKSB/LAP5 (Kim et al., 2010; Dobritsa et al., 2010), TETRAKETIDE α-PYRONE REDUCTASE1 (TKPR1/DRL1) and TKPR2/CCRL6 (Grienenberger et al., 2010; Tang et al., 2009), and two CYTOCHROME P450s (CYP703A2/DEX2 and CYP704B1 (Morant et al., 2007; Dobritsa et al., 2009b). Based on genetic and in vitro biochemical analyses, ACOS5, PKSA, PKSB, and TKPR1 are proposed to function together in the synthesis of hydroxylated tetraketide α-pyrones; polyketides which may form the major constituent of sporopollenin (Grienenberger et al., 2010). In support of this model, these enzymes are localized at the tapetum ER, interact in vivo and have been termed the ‘sporopollenin metabolon’ (Lallemand et al., 2013). CYP703A2 and CYP704B1, which hydroxylate medium- and long-chain fatty acids, may modify the fatty acid precursors of the polyketide produced by the sporopollenin metabolon for the synthesis of hydroxylated tetraketide α-pyrone(s) (Morant et al., 2007; Dobritsa et al., 2009b). In addition to the polyketide pathway, fatty alcohols produced by the fatty acyl-carrier protein reductase MS2 are required for normal sporopollenin formation (Chen et al., 2011).   The presence of phenolic constituents in sporopollenin is well documented (Ahlers et al., 1999; Wehling et al., 1989; Gubatz et al., 1993; Weng et al., 2010), but there is little information on the exact enzymes required for the biosynthesis of these constituents, or the specific nature of these compounds. Manipulation of the Arabidopsis monolignol biosynthetic pathway, resulting in enhanced levels of 5-hydroxyconiferyl alcohol subunits in lignin leads to abnormalities in pollen wall formation, revealing the joint contribution of certain phenylpropanoids to both lignin and sporopollenin formation (Weng et al., 2010). Mutations in the genes encoding TRANSPARENT TESTA4 (TT4) and 4-COUMARATE:COA LIGASE3 (4CL3), enzymes thought to be involved in flavonoid biosynthesis, and SPERMIDINE HYDROXYCINNAMOYL TRANSFERASE (SHT), an acyltransferase involved in conjugating spermidine to hydroxycinnamic acids, affect the composition of the Arabidopsis pollen wall (Dobritsa et al., 2011). However, as phenolic compounds are known constituents of both the sporopollenin   98 biopolymer and the pollen coat, it is unclear which layer of the mature pollen wall contributes the autofluorescence of these metabolites.   While our understanding of sporopollenin biosynthesis and composition has advanced rapidly over the last decade, mechanisms for sporopollenin trafficking from the tapetum, and exine assembly into the highly patterned pollen wall remain poorly understood (Ariizumi and Toriyama, 2011; Bedinger, 1992). The spatial distinction of tapetal cells from pollen grains in taxa with secretory tapeta (including Arabidopsis and rice) suggests a critical role for the export of sporopollenin during exine formation. ABCG26, an Arabidopsis ATP-binding cassette transport protein, is required for sporopollenin accumulation and has been proposed to traffic sporopollenin components out of tapetal cells following tetrad release (Quilichini et al., 2010; Kuromori et al., 2011a; Dou et al., 2011; Choi et al., 2011; Xu et al., 2010). In support of a role for ABCG26 in sporopollenin precursor export, ABCG26 function is required for exine formation and male fertility, and its expression in tapetal cells is maximal during tetrad formation and early free microspore stages when sporopollenin synthesis and deposition peak (Chapter 3; Quilichini et al., 2010). The apparent ortholog of ABCG26 in rice, ABCG15, is also required for exine formation, suggesting a conserved function for these transporters in sporopollenin export (Qin et al., 2013; Niu et al., 2013). Interestingly, additional defects in anther cuticle formation are observed in rice abcg15 mutants, despite the ABCG15 tapetum-specific expression, suggesting that ABCG15 may play additional roles in the trafficking of rice cuticular metabolites from tapetal cells (Qin et al., 2013). However, the substrates of these ABC transport proteins are unknown, and hypothetical links to sporopollenin traffic remain untested experimentally.   In addition to hypothesized sporopollenin transport proteins, orbicules, also called Ubisch bodies, are small granules formed along the locule side of tapetal cells that have been proposed to mediate sporopollenin traffic between the tapetum and developing microspores in some species (Wang et al., 2003). Orbicules are present almost exclusively in species with secretory tapeta, but are curiously absent in Arabidopsis (Huysmans et al., 1998). While the absence of orbicules in some taxa with secretory tapeta and the persistence of orbicules after pollen wall formation make their proposed function in sporopollenin transport subject to debate, the conspicuous presence of orbicules outside tapetal cells of some species suggests that additional   99 mechanisms for export of pollen wall constituents from the tapetum may be in place.  In this study, the biosynthesis and export of exine components was investigated using a novel approach to visualize living tapetal cells in intact anthers of Arabidopsis.  Using a variety of mutants with genes encoding the ABCG26 transporter and putative sporopollenin polyketide biosynthetic enzymes, we identified a genetic link between the polyketide sporopollenin precursor pathway and ABCG26, suggesting that the substrate of ABCG26 may be a product of the polyketide synthesis metabolon. Additionally, we identified novel hydroxycinnamoyl spermidine-containing orbicule-like bodies, which accumulate only in the absence of the polyketide pathway, and prior to tapetum programmed cell death. The accumulation of these orbicule-like bodies indicates that additional mechanisms of export for components of the exine exist, that co-trafficking of polyketides and hydroxycinnamoyl spermidines from tapetal cells may occur, and that hydroxycinnamoyl spermidines, previously known only as components of the pollen coat deposited after tapetum programmed cell death, are deposited simultaneously with sporopollenin framework from metabolically active tapetal cells.   4.2 Results  4.2.1 Tapetal cells of abcg26 anthers accumulate intrinsically fluorescent compounds in vacuole-like bodies   To test the hypothesis that the ABCG26 transporter exports sporopollenin precursors or other components required for exine formation from anther tapetal cells, abcg26 mutants were examined for the accumulation of exine components in the tapetum. Methods used to examine tapetal cells by transmission electron microscopy (TEM) have typically required tissue fixation, followed by sectioning and precise anther developmental staging. Due to the possibility of metabolite extraction from biological samples during fixation (Palade, 1952; Morgan and Huber, 1967; Bullock, 2011) and the challenges associated with anther sample preparation (including staging and plane of section for TEM), contrasting results for tapetum ultrastructure in abcg26 mutants have been reported (Quilichini et al., 2010; Choi et al., 2011; Dou et al., 2011; Kuromori et al., 2011a). While previous reports did not identify accumulations inside abcg26 tapetal cells,   100 some found accumulations behind tapetal cells or reported enlarged tapetal cells (Choi et al., 2011; Dou et al., 2011). To circumvent these challenges and the limitations of two-dimensional imaging, we devised a method for examining tapetal cells in intact anthers without fixation using multi-photon laser scanning microscopy.  The use of multi-photon microscopy enables deep imaging in living specimens through localized excitation (Zipfel et al., 2003) and it allowed for the visualization of developing microspores and tapetal cells throughout the three dimensional structure of the anther.    Based on current knowledge of sporopollenin composition, we predicted that sporopollenin precursors transported by ABCG26 from the tapetum are either lipidic or phenolic in nature, and would accumulate in tapetal cells as a result of defective transport in the abcg26 mutant. Given the strong intrinsic fluorescence of pollen exine and the numerous reports of phenolic compounds in the sporopollenin biopolymer (Wehling et al., 1989; Ahlers et al., 1999; Prahl et al., 1986; Rozema et al., 2001), an attractive hypothesis is that ABCG26 transports a phenolic compound, such as a hydroxycinnamic acid, coumarin or flavonol (Hutzler et al., 1998; Lichtenthaler et al., 1992; Buschmann et al., 2000), or another intrinsically fluorescent constituent of the sporopollenin biopolymer. We used two-photon (2-P) microscopy, a variation of multi-photon microscopy in which simultaneous excitation by two photons of red shifted light (700 – 1000 nm) is used to excite fluorophores in the ultraviolet (UV) range.  This enabled us to visualize the distribution of intrinsically fluorescent emissions in intact anthers.  Using this approach, all anther cell types from wild-type and mutant plants exhibited intrinsic fluorescence in both channels, which were pseudo-colored teal (420-460 nm) and red (495-540 nm) (Figure 4.1A-H). These wavelengths were selected for their ability to capture emission primarily from low wavelength-emitting phenolic compounds such as hydroxycinnamic acids and coumarins, and from higher wavelength-emitting compounds such as flavonols and chlorophyll, respectively. Emission differences were observed between wild-type (Col-0) and abcg26 (abcg26-1) developing microspore walls (Figure 4.1A-D) and tapetal cells (Figure 4.1E-H), and were first visible at the free uninucleate microspore stage of anther development (anther stage 8), following tetrad release. Wild-type microspores exhibited strong pollen wall fluorescence in both channels (Figure 4.1A, enlarged in B), while pollen wall fluorescence was absent from most abcg26 microspores (Figure 4.1C, enlarged in D). However, variation in the   101 abcg26 phenotype was common and some locules contained microspores with a thin layer of pollen wall fluorescence, which lacked reticulate patterning (Figure 4.2A,C).  Thus, this method was able to effectively capture, in live cells, the defects in abcg26 microspore walls that have been previously reported in fixed, sectioned material (Quilichini et al., 2010).   Figure 4.1: Vacuolar inclusions are present in abcg26 mutant tapetal cells.  In wild-type anthers, bright intrinsic fluorescence was emitted by microspore walls (A, enlarged in B) and diffusely by tapetal cells (E, enlarged in F) when imaged by two-photon microscopy. Emission was excluded from wild-type tapetum vacuoles, which appeared dark (arrows in B and F), however, the bright autofluorescence from underlying microspore walls could not be excluded (asterisk in F). In abcg26 anthers, locules were devoid of microspore wall emission (C) but showed bright autofluorescence at the locule periphery in tapetal cells (arrows in C, and enlarged in D). abcg26 tapetal cells contained autofluorescent vacuoles with brightly emitting puncta (G, enlarged in H) not seen in wild-type. When examined by TEM, wild-type tapetum vacuoles contained little to no debris (I, enlarged in J), while abcg26 vacuoles contained electron-dense circular core inclusions (arrows in L) within a debris-filled vacuole (K, enlarged in L). No consistent differences in emission from cell layers outside of the tapetum between wild-type and abcg26 were observed. Lo, locule, Msp, microspore, Tp, tapetum. Bars = 20 μm (A,C,E,G) and 5 μm (I, K).   102     103 Figure 4.2: Variation in the severity of the abcg26 anther phenotype. Varying levels of autofluorescence from abcg26 microspore walls and tapetum vacuoles were observed using two-photon microscopy. Correlated with increased levels of emission from microspores in some abcg26 locules, the emission from tapetal cell vacuoles appeared diminished (A-D). In some anthers, numerous autofluorescent structures were observed in each tapetal cell (E,F). Similarly, multiple cores in tapetum vacuoles could be detected by TEM (G,H). Autofluorescent inclusions were always observed in abcg26 tapetum vacuoles after tetrad release, however, the intensity of emission from the surrounding vacuole varied. Emission along the outer periclinal side of tapetal cells was additionally observed in many samples (A-F), which could not be precisely localized by two-photon imaging.  Using TEM, accumulations (indicated by arrows) were observed in abcg26 middle layer cells (I,J), which were not identified in WT. En, endothecium, ML, middle layer, Ta, tapetum, Vac, vacuole. Bars = 20 μm (A-F) and 1 μm (G-J).   In the tapetum, striking differences in intrinsic autofluorescence were observed as wild-type tapetal cells emitted diffuse fluorescence, with small non-fluorescent vacuoles (Figure 4.1A and E, arrows in B and F), while tapetal cells of abcg26 contained large spherical fluorescent vacuoles (Figure 4.1C and G, arrows in D and H) with one or more fluorescent ‘cores’. Fluorescence emission of the tapetum vacuole cores was strong in the lower wavelength range, and similar structures were not observed in any wild-type tapetum vacuoles. Some variation in the tapetum fluorescent inclusions was observed among anthers from the same abcg26 plant, for anthers at comparable stages of development (Figure 4.2). While some anthers exhibited a single, large fluorescent vacuole in each abcg26 tapetal cell, other anthers contained numerous fluorescent vacuoles per tapetal cell. Consistent with these observations, anther specimens with high levels of fluorescence in the tapetum vacuole lacked or had dim fluorescence around microspores, while specimens with low levels of tapetum vacuole fluorescence had brighter fluorescent emission from microspore walls (Figure 4.2B, D). In addition to tapetum vacuolar inclusions, bright fluorescent accumulations were often observed on the outer periclinal face of tapetal cells adjacent to the middle layer. Analysis of abcg26 developing anthers at a similar stage by TEM showed that these accumulations were inside cells of the middle layer, exhibited the same morphological and staining characteristics as the inclusions in tapetal cells (Figure 4.2I, J), and were never observed in wild-type middle layer cells (Figure 4.1I).  Previous TEM studies failed to identify enlarged vacuoles with associated inclusions in abcg26 tapetal cells. With the insight provided by live cell imaging data, careful examination of abcg26 tapetal cells by TEM revealed the presence of enlarged vacuoles in fixed abcg26   104 samples, aided by the knowledge that these vacuoles contain one or more cores. The abcg26 tapetum vacuole cores observed by TEM were visible as containing one or more circular electron-dense core structures (Figure 4.1K,L, 2G,H). Similar core structures were never observed in wild-type tapetal cell vacuoles (Figure 4.1I,J).  To test if the distribution of lipidic constituents might differ in abcg26 anthers relative to wild-type, anthers were stained with Nile Red, a dye that does not fluoresce in water but becomes fluorescent in hydrophobic environments and organic solvents (Greenspan et al., 1985). Penetration of the Nile Red dye into anther locules to levels that could be visualized required 20-30 minutes, and all images shown were captured within this time interval (Figure 4.3E-L). Unstained anthers emitted little to no fluorescence in the channel used to capture Nile Red fluorescence (Figure 4.3A-D).  In Nile Red-stained anthers, no differences in epidermis, endothecium and middle layer cell lipids were observed between wild-type and abcg26 samples (Figure 4.4). Wild-type microspores exhibited staining in the wall, but did not exhibit internal Nile Red fluorescence unless staining exceeded 30 minutes (Figure 4.4A, enlarged in C). In contrast, abcg26 microspores, when present, typically exhibited internal staining within 20 minutes, but lacked significant Nile Red fluorescence from microspore walls (Figure 4.4E, enlarged in G). These results are consistent with previous reports that a hydrophobic sporopollenin wall containing lipidic components is lacking in abcg26 mutants. Wild-type and abcg26 tapetal cells stained with Nile Red revealed abundant lipids in puncta, presumably elaioplasts and tapetosomes, throughout the cells (Figure 4.4B,D,F,H). Although differences in tapetum lipids were not observed between mutants and wild-type, the cores within abcg26 tapetum vacuoles stained with the hydrophobic dye (Figure 4.4F, enlarged in H). Wild-type tapetum vacuoles lacked Nile Red emission, were smaller than abcg26 vacuoles and lacked the associated stained inclusions (Figure 4.4B, enlarged in D).    105  Figure 4.3: Controls for the timed Nile Red staining of anthers.  Wild-type stamens immersed in 25% glycerol (A-D) or Nile Red staining solution (E-L) were imaged with a two-photon microscope within 20-30 minutes of submersion. The same anther stained with Nile Red and optically sectioned through the locule (E-H) and the tapetum (I-L) is shown. Nile Red fluorescence emission, false-coloured green, was captured from 575-630 nm (A,E,I).  Broader emission primarily from intrinsic fluorescence, falsely coloured red, was captured from 380-560 nm (B,F,J). Overlays for emission captured from both channels are shown (C,G,K) with magnified views in the far right panel (D,H,L). Bars = 20 μm.     106  Figure 4.4: abcg26 tapetum vacuole cores stain with Nile Red. Using two-photon microscopy, the lipid content of wild-type and abcg26 anthers was assayed by Nile Red staining. In wild-type (A,B, enlarged in C,D, respectively) and abcg26 (E,F, enlarged in G,H respectively), differences between anther lipids in cells external to the tapetum were not observed. As compared to the brightly staining wild-type microspore walls (C), abcg26 microspore walls (when present) did not stain or stained minimally (G). Microspore cytoplasm stained more readily in abcg26 than wild-type microspores stained for equivalent lengths of time. In tapetal cells, abcg26 enlarged vacuoles lacked Nile Red stain, except for the vacuole cores (F, arrows in H), which were not observed in wild-type tapetum vacuoles (B, arrows in D) . Msps, microspores; Tp, tapetum. Bars = 20 μm.     107  The development of these 2-P imaging methods for live anthers revealed that abcg26 mutants accumulate enlarged autofluorescent vacuoles with dense core structures. As sporopollenin is lacking on abcg26 microspores, these accumulations appear to be comprised of sporopollenin components in a pre-trafficked state prior to polymerization on the microspore surface.   4.2.2 The polyketide synthesis pathway is required for the accumulation of vacuolar inclusions in the tapetal cells of abcg26 mutants   The nature of the vacuolar material in the tapetum of abcg26 was investigated by examining double mutants affecting both the transporter and key enzymes required for sporopollenin biosynthesis. In a model proposed by Grienenberger et al. (2010), ACOS5, CYP703A2, CYP704B1, PKSA and PKSB, and TKPR1 work sequentially to synthesize a hydroxylated tetraketide α-pyrone(s), suggested to be a major building block of sporopollenin, since mutants in the genes encoding these enzymes are severely compromised in exine deposition. Although the products and intermediates included in this pathway are based on in vitro analyses of recombinant enzyme activities (Morant et al., 2007; Souza et al., 2009; Dobritsa et al., 2009b; Grienenberger et al., 2010; Kim et al., 2010), it represents the most well-characterized biosynthetic pathway of sporopollenin biosynthesis to date.  acos5, pksa pksb, and tkpr1 mutants exhibit highly similar phenotypes to abcg26, including the absence of pollen exine and strong reductions in male fertility. We hypothesized that the product(s) of this pathway, referred to herein as the polyketide pathway, could be exported by ABCG26. A prediction of this hypothesis is that the fluorescent putative ABCG26-trafficked wall metabolites that accumulate in abcg26 tapetal cell vacuoles are products of the polyketide pathway. To test this prediction, we examined anthers from double or triple mutants of abcg26 and acos5, pksa pksb, and tkpr1 using 2-P microscopy, and compared these images to the abcg26 single mutant. The fluorescent tapetum vacuoles observed in the abcg26 single mutant (Figure 4.1) were absent in the double or triple mutants abcg26 acos5 (Figure 4.5C,D), abcg26 pksa pksb (Figure 4.5E,F) and abcg26 tkpr1 (Figure 4.5G,H). These data indicate that fluorescent vacuolar material that accumulates in abcg26 requires the tapetum-localized polyketide pathway, supporting the hypothesis that the   108 proposed aliphatic polyketide(s) synthesized by the sequential action of ACOS5, PKSA PKSB, and TKPR1 is transported from tapetal cells by ABCG26. The tapetum vacuolar inclusions in abcg26 mutants persisted in double mutant combinations with other pollen wall mutants, including sht (see below, Figure 4.16), tt4, and 4cl3 (Figure 4.6). These data indicated that abcg26 vacuolar inclusions were lost only and specifically in polyketide pathway mutants, supporting the specific requirement for the polyketide pathway enzymes in the synthesis of these accumulating metabolites.     109    110 Figure 4.5: The enzymes ACOS5, PKSA PKSB, and TKPR1 of the polyketide synthesis pathway are required for formation of tapetum vacuolar inclusions in the abcg26 mutant. Two-photon microscopy of wild-type anther autofluorescence identified bright emission from developing microspore walls (A) and dim emission from tapetal cells (B). This technique revealed the absence of tapetum vacuolar inclusions (arrows), resembling those seen in abcg26, in the abcg26 acos5 double mutant (C,D), abcg26 pksa pksb triple mutant (E,F), and abcg26 tkpr1 double mutant (G,H). These double and triple mutants contained additional punctate fluorescence surrounding tapetal cells and some microspores (C-H) that was not observed in wild-type anthers. Bars = 20 μm.     Figure 4.6: Vacuolar inclusions are present in abcg26 4cl3 and abcg26 tt4 mutant tapetal cells. Using two-photon microscopy, tapetum vacuolar inclusions (arrows), resembling those seen in abcg26, persisted in abcg26 4cl3 (A,B) and abcg26 tt4 (C,D) double mutants. Varying levels of autofluorescence from abcg26 4cl3 and abcg26 tt4 microspore walls and tapetum vacuoles were observed, as previously shown for abcg26 single mutants. Bars = 20 μm.  4.2.3 Discrete, extracellular orbicule-like bodies surrounding tapetal cells and microspores accumulate in key sporopollenin biosynthetic mutants   When examined by 2-P microscopy under UV excitation, the double and triple mutants formed by abcg26 in combination with acos5, pksa pksb, and tkpr1 mutants exhibited unexpected fluorescent punctate surrounding tapetal cells (Figure 4.5D,F,H) and some   111 microspores (Figure 4.5C,E,G) that were not observed in wild-type (Figure 4.5A,B) or abcg26 single mutants (Figure 4.1C,D). This observation led us to examine the acos5 single mutant anthers by 2-P microscopy under UV excitation, to determine if these fluorescent bodies were similarly present in mutants of the polyketide pathway. As in the acos5 abcg26 double mutant, analysis of the acos5 single mutant revealed equivalent punctate fluorescent bodies on all sides of the tapetum and surrounding some microspores that were not observed in wild-type anthers (Figure 4.7A-F). These bodies appeared in acos5 anthers at the uninucleate microspore stage (known as anther stage 8) and became more abundant around tapeta over the course of microspore mitosis (anther stages 9-11). These orbicule-like bodies (ORBs) resembled the orbicules that surround tapetal cells in many other flowering plant species including rice, but are absent in Arabidopsis (Wang et al., 2003).  Autofluorescent ORBs were also observed in anthers of the other polyketide pathway mutants, cyp703a2, pksa pksb, and tkpr1 mutants (Figure 4.8C-H), but not in wild-type (Figure 4.8A,B), and have not been previously reported in descriptions of these mutants.     112  Figure 4.7: acos5 mutant exhibits tapetum-associated autofluorescent bodies, not observed in wild-type. Using two-photon microscopy, wild-type microspore walls emitted bright autofluorescence (A), while tapetal cells emitted diffusely (B) in developing, intact anthers. The sporopollenin biosynthetic mutant acos5 contained autofluorescent bodies around tapetal cells on all sides and around some microspores (C, enlarged in E). These bodies appeared as circular puncta (arrows in E) and were particularly abundant in sections through the tapetum along the outer periclinal face (D, enlarged in F). Lo, locule, Msp, microspore, Tp, tapetum. Bars = 20 μm.     113  Figure 4.8: cyp703a2, pksa pksb, and tkpr1 mutants exhibit tapetum-associated autofluorescent bodies, not observed in wild-type. Using two-photon microscopy, wild-type microspore walls emitted bright autofluorescence (A) and tapetal cells emitted diffuse autofluorescence (B) in developing, intact anthers. The sporopollenin biosynthetic mutants, cyp703a2 (C,D), pksa pksb (E,F) and tkpr1 (G,H), exhibited overlapping phenotypes, with autofluorescent bodies on all edges of tapetal cells and around some microspores (C,E,G). These orbicule-like bodies were particularly abundant in optical sections through the tapetum along the outer periclinal face (D,F,H). Bars = 20 μm.   114  TEM analysis confirmed the presence of ORBs and their precise locations within anthers of acos5 mutant, after the release of microspores from tetrads. These analyses revealed round, extracellular ORBs in acos5 anther locules (Figure 4.9C-F), which were absent from wild-type samples (Figure 4.9A,B). The ORBs were electron-translucent with a small fibrillar electron-dense core (Figure 4.9E). The ORBs appeared to be partially embedded in the tapetum plasma membrane on the inner periclinal (locule) side (Figure 4.9C,E), and were located between and behind tapetal cells on their outer periclinal faces. The bodies were additionally observed along the surface of some free microspores in place of exine (Figure 4.9D,F). We obtained cryo-SEM images of cryoplaned acos5 anthers (see Materials and Methods), which confirmed the abundance of these bodies, that are located extracellularly to tapetal cells and found in abundance along the outer periclinal side of tapetal cells at the vacuolated free microspore stage (Figure 4.9G). These bodies were also observed around some microspores, while others lacked them (Figure 4.9H). Using TEM, ORBs of similar electron-density and location to those in acos5 mutants were observed in the polyketide pathway mutants pksa pksb and tkpr1 (Figure 4.10) and in abcg26 acos5 double mutants (Figure 4.5I,J). The presence of extracellular ORBs in acos5, pksa pksb, and tkpr1 mutants, and the persistence of ORBs in the acos5 abcg26, pksa pksb abcg26, and tkpr1 abcg26 double and triple mutants suggested that fluorescent compounds in addition to those generated by the polyketide pathway, but of unknown composition, are able to exit tapetal cells in the absence of a functional ABCG26 transporter. The accumulation of these compounds at extracellular surfaces of tapetum and some microspore surfaces was observed only when polyketide product synthesis was directly affected, since the fluorescent bodies were not observed in the abcg26 mutant where transporter loss of function presumably impacts polyketide product transport, but not synthesis (Figure 4.1).      115    116 Figure 4.9: Ultrastructure of extracellular orbicule-like bodies in acos5 and abcg26 acos5 mutant anther locules. In TEM, wild-type tapetal cell edges appeared clear of debris or other bodies, with occasional invaginations (A). In contrast, the locule-facing plasma membrane of acos5 tapetal cells had deep invaginations that closely associated with bodies that appeared extracellular (C, and enlarged in E). The bodies were found on all faces of acos5 tapetal cells and appeared electron-translucent, with some electron-dense debris (E). The bodies identified in acos5 were also found around some microspores (D, enlarged in F), but were absent from wild-type microspore surfaces (B). Using cryo-SEM, the extracellular position of these bodies was confirmed, as their position outside the plasma membrane along the anther wall and between neighbouring tapetal cells could be seen clearly (G).  The accumulation of acos5 extracellular bodies around some microspores, but not others, was clearly visualized by cryo-SEM (H). Equivalent extracellular bodies were observed in acos5 abcg26 double mutant anthers, on all faces of the tapetum (I) and on some microspores (J). En, endothecium; Lo, locule; Msp, microspore; Tp, tapetum. Bars = 2 μm.   Figure 4.10: Extracellular bodies are observed in pksa pksb and tkpr1 anther locules. In TEM, wild-type tapetal cell plasma membranes appeared continuous with occasional invaginations, and were free of associated bodies (A). In pksa pksb double mutants and tkpr1 single mutants, bodies associated with the periphery of tapetal cells were observed on all sides of the plasma membrane, shown here on the inner periclinal face (arrows in B and C). These bodies were associated with deep tapetum plasma membrane invaginations and were often present in multiples (double arrows in B). Electron dense debris could be observed in these bodies (most notably in C). Lo, locule, Msp, microspore, Tp, tapetum. Bars = 2 μm.    No differences in lipid accumulation could be seen in tapetum lipids in acos5 and wild-type anthers stained with Nile Red (Figure 4.11). The extracellular ORBs did not stain with Nile Red (green color in Figure 4.11), but their presence in acos5 locules was confirmed by their autofluorescence (red color in Figure 4.11). As observed in abcg26 anthers, the cytoplasm of   117 developing acos5 microspore stained readily with Nile Red, but the walls, when present, only stained minimally, as compared to wild-type (Figure 4.11).   Figure 4.11: Extracellular autofluorescent bodies in acos5 anther locules do not stain with Nile Red. Using two-photon microscopy, the lipid content of wild-type and acos5 anthers was assayed by Nile Red staining. Nile Red fluorescence emission was captured from 575-630 nm (coloured green) and a broader emission primarily from intrinsic fluorescence was captured from 380-560 nm (coloured red). In wild-type (A,D) and acos5 (B,E, enlarged in C,F respectively), anther sporophytic cells showed no differences in lipids, with abundant staining bodies present in the tapetal cells. As compared to the brightly staining wild-type microspore walls, acos5 microspore walls (when present) did not stain or stained minimally with Nile Red (B, arrow in C). Microspore cytoplasm stained more readily than wild-type microspores stained for equivalent lengths of time. Surrounding acos5 tapetal cells, orbicule-like bodies were detected due to their intrinsic fluorescence (red puncta in F), which excluded Nile Red stain. Such bodies were absent in wild-type. Msp, microspore, Tp, tapetum. Bars = 20 μm.      118 4.2.4 Role of phenylpropanoid and flavonoid biosynthetic enzymes in sporopollenin biosynthesis    Double mutant analyses of abcg26 with key polyketide biosynthesis mutants (acos5, pksa pksb, and tkpr1) supported the hypothesis that ABCG26 exports a product of the polyketide biosynthetic pathway that is incorporated into sporopollenin. While the composition of the fluorescent ORBs observed in the polyketide pathway mutants remained unknown, we hypothesized that these bodies contain phenolic metabolites based on their autofluorescence and lack of staining with Nile Red. To test this hypothesis, we first examined the pollen wall phenotypes of Arabidopsis mutants defective in key enzymes required for phenolic compound biosynthesis.   Despite numerous reports that phenolic compounds, such as hydroxycinnamic acids, are present in sporopollenin (Wehling et al., 1989; Ahlers et al., 1999; 2003; Prahl et al., 1986; Rozema et al., 2001)), there is little information regarding the enzymes that may be involved in synthesizing such components. Thus, we targeted anthers of mutants in genes encoding enzymes in the phenylpropanoid and flavonoid pathways for analysis by 2-P. These included genes encoding 4-COUMARATE:COA LIGASE 3 (4CL3, At1g65060), as well as mutants in genes encoding CHALCONE SYNTHASE/TRANSPARENT TESTA 4 (CHS/TT4, At5g13930), and CYTOCHROME P450 84A1/FERULIC ACID 5-HYDROXYLASE 1 (CYP84A1/FAH1, At4g36220) and SPERMIDINE HYDROXYCINNAMOYL TRANSFERASE (SHT, At2g19070). SHT was of particular interest, since this enzyme is required for the biosynthesis of hydroxycinnamoyl spermidine conjugates (HC spermidines) that are constituents of the pollen coat in Arabidopsis (Grienenberger et al., 2009). Previous reports identified altered fluorescence emission from mature pollen grains of sht, 4cl3, and tt4 mutants (Dobritsa et al., 2011; Grienenberger et al., 2009). Here, we investigated the emission profiles of intrinsic fluorescence of developing microspores following tetrad release and preceding tapetal cell death, to differentiate between emissions from sporopollenin and pollen coat constituents.   To capture both the range of anther autofluorescence and the wavelength(s) where emission reaches maximum intensity, we performed spectral emission scans on intact developing mutant and wild-type anthers using 720 nm 2-P excitation and a 400-580 nm emission capture   119 range. Intensity projections over the range of wavelengths allowed emission spectra specific to traced microspore walls to be captured, while avoiding emission from surrounding sporophytic tissues (Figure 4.12). The mean emission intensities and SE values for wild-type, 4cl3-2, tt4-2, sht and fah1-2 (n=24 microspores per genotype) were normalized to the 460 nm peak, which reached its maximum intensity in wild-type samples, to correct for differences in emission intensity between samples. The resulting relative intensities from microspore emissions for wild-type, sht, 4cl3-2, tt4-2, and fah1-2 microspores are summarized in Figure 4.13. The wild-type microspore emission spectrum consisted of one broad peak with a maximum intensity at 460 nm. As reported previously for mature pollen grains, 4cl3 and tt4 mutant microspore walls exhibited broader emission surrounding the 460 nm peak relative to wild-type, with increased intensities below 460 nm and decreases above 460 nm. The fah1 and sht mutants showed a different trend, with spectral emission shifts towards wavelengths longer than 460 nm, peaking at 510 nm and 540 nm, respectively. Most notably, the emission profile from sht microspore walls prior to tapetal cell death differed dramatically from wild-type. The captured sht emissions were predominantly in the longer wavelength 495-550 nm channel, producing images with red-colored microspores, rather than teal microspores seen in wild-type (Figure 4.14A-D, F-I). These data indicate that the products synthesized by 4CL3, CHS/TT4, CYP84A1/FAH1 and SHT affect the fluorescent profile of the pollen exine, reflecting changes in the composition of the pollen cell wall in the stages when it consists predominantly of sporopollenin.  The dramatic fluorescence spectral profile differences between wild-type and sht microspore walls indicate that hydroxycinnamoyl spermidines are present in the microspore wall prior to tapetal cell death.     120  Figure 4.12: Microspore emission from traced regions of interest in developing anthers. Using two-photon microscopy, spectral emission scans in the 400-580 nm interval were obtained with a 10 nm step size under 720 nm laser excitation. For each genotype, four microspores per locule were traced, as shown for wild-type (A) and sht mutant (B) here. From pixels within traced regions, average emission intensity was plotted for each 10 nm wavelength interval captured for each anther sampled, with wild-type (C) and sht mutant (D) shown here.    121  Figure 4.13: Spectral emission profiles for wild-type, 4cl3, tt4, sht, and fah1 mutant microspores within developing anthers and prior to tapetal cell death. Under 720 nm laser excitation, spectral emission scans in the 400-580 nm interval were obtained with a 10 nm step size and normalized to the 460 nm peak emission intensity. For each genotype, four microspores per locule were traced from six anthers and mean +/- SE emission intensity at each 10 nm wavelength was plotted.   0	  0.5	  1	  1.5	  2	  2.5	  3	  400	  410	  420	  430	  440	  450	  460	  470	  480	  490	  500	  510	  520	  530	  540	  550	  560	  570	  580	  Relative	  Intensity	  Wavelength	  (nm)	  Wild-­‐type	  	  4cl3-­2	  tt4-­2	  sht	  fah1-­2	    122  Figure 4.14: Hydroxycinnamoyl spermidines constitute orbicule-like bodies in acos5. In the two-photon microscope, wild-type microspores emit bright autofluorescence preferentially in short wavelengths (420-460 nm, coloured teal), rather than longer wavelengths (495-540 nm, coloured red; A-D). In contrast, sht microspores emit bright autofluorescence predominantly in longer wavelengths (495-540 nm coloured red), rather than in short wavelengths (teal; F-I). Wild-type and sht anthers lack orbicule-like bodies surrounding tapetal cells (A-J). acos5 orbicule-like bodies surrounding tapetal cells and some microspores (K-N, arrows in O) are not observed in the sht acos5 double mutant by two-photon microscopy (P-S) and TEM (T). Bars = 20 μm (two-photon images), and 2 μm (TEM images).  4.2.5 Polyketide product-dependent extracellular fluorescent bodies (ORBs) contain hydroxycinnamoyl spermidines    Given our data on the biosynthetic origins of fluorescent material deposited on developing microspores, we tested the hypothesis that ORBs observed in polyketide pathway   123 mutants are composed of phenolic metabolites derived from phenylpropanoid metabolism by generating double mutants of phenylpropanoid biosynthetic genes and acos5.  In the sht acos5 double mutant, ORBs were completely absent, and were not found surrounding tapetal cells or microspores (Figure 4.14P-S). These results were confirmed by TEM observations, where abundant ORBs surrounded tapetal cells in acos5 mutants (Figure 4.14O), but were absent in sht acos5 double mutants (Figure 4.14T).  As negative controls, wild-type and sht mutant anthers lacked ORBs (Figure 4.14E and J).  These data suggest that the extracellular bodies observed in acos5 locules are composed of HC spermidines. In contrast, mutations in genes in the flavonoid pathway did not affect the ORBs surrounding tapetal cells and some microspores in acos5.  ORBs were observed in acos5 4cl3 and acos5 tt4 double mutants (Figure 4.15). These data suggest that the ORBs are not composed of flavonoid-derived compounds.      124 Figure 4.15: The acos5 4cl3 and acos5 tt4 double mutants exhibit the same phenotype as the acos5 single mutant. Two-photon imaging of anther autofluorescence revealed the presence of orbicule-like bodies, as seen in acos5 single mutants, in the 4cl3 acos5 (A,B) and tt4 acos5 double mutants (C,D). The orbicule-like bodies in 4cl3 acos5 and tt4 acos5 anthers surrounded tapetal cells on all sides and surrounded some microspores. These bodies appeared as circular puncta and were particularly abundant in sections through the tapetum along the outer periclinal face (D). Bars = 20 μm.   Taken together, these data suggest that the trafficking of HC spermidines to the locule and developing microspore wall occurs prior to tapetum programmed cell death, in conjunction with the apparent ABCG26-mediated export of polyketide pathway products.  Since the putative HC spermidine-containing bodies accumulate only when the polyketide pathway is disrupted, trafficking of the HC spermidines from the tapetum surface to developing microspores appears to require the polyketides.  One explanation for these data is that co-polymers of polyketides and HC spermidines traffic from the tapetum to the microspore wall, prior to deposition into the exine of developing microspores.   4.2.6 HC spermidines do not contribute to autofluorescence of vacuolar inclusions in tapetal cells of the abcg26 mutant    The presence of HC spermidine-containing ORBs in polyketide pathway mutants and in their respective double mutants with abcg26 indicated that HC spermidine trafficking from tapetal cells is independent on the ABCG26 transporter. However, the HC spermidine bodies did not accumulate in the abcg26 single mutant, for reasons that are not clear. To examine the possibility that HC spermidines accumulate inside tapetal cells in the absence of polyketide export, sht abcg26 double mutant anthers were examined in parallel to abcg26 anthers by 2-P microscopy. In sht abcg26 anthers, brightly fluorescent tapetum vacuolar inclusions similar to those observed in abcg26 were apparent (Figure 4.16A-H). The tapetum vacuolar inclusions observed in sht abcg26 and abcg26 appeared indistinguishable by TEM (Figure 4.16I,J). The persistence of tapetum vacuolar inclusions in sht abcg26 supports the model presented above that the vacuolar inclusions contain products of the polyketide biosynthetic pathway involving ACOS5, PKSA PKSB and TKPR1.    125    126 Figure 4.16: abcg26 and sht abcg26 tapetum vacuolar inclusions appear indistinguishable by two-photon microscopy and TEM.  In abcg26 (left panel) and sht abcg26 (right panel) anthers, bright intrinsic fluorescence was emitted at the locule periphery, in tapetum and middle layer cells (A and B, enlarged in C and D, respectively). abcg26 and sht abcg26 tapetal cells contained autofluorescent vacuoles with numerous brightly emitting puncta (E and F, enlarged in G and H, respectively). In TEM, tapetal cell vacuoles contained electron-dense circular cores in abcg26 (I) and sht abcg26 (J), which varied in number and size between samples. No consistent differences were observed between the inclusions in abcg26 and sht abcg26 anthers. Bars = 20 μm (A,B,E,F) and 1 μm (I, J).   4.2.7 The incorporation of HC spermidines in the pollen wall    While HC spermidines have been previously described as components of the pollen coat in Arabidopsis, their export from tapetal cells prior to programmed cell death, documented above, makes them strong candidates for incorporation into sporopollenin during exine formation. To determine whether the spermidine conjugates are incorporated into the sporopollenin biopolymer or remain more loosely attached as components of the pollen coat, the chemical recalcitrance of wild-type and sht mutant sporopollenin of mature pollen grains after removal of the overlying pollen coat was assessed. To test the chemical integrity of sht pollen grains, pollen were subjected to acetolysis, a standard test for sporopollenin chemical integrity (representative images in Figure 4.17 for three trials). However, these analyses failed to reveal any differences between wild-type and sht mutant pollen walls, which were both insensitive to acetolysis, with 11.3% wild-type exines, and 5.3% sht exines showing visible damage (n=600 pollen grains per genotype).     127  Figure 4.17: The chemical recalcitrance of the sporopollenin polymer in wild-type and sht mutant pollen is indistinguishable. Bright-field image of wild-type (A,C) and sht (B,D) exines after acetolysis treatment. The majority of pollen grains exhibited no visible damage to their exines after acetolysis treatment (A,B), with a minor fraction lysed or exhibiting visible minor damage (C,D). Most pollen exine remained intact (A,B), but a small percentage (11% of wild-type and 5% of sht) exines exhibited damage after treatment (C,D). Bars = 10 μm (A-D).   4.3 Discussion   In this study, we provide new insights into the nature of sporopollenin and pollen wall exine formation. The genetic link found between the ABCG26 transporter and the polyketide pathway involved in sporopollenin biosynthesis provides evidence that the substrate transported by ABCG26 is a polyketide product.  Furthermore, our analyses showed that HC spermidine conjugates known to be present in the Arabidopsis pollen coat are trafficked from tapetal cells prior to programmed cell death, in an ABCG26-independent fashion. Although polyketide components can assemble in the absence of HC spermidines, spermidine conjugates required the polyketide component of sporopollenin for normal trafficking and deposition on microspores. In addition to revealing mechanisms of pollen exine traffic previously not understood, the novel two-photon microscopy approach employed here has opened the doors to the study of intact anthers and provides a useful tool for future studies.   128  4.3.1 In planta characterization of the putative substrate of ABCG26   Previous studies that relied on examination of fixed and sectioned tapetal cells were unable to identify accumulations in abcg26 mutant tapeta that could be indicative of putative ABCG26 substrates.  In contrast, two-photon microscopy identified inclusions in abcg26 tapeta, in the form of large autofluorescent vacuoles.  This information facilitated a targeted genetic approach to determine the biosynthetic origin of these inclusions as polyketide pathway products.  Based on the first appearance of these vacuolar inclusions in abcg26 tapetal cells immediately following the tetrad stage, followed by their continued enlargement over the course of microspore development, the ABCG26 transporter appears to be active over this developmental period, consistent with the timing of its expression in tapetal cells (Quilichini et al., 2010).    Analysis of lipidic metabolite accumulation in live tapetal cells, using Nile Red, revealed similar abundances of lipids in wild-type and abcg26 cells.  Thus, the compounds accumulating in abcg26 tapetal cells differ dramatically in nature from the putative cuticular substrates that accumulate in abcg11 and abcg12 mutants, two half-size ABCG transporters in the same subfamily (Pighin et al., 2004; Bird et al., 2007). Accumulations in abcg11 and abcg12 epidermal cells stain with Nile Red, appear lamellar and are associated with the ER (Pighin et al., 2004; Bird et al., 2007; McFarlane et al., 2010).  Therefore, the ABCG26 substrate is likely a unique polyketide-derived, autofluorescent compound translocated from tapetal cells to the microspores in the locule, distinct from the apparent lipidic cargo of ABCG11 and ABCG12.   4.3.2 The polyketide biosynthetic pathway and transport by ABCG26 are linked    Only a handful of enzymes required for sporopollenin synthesis in Arabidopsis have been characterized biochemically. Among these, ACOS5, PKSA, PKSB, and TKPR1 are proposed to function sequentially in the production of tetraketide α-pyrones (Grienenberger et al., 2009). The strong phenotypes of acos5, pksa pksb, and tkpr1 mutants that, like abcg26, are male sterile and lack sporopollenin deposition, as well as the overlapping spatial and temporal expression   129 patterns shared by the biosynthetic genes and ABCG26, suggested that polyketides produced by these enzymes could be exported by ABCG26 (Quilichini et al., 2010; Grienenberger et al., 2010). We directly tested this hypothesis by examining the phenotypes of double and triple mutant combinations affecting each of the enzymes in the pathway in the abcg26 mutant background. Two-photon imaging and TEM analyses of the abcg26 acos5, abcg26 pksa pksb, and abcg26 tkpr1 mutants revealed phenotypes identical to the acos5, pksa pksb and tkpr1 mutants, respectively, including the lack of autofluorescent vacuolar inclusions in tapetal cells. The suppression of the abcg26 tapetum vacuolar inclusion phenotype in each of these mutant backgrounds provides direct evidence that the polyketide biosynthetic pathway is required for these ABCG26-dependent tapetal cell inclusions. The simplest interpretation is thus that the tetraketide α-pyrones or their derivatives are in fact the ABCG26 substrates, although transport assays will be required to precisely define the ABCG26 substrate specificity.   The finding that tapetum inclusions in abcg26 likely represent products of the polyketide pathway provides a better understanding of the roles played by polyketide pathway product(s) in exine formation in planta. The bright fluorescence of the abcg26 tapetum-trapped metabolites suggests the presence of aromatic rings or conjugated pi bonds in the metabolites. While the proposed product of the sporopollenin polyketide metabolon is a tetraketide α-pyrone, there is limited literature on the intrinsic fluorescent properties of α-pyrone rings. It is also possible that the tetraketide product of this polyketide pathway may be further modified or cyclized prior to export from the tapetum. Polyketide cyclization by type III polyketide synthases, such as PKSA and PKSB, is known to vary, encompassing heterocyclic lactone formation, intramolecular condensations either by aldol or Claisen carbon-carbon bond formation, or to be absent altogether (Austin and Noel, 2003; Funa et al., 2006; Cook et al., 2010; Tropf et al., 1994; Goyal et al., 2008). In Cannabis, the cooperative activity of two proteins, a type III PKS and an accessory protein called olivetolic acid cyclase, is required for the correct cyclization of polyketides in the production of a key intermediate in THC synthesis (Gagne et al., 2012). Without olivetolic acid cyclase, the polyketide enzyme forms olivetol and α-pyrone by-products.  The occurrence and function of α-pyrones in plants is minimally understood, and some reports have suggested they are derailment by-products of in vitro assays (Yamaguchi et al., 1999; Akiyama et al., 1999; Abe et al., 2004). However, novel pathways capable of pyrone formation   130 in plants have recently been described, and their presence in the sporopollenin biopolymer is plausible (Weng et al., 2012; Eckermann et al., 1998). Thus, the exact chemical nature of the autofluorescent polyketide pathway-dependent product that appears to be the ABCG26 substrate for export into the locule remains to be determined.   4.3.3 Orbicule-like bodies in polyketide mutants provide insight into trafficking of novel exine components    The suppression of ABCG26-dependent tapetum vacuolar inclusions by polyketide biosynthetic mutants provides evidence for a functional link between the synthesis and export of polyketides, both of which are required for sporopollenin formation. However, a second and more puzzling phenotype was observed in mutants blocked in polyketide biosynthesis. Locule-localized autofluorescent ORBs were observed in acos5, pksa pksb, tkpr1 and in their respective double mutants in the abcg26 background by two-photon microscopy. These had obvious similarity to the orbicules observed in the locules of some species in association with exine deposition (Wang et al., 2003; Huysmans et al., 1998). The presence of ORBs in the absence of polyketide biosynthesis suggested the existence of a second pathway that requires the polyketide component of sporopollenin in order to be assembled into the exine, and that these metabolites are exported into the locule independently of ABCG26.  4.3.4 HC spermidines are exported from tapetal cells prior to programmed cell death and are incorporated into exine   The ORBs observed in polyketide biosynthetic pathway mutants provide information that extends current understanding of exine assembly. The autofluorescent nature of ORBs supports the presence of aromatic rings and their exclusion of Nile Red stain suggests a non-hydrophobic composition, while their dependence on a functional polyketide pathway suggests interaction with these metabolites. In the acos5 mutant background, the sht mutant, blocked in HC spermidine synthesis, suppressed ORB formation (Figure 4.14), but mutants blocking other   131 phenylpropanoid branch pathways leading to flavonoid and monolignol biosynthesis had no effect.   While sht mutants produce fertile, morphologically normal pollen (Grienenberger et al., 2009), the exine of developing sht microspores exhibited significantly shifted emission spectra relative to wild-type microspores prior to tapetum programmed cell death based on two-photon microscopy, indicating a change in exine composition in the sht mutant early in microspore wall development.  These data indicate that HC spermidines, previously characterized components of the pollen coat, are not contributed solely after tapetum programmed cell death, but are exported from tapetal cells with similar timing to the sporopollenin components contributed by the polyketide pathway. This provides novel evidence for the pre-mortem trafficking of pollen coat components from the tapetum and suggests that, because HC spermidines are present in the locule at the time of sporopollenin assembly, these compounds could be incorporated into the growing polymer.  The dependence of HC spermidine accumulation in ORBs upon polyketide biosynthesis raises intriguing questions regarding the functional link between these two apparent exine components.  The polyketide component of sporopollenin is capable of assembly in the absence of HC spermidines, as seen from the sht mutant, while the HC spermidines require polyketide synthesis to assemble on the pollen wall. One plausible interpretation of these data is that polyketides form an exine framework by self-assembly, to which HC spermidines may adhere during the phase of rapid exine development on microspores released form tetrads.  Since HC spermidines coalesce into ORBs in the absence of polyketide biosynthesis, co-trafficking and potential physical interaction of HC spermidines with polyketides upon their release into the locule is an attractive model. The presence of HC spermidine-containing ORBs around some microspores, and their absence around others, is puzzling and suggests that unknown anchoring or targeting mechanisms may also be in place. Finally, an alternate explanation for ORB formation in polyketide mutants is that the metabolic block in the polyketide pathway introduced by acos5 and other mutants in the pathway leads to increased metabolic flux into the HC spermidine pathway (Weng et al., 2010).   Since HC spermidine-containing ORBs still formed when polyketide biosynthetic mutations were placed in the abcg26 mutant background, their export from tapetal cells appears   132 to be ABCG26 independent. Although the mechanism for HC spermidine export from tapetal cells is unknown, it is interesting that these bodies were observed on all tapetum faces in polyketide mutants, suggesting that the tapetal cells may not be polarized in their secretion to the locule.   At present, we have no explanation for the lack of HC spermidine-containing ORBs in the abcg26 single mutant, which, according to our model, should lack polyketide accumulation in locules similarly to acos5, acos5 abcg26, and other polyketide mutants in wild-type or abcg26 backgrounds in which ORBs are prominent. If only transport of polyketides, and not their biosynthesis, is affected in the abcg26 transporter mutant, then residual transport of some polyketide product into the locule could explain these results.  Residual transport could occur from the presence of several other ABCG transporters, whose expression in tapetum has been demonstrated (Ariizumi and Toriyama, 2011).  Alternatively, the export of HC spermidines from intact tapetal cells could be inhibited by the accumulation of polyketides in tapetal cells brought about by their blocked transport. This in turn would suggest that abnormal interaction between polyketides and HC spermidines within tapetal cells, or the existence of a feedback mechanism, represses HC spermidine biosynthesis upon abnormal polyketide accumulation in tapetal cells.   Finally, the functions of HC spermidines that appear to be deposited into the exine in parallel with polyketide α-pyrones remains to be established. Mutants lacking SHT function are fertile (Grienenberger et al., 2009), and we found no difference in the sensitivity of sht vs. wild-type pollen grains to acetolysis (Figure 4.15), suggesting that the robustness of pollen wall exine is not compromised in the absence of this component. However, it is possible that HC spermidine components play more subtle roles in resistance to environmental stresses such as desiccation or UV irradiance.    In summary, our results from live cell imaging of Arabidopsis anthers reveal that the cargo transported by ABCG26 includes polyketide α-pyrone components of sporopollenin, that HC spermidines are transported from tapetal cells into the anther locule prior to tapetum programmed cell death, and that HC spermidines are components of the exine. A complex and as of yet poorly understood interplay between polyketide products and HC spermidine biosynthesis appears to govern their trafficking and deposition, and will be the basis for future studies.      133 4.4 Materials and methods  4.4.1 Plant growth   Arabidopsis seeds were germinated on Murashige and Skoog medium plates, pH 5.7. Seeds were imbibed in the dark at 4ºC, grown under continuous light for 7 to 10 days and then transplanted to soil (Sunshine Mix 4; Sungrow Horticulture). Seedlings were grown to maturity at 20ºC in 18 hour light and 6 hour dark cycles.    4.4.2 Two-photon laser scanning microscopy analysis   For stamen dissection, unopened flower buds were excised from plants using an Olympus SZX10 research stereomicroscope. Buds measuring 1-1.2mm in the free microspore stage were used for stamen collection. Stamens from one bud were immersion in distilled water for intrinsic fluorescence analysis or Nile Red staining solution for lipid analysis. Air bubbles were removed by gentle manipulation and samples were covered with a 1.5mm coverslip. Stamens were examined by 2-P microscopy immediately following dissection (and staining, when applicable) using the Olympus Multi-photon Laser Scanning Microscope FV1000MPE and the Olympus XLPLN 25X WMP dedicated objective.   Anther intrinsic fluorescence was examined using 720 nm excitation and the BFP/GFP/RFP/DsRed filter cube for four-channel imaging of stamens immersed in distilled water. Emission was captured in two channels, RXD1 (420-460 nm, coloured teal) and RXD2 (495-540 nm, coloured red).   Nile Red dye (9-diethylamino-5H-benzo[alpha]phenoxazine-5-one) stock solution was prepared in acetone (0.1mg/mL) and fresh working solutions were made in sterile 25% glycerol (1µg/mL) (Greenspan et al., 1985). Microscope settings optimal for intracellular Nile Red fluorescence emission detection were determined by comparing the fluorescence emission profiles for wild-type anthers in 25% glycerol and 25% glycerol with 1µg/mL Nile Red. 810 nm excitation and the CFP/YFP/RFP/DsRed filter cube (Olympus) were used for two-channel imaging, capturing emission from RXD3 (380-560 nm, coloured red) for intrinsic fluorescence   134 and RXD4 (575-630 nm, coloured green) for Nile Red. Unstained anthers fluoresced in the RXD3 channel, with only traces of fluorescence emission in the RXD4 channel. Wild-type anthers immersed in Nile Red staining solution emitted fluorescence due to Nile Red fluorescence initially along the anther cuticle, and microspore wall staining became visible after 20-30 minutes. Olympus FV10-ASW ver03.01 and Volocity Version 6.1.1 software packages were used for image analyses.  4.4.3 Spectral analysis (lambda scans)   After orienting samples with the UV intrinsic fluorescence settings described above, lambda analyses were performed with 15% laser power (with the exception of tt4-2 mutants) at 720 nm excitation, with 800 HV, 8 s pixel dwell time per µm, and a 256x256 pixel density, with RDM690, VBF and PMT1 detector selections in place. Emission was collected between 400-600 nm, with a 10 nm bandwidth and step size. Samples that burst during lambda scans were not included in analyses. Following spectral scans, emission from traced regions of interest (from the intensity projection over the Lambda axis) were plotted using the Olympus FV10-ASW ver03.01. Intensity projection over lambda axis was used to trace microspore wall fluorescence (n=200 pollen grains, per trial).   4.4.4 Transmission electron microscopy analysis   Unopened flower buds were excised from plants, sepals and petals excised, and specimens were submerged in 0.2M sucrose in uncoated copper type B sample holders (Ted Pella, Inc.). Samples were frozen at high pressure with a Leica EM HPM 100 high pressure freezer, followed by freeze substitution and embedding in Spurr’s resin (Spurr, 1969) as described by McFarlane et al. (2008). Freeze substitution medium contained 2% (w/v) osmium tetroxide in acetone with 8% (v/v) dimethoxypropane.      135 4.4.5 Acetolysis   A pellet of pollen from ten open flowers was washed with glacial acetic acid, then placed in acetolysis solution (9 parts acetic anhydride : 1 part concentrated sulfuric acid), sealed and boiled for 10 minutes.  Remaining exines were washed in glacial acetic acid, then three times with water and imaged with a light microscope (Leica DMR microscope). Three trials, n = 200 exines counted per trial.   136 Chapter 5: The putative role of lipid transfer proteins in sporopollenin transport  5.1 Introduction   Lipid transfer proteins (LTPs), named for their in vitro ability to move phospholipids and fatty acids between membranes, form a large group of small, basic proteins in land plants (Edstam et al., 2011). The characteristic presence of eight cysteines distributed over the backbone of these proteins enable the formation of a lipid-binding hydrophobic pocket through disulphide bonds (Douliez et al., 2000; José-Estanyol et al., 2004). The majority of LTPs also have a signal peptide that directs these proteins to the apoplast, suggesting that the function of LTPs may extend beyond mediating transfers between membranes within the cell (Kader, 1996). Thus, a number of reports have implicated LTPs in secretory roles for the formation of plant hydrophobic barriers, including the cuticle (DeBono et al., 2009; Sterk et al., 1991) and the pollen wall (Zhang et al., 2008; Ariizumi and Toriyama, 2011; Huang et al., 2009).  However, the physiological function of LTPs in planta is largely unknown (Boutrot et al., 2008).   In support of their proposed function in male reproductive development and/or pollen exine traffic, a large number of LTPs have been noted for their anther- or tapetum-specific expression patterns (Huang et al., 2009). OsC6, encoding a type VII LTP in rice, is preferentially expressed in the tapetum and is required for normal exine and orbicule formation (Boutrot et al., 2008; Zhang et al., 2010). However, as with a number of rice proteins required for sporopollenin synthesis or deposition, OsC6 function appears to extend to anther cuticle formation (Zhang et al., 2010). In Arabidopsis, ARABIDOPSIS THALIANA ANTHER7 (ATA7/LTPH1) exhibits tapetum-specific expression in the free microspore stage by in situ hybridization (Rubinelli et al., 1998). In addition, the expression of three genes encoding type III LTPs (LTPC6, LTPC9, and LTPC14) is restricted to the tapetum, and plants carrying the dual knockdown of LTPC6 and LTPC14 exhibited abnormal intine and dehydration-sensitive pollen (Huang et al., 2013b). Interestingly, the OsC6 and AtC6, C9 and C14 LTPs exhibit different localizations from their encoding transcripts, as they were localized to the locule and anther epidermis in rice, or in the locule and in association with the pollen exine in Arabidopsis (Zhang et al., 2010; Huang et al.,   137 2013b). Altogether, a number of LTPs appear to exhibit spatially and developmentally restricted expression patterns within anthers, and some have roles that support their function in pollen wall (or sporopollenin trafficking). Despite numerous proposals, however, LTP(s) required for sporopollenin traffic and/or deposition on developing microspores have not been identified in Arabidopsis.  This study focused on testing the role of LTPs, if any, in sporopollenin transfer from tapetal cells in Arabidopsis. Based on co-expression analysis with ACYL-COA SYNTHASE5 (ACOS5), whose enzymatic product appears to produce a central precursor of sporopollenin (Souza et al., 2009), candidate LTPs with expression patterns highly correlated with ACOS5 were selected for analysis by reverse genetics, to test the hypothesis that they encode proteins with roles in sporopollenin trafficking. Among the co-expressed genes, plants that were heterozygous for the mutation in the SALK_038995 mutant line, with putative insertion in the promoter of ATA7/LTPH1, exhibited reduced fertility and abnormalities in pollen. No pollen wall defects were identified, but post-meiotic defects in pollen development were identified in locules containing bicellular pollen grain, suggesting that this gene warrants further study.   5.2 Results  5.2.1 Lipid transfer protein candidate selection   To test the hypothesis that one or more LTPs are involved in transferring sporopollenin components between tapeta and microspores, we searched for genes encoding annotated LTPs whose expression correlated with the tapetum-expressed ACYL-COA SYNTHASE5 (ACOS5; Souza et al., 2009). Using the Platform for RIKEN Metabolomics database (PRIMe,; tissue and development v.1), a correlated gene search with ACOS5 identified 10 LTPs with r2 > 0.50 (Table 5.1). Candidate LTPs selected for functional characterization had a high correlation coefficient with ACOS5 (r2 > 0.90), an expression pattern restricted to stage 9-11 flower buds (Winter et al., 2007), and a minimum of one T-DNA insertion line available through the Arabidopsis Biological Resource Centre (Alonso, 2003).    138  Table 5.1: Arabidopsis thaliana LIPID TRANSFER PROTEIN (LTP) genes co-expressed with ACOS5    Using a reverse genetic approach, we sought to functionally characterize the LTPs tightly co-expressed with ACOS5 based on the putative involvement of their encoded proteins in sporopollenin transfer between tapeta and microspores. Mutant plants were examined for phenotypic differences relative to wild-type Arabidopsis grown under the same conditions, with an emphasis on plant fertility and pollen production. T-DNA insertions were confirmed using gene-specific primers for all lines (Table 5.2). Plants that were homozygous for T-DNA insertions were identified for SALK_065158, SAIL_294D02, SALK_078886 seed lines, with putative insertion in or upstream of At5g07230/LTPC6, At3g52130/LTPC3 and At4g28395/ATA7/LTPH1, respectively (Table 5.1). None of these plants exhibited fertility reductions or visible pollen abnormalities, as determined by Alexander stain for viability, SEM for morphology and Auramine O stain for exine, and thus were not studied further.   Table 5.2: Sequences of primers used for PCR.     139  5.2.2 SALK_038995 heterozygous plants exhibit reduced fertility and pollen defects   For a fourth insertion line studied, SALK_038995, with a putative insertion upstream of At4g28395/LTPH1/ATA7 (Figure 5.1A), heterozygous plants exhibited a visible reduction in fertility (Figure 5.1B). As no homozygous plants were identified, plants heterozygous for the SALK_038995 T-DNA insertion were phenotypically characterized further. Fertility reduction in the mutated plants was evident from their short siliques with an average length of 0.89cm +/- 0.1, compared to 1.2cm +/- 0.2 in wild-type (n=15 siliques from the main stem, per genotype; Figure 5.1C). Seed set was additionally reduced from an average of 43 +/- 7 seeds per silique  in wild-type to 13 +/- 2 in SALK_038995 plants. The height of the SALK_038995 mutants was increased, measuring an average of 53.3 +/- 2cm along the main stem (n=6), in comparison to wild-type (34.5 +/- 3cm, n=8). Pollen was visible on the dehisced anthers of wild-type and SALK_038995 flowers, but appeared to adhere more readily to the anthers of the latter (Figure 5.1D,E).     140  Figure 5.1: Identification of Arabidopsis LTPH1 T-DNA insertion alleles. A) Scheme representing LTPH1, where black boxes denote exons (with grey segments representing untranslated regions), separated by horizontal black line denoting two introns.  Positions of T-DNA insertion in the SALK_078886 and SALK_038995 mutant lines are indicated by triangles. B) Wild-type (left) and SALK 038995 heterozygous (het, right) plant morphology at maturity. C) Siliques removed from wild-type (far left; 2.1cm in length) and SALK 038995 (right three; 1.8cm, 1.5cm, 0.5cm in length). Mature wild-type (D) and heterozygous SALK 038995 (E) flowers. Bars = 200 μm.     141  The reduced fertility in SALK_038995 and its co-expression with ACOS5 led us to examine the pollen produced by these plants for exine defects. Scanning electron microscopy (SEM) revealed that a large portion of mutant pollen was collapsed and shrunken, despite the application of critical point drying to minimize the damaging effects of surface tension during sample dehydration (Figure 5.2A,C). However, a portion of pollen in the SALK_038995 mutant plants was indistinguishable from wild-type, with three apertures, a near-spherical shape and reticulate exine (Figure 5.2A-C). Alexander stain for pollen viability supported the SEM results, with viable cytoplasm, indicated by pink staining, in wild-type and some SALK_038995 pollen, and numerous green pollen that lacked visible cytoplasmic contents specifically produced by SALK_038995 heterozygous plants (Figure 5.2D,E). Pollen exine was further examined with Auramine O stain, and no differences from wild-type were observed in the reticulate pattern of exine of wild-type and SALK_038995 pollen, even in the fraction of mutant pollen that collapsed (Figure 5.2F,G). These data suggested that the cytological development of the gametophyte, rather than wall production, of numerous pollen grains was perturbed in the SALK_038995 heterozygous mutant.      142  Figure 5.2: Pollen and pollen exine structure in wild-type and SALK 038995 heterozygous mutant plants. A-C) Scanning electron micrographs of critical point dried pollen from wild-type (B) and SALK 038995 mutant plants (A,C). Light microscope image of Alexander stained pollen from wild-type (D) and SALK 038995 (E). Confocal microscope image of Auramine O stained pollen from wild-type (F) and SALK 038995 (G). Bars = 10 μm (A), 5 μm (B,C).    143  To determine the stage at which the phenotypic defect arises in select SALK_038995 pollen, we examined pollen development in mutant anthers relative to wild-type. Microspores in the SALK_038995 plants were undifferentiable from wild-type in the callose-encased tetrad (Figure 5.3A,B) and early free microspore stages of development (Figure 5.3C,D). These data suggest normal meiotic divisions, callose production, callose degradation and exine formation in the SALK_038995 mutant. After the uninucleate stage of pollen development, two populations of microspores were evident in anther locules of the mutant. As in wild-type, some pollen in the locules of SALK_038995 anthers at a comparable stage were bicellular, contained densely staining cytoplasm and small vacuoles (Figure 5.3E, black arrowheads in F). However, a second group of pollen with lightly staining (or collapsed) cytoplasm and large vacuoles resembling an earlier ring-vacuolate stage of microspore development were specifically identified in SALK_038995 anthers (white arrowheads in Fig 5.3F). By the tricellular stage of pollen formation, differences between the two populations of pollen grains in SALK_038995 locules were dramatic (Figure 5.3G,H). While one population of pollen appeared fully developed and indistinguishable from the cytoplasmically rich tricellular pollen of wild-type (Figure 5.3G,H), the other population of pollen appeared to be empty, folded exine shells lacking cytoplasmic contents (white arrowheads in Fig 5.3H).     144    145 Figure 5.3: Microspore development in wild-type and SALK 038995 heterozygous mutant anthers. Light microscope images of wild-type (left panel) and SALK 038995 heterozygous mutant anthers (right panel). Microspore development appeared identical in wild-type and SALK 038995 mutant anthers when a callose wall encased tetrads of microspores (A,B) and after the release of free uninucleate microspores from tetrads (C,D). E) Anthers containing bicellular pollen grains (E,F) or tricellular pollen grains (G,H) exhibited differences in pollen morphology in wild-type and SALK 038995. E) All wild-type pollen contained densely staining cytoplasm and small vacuoles. F) SALK 038995 anther locules contained some pollen resembling wild-type (black arrowheads) and some pollen with lightly staining cytoplasm, large vacuoles or collapsed cytoplasm (white arrowheads). H) Tricellular pollen grains with dense cytoplasmic contents were present in wild-type anthers (G) and SALK 038995 mutant anthers (H). Tricellular pollen grains in SALK 038995 were additionally surrounded by exine fragments with minimal or no cytoplasmic contents (H), that were not present in wild-type locules (G). msps, microspores. Bars = 10 μm.     Further inspection of anthers at the bicellular stage supported the presence of two populations of pollen in the mutant line (Figure 5.4). The aberrant pollen differed from the expected appearance of pollen at the bicellular stage, due to its altered cytoplasmic staining and vacuole size. Interestingly, some pollen grains contained abnormal vesicles in the vacuole, consistent with autophagy and cell death (boxed in Figure 5.4), and were present in numerous samples (arrows in Figure 5.5). Transmission electron microscopy (TEM) showed that abnormalities were restricted to the autophagic bodies in the cytoplasm, and pollen walls exhibited intine and exine with no morphological abnormalities (Figure 5.5B). Also, some pollen in the mutants appeared collapsed at the bicellular stage (red arrowheads in Fig 5.4 and Fig 5.5). No differences in tapetal cell content, morphology or timing of programmed cell death were observed between wild-type and SALK_038995 specimens viewed by light microscopy or TEM. Altogether, these data indicate that the insertion in SALK_038995 lines causes decreased plant fertility and a severe reduction in viable pollen production, with defects arising in post-meiotic stages of pollen development. As pollen abortion was visible in approximately half of the pollen grains formed, and no plants homozygous for the SALK 038995 insertion were identified, the mutation in SALK_038995 seeds appears to affect a gene expressed in the gametophytic generation, which is required for pollen survival past the bicellular stage.     146  Figure 5.4: Anther and pollen morphology at the bicellular pollen stage in SALK 038995 heterozygous mutant anthers. Three assembled light microscope images of one SALK 038995 heterozygous mutant anther locule. The anther locule exhibited pollen with a variety of morphologies. Locules were determined to be in the bicellular stage of pollen development, based on the identification of two nuclei within one pollen grain (see dual arrow). This SALK 038995 anther locule contained some pollen with typical morphology exhibited at this stage (black arrowheads). However, some pollen exhibited decreased cytoplasmic staining and large vacuoles (white arrowheads), autophagic bodies (white boxes), or collapsed cytoplasm (red arrowhead). Bar = 10 μm.   147   Figure 5.5: Autophagic bodies in pollen from SALK 038995 heterozygous mutant plants. Light microscope images (A) and transmission electron micrographs (B) of pollen from SALK 038995 plants (heterozygous for the mutation). All pollen shown contains bodies resembling autophagic vacuoles filled with cellular components (black arrows) or collapsed cytoplasmic contents (red arrowhead). Cy, cytoplasm; Ex, exine; In, intine; Lo, locule; N, nucleus; Tp, tapetum. All images in (A) are of the same magnification. Bars = 2 μm (B).   5.3 Discussion   In this study we used co-expression analysis as a candidate selection tool, together with a reverse genetic approach, to test the role of LTPs in sporopollenin formation. Co-expression was successful in identifying a gene required for post-meiotic development of pollen. From this   148 work, it is clear that the abundance of LTPs co-expressed with ACOS5 warrants the attention of pollen wall researchers, as these LTPs provide a putative mechanism for the movement of sporopollenin precursors from tapetal cells to microspores. However, data from this study suggest that the function of LTPs may extend beyond pollen wall formation, to the development of pollen through microgametogenesis. However, further work is required, as it was not clear from these data whether the gene of interest (At4g28395/ATA7/LTPH1) was affected in the SALK 038995 mutant line, as only one of the two mutant lines with insertions upstream of the gene of interest exhibited a phenotype. Since this work, an additional insertion in the second exon of ATA7/LTPH1 has become available (GabiKat_288A12), and follow-up analyses will clarify the relationship between the mutant phenotype observed and the gene affected.   Previous data identified a tapetum-specific expression pattern for ATA7/LTPH1 by in situ hybridization (Rubinelli et al., 1998). However, SALK_038995 plants that were heterozygous for the T-DNA insertion exhibited a visible phenotype in approximately half of the pollen produced. Thus, these data suggest that mutation upstream of ATA7/LTPH1 affects the expression of a gene that is expressed by male gametophytes, rather than the sporophytic tapetum. In support of gametophytic expression of ATA7/LTPH1, the tissue-specific dataset from the Arabidopsis eFP browser identifies ATA7/LTPH1 as a gene with elevated expression levels in uninucleate microspores, tricellular pollen, and the globular, heart and torpedo stages of embryo development (Winter et al., 2007). Thus, if ATA7/LTPH1 is the mutated gene that produces the phenotype described herein, the tissue-specific expression of this gene must be revisited experimentally and embryo development in the SALK _038995 mutant line should be examined.   Here, we identified a gene that is required for pollen microgametogenesis, and results in the developmental arrest of pollen when mutated. This severe phenotype is surprising for the mutation of a single LTP, as previously characterized mutations affecting LTPs typically exhibit subtle abnormalities (Kim et al., 2012). Therefore, it is possible that the annotation for ATA7/LTPH1 is incorrect and that the encoded protein does not function in lipid transfer in planta. Although ATA7/LTPH1 sequence encodes the classic cysteine motif backbone (C-Xn-C-Xn-CC-Xn-CXC-Xn-C-Xn-C), based on a phylogenetic analyses performed by Rubinelli et al., (1997), this gene shows limited similarity to other annotated LTPs. Specifically, ATA7/LTPH1 encodes a protein with a lower isoelectric point (pI of 6.45) relative to other more basic LTP   149 proteins (pI 9-10), encodes a larger processed protein (15.6 kDa in contrast to 9-14kDa for most LTPs), lacks three of the conserved basic residues thought to bind polar lipids, and has two introns rather than one or none found in other LTP genomic DNA sequences (Rubinelli et al., 1998; Boutrot et al., 2008). Altogether, these data suggest that ATA7/LTPH1 may encode a protein required for pollen microgametogenesis that does not function as an LTP.  Future studies on ATA7/LTPH1 will require in vitro and/or in vivo characterization of the proteins’ function.   In this study, none of the T-DNA insert lines examined produced plants bearing exine defective pollen, and thus provided no evidence supporting the hypothesis that LTPs highly co-expressed with ACOS5 function in sporopollenin transport. However, the hypothesis that LTPs function in pollen wall transfer or assembly should not be discarded, given the challenges in studying these genes. As demonstrated here, obtaining mutation(s) in the coding sequence and/or promoter of LTPs of interest can be difficult, as these genes are small (encoding proteins typically 9-14kDa), and infrequently associated with T-DNA insertions. Also, when mutant ltp T-DNA lines are available, the associated phenotypes are often subtle (Kim et al., 2012). The abundance of LTPs also makes their study challenging, as multiple LTPs from within the same type may function redundantly. Thus, multiple mutations affecting more than one LTP are often required for phenotypic abnormalities to be detected, as in ltpc6 ltpc14 in pollen intine wall formation (Huang et al., 2013b). Future reverse genetic studies capable of generating plants with mutations affecting multiple co-expressed LTPs, particularly of the same LTP type, may reveal exine abnormalities consistent with LTP-mediated sporopollenin translocation.   5.4 Materials and methods    Plant growth conditions, and methods of sample preparation for light and TEM analysis used are identical to that described in Chapter 2 of this thesis. Critical point drying and SEM analysis    Twenty open flowers per genotype were fixed in 0.1M phosphate buffer with 1% paraformaldehyde and 2% glutaraldehyde at pH 7.4. Samples from each genotype were prepared   150 in parallel, using a multi-chamber sample holder. After vacuum infiltration and fixation overnight, flowers were rinsed in 0.1M phosphate buffer then dehydrated with an ethanol series (increasing ethanol concentration by 10% every thirty minutes, with an extra 100% ethanol change). Samples were dried with the critical point dryer (Autosamdri 815B, Tousimis), and stamens were gently removed, positioned on aluminum SEM stubs and coated with 8 nm gold (Cressington 208C high resolution sputter coater). SEM imaging was performed within 24 hours using a variable pressure SEM device (Hitachi S-2600N) at an accelerating voltage of 8kV. Pollen stains   Alexander stain was used for pollen viability analysis, as described previously (Johnson-Brousseau and McCormick, 2004) and stained pollen was examined with a light microscope (Leica DMR).  For exine analysis, pollen was stained with Auramine O. A pellet of pollen in water was obtained from ten open flowers by centrifugation. Water was exchanged with freshly made 0.001% Auramine O in 50mM TrisHCl, pH 7.5. Pellets were resuspended and left to stain for 1-2 minutes, then pelleted and rinsed three times with distilled water to remove residual stain. Pollen was mounted in water for observation by spinning disk confocal microscopy (Perkin-Elmer) using 491 nm excitation and a 528/38 nm emission filter. Images were processed with Volocity 4.3.2 (Improvision).       151 Chapter 6: Conclusions  6.1 Major findings of this dissertation   The dense cytoplasm and multinucleate state of tapetal cells, together with their proximity to developing pollen grains, has led to over a century of research on their function (Lantis, 1912; Frye, 1901; Witkus, 1945). The importance of the tapetum as a secretory entity specifically involved in the formation of the pollen wall has also been of long-standing interest to pollen scientists (Ubisch, 1927; Taylor, 1959; Heslop-Harrison and Mackenzie, 1967). Through numerous studies, particularly employing genetic and molecular tools to characterize genes required for pollen wall formation, the function of the tapetum in the synthesis of sporopollenin and pollen coat constituents has been demonstrated (Ariizumi and Toriyama, 2011). However, a mechanism for the export and translocation of tapetum-derived components of the pollen wall prior to tapetal cell death has remained an area of great uncertainty. More specifically, the mechanism enabling the high efflux of sporopollenin from tapetal cells, required to form pollen walls, was previously not known. The body of research detailed in this thesis addressed this gap in our understanding.  This research provided novel insight into sporopollenin traffic by combining leading-edge imaging technologies with the genetic and molecular biology tools available in Arabidopsis thaliana (Arabidopsis). As described in Chapter 2, this research began by examining the ultrastructure of the tapetum and microspore wall at all stages of pollen wall development in this model plant. The obtained data clarified and revised the morphological features associated with sporopollenin formation, highlighting the uninucleate stage of pollen development as the key stage for sporopollenin formation, in which the microspore wall rapidly assumes a highly structured sporopollenin backbone.  While this ultrastructural information demonstrated the high levels of sporopollenin present in association with uninucleate microspores, the cell biology of tapeta provided little information on the processes that move sporopollenin constituents from the tapetum to microspores. In particular, the absence of vesicles or other bodies that could assist sporopollenin transport was not observed within the medium of the locule by TEM. Thus, despite the highly active nature of tapetal cells and the appearance of sporopollenin on microspores at   152 the same uninucleate microspore stage, the mode of sporopollenin export and traffic was not apparent.   The absence of visible traffic between the tapetum and microspores led us to test the hypothesis that sporopollenin precursors are exported from tapetal cells by the action of one or more transport proteins, as described in Chapter 3. Aided by the previous identification of genes encoding putative sporopollenin biosynthetic machinery, the ABCG26 gene, encoding an annotated putative ATP-binding cassette transport protein, was identified through co-expression analysis as a top candidate for the export of sporopollenin precursors. This hypothesis was supported by the tapetum-preferred expression pattern exhibited by ABCG26, which peaked at the uninucleate stage of development. Using a reverse genetic approach, the defects in microspore wall formation precisely at the uninucleate stage of abcg26 anther development supported a role for ABCG26 in sporopollenin export. However, the limited understanding of sporopollenin composition and lack of abnormalities observed in tapetal cells by TEM led us to investigate the putative substrate of ABCG26 by alternate methods.   I hypothesized that plants lacking ABCG26 export activity, such as in abcg26 mutants, would accumulate the substrate of this transporter in tapetal cells (Chapter 4). Numerous challenges associated with studying the tapetum, including tissue limitations for metabolite profiling analyses and their depth within the developing anther, initially posed the greatest obstacle towards identifying accumulations in abcg26 mutants. Thus, the development of a new approach to study tapetal cells, and thereby test putative substrates of ABCG26, was necessary. The application of two-photon (2-P) microscopy enabled the visualization of different metabolite classes within the tapetum, locule and microspores of live anther specimens and detection of accumulating metabolites, potentially trafficked by ABCG26, in tapetal cell vacuoles of the abcg26 mutant. With the spatial and temporal resolution provided by this technique, the appearance of these accumulations was found to parallel the appearance of sporopollenin on wild-type free microspores and subsequent pollen development stages. Although the intrinsic fluorescence and hydrophobic nature of these inclusions gave an indication of the properties of the accumulating metabolites, the identity of these putative ABCG26 substrates was unknown.  Armed with a toolbox of previously characterized biosynthetic mutants altered in exine formation, the application of 2-P microscopy, and subsequent confirmation by TEM, allowed the   153 analysis of double mutants affecting the transporter and the synthesis of select metabolites present in the exine, to determine the class of compounds accumulating in abcg26 tapeta. With these tools, the putative substrate of ABCG26 was shown to be a polyketide product of the sporopollenin metabolon formed by ACOS5, PKSA, PKSB and TKPR1.   2-P microscopy of various exine-deficient mutants additionally revealed information on the pre-mortem trafficking of tapetum-derived hydroxycinnamoyl spermidines, previously known to be constituents of the pollen coat.  Together with the proposed ABCG26-mediated polyketide export, the dual traffic of pollen exine constituents from tapetal cells indicates that multiple routes of export from tapetal cells are possible. These data additionally raise the possibility that sporopollenin is a heteropolymer of constituents that traffic from tapetal cells and assemble on the microspore surface.    In summary, through the co-application of high quality imaging techniques and genetic tools, this dissertation informs the biochemistry, transport and assembly of sporopollenin. This work characterized a novel gene required for sporopollenin formation in Arabidopsis, and its annotation as a transport protein, together with its mutant phenotype support its role in sporopollenin export from tapetal cells. This research extended beyond the functional characterization of a single gene, to determine the functional relationships between a set of known sporopollenin related genes in planta.  The tools employed in these analyses hold tremendous potential for future studies on the interplay of processes governing sporopollenin formation. In particular, progress in understanding the mechanism of sporopollenin traffic through the locule and mechanisms of sporopollenin assembly in planta are poorly understood, but previously characterized and future mutants related to these processes could benefit from the application of these tools.   6.2 Questions arising from this research and future directions    By providing insight into the transport of sporopollenin and pollen coat constituents, this research has opened the door to a plethora of new questions on sporopollenin and the processes governing pollen wall traffic and assembly.    154 6.2.1 What is the substrate of ABCG26?    My data suggest a polyketide, produced by the sporopollenin metabolon, as the substrate of ABCG26. While enzyme assays using ACOS5, PKSA PKSB and TKPR1 sequentially produce tetraketide α-pyrones in vitro (Grienenberger et al., 2010), the in planta sporopollenin precursor transported by ABCG26 was not known. Although our understanding of the function of α-pyrones in plants is limited, the possibility that the α-pyrones produced by the sporopollenin metabolon represent the in planta products of this pathway should be seriously considered, as a few reports have demonstrated that plants are capable of α-pyrone synthesis (Eckermann et al., 1998; Weng et al., 2012). However, it is also possible that the α-pyrones observed previously are by-products of in vitro assays (Yamaguchi et al., 1999; Akiyama et al., 1999; Abe et al., 2004). In particular, type III polyketide synthases, such as PKSA and PKSB, are known to vary in their polyketide product cyclizations, with different intramolecular or heterocyclic condensations capable of producing aromatic rings or lactones, or lacking cyclization activity altogether (Tropf et al., 1994; Austin and Noel, 2003; Funa et al., 2006). The PKSA and/or PKSB enzymes may also function together with an accessory protein to alter the cyclization of polyketides as they are synthesized, as in the production of olivetolic acid in Cannabis (Gagne et al., 2012). Interestingly, the tapetum-trapped polyketide pathway-dependent metabolites in abcg26 were fluorescent under UV excitation, suggesting that conjugated pi bonds such as those present in aromatic rings, may be present in the metabolites produced by the sporopollenin metabolon. Thus, it is possible that the tetraketide product of the sporopollenin-related polyketide pathway may be further modified or cyclized prior to export from the tapetum and future experiments aimed at identifying accessory proteins capable of cyclization activity would be of value.   A second approach to determine the in planta product of the polyketide pathway and putative substrate of ABCG26 could employ direct metabolic profiling of abcg26 immature anthers or tapeta, relative to wild-type. Sensitive biochemical analyses could enable the identification of tapetum polyketides, predicted to be specific or at elevated levels in abcg26 anther extracts, as their incorporation into sporopollenin in wild-type pollen walls would prevent their extraction. However, this approach is likely to pose significant challenges in data analysis,   155 as the metabolites of interest are unknown and difficult to identify from the complex mixture of an anther extract. To circumvent these challenges, microanalysis methods to profile tapetum metabolites could be more informative. To employ micrometabolomic approaches on tapetal cells, sample preparation would require their isolation, such as by laser microdissection, or their exposure to ionization, such as by cryo-fracturing immature anthers. The application of laser microdissection for metabolic profiling has been prevalent in biomedical research for the last decade and this technology has been successfully applied more recently for the analysis of specialized plant cells (Reviewed by Moco et al., 2009). Laser microdissection has been successfully applied for the extraction of transcripts from tapetal cells or anther sporophytic tissues in Arabidopsis, rice and maize (Reviewed by Huang et al., 2011). However, the high levels of required dissections, inability to amplify metabolites, and sensitive metabolic profiling techniques required, suggest that further development of these techniques will be necessary before they can be successfully applied to Arabidopsis tapetum metabolome studies (Taylor-Teeples et al., 2011; Huang et al., 2011). As an alternative to tapetum dissection, cryo-fractured or sectioned anthers mounted for nano time-of-flight (ToF) secondary ionization mass spectrometry (SIMS), or in situ matrix-assisted laser desorption ionization (MALDI) tissue imaging using quadrupole ion mobility ToF mass spectrometry (Moco et al., 2009; Svatoš, 2011) could be used to examine the distribution of metabolites in anthers, and to “hone in” on compounds localized to abcg26 tapetal cells. Although these approaches will require biochemical expertise and optimized sample preparation, these analyses could enable the first identification of sporopollenin precursors, in planta, prior to their export from tapetal cells and polymerization on microspores.   Finally, the development of assays to test the transport activity and specificity of ABCG26, particularly against polyketide metabolite(s), would be an invaluable tool for testing putative in planta substrates of this protein. As most genes encoding enzymes characterized in sporopollenin biosynthesis are specifically expressed in tapeta, cells expressing ABCG26 for transport assays would also require the expression of ACOS5, PKSA, PKSB and TKPR1, for the sequential synthesis of the putative ABCG26 substrates of interest. In considering the challenges previously identified with the expression, glycosylation and targeting of ABC proteins in heterologous expression systems including Saccharomyces cerevisiae (Yang and Murphy, 2009),   156 the experimental design proposed is likely to require significant optimization or the transition to plant cells, such as Arabidopsis leaf protoplasts. Altogether, the in planta chemical identity of the polyketide pathway-dependent product and ABCG26 substrate remains to be determined and ABCG26-mediated transport remains to be demonstrated. Future experiments aimed at identifying and testing the substrate of ABCG26 in planta are required.   6.2.2 What is the dimerization partner(s) of ABCG26 exhibit in planta?   ABC transporters in the G subfamily, including ABCG26, are half-molecule transporters that require a partner for transport (Higgins and Linton, 2001). The dimerization form of ABCG26 proteins in planta is not currently known, but would inform its function and should be tested in future studies. Although the phenotype observed in abcg26 mutant plants was severe, it is not clear why there is variation among abcg26 silique fertility elongation and seed production. One possible explanation is that ABCG26 has the ability to heterodimerize with one or more ABCGs in the tapetum, to form a functional transport channel for the export of some pollen wall components. Interestingly, the ABCG11 and ABCG12 transporters required for diverse cuticular lipid export are capable of dimerizing in different combinations and have been proposed to function as a generalist and specialist in their export of cuticular lipids, respectively (McFarlane et al., 2010). This model is based primarily on the apparent broad substrate specificity of ABCG11, as suggested by the decrease in all cutin and wax constituents of abcg11 cuticles, and its ability to homodimerize or heterodimerize with ABCG12 in planta, in contrast to ABCG12’s predicted wax-specific export function and obligate heterodimerization with ABCG11. Based on the model for the dimerization and varied substrate export capabilities of these ABCG transporters, it is tempting to speculate that a similar mechanism for specifying different exine constituents from tapeta may be in place. For example, ABCG26 heterodimerization with another ABCG protein may function to partially compensate for the loss of function mutation in the abcg26 mutant. ABCG11 is a strong candidate for heterodimerization with ABCG26, as ABCG11 is expressed in anthers and abcg11 mutants exhibit defects in pollen coat lipids and acetolysis-sensitive pollen (Bird et al., 2007; Panikashvili et al., 2007). The ability of ABCG26   157 to form homo- or heterodimers could be tested using bimolecular fluorescence complementation or similar methods to test protein-protein interaction.  6.2.3 ABCG26 subcellular localization: tapetum polarized secretion?    A second interesting area of research extending from this research on ABCG26 will be the subcellular localization of this protein in tapetal cells. With 2-P imaging capabilities, studies on the subcellular localization of GFP-tagged ABCG26 in tapetal cells will be informative for a number of reasons. In particular, the localization pattern of ABCG26 would inform the polarity of tapetal cell secretion. For example, if ABCG26 localizes to the tapetum plasma membrane, its localization on all faces of the tapetal cells would support non-polarized secretion from tapetal cells, while its preferential localization at sub-domains of the tapetum plasma membrane would support polarity in the export of sporopollenin. Also, ABCG26 subcellular localization in protoplasts revealed preferential plasma membrane localization, with occasional ER localization signals. The high level of ABCG26 transgene in these cells was predicted to alter ABCG26 subcellular traffic to the plasma membrane, resulting in some ER localization.  Thus, the expression of this gene driven by its native promoter is predicted to produce a plasma membrane localization of ABCG26 in tapetal cells. The surprising observation of vacuolar inclusions in abcg26 middle layer cells, which were not differentiable from those in tapetum vacuoles, suggest that ABCG26 may function in both cell types. Thus, subcellular localization of ABCG26 observed in middle layer cells could indicate a supportive role for middle layer cells for the previously expected tapetum-specific role in the export of sporopollenin precursors.   6.2.4 Are hydroxycinnamoyl spermidines present in the sporopollenin biopolymer?   Data presented in Chapter 4 indicate the movement of hydroxycinnamoyl (HC) spermidines, known pollen coat constituents, from tapetal cells to the anther locule prior to tapetum programmed cell death in Arabidopsis. These data implicate HC spermidines as candidates for incorporation into the sporopollenin biopolymer, in addition to their loose attachment as components of the pollen coat. However, no evidence for the presence of HC   158 spermidines in sporopollenin was obtained, as wild-type and spermidine hydroxycinnamoyl transferase (sht) mutant pollen exhibited no significant difference in sporopollenin recalcitrance, as determined by acetolysis. Currently, acetolysis is the only method applied as an indicator of sporopollenin integrity and chemical recalcitrance. Although this method provides a valuable tool in sporopollenin analyses, it is limited in its chemical information, as sporopollenin walls that have been altered or are deficient in their formation are destroyed by acetolysis treatment. It is also unclear whether minor modifications to the sporopollenin biopolymer produce observable differences in acetolysis-sensitivities. Further, acetolysis specifically tests the chemical integrity of the sporopollenin, but does not provide information on the physical strength or rigidity of the biopolymer. The application of methods such as atomic force microscopy, to test the physical properties of sporopollenin walls, may enable more subtle alterations to the sporopollenin biopolymer resulting from gene mutations, to be identified. While Auramine O is commonly used as an exine stain, the constituents it binds in the wall have not been characterized. This stain may prove to be a useful tool in identifying subtle chemical alterations in the sporopollenin polymer that could be undetectable by acetolysis. Thus, future methods for the analysis of sporopollenin chemical and physical integrity, together with informative indicators for sporopollenin are needed.   6.2.5 How does sporopollenin traffic through the locule and assemble on the primexine of developing microspores?   While most cell layers of the anther wall have defined roles in pollen development and pollen wall formation, the function of the medium that fills the locule and is in direct contact with both the tapetal cells and developing pollen grains, is poorly understood. The locule fluid is of central importance to studies in pollen wall formation, serving as the apparent medium in which sporopollenin constituents traffic, and in which baculae and tecta assemble. From our analyses, the locule fluid appears abundant around microspore tetrads through to bicellular pollen grains, in parallel to the predicted time frame in which sporopollenin or its components may be in the locule. Although there is little to no information on the composition of locule fluid, it was interesting to discover its homogeneous appearance in sections of embedded anthers   159 viewed by light microscopy and transmission electron microscopy, in contrast to its lattice-like appearance in freeze-fractured anthers (as detailed in Chapter 2). The observation of structured locule fluid of anthers is likely due to the formation of ice crystals upon freezing; however, the ordered nature of these ice crystals may point to structural organization within the locule that has not been previously identified. If this interpretation is correct, the substructural organization within the locule in Arabidopsis or other flowering species with secretory tapeta could provide a mechanism to connect tapeta and microspores and could aid traffic between them. Future work on the composition of the locule, particularly on its proteinaceous content, including tapetum-secreted proteins such as LTPs (Huang et al., 2013b; 2009), holds tremendous potential to advance our understanding of the traffic and assembly of sporopollenin, as it provides the medium in which these processes occur.  The mechanism of sporopollenin assembly on the surface of microspores is unknown (Scott, 1994). One approach towards greater understanding of the mechanisms governing sporopollenin deposition and polymerization could entail the use of mutants affected in sporopollenin pattern formation, for example those identified by Dobritsa et al., (2011) in a comprehensive screen for mutants affected in exine production. In Chapter 2, Arabidopsis pollen wall ultrastructure over the stages of pollen formation was defined using cryo-fixation techniques, and revealed lamellae within the exine wall on early uninucleate microspores. The appearance of lamellae in exines has previously been reported and is hypothesized to indicate an early-assembling sporopollenin framework that transitions into a homogenous acetolysis-resistant form of polymerized sporopollenin (Rowley, 1962; Dickinson and Heslop-Harrison, 1968). In support of the compositional homology between sporopollenin and other plant biopolymers including cutin, suberin and lignin proposed previously (Schulze Osthoff and Wiermann, 1987; Scott, 1994), the lamella observed forming cutin, suberin and sporopollenin polymers suggest that a conserved mechanism for polymerization may exist (Bernards, 2002; Scott, 1994). Despite a limited understanding of cutin, suberin and lignin, knowledge surrounding the biosynthesis, transport and polymerization of these biopolymers will continue to provide analogous information for the study of sporopollenin.    160 References Aarts, M.G.M., Hodge, R., Kalantidis, K., Florack, D., Wilson, Z.A., Mulligan, B.J., Stiekema, W.J., Scott, R., and Pereira, A. (1997). The Arabidopsis MALE STERILITY 2 protein shares similarity with reductases in elongation/condensation complexes. The Plant Journal 12: 615–623. Abe, I., Watanabe, T., and Noguchi, H. (2004). Enzymatic formation of long-chain polyketide pyrones by plant type III polyketide synthases. Phytochemistry 65: 2447–2453. Ahlers, F., Bubert, H., Steuernagel, S., and Wiermann, R. (2000). The nature of oxygen in sporopollenin from the pollen of Typha angustifolia L. Z. Naturforsch., C, J. Biosci. 55: 129–136. Ahlers, F., Lambert, J., and Wiermann, R. (2003). Acetylation and silylation of piperidine solubilized sporopollenin from pollen of Typha angustifolia L. Z. Naturforsch., C, J. Biosci. 58: 807–811. Ahlers, F., Thom, I., Lambert, J., Kuckuk, R., and Rolf Wiermann (1999). 1H NMR analysis of sporopollenin from Typha angustifolia. Phytochemistry 50: 1095–1098. Akiyama, T., Shibuya, M., Liu, H.-M., and Ebizuka, Y. (1999). p-Coumaroyltriacetic acid synthase, a new homologue of chalcone synthase, from Hydrangea macrophylla var. thunbergii. Eur J Biochem 263: 834–839. Alonso, J.M. (2003). Genome-wide insertional mutagenesis of Arabidopsis thaliana. Science 301: 653–657. Ariizumi, T. and Toriyama, K. (2011). Genetic regulation of sporopollenin synthesis and pollen exine development. Annu Rev Plant Biol 62: 437–460. Ariizumi, T., Amagai, M., Shibata, D., Hatakeyama, K., Watanabe, M., and Toriyama, K. (2002). Comparative study of promoter activity of three anther-specific genes encoding lipid transfer protein, xyloglucan endotransglucosylase/hydrolase and polygalacturonase in transgenic Arabidopsis thaliana. Plant Cell Rep. 21: 90–96. Ariizumi, T., Hatakeyama, K., Hinata, K., Inatsugi, R., Nishida, I., Sato, S., Kato, T., Tabata, S., and Toriyama, K. (2004). Disruption of the novel plant protein NEF1 affects lipid accumulation in the plastids of the tapetum and exine formation of pollen, resulting in male sterility in Arabidopsis thaliana. The Plant Journal 39: 170–181. Ariizumi, T., Hatakeyama, K., Hinata, K., Sato, S., Kato, T., Tabata, S., and Toriyama, K. (2003). A novel male-sterile mutant of Arabidopsis thaliana, faceless pollen-1, produces pollen with a smooth surface and an acetolysis-sensitive exine. Plant Mol. Biol. 53: 107–116.   161 Ariizumi, T., Hatakeyama, K., Hinata, K., Sato, S., Kato, T., Tabata, S., and Toriyama, K. (2005). The HKM gene, which is identical to the MS1 gene of Arabidopsis thaliana, is essential for primexine formation and exine pattern formation. Sex. Plant Reprod. 18: 1–7. Ariizumi, T., Kawanabe, T., Hatakeyama, K., Sato, S., Kato, T., Tabata, S., and Toriyama, K. (2008). Ultrastructural characterization of exine development of the transient defective exine 1 mutant suggests the existence of a factor involved in constructing reticulate exine architecture from sporopollenin aggregates. Plant Cell Physiol. 49: 58–67. Austin, M.B. and Noel, J.P. (2003). The chalcone synthase superfamily of type III polyketide synthases. Nat. Prod. Rep. 20: 79–110. Bedinger, P. (1992). The remarkable biology of pollen. The Plant Cell 4: 879. Bernard, A. and Joubès, J. (2012). Arabidopsis cuticular waxes: Advances in synthesis, export and regulation. Prog. Lipid Res. 52: 110–129. Bernards, M.A. (2002). Demystifying suberin. Can. J. Bot. 80: 227–240. Bird, D.A. (2008). The role of ABC transporters in cuticular lipid secretion. Plant Science 174: 563–569. Bird, D.A., Beisson, F., Brigham, A., Shin, J., Greer, S., Jetter, R., Kunst, L., Wu, X., Yephremov, A., and Samuels, L. (2007). Characterization of Arabidopsis ABCG11/WBC11, an ATP binding cassette (ABC) transporter that is required for cuticular lipid secretion. The Plant Journal 52: 485–498. Blackmore, S., Wortley, A.H., Skvarla, J.J., and Rowley, J.R. (2007). Pollen wall development in flowering plants. New Phytol. 174: 483–498. Bolte, S., Talbot, C., Boutte, Y., Catrice, O., Read, N.D., and Satiat-Jeunemaitre, B. (2004). FM-dyes as experimental probes for dissecting vesicle trafficking in living plant cells. Journal of Microscopy 214: 159–173. Boutrot, F., Chantret, N., and Gautier, M.-F. (2008). Genome-wide analysis of the rice and Arabidopsis non-specific lipid transfer protein (nsLtp) gene families and identification of wheat nsLtp genes by EST data mining. BMC Genomics 9: 86. Brooks, J. and Shaw, G. (1968). Chemical structure of the exine of pollen walls and a new function for carotenoids in nature. Nature 219: 532–533. Bullock, G.R. (2011). The current status of fixation for electron microscopy: A review. Journal of Microscopy 133: 1–15. Buschmann, C., LANGSDORF, G., and Lichtenthaler, H.K. (2000). Imaging of the blue, green, and red fluorescence emission of plants: an overview. Photosynthetica 38: 483–491.   162 Chang, H.-S., Zhang, C., Chang, Y.-H., Zhu, J., Xu, X.-F., Shi, Z.-H., Zhang, X.-L., Xu, L., Huang, H., Zhang, S., and Yang, Z.-N. (2012). No primexine and plasma membrane undulation is essential for primexine deposition and plasma membrane undulation during microsporogenesis in Arabidopsis. Plant Physiol. 158: 264–272. Chebli, Y., Kaneda, M., Zerzour, R., and Geitmann, A. (2012). The cell wall of the Arabidopsis pollen tube—spatial distribution, recycling, and network formation of polysaccharides. Plant Physiol. 160: 1940–1955. Chen, W., Yu, X.H., Zhang, K., Shi, J., De Oliveira, S., Schreiber, L., Shanklin, J., and Zhang, D. (2011). MALE STERILE2 encodes a plastid-localized fatty acyl carrier protein reductase required for pollen exine development in Arabidopsis. Plant Physiol. 157: 842–853. Chen, X., Goodwin, S.M., Boroff, V.L., Liu, X., and Jenks, M.A. (2003). Cloning and characterization of the WAX2 gene of Arabidopsis involved in cuticle membrane and wax production. The Plant Cell 15: 1170–1185. Choi, H., Jin, J.-Y., Choi, S., Hwang, J.-U., Kim, Y.-Y., Suh, M.C., and Lee, Y. (2011). An ABCG/WBC-type ABC transporter is essential for transport of sporopollenin precursors for exine formation in developing pollen. The Plant Journal 65: 181–193. Colpitts, C.C., Kim, S.S., Posehn, S.E., Jepson, C., Kim, S.Y., Wiedemann, G., Reski, R., Wee, A.G.H., Douglas, C.J., and Suh, D.-Y. (2011). PpASCL, a moss ortholog of anther-specific chalcone synthase-like enzymes, is a hydroxyalkylpyrone synthase involved in an evolutionarily conserved sporopollenin biosynthesis pathway. New Phytol. 192: 855–868. Cook, D., Rimando, A.M., Clemente, T.E., Schröder, J., Dayan, F.E., Nanayakkara, N.P.D., Pan, Z., Noonan, B.P., Fishbein, M., Abe, I., Duke, S.O., and Baerson, S.R. (2010). Alkylresorcinol synthases expressed in Sorghum bicolor root hairs play an essential role in the biosynthesis of the allelopathic benzoquinone sorgoleone. The Plant Cell 22: 867–887. Cronk, Q. (2009). The molecular organography of plants (Oxford University Press: New York). de Azevedo Souza, C., Kim, S.S., Koch, S., Kienow, L., Schneider, K., McKim, S.M., Haughn, G.W., Kombrink, E., and Douglas, C.J. (2009). A novel fatty Acyl-CoA Synthetase is required for pollen development and sporopollenin biosynthesis in Arabidopsis. The Plant Cell 21: 507–525. DeBono, A., Yeats, T.H., Rose, J.K.C., Bird, D.A., Jetter, R., Kunst, L., and Samuels, L. (2009). Arabidopsis LTPG is a glycosylphosphatidylinositol-anchored lipid transfer protein required for export of lipids to the plant surface. The Plant Cell 21: 1230–1238. Dickinson, H. (1995). Dry stigmas, water and self-incompatibility in Brassica. Sex. Plant Reprod. 8: 1–10.   163 Dickinson, H.G. and Heslop-Harrison, J. (1968). Common mode of deposition for the sporopollenin of sexine and nexine. Nature 220: 926–927. Dickinson, H.G. and Sheldon, J.M. (1986). The generation of patterning at the plasma membrane of the young microspore of Lilium. Linnean Society symposium series: 1–17. Doan, T.T.P., Carlsson, A.S., Hamberg, M., Bülow, L., Stymne, S., and Olsson, P. (2009). Functional expression of five Arabidopsis fatty acyl-CoA reductase genes in Escherichia coli. Journal of Plant Physiology 166: 787–796. Dobritsa, A.A., Geanconteri, A., Shrestha, J., Carlson, A., Kooyers, N., Coerper, D., Urbanczyk-Wochniak, E., Bench, B.J., Sumner, L.W., Swanson, R., and Preuss, D. (2011). A large-scale genetic screen in Arabidopsis to identify genes involved in pollen exine production. Plant Physiol. 157: 947–970. Dobritsa, A.A., Lei, Z., Nishikawa, S.-I., Urbanczyk-Wochniak, E., Huhman, D.V., Preuss, D., and Sumner, L.W. (2010). LAP5 and LAP6 encode anther-specific proteins with similarity to chalcone synthase essential for pollen exine development in Arabidopsis. Plant Physiol. 153: 937–955. Dobritsa, A.A., Nishikawa, S.-I., Preuss, D., Urbanczyk-Wochniak, E., Sumner, L.W., Hammond, A., Carlson, A.L., and Swanson, R.J. (2009a). LAP3, a novel plant protein required for pollen development, is essential for proper exine formation. Sex. Plant Reprod. 22: 167–177. Dobritsa, A.A., Shrestha, J., Morant, M., Pinot, F., Matsuno, M., Swanson, R., Møller, B.L., and Preuss, D. (2009b). CYP704B1 is a long-chain fatty acid omega-hydroxylase essential for sporopollenin synthesis in pollen of Arabidopsis. Plant Physiol. 151: 574–589. Dong, X., Hong, Z., Sivaramakrishnan, M., Mahfouz, M., and Verma, D.P.S. (2005). Callose synthase (CalS5) is required for exine formation during microgametogenesis and for pollen viability in Arabidopsis. The Plant Journal 42: 315–328. Dou, X.-Y., Yang, K.-Z., Zhang, Y., Wang, W., Liu, X.-L., Chen, L.-Q., Zhang, X.-Q., and Ye, D. (2011). WBC27, an adenosine tri-phosphate-binding cassette protein, controls pollen wall formation and patterning in Arabidopsis. Journal of Integrative Plant Biology 53: 74–88. Doughty, J., Hedderson, F., McCubbin, A., and Dickinson, H. (1993). Interaction between a coating-borne peptide of the Brassica pollen grain and stigmatic S (self-incompatibility)-locus-specific glycoproteins. Proc. Natl. Acad. Sci. U.S.A. 90: 467–471. Douliez, J.-P., Michon, T., Elmorjani, K., and Marion, D. (2000). Mini Review: Structure, biological and technological functions of lipid transfer proteins and indolines, the major lipid binding proteins from cereal kernels. Journal of Cereal Science 32: 1–20.   164 Echlin, P. and Godwin, H. (1968). The ultrastructure and ontogeny of pollen in Helleborus foetidus L. J. Cell. Sci. 3: 161–174. Eckermann, S., Schröder, G., Schmidt, J., Strack, D., Edrada, R.A., Helariutta, Y., Elomaa, P., Kotilainen, M., Kilpeläinen, I., and Proksch, P. (1998). New pathway to polyketides in plants. Nature 396: 387–390. Edlund, A.F. (2004). Pollen and stigma structure and function: the role of diversity in pollination. The Plant Cell 16: S84–S97. Edstam, M.M., Viitanen, L., Salminen, T.A., and Edqvist, J. (2011). Evolutionary history of the non-specific lipid transfer proteins. Molecular Plant 4: 947–964. El-Ghazaly, G. and Jensen, W.A. (1986). Studies of the development of wheat (Triticum aestivum) pollen. Grana 25: 1–29. Elleman, C.J., FRANKLIN-TONG, V., and Dickinson, H.G. (1992). Pollination in species with dry stigmas: the nature of the early stigmatic response and the pathway taken by pollen tubes. New Phytol. 121: 413–424. Falasca, G., D'Angeli, S., Biasi, R., and Fattorini, L. (2013). Tapetum and middle layer control male fertility in Actinidia deliciosa. Annals of Botany 112: 1045–1055. Fiebig, A., Mayfield, J.A., Miley, N.L., Chau, S., Fischer, R.L., and Preuss, D. (2000). Alterations in CER6, a gene identical to CUT1, differentially affect long-chain lipid content on the surface of pollen and stems. The Plant Cell 12: 2001–2008. Fitzgerald, M.A. and Knox, R.B. (1995). Initiation of primexine in freeze-substituted microspores of Brassica campestris. Sex. Plant Reprod. 8: 99–104. Foster, G.D., Robinson, S.W., Blundell, R.P., Roberts, M.R., Hodge, R., Draper, J., and Scott, R.J. (1992). A Brassica napus mRNA encoding a protein homologous to phospholipid transfer proteins, is expressed specifically in the tapetum and developing microspores. Plant Science 84: 187–192. Frye, T.C. (1901). Development of the pollen in some Asclepiadaceae. Botanical Gazette: 325–331. Funa, N., Ozawa, H., Hirata, A., and Horinouchi, S. (2006). Phenolic lipid synthesis by type III polyketide synthases is essential for cyst formation in Azotobacter vinelandii. Proc. Natl. Acad. Sci. U.S.A. 103: 6356–6361. Gabarayeva, N.I. and Grigorjeva, V.V. (2013). Experimental modelling of exine-like structures. Grana 4: 251-257. Gagne, S.J., Stout, J.M., Liu, E., Boubakir, Z., Clark, S.M., and Page, J.E. (2012). Identification of olivetolic acid cyclase from Cannabis sativa reveals a unique catalytic route   165 to plant polyketides. Proceedings of the National Academy of Sciences 109: 12811–12816. Galati, B.G., Monacci, F., Gotelli, M.M., and Rosenfeldt, S. (2006). Pollen, tapetum and orbicule development in Modiolastrum malvifolium (Malvaceae). Annals of Botany 99: 755–763. Gilkey, J.C. and Staehelin, L.A. (1986). Advances in ultrarapid freezing for the preservation of cellular ultrastructure. J. Elec. Microsc. Tech. 3: 177–210. Goldberg, R.B., Beals, T.P., and Sanders, P.M. (1993). Anther development: basic principles and practical applications. The Plant Cell 5: 1217–1229. Goldberg, R.B., Sanders, P.M., and Beals, T.P. (1995). A novel cell-ablation strategy for studying plant development. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 350: 5–17. Goyal, A., Saxena, P., Rahman, A., Singh, P.K., Kasbekar, D.P., Gokhale, R.S., and Sankaranarayanan, R. (2008). Structural insights into biosynthesis of resorcinolic lipids by a type III polyketide synthase in Neurospora crassa. Journal of Structural Biology 162: 411–421. Graf, G.A. (2003). ABCG5 and ABCG8 Are obligate heterodimers for protein trafficking and biliary cholesterol excretion. Journal of Biological Chemistry 278: 48275–48282. Greenspan, P., Mayer, E.P., and Fowler, S.D. (1985). Nile red: a selective fluorescent stain for intracellular lipid droplets. J. Cell Biol. 100: 965–973. Grienenberger, E., Besseau, S., Geoffroy, P., Debayle, D., Heintz, D., Lapierre, C., Pollet, B., Heitz, T., and Legrand, M. (2009). A BAHD acyltransferase is expressed in the tapetum of Arabidopsis anthers and is involved in the synthesis of hydroxycinnamoyl spermidines. The Plant Journal 58: 246–259. Grienenberger, E., Kim, S.S., Lallemand, B., Geoffroy, P., Heintz, D., Souza, C. de A., Heitz, T., Douglas, C.J., and Legrand, M. (2010). Analysis of TETRAKETIDE α-PYRONE REDUCTASE function in Arabidopsis thaliana reveals a previously unknown, but conserved, biochemical pathway in sporopollenin monomer biosynthesis. The Plant Cell 22: 4067–4083. Guan, Y.-F., Huang, X.-Y., Zhu, J., Gao, J.-F., Zhang, H.-X., and Yang, Z.-N. (2008). RUPTURED POLLEN GRAIN1, a member of the MtN3/saliva gene family, is crucial for exine pattern formation and cell integrity of microspores in Arabidopsis. Plant Physiol. 147: 852–863. Gubatz, S., Arendt, S., Rittscher, M., and Wiermann, R. (1992). Recent aspects of sporopollenin biosynthesis. In Angiosperm Pollen and Ovules (Springer New York: New York, NY), pp. 187–190. Gubatz, S., Rittscher, M., Meuter, A., Nagler, A., and Wiermann, R. (1993). Tracer   166 experiments on sporopollenin biosynthesis. Grana 32: 12–17. Guilford, W.J., Schneider, D.M., Labovitz, J., and Opella, S.J. (1988). High resolution solid state 13C NMR spectroscopy of sporopollenins from different plant taxa. Plant Physiol. 86: 134–136. Hamberger, B., Ellis, M., Friedmann, M., Souza, C. de A., Barbazuk, B., and Douglas, C.J. (2007). Genome-wide analyses of phenylpropanoid-related genes in Populus trichocarpa, Arabidopsis thaliana, and Oryza sativa: the Populus lignin toolbox and conservation and diversification of angiosperm gene families. Botany 85: 1182–1201. Hemsley, A.R., Chaloner, W.G., Scott, A.C., and GROOMBRIDGE, C.J. (1992). Carbon-13 solid-state nuclear magnetic resonance of sporopollenins from modern and fossil plants. Annals of Botany 69: 545–549. Hemsley, A.R., Collinson, M.E., Kovach, W.L., Vincent, B., and Williams, T. (1994). The role of self-assembly in biological systems: evidence from iridescent colloidal sporopollenin in Selaginella megaspore walls. Philosophical Transactions of the Royal Society B: Biological Sciences 345: 163–173. Hemsley, A.R., Jenkins, P.D., E Collinson, M., and Vincent, B. (1996). Experimental modelling of exine self‐assembly. Botanical Journal of the Linnean Society 121: 177–187. Herminghaus, S., Arendt, S., Gubatz, S., Rittscher, M., and Wiermann, R. (1988). Aspects of sporopollenin biosynthesis: phenols as integrated compounds of the biopolymer. In Sexual Reproduction in Higher Plants, M. Cresti, ed (Springer Berlin Heidelberg: Berlin, Heidelberg), pp. 169–174. Heslop-Harrison, J. (1968a). Pollen wall development. The succession of events in the growth of intricately patterned pollen walls is described and discussed. Science 161: 230–237. Heslop-Harrison, J. (1968b). Tapetal origin of pollen‐coat substances in Lilium. New Phytol. 67: 779–786. Heslop-Harrison, J. (1968c). Wall development within the microspore tetrad of Lilium longiflorum. Can. J. Bot. 46: 1185–1192. Heslop-Harrison, J. and Mackenzie, A. (1967). Autoradiography of soluble [2-14C] thymidine derivatives during meiosis and microsporogenesis in Lilium anthers. Hess, M.W. (1993). Cell-wall development in freeze-fixed pollen: intine formation of Ledebouria socialis (Hyacinthaceae). Planta. Higgins, C.F. and Linton, K.J. (2001). Structural biology. The xyz of ABC transporters. Science 293: 1782–1784.   167 Hsieh, K. and Huang, A.H. (2005). Lipid‐rich tapetosomes in Brassica tapetum are composed of oleosin‐coated oil droplets and vesicles, both assembled in and then detached from the endoplasmic reticulum. The Plant Journal 43: 889–899. Hsieh, K. and Huang, A.H. (2007). Tapetosomes in Brassica tapetum accumulate endoplasmic reticulum-derived flavonoids and alkanes for delivery to the pollen surface. The Plant Cell 19: 582–596. Hsieh, K. and Huang, A.H.C. (2004). Endoplasmic reticulum, oleosins, and oils in seeds and tapetum cells. Plant Physiol. 136: 3427–3434. Hsieh, K., Wu, S.S., Ratnayake, C., and Huang, A.H. (2003). Tapetosomes and elaioplasts in Brassica and Arabidopsis floral tapetum. In Advanced Research on Plant Lipids (Springer: Netherlands), pp. 215–218. Huang, C.Y., Chen, P.-Y., Huang, M.-D., Tsou, C.-H., Jane, W.-N., and Huang, A.H.C. (2013a). Tandem oleosin genes in a cluster acquired in Brassicaceae created tapetosomes and conferred additive benefit of pollen vigor. Proc. Natl. Acad. Sci. U.S.A. 110: 14480–14485. Huang, M.-D., Chen, T.-L.L., and Huang, A.H.C. (2013b). Abundant type III lipid transfer proteins in Arabidopsis tapetum are secreted to the locule and become a constituent of the pollen exine. Plant Physiol. 163: 1218–1229. Huang, M.-D., Wei, F.-J., Wu, C.-C., Hsing, Y.-I.C., and Huang, A.H.C. (2009). Analyses of advanced rice anther transcriptomes reveal global tapetum secretory functions and potential proteins for lipid exine formation. Plant Physiol. 149: 694–707. Huang, M.D., Hsing, Y.I.C., and Huang, A.H.C. (2011). Transcriptomes of the anther sporophyte: availability and uses. Plant Cell Physiol. 52: 1459–1466. Hutzler, P., Fischbach, R., Heller, W., Jungblut, T.P., Reuber, S., Schmitz, R., Veit, M., Weissenböck, G., and Schnitzler, J.P. (1998). Tissue localization of phenolic compounds in plants by confocal laser scanning microscopy. J. Exp. Bot. 49: 953–965. Huysmans, S., El-Ghazaly, G., and Smets, E. (1998). Orbicules in angiosperms: morphology, function, distribution, and relation with tapetum types. Bot. Rev 64: 240–272. Huysmans, S., El-Ghazaly, G., Smets, E., Nordenstam, B., El-Ghazaly, G., and Kassas, M. (2000). Orbicules: still a well-hidden secret of the anther. In Plant systematics for the 21st century, B. Nordenstam, G. El-Ghazaly, and M. Kassas, eds (Portland Press: London), pp. 201–212. Huysmans, S., Verstraete, B., Smets, E., and Chatrou, L.W. (2010). Distribution of orbicules in Annonaceae mirrors evolutionary trend in angiosperms. Plecevo 143: 199–211.   168 Hülskamp, M., Kopczak, S.D., Horejsi, T.F., Kihl, B.K., and Pruitt, R.E. (1995). Identification of genes required for pollen-stigma recognition in Arabidopsis thaliana. The Plant Journal 8: 703–714. Ito, T. and Shinozaki, K. (2002). The MALE STERILITY1 gene of Arabidopsis, encoding a nuclear protein with a PHD-finger motif, is expressed in tapetal cells and is required for pollen maturation. Plant Cell Physiol. 43: 1285–1292. Ito, T., Nagata, N., Yoshiba, Y., Ohme-Takagi, M., Ma, H., and Shinozaki, K. (2007). Arabidopsis MALE STERILITY1 encodes a PHD-type transcription factor and regulates pollen and tapetum development. The Plant Cell 19: 3549–3562. Johnson, D.A. and Thomas, M.A. (2007). The monosaccharide transporter gene family in Arabidopsis and rice: a history of duplications, adaptive evolution, and functional divergence. Molecular Biology and Evolution 24: 2412–2423. Johnson, D.A., Hill, J.P., and Thomas, M.A. (2006). The monosaccharide transporter gene family in land plants is ancient and shows differential subfamily expression and expansion across lineages. BMC Evolutionary Biology 6: 64. Johnson-Brousseau, S.A. and McCormick, S. (2004). A compendium of methods useful for characterizing Arabidopsispollen mutants and gametophytically- expressed genes. The Plant Journal 39: 761–775. José-Estanyol, M., Gomis-Rüth, F.X., and Puigdomènech, P. (2004). The eight-cysteine motif, a versatile structure in plant proteins. Plant Physiology and Biochemistry 42: 355–365. Kader, J.-C. (1996). Lipid-transfer proteins in plants. Annu. Rev. Plant. Physiol. Plant. Mol. Biol. 47: 627–654. Kawase, M. and Takahashi, M. (1995). Chemical composition of sporopollenin in Magnolia grandiflora (Magnoliaceae) and Hibiscus syriacus (Malvaceae). Grana 34: 242–245. Keijzer, C.J. (1987). The processes of anther dehiscence and pollen dispersal. I. The opening mechanism of longitudinally dehiscing anthers. New Phytol. 105: 487–498. Kenrick, P. and Crane, P.R. (1997). The origin and early evolution of plants on land. Nature 389: 33–39. Kienow, L., Schneider, K., Bartsch, M., Stuible, H.P., Weng, H., Miersch, O., Wasternack, C., and Kombrink, E. (2008). Jasmonates meet fatty acids: functional analysis of a new acyl-coenzyme A synthetase family from Arabidopsis thaliana. J. Exp. Bot. 59: 403–419. Kim, H., Lee, S.B., Kim, H.J., Min, M.K., Hwang, I., and Suh, M.C. (2012). Characterization of glycosylphosphatidylinositol-anchored lipid transfer protein 2 (LTPG2) and overlapping function between LTPG/LTPG1 and LTPG2 in cuticular wax export or accumulation in   169 Arabidopsis thaliana. Plant Cell Physiol. 53: 1391–1403. Kim, H.U. (2002). A novel group of oleosins is present inside the pollen of Arabidopsis. Journal of Biological Chemistry 277: 22677–22684. Kim, H.U. and Huang, A.H. (2003). Oleosins and plastid-lipid-associated proteins in Arabidopsis. In Advanced Research on Plant Lipids, N. Murata, ed (Springer), pp. 147–150. Kim, H.U., Wu, S.S., Ratnayake, C., and Huang, A.H. (2001). Brassica rapa has three genes that encode proteins associated with different neutral lipids in plastids of specific tissues. Plant Physiol. 126: 330–341. Kim, S.S., Grienenberger, E., Lallemand, B., Colpitts, C.C., Kim, S.Y., Souza, C. de A., Geoffroy, P., Heintz, D., Krahn, D., Kaiser, M., Kombrink, E., Heitz, T., Suh, D.-Y., Legrand, M., and Douglas, C.J. (2010). LAP6/POLYKETIDE SYNTHASE A and LAP5/POLYKETIDE SYNTHASE B encode hydroxyalkyl α-pyrone synthases required for pollen development and sporopollenin biosynthesis in Arabidopsis thaliana. The Plant Cell 22: 4045–4066. Kirkpatrick, A.B. and Owen, H.A. (2013). Observation of early pollen exine patterning by scanning electron microscopy. Microscopy and Microanalysis 19: 134–135. Kiss, J.Z. and Staehelin, L.A. (1995). High pressure freezing. In Rapid freezing, freeze fracture and deep etching (Wiley-Liss: New York), pp. 89–104. Knox, R.B. and Heslop-Harrison, J. (1970). Pollen-wall proteins: localization and enzymic activity. J. Cell. Sci. 6: 1–27. Knox, R.B. and Heslop-Harrison, J. (1971). Pollen-wall proteins: the fate of intine-held antigens on the stigma in compatible and incompatible pollinations of Phalaris tuberosa L. J. Cell. Sci. 9: 239–251. Koduri, P.K.H., Gordon, G.S., Barker, E.I., Colpitts, C.C., Ashton, N.W., and Suh, D.-Y. (2010). Genome-wide analysis of the chalcone synthase superfamily genes of Physcomitrella patens. Plant Mol. Biol. 72: 247–263. Koltunow, A.M., Truettner, J., Cox, K.H., Wallroth, M., and Goldberg, R.B. (1990). Different temporal and spatial gene expression patterns occur during anther development. The Plant Cell 2: 1201–1224. Koornneef, M., Hanhart, C.J., and Thiel, F. (1989). A genetic and phenotypic description of eceriferum (cer) mutants in Arabidopsis thaliana. Journal of Heredity 80: 118–122. Kurata, T., Kawabata Awai, C., Sakuradani, E., Shimizu, S., Okada, K., and Wada, T. (2003). The YORE‐YORE gene regulates multiple aspects of epidermal cell differentiation in Arabidopsis. The Plant Journal 36: 55–66.   170 Kuromori, T., Ito, T., Sugimoto, E., and Shinozaki, K. (2011a). Arabidopsis mutant of AtABCG26, an ABC transporter gene, is defective in pollen maturation. Journal of Plant Physiology 168: 2001–2005. Kuromori, T., Miyaji, T., Yabuuchi, H., Shimizu, H., Sugimoto, E., Kamiya, A., Moriyama, Y., and Shinozaki, K. (2010). ABC transporter AtABCG25 is involved in abscisic acid transport and responses. Proceedings of the National Academy of Sciences 107: 2361–2366. Kuromori, T., Sugimoto, E., and Shinozaki, K. (2011b). Arabidopsis mutants of AtABCG22, an ABC transporter gene, increase water transpiration and drought susceptibility. The Plant Journal 67: 885–894. Lallemand, B., Erhardt, M., Heitz, T., and Legrand, M. (2013). Sporopollenin biosynthetic enzymes interact and constitute a metabolon localized to the endoplasmic reticulum of tapetum cells. Plant Physiol. 162: 616–625. Lantis, V. (1912). Development of the microsporangia and microspores of Abutilon theophrasti. Botanical Gazette: 330–335. Lauga, B., Charbonnel-Campaa, L., and Combes, D. (2000). Characterization of MZm3-3, a Zea mays tapetum-specific transcript. Plant Sci. 157: 65–75. Le Hir, R., Sorin, C., Chakraborti, D., Moritz, T., Schaller, H., Tellier, F., Robert, S., Morin, H., Bako, L., and Bellini, C. (2013). ABCG9, ABCG11 and ABCG14 ABC transporters are required for vascular development in Arabidopsis. The Plant Journal 76: 811–824. Li, N., Zhang, D.-S., Liu, H.-S., Yin, C.-S., Li, X.-X., Liang, W.-Q., Yuan, Z., Xu, B., Chu, H.-W., Wang, J., Wen, T.-Q., Huang, H., Luo, D., Ma, H., and Zhang, D.-B. (2006). The rice tapetum degeneration retardation gene is required for tapetum degradation and anther development. The Plant Cell 18: 2999–3014. Lichtenthaler, H.K., Stober, F., and Lang, M. (1992). The nature of the different laser-induced fluorescence signatures of plants. EARSeL Adv. Remote Sens 1: 20–30. Liu, L. and Fan, X.-D. (2013). Tapetum: regulation and role in sporopollenin biosynthesis in Arabidopsis. Plant Mol. Biol. 83: 165–175. Luo, B., Xue, X.-Y., Hu, W.-L., Wang, L.-J., and Chen, X.-Y. (2007). An ABC transporter gene of Arabidopsis thaliana, AtWBC11, is involved in cuticle development and prevention of rgan fusion. Plant Cell Physiol. 48: 1790–1802. Ma, D.-P., Tan, H., Si, Y., Creech, R.G., and Jenkins, J.N. (1995). Differential expression of a lipid transfer protein gene in cotton fiber. Biochimica et Biophysica Acta (BBA) - Lipids and Lipid Metabolism 1257: 81–84. Mariani, C., Beuckeleer, M.D., Truettner, J., Leemans, J., and Goldberg, R.B. (1990).   171 Induction of male sterility in plants by a chimaeric ribonuclease gene. Nature 347: 737–741. Mayfield, J.A. (2001). Gene families from the Arabidopsis thaliana pollen coat proteome. Science 292: 2482–2485. Mayfield, J.A. and Preuss, D. (2000). Rapid initiation of Arabidopsis pollination requires the oleosin-domain protein GRP17. Nat Cell Biol 2: 128–130. McCormick, S. (1993). Male gametophyte development. The Plant Cell 5: 1265. McFarlane, H.E., Shin, J.J.H., Bird, D.A., and Samuels, A.L. (2010). Arabidopsis ABCG transporters, which are required for export of diverse cuticular lipids, dimerize in different combinations. The Plant Cell 22: 3066–3075. McFarlane, H.E., Young, R.E., Wasteneys, G.O., and Samuels, A.L. (2008). Cortical microtubules mark the mucilage secretion domain of the plasma membrane in Arabidopsis seed coat cells. Planta 227: 1363–1375. McNeil, K.J. and Smith, A.G. (2009). A glycine-rich protein that facilitates exine formation during tomato pollen development. Planta 231: 793–808. Mentewab, A. and Stewart, C.N. (2005). Overexpression of an Arabidopsis thaliana ABC transporter confers kanamycin resistance to transgenic plants. Nat Biotechnol 23: 1177–1180. Millar, A.A., Clemens, S., Zachgo, S., Giblin, E.M., Taylor, D.C., and Kunst, L. (1999). CUT1, an Arabidopsis gene required for cuticular wax biosynthesis and pollen fertility, encodes a very-long-chain fatty acid condensing enzyme. The Plant Cell 11: 825–838. Mitsuda, N., Seki, M., Shinozaki, K., and Ohme-Takagi, M. (2005). The NAC transcription factors NST1 and NST2 of Arabidopsis regulate secondary wall thickenings and are required for anther dehiscence. The Plant Cell 17: 2993–3006. Mizuuchi, Y., Shimokawa, Y., Wanibuchi, K., Noguchi, H., and Abe, I. (2008). Structure function analysis of novel type III polyketide synthases from Arabidopsis thaliana. Biol. Pharm. Bull. 31: 2205–2210. Moco, S., Schneider, B., and Vervoort, J. (2009). Plant micrometabolomics: the analysis of endogenous metabolites present in a plant cell or tissue. J. Proteome Res. 8: 1694–1703. Moore, S.E.M., Hemsley, A.R., French, A.N., Dudley, E., and Newton, R.P. (2006). New insights from MALDI-ToF MS, NMR, and GC-MS: mass spectrometry techniques applied to palynology. Protoplasma 228: 151–157. Morant, M., Jørgensen, K., Schaller, H., Pinot, F., Møller, B.L., Werck-Reichhart, D., and Bak, S. (2007). CYP703 is an ancient cytochrome P450 in land plants catalyzing in-chain hydroxylation of lauric acid to provide building blocks for sporopollenin synthesis in pollen.   172 The Plant Cell 19: 1473–1487. Morgan, T.E. and Huber, G.L. (1967). Loss of lipid during fixation for electron microscopy. J. Cell Biol. 32: 757–760. Murphy, D.J. (2006). The extracellular pollen coat in members of the Brassicaceae: composition, biosynthesis, and functions in pollination. Protoplasma 228: 31–39. Murphy, D.J. and Ross, J. (1998). Biosynthesis, targeting and processing of oleosin‐like proteins, which are major pollen coat components in Brassica napus. The Plant Journal 13: 1–16. Nishikawa, S.-I., Zinkl, G.M., Swanson, R.J., Maruyama, D., and Preuss, D. (2005). Callose (β-1,3 glucan) is essential for Arabidopsis pollen wall patterning, but not tube growth. BMC Plant Biol. 5: 1–9. Niu, B.-X., He, F.-R., He, M., Ren, D., Chen, L.-T., and Liu, Y.-G. (2013). The ATP-binding cassette transporter OsABCG15 is required for anther development and pollen fertility in rice. Journal of Integrative Plant Biology 55: 710–720. Owen, H.A. and Makaroff, C.A. (1995). Ultrastructure of microsporogenesis and microgametogenesis in Arabidopsis thaliana (L.) Heynh. ecotype Wassilewskija (Brassicaceae). Protoplasma 185: 7–21. Pacini, E. (2010). Relationships between tapetum, loculus, and pollen during development. Int. J Plant Sci. 171: 1–11. Pacini, E. and Hesse, M. (2005). Pollenkitt–its composition, forms and functions. Flora - Morphology, Distribution, Functional Ecology of Plants 200: 399–415. Palade, G.E. (1952). A study of fixation for electron microscopy. Journal of Experimental Medicine 95: 285–298. Panikashvili, D., Savaldi-Goldstein, S., Mandel, T., Yifhar, T., Franke, R.B., Höfer, R., Schreiber, L., Chory, J., and Aharoni, A. (2007). The Arabidopsis DESPERADO/AtWBC11 transporter is required for cutin and wax secretion. Plant Physiol. 145: 1345–1360. Panikashvili, D., Shi, J.X., Bocobza, S., Franke, R.B., Schreiber, L., and Aharoni, A. (2010). The Arabidopsis DSO/ABCG11 transporter affects cutin metabolism in reproductive organs and suberin in roots. Molecular Plant 3: 563–575. Panikashvili, D., Shi, J.X., Schreiber, L., and Aharoni, A. (2011). The Arabidopsis ABCG13 transporter is required for flower cuticle secretion and patterning of the petal epidermis. New Phytologist 190: 113–124. Parish, R.W. and Li, S.F. (2010). Death of a tapetum: A programme of developmental altruism.   173 Plant Science 178: 73–89. Paxson-Sowders, D.M., Dodrill, C.H., Owen, H.A., and Makaroff, C.A. (2001). DEX1, a novel plant protein, is required for exine pattern formation during pollen development in Arabidopsis. Plant Physiol. 127: 1739–1749. Paxson-Sowders, D.M., Owen, H.A., and Makaroff, C.A. (1997). A comparative ultrastructural analysis of exine pattern development in wild-type Arabidopsis and a mutant defective in pattern formation. Protoplasma 198: 53–65. Persson, S., Paredez, A., Carroll, A., Palsdottir, H., Doblin, M., Poindexter, P., Khitrov, N., Auer, M., and Somerville, C.R. (2007). Genetic evidence for three unique components in primary cell-wall cellulose synthase complexes in Arabidopsis. Proc. Natl. Acad. Sci. U.S.A. 104: 15566–15571. Phan, H.A., Li, S.F., and Parish, R.W. (2011). MYB80, a regulator of tapetal and pollen development, is functionally conserved in crops. Plant Mol. Biol. 78: 171–183. Piffanelli, P. and Murphy, D.J. (1998). Novel organelles and targeting mechanisms in the anther tapetum. Trends in Plant Science 3: 1–4. Piffanelli, P., Ross, J.H., and Murphy, D.J. (1998). Biogenesis and function of the lipidic structures of pollen grains. Sex. Plant Reprod. 11: 65–80. Pighin, J.A., Zheng, H., Balakshin, L.J., Goodman, I.P., Western, T.L., Jetter, R., Kunst, L., and Samuels, A.L. (2004). Plant cuticular lipid export requires an ABC transporter. Science 306: 702–704. Platt, K.A., Huang, A.H., and Thomson, W.W. (1998). Ultrastructural study of lipid accumulation in tapetal cells of Brassica napus L. cv. Westar during microsporogenesis. Int. J Plant Sci.: 724–737. Prahl, A.K., Rittscher, M., and Wiermann, R. (1986). New aspects of sporopollenin biosynthesis. In Biotechnology and Ecology of Pollen, D.L. Mulcahy, ed (Springer-Verlag: New York), pp. 313–318. Prahl, A.K., Springstubbe, H., Grumbach, K., and Wiermann, R. (1985). Studies on sporopollenin biosynthesis: the effect of inhibitors of carotenoid biosynthesis on sporopollenin accumulation. Zeitschrift fur Naturforschung. Section C, Biosciences 40: 621–626. Preuss, D., Lemieux, B., Yen, G., and Davis, R.W. (1993). A conditional sterile mutation eliminates surface components from Arabidopsis pollen and disrupts cell signaling during fertilization. Genes & Development 7: 974–985. Qin, P., Tu, B., Wang, Y., Deng, L., Quilichini, T.D., Li, T., Wang, H., Ma, B., and Li, S. (2013). ABCG15 encodes an ABC transporter protein, and is essential for post-meiotic   174 anther and pollen exine development in rice. Plant Cell Physiol. 54: 138–154. Quilichini, T.D., Friedmann, M.C., Samuels, A.L., and Douglas, C.J. (2010). ATP-binding cassette transporter G26 is required for male fertility and pollen exine formation in Arabidopsis. Plant Physiol. 154: 678–690. Rea, P.A. (2007). Plant ATP-binding cassette transporters. Annu Rev Plant Biol 58: 347–375. Ríos, G., Tadeo, F.R., Leida, C., and Badenes, M.L. (2013). Prediction of components of the sporopollenin synthesis pathway in peach by genomic and expression analyses. BMC Genomics 14: 40. Ross, J. and Murphy, D.J. (1996). Characterization of anther‐expressed genes encoding a major class of extracellular oleosin‐like proteins in the pollen coat of Brassicaceae. The Plant Journal 9: 625–637. Rowland, O., Lee, R., Franke, R., Schreiber, L., and Kunst, L. (2007). The CER3 wax biosynthetic gene from Arabidopsis thaliana is allelic to WAX2/YRE/FLP1. FEBS Letters 581: 3538–3544. Rowley, J.R. (1962). Stranded arrangement of sporopollenin in the exine of microspores of Poa annua. Science 137: 526–528. Rowley, J.R. and Morbelli, M.A. (2009). Connective structures between tapetal cells and spores in Lycophyta and pollen grains in angiosperms — A review. Review of Palaeobotany and Palynology 156: 157–164. Rozema, J., Broekman, R.A., Blokker, P., Meijkamp, B.B., de Bakker, N., van de Staaij, J., van Beem, A., Ariese, F., and Kars, S.M. (2001). UV-B absorbance and UV-B absorbing compounds (para-coumaric acid) in pollen and sporopollenin: the perspective to track historic UV-B levels. Journal of Photochemistry and Photobiology B: Biology 62: 108–117. Rubinelli, P., Hu, Y., and Ma, H. (1998). Identification, sequence analysis and expression studies of novel anther-specific genes of Arabidopsis thaliana. Plant Mol. Biol. 37: 607–619. Sanchez-Fernandez, R. (2001). The Arabidopsis thaliana ABC protein superfamily, a complete inventory. Journal of Biological Chemistry 276: 30231–30244. Sanchez-Fernandez, R., Rea, P.A., Davies, T.G., and Coleman, J.O. (2001). Do plants have more genes than humans? Yes, when it comes to ABC proteins. Trends in Plant Science 6: 347–348. Sanders, P.M., Bui, A.Q., Weterings, K., McIntire, K.N., Hsu, Y.C., Lee, P.Y., Truong, M.T., Beals, T.P., and Goldberg, R.B. (1999). Anther developmental defects in Arabidopsis thaliana male-sterile mutants. Sex. Plant Reprod. 11: 297–322. Schilmiller, A.L., Stout, J., Weng, J.-K., Humphreys, J., Ruegger, M.O., and Chapple, C.   175 (2009). Mutations in the cinnamate 4-hydroxylase gene impact metabolism, growth and development in Arabidopsis. The Plant Journal 60: 771–782. Schulze Osthoff, K. and Wiermann, R. (1987). Phenols as integrated compounds of sporopollenin from Pinus pollen. Journal of Plant Physiology 131: 5–15. Scott, R.J. (1994). Pollen exine: The sporopollenin enigma and the physics of pattern. In Molecular and Cellular Aspects of Plant Reproduction (Cambridge University Press: Cambridge, UK), pp. 49–81. Scott, R.J., Spielman, M., and Dickinson, H.G. (2004). Stamen structure and function. The Plant Cell 16 Suppl: S46–60. Sheldon, J.M. and Dickinson, H.G. (1983). Determination of patterning in the pollen wall of Lilium henryi. J. Cell. Sci. 63: 191–208. Sossountzov, L., Ruiz-Avila, L., Vignols, F., Jolliot, A., Arondel, V., Tchang, F., Grosbois, M., Guerbette, F., Miginiac, E., and Delseny, M. (1991). Spatial and temporal expression of a maize lipid transfer protein gene. The Plant Cell 3: 923–933. Southworth, D. (1974). Solubility of pollen exines. Am. J. Bot.: 36–44. Spurr, A.R. (1969). A low-viscosity epoxy resin embedding medium for electron microscopy. Journal of Ultrastructure Research 26: 31–43. Sterk, P., Booij, H., Schellekens, G.A., Van Kammen, A., and De Vries, S.C. (1991). Cell-specific expression of the carrot EP2 lipid transfer protein gene. The Plant Cell 3: 907–921. Stieglitz, H. (1977). Role of β-1,3-glucanase in postmeiotic microspore release. Developmental Biology 57: 87–97. Suarez-Cervera, M., Marquez, J., and Seoane-Camba, J. (1995). Pollen grain and Ubisch body development in Platanus acerifolia. Review of Palaeobotany and Palynology 85: 63–84. Suh, M.C. (2005). Cuticular lipid composition, surface structure, and gene expression in Arabidopsis stem epidermis. Plant Physiol. 139: 1649–1665. Sun, M.-X., Huang, X.-Y., Yang, J., Guan, Y.-F., and Yang, Z.-N. (2013). Arabidopsis RPG1 is important for primexine deposition and functions redundantly with RPG2 for plant fertility at the late reproductive stage. Plant Reprod 26: 83–91. Suzuki, T., Masaoka, K., Nishi, M., Nakamura, K., and Ishiguro, S. (2008). Identification of kaonashi mutants showing abnormal pollen exine structure in Arabidopsis thaliana. Plant Cell Physiol. 49: 1465–1477. Svatoš, A. (2011). Single-cell metabolomics comes of age: new developments in mass   176 spectrometry profiling and imaging. Analytical chemistry 83: 5037–5044. Tang, L.K., Chu, H., Yip, W.K., Yeung, E.C., and Lo, C. (2009). An anther-specific dihydroflavonol 4-reductase-like gene (DRL1) is essential for male fertility in Arabidopsis. New Phytol. 181: 576–587. Taylor, J.H. (1959). Autoradiographic studies of nucleic acids and proteins during meiosis in Lilium longiflorum. Am. J. Bot. 46: 477–484. Taylor, J.S. and Raes, J. (2004). Duplication and divergence: the evolution of new genes and old ideas. Annu. Rev. Genet. 38: 615–643. Taylor-Teeples, M., Ron, M., and Brady, S.M. (2011). Novel biological insights revealed from cell type-specific expression profiling. Curr. Opin. Plant Biol. 14: 601–607. Theodoulou, F.L. (2000). Plant ABC transporters. Biochimica et Biophysica Acta (BBA)-Biomembranes 1465: 79–103. Thoma, S., Hecht, U., Kippers, A., Botella, J., De Vries, S., and Somerville, C. (1994). Tissue-specific expression of a gene encoding a cell wall-localized lipid transfer protein from Arabidopsis. Plant Physiol. 105: 35–45. Ting, J.T., Wu, S.S., Ratnayake, C., and Huang, A.H. (1998). Constituents of the tapetosomes and elaioplasts in Brassica campestris tapetum and their degradation and retention during microsporogenesis. The Plant Journal 16: 541–551. Tropf, S., Lanz, T., Rensing, S.A., Schr der, J., and Schr der, G. (1994). Evidence that stilbene synthases have developed from chalcone synthases several times in the course of evolution. J Mol Evol 38: 610–618. Ubisch, G. (1927). Zur Entwicklungsgeschichte der Antheren. Planta 3: 490–495. Ukitsu, H., Kuromori, T., Toyooka, K., Goto, Y., Matsuoka, K., Sakuradani, E., Shimizu, S., Kamiya, A., Imura, Y., Yuguchi, M., Wada, T., Hirayama, T., and Shinozaki, K. (2007). Cytological and biochemical analysis of COF1, an Arabidopsis mutant of an ABC transporter gene. Plant Cell Physiol. 48: 1524–1533. Van Bergen, P.F., Blokker, P., Collinson, M.E., Damsté, J.S.S., and De Leeuw, J.W. (2004). Structural biomacromolecules in plants: what can be learnt from the fossil record. In Evolution of Plant Physiology (Elsevier: Amsterdam), pp. 133–154. Van Bergen, P.F., Collinson, M.E., Briggs, D., De Leeuw, J.W., Scott, A.C., Evershed, R.P., and Finch, P. (1995). Resistant biomacromolecules in the fossil record. Acta Bot. Neerl 44: 319–342. van Meer, G., Halter, D., Sprong, H., Somerharju, P., and Egmond, M.R. (2006). ABC lipid transporters: extruders, flippases, or flopless activators? FEBS Letters 580: 1171–1177.   177 Velamakanni, S., Wei, S.L., Janvilisri, T., and Veen, H.W. (2007). ABCG transporters: structure, substrate specificities and physiological roles. J Bioenerg Biomembr 39: 465–471. Verrier, P.J., Bird, D.A., Burla, B., Dassa, E., Forestier, C., Geisler, M., Klein, M., Kolukisaoglu, Ü., Lee, Y., Martinoia, E., Murphy, A., Rea, P.A., Samuels, L., Schulz, B., Spalding, E.P., Yazaki, K., and Theodoulou, F.L. (2008). Plant ABC proteins – a unified nomenclature and updated inventory. Trends in Plant Science 13: 151–159. Vinckier, S., Cadot, P., and Smets, E. (2005). The manifold characters of orbicules: structural diversity, systematic significance, and vectors for allergens. Grana 44: 300–307. Wallace, S., Fleming, A., Wellman, C.H., and Beerling, D.J. (2011). Evolutionary development of the plant and spore wall. AoB Plants 2011: plr027. Wang, A., Xia, Q., Xie, W., Datla, R., and Selvaraj, G. (2003). The classical Ubisch bodies carry a sporophytically produced structural protein (RAFTIN) that is essential for pollen development. Proc. Natl. Acad. Sci. U.S.A. 100: 14487–14492. Wehling, K., Niester, C., Boon, J.J., Willemse, M., and Wiermann, R. (1989). p-Coumaric acid—a monomer in the sporopollenin skeleton. Planta 179: 376–380. Weng, J.-K., Li, Y., Mo, H., and Chapple, C. (2012). Assembly of an evolutionarily new pathway for α-pyrone biosynthesis in Arabidopsis. Science 337: 960–964. Weng, J.-K., Mo, H., and Chapple, C. (2010). Over-expression of F5H in COMT-deficient Arabidopsis leads to enrichment of an unusual lignin and disruption of pollen wall formation. The Plant Journal 64: 898–911. Wiermann, R., Ahlers, F., and Schmitz-Thom, I. (2005). Sporopollenin. In Biopolymers, A. Steinbüchel and M. Hofrichter, eds (Wiley-VCH Verlag: Weinheim, Germany), pp. 209–227. Wilmesmeier, S. and Wiermann, R. (1995). Influence of EPTC (S-ethyl-dipropyl-thiocarbamate) on the composition of surface waxes and sporopollenin structure in Zea mays. Journal of Plant Physiology 146: 22–28. Wilson, Z.A. and Zhang, D.-B. (2009). From Arabidopsis to rice: pathways in pollen development. J. Exp. Bot. 60: 1479–1492. Wilson, Z.A., Morroll, S.M., Dawson, J., Swarup, R., and Tighe, P.J. (2001). The Arabidopsis MALE STERILITY1 (MS1) gene is a transcriptional regulator of male gametogenesis, with homology to the PHD-finger family of transcription factors. The Plant Journal 28: 27–39. Winter, D., Vinegar, B., Nahal, H., Ammar, R., Wilson, G.V., and Provart, N.J. (2007). An “Electronic Fluorescent Pictograph” browser for exploring and analyzing large-scale biological data sets. PLoS ONE 2: e718.   178 Witkus, E.R. (1945). Endomitotic tapetal cell divisions in Spinacia. Am. J. Bot. 32: 326–330. Worrall, D., Hird, D.L., Hodge, R., Paul, W., Draper, J., and Scott, R. (1992). Premature dissolution of the microsporocyte callose wall causes male sterility in transgenic tobacco. The Plant Cell 4: 759–771. Wu, S.S., Platt, K.A., Ratnayake, C., Wang, T.W., Ting, J.T., and Huang, A.H. (1997). Isolation and characterization of neutral-lipid-containing organelles and globuli-filled plastids from Brassica napus tapetum. Proc. Natl. Acad. Sci. U.S.A. 94: 12711–12716. Wu, S.S.H., Moreau, R.A., Whitaker, B.D., and Huang, A.H.C. (1999). Steryl esters in the elaioplasts of the tapetum in developing Brassica anthers and their recovery on the pollen surface. Lipids 34: 517–523. Xu, J., Yang, C., Yuan, Z., Zhang, D., Gondwe, M.Y., Ding, Z., Liang, W., Zhang, D., and Wilson, Z.A. (2010). The ABORTED MICROSPORES regulatory network is required for postmeiotic male reproductive development in Arabidopsis thaliana. The Plant Cell 22: 91–107. Yamaguchi, T., Kurosaki, F., Suh, D.Y., Sankawa, U., Nishioka, M., Akiyama, T., Shibuya, M., and Ebizuka, Y. (1999). Cross-reaction of chalcone synthase and stilbene synthase overexpressed in Escherichia coli. FEBS Letters 460: 457–461. Yang, C., Vizcay-Barrena, G., Conner, K., and Wilson, Z.A. (2007). MALE STERILITY1 is required for tapetal development and pollen wall biosynthesis. The Plant Cell 19: 3530–3548. Yang, H. and Murphy, A.S. (2009). Functional expression and characterization of Arabidopsis ABCB, AUX 1 and PIN auxin transporters in Schizosaccharomyces pombe. The Plant Journal 59: 179–191. Yazaki, K. (2006). ABC transporters involved in the transport of plant secondary metabolites. FEBS Letters 580: 1183–1191. Zhang, D., Liang, W., Yin, C., Zong, J., Gu, F., and Zhang, D. (2010). OsC6, encoding a lipid transfer protein, is required for postmeiotic anther development In rice. Plant Physiol. 154: 149–162. Zhang, D.-S., Liang, W.-Q., Yuan, Z., Li, N., Shi, J., Wang, J., Liu, Y.-M., Yu, W.-J., and Zhang, D.-B. (2008). Tapetum degeneration retardation is critical for aliphatic metabolism and gene regulation during rice pollen development. Molecular Plant 1: 599–610. Zhang, Z.-B., Zhu, J., Gao, J.-F., Wang, C., Li, H., Li, H., Zhang, H.-Q., Zhang, S., Wang, D.-M., Wang, Q.-X., Huang, H., Xia, H.-J., and Yang, Z.-N. (2007). Transcription factor AtMYB103 is required for anther development by regulating tapetum development, callose dissolution and exine formation in Arabidopsis. The Plant Journal 52: 528–538.   179 Zimmermann, P., Hennig, L., and Gruissem, W. (2005). Gene-expression analysis and network discovery using Genevestigator. Trends in Plant Science 10: 407–409. Zinkl, G.M., Zwiebel, B.I., Grier, D.G., and Preuss, D. (1999). Pollen-stigma adhesion in Arabidopsis: a species-specific interaction mediated by lipophilic molecules in the pollen exine. Development 126: 5431–5440. Zipfel, W.R., Williams, R.M., and Webb, W.W. (2003). Nonlinear magic: multiphoton microscopy in the biosciences. Nat Biotechnol 21: 1369–1377. 


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