Crassulacean Acid Metabolism in Tropical Orchids: Integrating Phylogenetic, Ecophysiological and Molecular Genetic Approaches

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University of Nevada, Reno

Crassulacean acid metabolism in tropical orchids: integrating phylogenetic, ecophysiological and molecular genetic approaches

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biochemistry and Molecular Biology

Katia I. Silvera

Dr. John C. Cushman/ Dissertation Advisor

May 2010

THE GRADUATE SCHOOL

We recommend that the dissertation prepared under our supervision by

KATIA I. SILVERA

entitled

Crassulacean Acid Metabolism In Tropical Orchids: Integrating Phylogenetic, Ecophysiological And Molecular Genetic Approaches

be accepted in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

John C. Cushman, Ph.D., Advisor

Jeffrey F. Harper, Ph.D., Committee Member

Robert S. Nowak, Ph.D., Committee Member

David K.Shintani, Ph.D., Committee Member

David W. Zeh, Ph.D., Graduate School Representative

Marsha H. Read, Ph. D., Associate Dean, Graduate School

May, 2010

ABSTRACT

Crassulacean Acid Metabolism (CAM) is a water-conserving mode of photosynthesis present in approximately 7% of vascular plant species worldwide. CAM photosynthesis minimizes water loss by limiting CO2 uptake from the atmosphere at night, improving the ability to acquire carbon in water and CO2-limited environments. In neotropical orchids, the CAM pathway can be found in up to 50% of species. To better understand the role of

CAM in species radiations and the molecular mechanisms of CAM evolution in orchids, we performed carbon stable isotopic composition of leaf samples from 1,102 species native to Panama and Costa Rica, and character state reconstruction and phylogenetic trait analysis of CAM and epiphytism. When ancestral state reconstruction of CAM is overlain onto a phylogeny of orchids, the distribution of photosynthetic pathways shows that C3 photosynthesis is the ancestral state and that CAM has evolved independently several times within the Orchidaceae. Using phylogenetic trait analysis, we found that divergences in photosynthetic pathway and epiphytism are consistently correlated through evolutionary time and are related to the prevalence of CAM epiphytes in lower elevations and abundant species diversification of high elevation epiphytes. The multiple independent evolutionary origins of CAM in orchids suggest that evolution from C3 to weak and strong CAM might involve relatively few genetic changes.

In plants performing CAM, phosphoenolpyruvate carboxylase (PEPC) catalyzes the initial fixation of atmospheric CO2 into C4-dicarboxylic acids forming oxaloacetate and inorganic phosphate as a product. PEPC is a ubiquitous enzyme that belongs to a ii multigene family with each gene encoding a function- and tissue-specific isoform of the enzyme. CAM-specific PEPC isoforms might have evolved from ancestral non- photosynthetic C3 isoforms by gene duplication and acquired transcriptional control sequences that mediate increased mRNA expression and leaf-specific or leaf-preferential expression patterns. In order to understand patterns of PEPC family diversification over evolutionary times, PEPC genes families in ten closely-related orchid species from the

Subtribe Oncidiinae with a range of photosynthetic pathways from C3-photosynthesis

(Oncidium maduroi, Ticoglossum krameri, and Oncidium sotoanum) to weak CAM

(Oncidium panamense, Oncidium sphacelatum, Gomesa flexuosa and Rossioglossum insleayi) to strong CAM (Rossioglossum ampliatum, Trichocentrum nanum, and

Trichocentrum carthaginense) were characterized. At least three major changes are hypothesized to have occurred during evolution to adapt the CAM progenitor genes for function in CAM plants: 1) CAM isoform genes in orchids have evolved highly expressed mRNA expression patterns; 2) leaf preferential (or specific) expression patterns; and 3) circadian clock control expression patterns. We found that up to five

PEPC isoforms are present in orchids, with one putative CAM-specific PEPC isogene with discrete amino acid changes identified in CAM species based on cDNA clone sampling, and an evident shift in PEPC isoform number from 2-3 isoforms in C3 species, to 3-4 isoforms in weak CAM species, to 4-5 isoforms in strong CAM species. Validation of the isotopic analysis and the molecular genetic analysis of PEPC gene family using 24- hour gas exchange showed that weak CAM species exhibit limited amounts of nocturnal

CO2 uptake and fixation when compared to strong CAM species. iii

To understand the molecular mechanisms responsible for the recruitment of

CAM-specific genes, 454 sequencing of cDNA prepared from RNA of the strong CAM species Rossioglossum ampliatum was conducted, and resulted in 189 Mb of DNA sequence, 41,115 contigs, and 100,889 singletons. A NimbleGen microarray constructed and used in a C3 species (Oncidium maduroi), a weak CAM species (Oncidium panamense) and a strong CAM species (Rossioglossum ampliatum), showed that C3 and weak CAM species had average hybridization intensities that diverged from the strong

CAM species by 2 and 3 percent, respectively. From 13,566 genes that showed a significant 4.6-fold difference in expression levels from the comparisons between CAM,

C3 and weak CAM, 4,520 genes showed a greater than 4.6-fold increase in the ratio of

CAM/C3 relative transcript abundance, whereas 3,745 genes showed a greater than 4.6- fold decrease in the ratio of CAM/C3 relative transcript abundance. A maximal increase or decrease in relative transcript abundance of more than 1,000- and 500-fold, respectively, was observed. The results of the microarray analysis will serve as a catalogue of gene expression patterns available for future work aimed at understanding

CAM specific expression patterns, and can be used to further understand gene regulation by in-depth analysis of the transcriptional control regions responsible for altered gene expression patterns associated with CAM evolution. Several patterns of CAM evolution have been demonstrated in orchids, thus improving our understanding of the functional significance and evolutionary origins of CAM. The results of this project will aid in understanding photosynthetic plasticity in plants.

Dedication

I dedicate this dissertation to my parents Flor Maria M. de Silvera and Gaspar A. Silvera. They taught me the love for orchids, the patience to cultivate them, and the wisdom to understand them. Beyond all, they gave me the confidence and encouragement to always follow my dreams. I am who I am because of them. v

ACKNOWLEDGMENTS

I would like to thank my advisor John Cushman for his invaluable guidance throughout these five years. John has been an extraordinary mentor; he gave me the freedom to explore my own ideas, and supported me especially when I was doing research in Riverside and Panama. He always cared about my development as a scientist, he helped me write proposals for funding and he took the time to teach me new techniques. I will always be grateful for your patience, encouragement, and the care you put into reading and editing my writing. I am a much better scientist because of you!

I thank the members of my dissertation committee Jeff Harper, Bob Nowak, Dave

Shintani and David Zeh for all their helpful advice. They were always willing to meet with me to chat about my project and take the time to work with me on developing new ideas, especially during my qualifying exam.

Thank you Klaus Winter for putting me in touch with John Cushman, and for introducing me to the world of CAM. You have been an amazing mentor throughout these years, and have always provided support and guidance while working at STRI in

Panama. You have helped me maintain confidence in my research and have always provided me with valuable career advice. Working in your lab is inspiring.

I thank the members of the Cushman lab. I have learned so much from each of you! Thank you Becky Albion, you have been an amazing friend and colleague, you are always happy to discuss ideas with me, teach me techniques, and take care of the orchids!

You are a wonderful person to have in the lab! Thank you Letty Rodriguez, for working with me in the project and for sharing my passion for orchids. Thank you for always vi being positive and optimistic, even when research was difficult. Thank you JR Tillett for introducing me to so many fun places and people in Reno, and for exchanging music with me. Thank you Patricia Berninsone for opening your home to me and treating me like family! Thanks Leyla and Upul for your friendship. Thanks Miguel Rodriguez for training me when I first arrived to the lab. Thanks Abou, Bahay, Jill, Gadi, Gouthu,

Jerome, Lina, Matt, Mark, Mustafa, Sage, Sabine, Sangho, and Tatiana for being such wonderful labmates and friends. Thank you to the supportive staff at UNR, Marianne

Davis, Ron Robards, Becky Hess, and Jessica Corey for taking care of my fellowship paperwork every semester. Thanks Tim and Corene for your friendship and for helping me keep things in perspective. Thanks to Tim Close and Ray Fenton for welcoming me in their lab in Riverside. Thank you Mark Whitten for always providing good advice, and for continuing collaborating in orchid projects with me. Thank you Kent Perkins, Kurt

Neubig, Norris Williams, Bruce Holst, Jim Solomon, Mireya Correa, and Carmen

Galdames for access and assistance with herbarium collections. Thank you Orquideas

Tropicales for donating orchids for the project. Thank you Todd Dawson and Stefania

Mambelli for isotopic analysis. Thank you Orelis Arosemena for taking care of permits.

Special thank you to my parents Flor Maria and Gaspar for your unconditional love and support. To my brothers, and all my family and friends in Panama, who have always cared for me, and taken pride in all my achievements. A very special thank you to my amazing husband Lou Santiago, you are wonderful! Not only are you my husband and best friend, you are also my collaborator! Life with you is exciting and always an adventure. I would have not been able to complete this PhD without you, thank you for your encouragement, unconditional love and support. vii

TABLE OF CONTENT

LIST OF FIGURES AND TABLES…………………………………………………….x

CHAPTER ONE- INTRODUCTORY CHAPTER: Evolution along the crassulacean acid metabolism continuum……...... …………………………………1 Abstract...... …………………………………….……………………………...…..2 Introduction..………………………………….……………………………..….....3 Phases of CAM………………………….………………………………...3 Permutations of CAM…………………………………………………...... 5 CAM plasticity………………………………………………...... 6 Requirements for CAM…………………………………………………………....7 Convergence of leaf succulence in CAM species………………………....8 Evolutionary drivers of CAM……………………………………….…...11 Taxonomic distribution of CAM………………………………………………...12 Estimating the prevalence of CAM……………………………………....13 Orchids as a model for the study of CAM evolution………………………….....16 Mapping the occurrence of CAM within the Orchidaceae……………....17 Molecular evolution of CAM……………………………………………...... 19 Molecular markers for studying CAM evolution………………………...19 Circadian clock regulated markers………………………………...... 23 Circadian clock specialization during CAM evolution…………………..25 Acknowledgments………………………………………………………………..26 Literature cited…………………………………………………………...... 28

CHAPTER TWO: Crassulacean acid metabolism and epiphytism linked to adaptive radiations in the Orchidaceae………………...... …………....……….....48 Abstract……………………………………………………………………...... 49 Introduction…………………………………………………………………...... 50 Material and methods………………………………………………………….....52 Site description………………………………………………………...... 52 Carbon isotope analysis……………………………………...... 53 Climatic data……………………………………………………...... 54 Orchid phylogeny and species nomenclature……………………...... 54 Phylogenetic and statistical analysis…………………...... 55 Results……………………………………………………………………………58 Discussion………………………………………………………………………..60 Conclusions………………………………………………………………………64 Acknowledgments……………………………………………………………...... 65 Literature cited…………………………………………………………...... 67

CHAPTER THREE: Incidence of crassulacean acid metabolism in the Orchidaceae derived from carbon isotope ratios: a checklist of the flora of Panama and Costa Rica……………………………………………………………...... 77 Abstract…………………………………………………………………………..78 viii

Introduction………………………………………………………………………78 Material and methods………………………………………………………….....80 Site description…………………………………………………………...80 Carbon isotope analysis………………………………………...... 81 Results……………………………………………………………………………83 Discussion………………………………………………………………………..84 Acknowledgments……………………………………………………………...... 87 Literature cited…………………………………………………………………...88

CHAPTER FOUR: Dynamics of the phosphoenolpyruvate carboxylase (Ppc) gene family in the Orchidaceae (Subtribe Oncidiinae) during the evolution of crassulacean acid metabolism…...... ……………………………...122 Abstract…………………………………………………………………………122 Introduction……………………………………………………………………..123 Material and methods…………………………………………………………...127 Plant material…………………………………………………………...127 Gas exchange measurements…………………………………………...127 Extraction of RNA……………………………………………………...128 RT-PCR amplification and cloning…………………………………….128 Sequence analysis………………………………………………………130 Race amplification……………………………………………………...130 Results and discussion………………………………………………………….131 Oncidiinae species and characterization………………………………..131 Photosynthetic pattern of Oncidiinae species…………………………..131 Identification of multiple PEPC isogenes in the Oncidiinae species...... 135 Conclusions……………………………………………………………………..141 Acknowledgments………………………………………………………………142 Literature cited………………………………………………………………….143

CHAPTER FIVE: Large-scale microarray gene expression profiling in Oncidiinae orchids along the C3 to CAM evolutionary continuum…………...... 210 Abstract…………………………………………………………………………210 Introduction……………………………………………………………………..211 Material and methods…………………………………………………………...214 Plant material…………………………………………………………...214 RNA isolation…………………………………………………………..215 454 Life Sciences (Roche) sequencing technology…………………….215 NimbleGen microarray fabrication and analysis……………………….216 Results and discussion………………………………………………………….217 Categorization of CAM-related genes………………………………….219 Conclusions……………………………………………………………………..222 Acknowledgments………………………………………………………………223 Literature cited………………………………………………………………….224

CHAPTER SIX: Concluding remarks…………………………...…………………. 254 Literature cited………………………………………………………………….258 x

LIST OF FIGURES AND TABLES

CHAPTER ONE

Table 1. !13C values, nocturnal fluctuation in titratable acidity, and leaf traits from 173 orchid species………...………………………………………………..41

Table 2. Taxonomic distribution of CAM plants, including family, order and exemplar species………………………………………………………………....42

Figure 1. CAM metabolite fluxes during nighttime (left panel) and daytime (right panel)…………….……………………..………………………………….43

Figure 2. Major requirements hypothesized for the evolution of CAM along the evolutionary progression from C3 photosynthesis to weak CAM and strong CAM...... 44

Figure 3. Summary tree for the classification of the family Orchidaceae………………………………………………………………………45

Figure 4. Summary tree for the appearance of weak and strong CAM among Oncidiinae species………………………………………………….…….46

Figure 5. Evolution of PEPC genes (Ppc) in the Orchidaceae…………………..47

CHAPTER TWO

Figure 1. A, Percentage of CAM species as a function of altitude. B, Total number of species as a function of altitude…...……………………...... 72

Figure 2. Relationship between !13C and climatic variables………………….….73

Figure 3. Orchidaceae tree showing the relationship among 147 tropical genera…………………………………………………………………………….74

Figure 4. Orchidaceae tree showing the relationship among 53 subtribes………75

Figure 5. Pairwise correlation between !13C and epiphytism (A), and divergence in !13C and epiphytism among radiations in the tree for orchid genera (B)………………………………………………………………………...76

CHAPTER THREE

Figure 1. Frequency of leaf !13C values for 1,002 Panamanian and Costa Rican orchid species………………..……………………………………..90

Table 1. Leaf carbon isotopic values (‰) and voucher information from 1,002 orchid species of Panama and Costa Rica, including current taxonomic name and herbarium accession number……………………...91

Table 2. List of orchid genera containing species with CAM…………….……119

CHAPTER FOUR

Table 1. !13C values, leaf thickness and titratable acidity for ten Oncidiinae species…………………………………………………………………………..146

Table 2. Gene specific primer sets (GSP 5’– 3’ position) used for 3’RACE amplification of the cDNA fragment coding for Ppc isoform for ten Oncidiinae species and 36 isoforms...... 147

Table 3. Different functional PEPC isoforms from ten Oncidiinae species based on relative abundance of clone sampling………………………………...148

Table 4. Multiple sequence similarity scores (%) among PEPC contigs (36974, 35735, 10159) obtained from 454 pyrosequencing of a strong CAM species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (weak CAM), and Oncidium maduroi (C3)..…………………………………………………...149

Figure 1. Phylogeny for ten Oncidiinae species……………………...……….150

Figure 2. Continuous net CO2 uptake by C3 orchid species during a 12 h light (white bar) and 12 h dark period (black bar).…………………………...... 151

Figure 3. Continuous net CO2 uptake by weak CAM orchid species during a 12 h light (white bar) and 12 h dark period (black bar)………….……152

Figure 4. Continuous net CO2 uptake by strong CAM orchid species during a 12 h light (white bar) and 12 h dark period (blackbar)………………..153

Figure 5. Aligned partial nucleotide sequence of 1,100 base pairs from 35 PEPC isoforms from ten Oncidiinae species: Trichocentrum nanum xii

(CAM), Rossioglossum ampliatum (CAM), Trichocentrum carthaginense (CAM), Gomesa flexuosa (Weak CAM), Oncidium sphacelatum (Weak CAM), Oncidium panamense (Weak CAM), Rossioglossum insleayi (Weak CAM), Oncidium sotoanum (C3), Oncidium maduroi (C3), Rossioglossum krameri (C3)……………………...……………………...……..154

Figure 7. Phylogenetic tree derived from neighbor joining analysis of 1,100-bp alignment of Ppc nucleotides sequences from ten closely-related Oncidiinae species……………………………...………………196

Figure 8. Phylogenetic tree derived from neighbor joining analysis of 363 amino acid alignment of Ppc from ten closely-related Oncidiinae species…………………………………………………………………………..197

Figure 10. Protein alignment for six isoforms from three Oncidiinae species, including the 3’UTR region recovered using 3’RACE…………………………199

Figure 11. Aligned partial nucleotide sequence of 876 base pairs between PEPC contig 35735 obtained from 454 pyrosequencing of a strong CAM species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (Weak CAM), and Oncidium maduroi (C3)…………………...... ……………………………………………..200

Figure 12. Aligned partial nucleotide sequence of 221 base pairs between PEPC contig 10159 obtained from 454 pyrosequencing of a strong CAM species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (Weak CAM), and Oncidium maduroi (C3)……………………………………………………………………206

Figure 13. Aligned partial nucleotide sequence of 224 base pairs between PEPC contig 36974 obtained from 454 pyrosequencing of a strong CAM xiii

species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (Weak CAM), and Oncidium maduroi (C3)……………………………………………………………………208

CHAPTER FIVE

Table 1. List of genes with CAM-related function derived from microarray data………………………………………………………………….…………..226

Table 2. List of the 30 genes exhibiting the most decrease in transcript abundance between C3 and weak CAM species………………………………..231

Table 3. List of the 30 genes exhibiting the most decrease in transcript abundance between CAM and C3 species………………………………………233

Table 4. List of the 30 genes exhibiting the most decrease in transcript abundance between CAM and weak CAM species……………………….....…235

Table 5. List of the 30 genes exhibiting the most increase in transcript abundance between C3 and weak CAM species………………………………..237

Table 6. List of the 30 genes exhibiting the most increase in transcript abundance between CAM and C3 species. …………………………………….238

Table 7. List of the 30 genes exhibiting the most increase in transcript abundance between CAM and weak CAM species. …………………………...239

Figure 1. Principal component analysis (PCA) of mRNA expression profiles of three Oncidiinae species and four biological replicates per species…………240

Figure 2. Normalized mRNA expression for the comparison between weak CAM and CAM (A); C3 and CAM (B) and weak CAM and C3 (C)…………...241

Figure 3 Plot of log2-transformed intensity ratios showing increased or decreased mRNA abundance values of probe sets between weak CAM

and CAM, and between CAM and C3…………………………………………. 243

Figure 4. Average mRNA expression (±SD) of selected genes with CAM-related functions that exhibit shifts in expression patterns between CAM, weak

CAM and C3 species……………………………………………………………244

Chapter I

Introductory Chapter

Evolution Along the Crassulacean Acid Metabolism Continuum

Katia Silvera1, Kurt M Neubig2, W. Mark Whitten2, Norris H. Williams2, Klaus Winter3 , and John C. Cushman1,*

1Department of Biochemistry & Molecular Biology, MS200, University of Nevada,

Reno, NV 89557-0200, USA.

2Florida Museum of Natural History, University of Florida, Gainesville, FL 32611-7800,

USA.

3Smithsonian Tropical Research Institute, P.O. Box 0843-03092, Balboa, Ancón,

Republic of Panama.

*Corresponding author;

John C. Cushman

E-mail address: [email protected]

Functional Plant Biology (Invited review article, Submitted) 2

ABSTRACT

Crassulacean acid metabolism (CAM) is a specialized mode of photosynthesis that improves atmospheric CO2 assimilation in water-limited terrestrial and epiphytic habitats, and in CO2-limited aquatic habitats. In contrast to C3 and C4 plants, CAM plants take up

CO2 from the atmosphere partially or predominantly at night. CAM is taxonomically widespread among vascular plants and is present in many succulent species that occupy semi-arid regions, as well as in tropical epiphytes and in some aquatic macrophytes. This water-conserving photosynthetic pathway has evolved multiple times, and is found in up to an estimated 10% of vascular plant species from more than 35 families of angiosperms. Although many aspects of CAM molecular biology, biochemistry and ecophysiology are well understood, relatively little is known about the evolutionary origins of CAM. This review focuses on four main topics: 1) drivers of CAM evolution,

2) co-requisites of CAM evolution, 3) the prevalence and taxonomic distribution of CAM among vascular plants with an emphasis on the Orchidaceae, and 4) the molecular underpinnings of CAM evolution.

Abbreviations: CAM, Crassulacean acid metabolism; !13C, carbon stable isotopic composition; Gpt2, glucose-6-phosphate/Pi translocator; Glc-6-P, glucose-6-phosphate

PEPC, phosphoenolpyruvate carboxylase; PPCK, PEPC kinase.

Keywords: Crassulacean acid metabolism, evolution, !13C, phosphoenolpyruvate carboxylase, photosynthesis.

INTRODUCTION

Crassulacean acid metabolism (CAM) is one of three modes of photosynthetic assimilation of atmospheric CO2, along with C3 and C4 photosynthesis. The net result of

CAM is an improvement in water use efficiency (WUE; CO2 fixed per unit water lost) generally six-fold higher than for C3 plants and three-fold higher than for C4 plants under comparable conditions (Nobel 1996). Thus, CAM is an important ecophysiological metabolic adaptation that permits plants to thrive in semi-arid habitats and habitats with intermittent or seasonal water availability (Cushman 2001; Winter and Smith 1996). In this review, we examine the permutations and plasticity of CAM in the context of evolution, the metabolic and genetic requirements for CAM, including leaf succulence, and the likely evolutionary drivers of CAM. We next examine the current surveys of the taxonomic distribution of CAM species and the several survey methods used to estimate the prevalence of CAM. Lastly, we discuss molecular evolution of CAM, including the origins of CAM, molecular markers used to study the evolutionary progression of gene family changes, circadian clock control, and future research directions.

Phases of CAM

The physiological and biochemical temporal sequence of events that constitute CAM have been described in detail as four discrete phases (Griffiths 1988; Lüttge 1987;

Osmond 1978; Winter 1985). Phase I is typically characterized by nocturnal stomatal opening and transpiration, CO2 uptake and fixation by phosphoenolpyruvate carboxylase

(PEPC) in the cytosol and the formation of C4 organic acids (usually malic acid), which are stored in the vacuole (Fig. 1). The rate of nocturnal CO2 assimilation is governed by 4 mesophyll processes, such as carbohydrate storage reserves for carboxylation (Cushman et al. 2008) or vacuolar storage capacity, rather than stomatal conductance (Winter 1985).

Depending on the CAM species, a variety of storage carbohydrates (e.g., starch, glucans, soluble hexoses) might be metabolized to produce phosphoenolpyruvate (PEP), the substrate for carboxylation (Christopher and Holtum 1996; Christopher and Holtum

1998; Holtum et al. 2005). Phase I reflects the fundamental adaptation of CAM resulting in reduced transpiration and improved water economy (Griffiths 1988). Phase II describes the transition from PEP carboxylase to ribulose-1,5-bisphosphate carboxylase/oxygenase

(RUBISCO) mediated carboxylation during the early light period leading to carbohydrate production. Thus, CO2 fixation during this phase is a mixture of CO2 derived from organic acid decarboxylation and direct uptake from the atmosphere. Phase III encompasses the period of major efflux of organic acids from the vacuole with subsequent decarboxylation resulting in internal leaf CO2 partial pressures of greater than

100 times atmospheric levels (Cockburn et al. 1979; Spalding et al. 1979), decline in stomatal opening and transpiration, and even CO2 release from the leaf despite low stomatal conductance (Frimert et al. 1986). Decarboxylation is catalyzed by either cytosolic PEP carboxykinase (PEPCK) or cytosolic NADP+-/mitrochondrial NAD+-malic enzymes (ME) (Christopher and Holtum 1996; Holtum et al. 2005; Smith and Bryce

1992). This CO2 concentrating mechanism or “CO2 pump” effectively suppresses photorespiration during this phase. Phase IV is a second transitional phase marked by the depletion of organic acid stores, slower rates of decarboxylation, declines in internal partial pressure of CO2, and increases in stomatal conductance depending on the prevailing environmental conditions. CO2 fixation during the early phase IV is a mixture 5

of CO2 assimilation derived mainly from organic acid decarboxylation and direct atmospheric uptake and assimilation via RUBISCO, however, carboxylation into C4 acids by PEPC increases as the dark period approaches (Ritz et al. 1986). Because CAM plants perform both CO2 fixation steps within the same cell, futile cycling of CO2 is minimized by temporal control of the kinetic properties of PEPC in response to malic acid (Winter

1982) and RUBISCO in response to light (Maxwell et al. 1999; Griffiths et al. 2002). The diel change in kinetic properties of PEPC (Winter, 1982) are triggered by reversible phosphorylation events catalyzed by a dedicated protein kinase (Hartwell et al. 1996;

Hartwell et al. 1999; Taybi et al. 2000), whose expression is controlled by the circadian clock (Hartwell 2005a; Hartwell 2005b).

Permutations of CAM

Although the four CAM phases definition appear adequate to describe all observed acid metabolism phenomena (Lüttge 1987), additional terminologies have been suggested to describe CAM in some astomatal aquatic species or in the astomatal green aerial roots of epiphytic orchids (Cockburn 1985). Furthermore, environmental conditions can modulate the extent to which each phase is manifested (see also next section) (Cushman 2001;

Cushman and Borland 2002). For example, water deficit stress can reduce or eliminate phase IV and light and temperature can regulate the appearance or onset of phases II and

III (Griffiths 1988). Under severe water deficit stress, phase I net nocturnal CO2 uptake can be eliminated completely along with virtually all stomatal conductance across the four phases. This phenomena, termed “CAM idling”, results in small, sustained diel fluctuations in organic acids with 100% of the CO2 fixed into malate being derived from 6

internally recycled respiratory CO2 (Szarek et al. 1973; Ting 1985). CAM idling might play an important role in the prevention of photoinhibition by maintaining photosystem stability (Osmond 1982). The phenomena of “CAM-cycling” or “weak CAM” have also been described, wherein organic acid fluctuations are observed, but with little or no net nocturnal CO2 fixation by PEPC (Sipes and Ting 1985; Ting 1985). In the context of evolution, CAM-cycling has been interpreted to be a basal form of CAM with increasing nocturnal CO2 fixation being associated with an increasingly advanced state among the

Crassulaceae (Teeri 1982; Terri 1982) and the Bromeliaceae (Smith et al. 1986). The ecophysiological significance of CAM-idling might be to keep plants poised to engage in full CAM once drought conditions end by maintaining the capacity for organic acid fluctuation (Ting 1985). Similarly, the evolutionary importance of weak CAM might serve as a genetic reservoir for CAM radiations in the context of changing environmental conditions or habitat exploitation, such as epiphytism (Silvera et al. 2005; Silvera et al.

2009). Finally, the term “latent CAM” has been used to describe an intermediate form of

CAM wherein organic acid concentrations remain high, but constant throughout the diel cycle (Schuber and Kluge 1981). As with CAM-cycling or weak CAM, latent CAM might be regarded as a step along the progression from C3 to CAM (Lee and Griffiths

1987).

CAM Plasticity

The degree to which CAM operates can vary greatly depending on the evolutionary history of a given species and its environmental conditions with the end result being a continuum of differences in the degree to which nocturnal net uptake of CO2 occurs in 7

relation to daytime net CO2 uptake (Cushman and Bohnert 1999; Cushman 2001;

Cushman and Borland 2002; Dodd et al. 2002). For example, many CAM species engage in “obligate” or “constitutive” CAM in fully mature photosynthetic organs (e.g. leaves and stems), although the extent of gas exchange and nocturnal acidification might be modulated by prevailing environmental condition (Griffiths 1988). Many members of the

Cactaceae and Crassulaceae provide excellent examples of this type of CAM. In contrast, “facultative”, “inducible”, or “optional” CAM or C3-CAM intermediate species engage in CAM in response to environmental stimuli such as drought stress (Winter

1985; Griffiths 1988; Winter et al. 2008). The expression of CAM in such C3-CAM species varies dynamically with experimental manipulating conditions, such as photoperiod (Brulfert and Queiroz 1982), water status, light or CO2 availability, temperature, nutritional status, salinity, anoxia, or atmospheric CO2 levels (Winter 1985;

Lüttge 1987; Griffiths 1988; Roberts et al. 1997). The common ice plant,

Mesembryanthemum crystallinum, a member of the Aizoaceae, is a well-studied example of inducible CAM, now known to be under strict environmental control (Winter 1985;

Winter and Holtum 2007; Cushman et al. 2008).

Requirements for CAM

The basic enzymatic machinery essential for CAM operation is present in the mesophyll cells of all plant species, so what evolutionary changes must occur in order for CAM to function? The first and foremost diagnostic indicator of CAM is nocturnal CO2 uptake

(Fig. 2). Second, diel fluctuations in organic acids and reciprocal diurnal fluctuations of storage carbohydrates such as starch, glucans, or soluble hexoses are typical features of 8 the CAM cycle (Ting 1985). Third, enhanced expression of PEPC and decarboxylating

(e.g., PEPCK or NADP+-/ NAD+-ME) enzymes and associated transporters across the tonoplast (e.g., vacuolar H+-ATPase) and mitochondria are needed (Holtum et al. 2005;

Cushman et al. 2008). Fourth, enhanced expression of enzymes of both the glycolytic and gluconeogenic pathways are required to support large (typically 40-60% of available reserves), reciprocating pools of carbohydrates (Paul et al. 1993; Borland and Dodd

2002; Dodd et al. 2003) and associated transport activities across the chloroplast envelope (Häusler et al. 2000; Kore-eda et al. 2005). Discrete isogenes appear to be recruited selectively in order to fulfill the activities necessary for CAM function (Kore- eda et al. 2005; Cushman et al. 2008). Fifth, some degree of leaf succulence characterized by mesophyll cells with increased cell size, increased mesophyll tissue and leaf thickness due to large storage vacuoles. Such large cell volume per unit leaf or stem area ensure a high capacity for nocturnal organic acid storage as well as water storage, whereas chloroplasts supply the requisite daytime carbohydrate biosynthesis (Gibson

1982; Nelson et al. 2005). Finally, circadian clock control of mRNA, translational, and posttranslational regulatory outputs are required to ensure that synthesis and transport of reciprocating organic acid and carbohydrate pools are properly synchronized along the diel CAM cycle (Borland et al. 1999; Hartwell et al. 1999; Taybi et al. 2000; Dodd et al.

2003; Cushman et al. 2008).

Convergence of Leaf Succulence in CAM species

A general anatomical feature of CAM plants and apparent evolutionary co-requisite for

CAM is leaf succulence (Fig. 2) with vacuoles occupying 90-95% of cell volume with 9 cell dimensions of 100 "m or larger (Gibson 1982; Smith 1984). A tight correlation between greater leaf succulence or thickness and increased magnitude of CAM has been observed within the Crassulaceae (Teeri et al. 1981; Kluge et al. 1991; Kluge et al.

1993), the Orchidaceae (Winter et al. 1983; Silvera et al. 2005), as well as many other diverse CAM families (Nelson et al. 2005; Nelson and Sage 2008). Large cell size leads to a tightly packed chlorenchyma with reduced intercellular air spaces (IAS) and reduced surface area exposure of mesophyll cells to IAS (Lmes/area), which likely results in low internal conductance of CO2 (gi) and a restriction of CO2 efflux that presumably enhances

CAM efficiency (Maxwell et al. 1997; Osmond et al. 1999; Borland et al. 2000; Nelson et al. 2005; Nelson and Sage 2008). In C3 plants, however, low (gi) is known to limit C3 photosynthesis (Evans and von Caemmerer 1996; Evans and Loreto 2000) and also likely limits CO2 uptake during phase IV in CAM plants (Maxwell et al. 1999; Maxwell 2002).

Thus, a trade-off exists between carbon gain in C3 and CAM as a result of leaf succulence. In CAM species, the diffusional barrier created by large, tightly packed cells limits the loss of internal CO2 leakage during all four phases of CAM, but particularly during phase III, thereby enhancing carbon economy. In contrast, access to CO2 is restricted during the C3-dominated phase IV of CAM (Nelson and Sage 2008). CAM species with greater leaf succulence show a higher degree of nocturnal CO2 fixation and exhibit an overall stronger reliance on CAM due to low mesophyll CO2 conductance

(Griffiths et al. 2008).

If the above anatomical traits associated with leaf succulence enhance the degree of CAM photosynthesis and limit the degree of C3 photosynthesis, then all CAM species, regardless of their evolutionary lineage, would be expected to converge to a common 10 succulent leaf anatomy. Supporting evidence for this hypothesis has been obtained by comparing the degree of leaf succulence, indicated by leaf thickness, to leaf !13C values

(Winter et al. 1983; Zotz and Ziegler 1997). For example, a survey of leaf thickness and leaf !13C values in 173 tropical orchid species (Table 1) revealed that in species with leaf

13 ! C values commonly observed for C3 plants (–33 to –22‰), leaf thickness averaged 0.5

± 0.4 mm, whereas in species with !13C values typical of CAM plants (–20 to –12‰), leaf thickness averaged 2.2 ± 2.1 mm. In species with intermediate !13C values (–22 to –

20‰), leaf thickness averaged 1.2 mm ± 0.7 mm (Silvera et al. 2005). Thus, relative leaf thickness might serve as a useful surrogate indicator for the presence of CAM activity provided that hydrenchyma (without chloroplast containing cells) is not the major contributor of leaf thickness (Fig. 2). Just as increased leaf thickness confers a selective advantage for committing fully to CAM, increased leaf succulence is deleterious for CO2 fixation via the C3 pathway and overall photosynthetic productivity (Borland et al. 1994;

Borland and Dodd 2002). Thus, evolutionary progression from the C3 to CAM state would appear to favor either retention of C3 photosynthesis or full conversion to CAM, but not the intermediate state. Indeed, such a pattern is reflected in the bimodal distribution of high and low IAS and Lmes/area values of weak and strong CAM species

(Nelson and Sage 2008), as well as the bimodal distribution of !13C values observed in large surveys of plant families with mixtures of C3/weak CAM and CAM species (see below) (Zotz and Ziegler 1997; Pierce et al. 2002; Crayn et al. 2004; Holtum et al. 2004;

Silvera et al. 2005; Silvera et al. 2009; Silvera et al. 2010).

Evolutionary drivers of CAM

Numerous reports have postulated that C3 photosynthesis is the evolutionary ancestral or progenitor state for CAM with the progression towards CAM involving several incremental steps (Teeri 1982; Pilon-Smits et al. 1996; Crayn et al. 2004; Silvera et al.

2009). Reversion of CAM to the C3 state is also possible (Teeri 1982) and evidence for likely reversal events, associated with radiations into less xeric habitats, has come from large-scale isotopic surveys within the Bromeliaceae (Crayn et al. 2004) and the

Orchidaceae (Silvera et al. 2009). The reversion of CAM to C3 photosynthesis points to the complex evolutionary histories within these taxa. Although the main driver for this progression to CAM evolution remains unclear, several hypotheses have been put forward. Water limitation and the resulting limitation of CO2 brought about by stomatal closure and reductions in atmospheric CO2 concentrations during the late Tertiary period

(Pearson and Palmer 2000) may have provided the selective pressures for the evolution of

CAM over the last 40-100 million years (Monson 1989; Ehleringer and Monson 1993;

Raven and Spicer, 1996). It is difficult, however, to determine the first origin of CAM in plants, especially because the majority of families in which CAM is present originated recently and fossil evidence of CAM has not been discovered (Raven and Spicer 1996).

Interestingly, Dendrobium and Earina (Epidendroideae) macrofossils of orchid specimens from early Miocene (23-20 MYA) have been described, however, these were not investigated for the presence of CAM (Conran et al. 2009). Based on the broad diversity of taxa showing CAM compared to C4 species, CAM likely evolved first, and based on the presence of CAM in ancient groups such as the isoetids and cycads, CAM may have appeared as early as the Triassic (Griffiths 1992; Ehleringer and Monson 12

1993). In any event, CAM likely evolved in response to selection for increased carbon gain and increased water use efficiency (Ehleringer and Monson 1993) after global reduction in atmospheric CO2 concentration during the late Miocene and early

Pleistocene climate. Notably, a large CAM radiation event in the most species rich epiphytic clade in orchids (Fig. 3), the Epidendroideae, was predicted to have originated about 65 MYA and linked to the decline of atmospheric CO2 during the Tertiary (Fig. 3)

(Silvera et al. 2009). CAM has contributed to the exploitation of wider epiphytic habitat ranges, from low elevation sites where CAM orchids are mostly present, to mid-elevation tropical forest sites of around 1,000 m, were moist suitable microenvironments exist for epiphytic orchid colonization (Silvera et al. 2009).

Taxonomic distribution of CAM

Numerous past studies have estimated the distribution of CAM (Moore 1982; Winter

1985; Lüttge 1987; Griffiths 1988; Ehleringer and Monson 1993). CAM is widespread within the plant kingdom in at least 343 genera in 36 plant families comprising approximately 6.5% of flowering plant species (Table 2; JAC Smith, unpublished data)

(Griffiths 1989; Smith and Winter 1996; Holtum et al. 2007). In contrast, C4 photosynthesis occurs in 19 families and account for approximately 3% of plant species comprising mainly grasses and sedges and some dicots (Sage 2001; Sage 2004). CAM is generally not found in more primitive orders (with a few exceptions). The oldest lineage with CAM is represented by Isoetes, a mostly aquatic or semi-aquatic group distributed in oligotrophic lakes or mesotrophic shallow seasonal pools (Keeley 1998). The retention of

CAM in this group is linked to the chronically low or daytime decline in levels of 13

dissolved CO2 in these aquatic environments (Keeley 1996). CAM has also been documented within the Cycadales, Welwitschia mirabilis (Welwischiaceae) (von Willert et al. 2005) and the Gnetales, Dioon edule (Zamiaceae) (Vovides et al. 2002), and several epiphytic families of ferns within the Polypodiaceae (Holtum and Winter 1999) and the

Vittariaceae (Martin et al. 2005).

The widespread taxonomic distribution of both C4 and CAM plants indicates that

C4 and CAM plants must have evolved independently multiple times, even within a single genus (Monson 1989; Ehleringer and Monson 1993; Kellogg 1999; Monson 1999;

Silvera et al. 2009). Studies with limited taxon sampling by !13C analysis have been reported for the Crassulaceae (Kalanchoë) (Kluge et al. 1991), Sedum and Aeonium

(Pilon-Smits et al. 1996), Clusiaceae (Gehrig et al. 2003), and Orchidaceae (Cymbidium)

(Motomura et al. 2008). More extensive combined taxon and isotopic sampling has been completed only within the Bromeliaceae (Crayn et al. 2004), and the Orchidaceae

(Silvera et al. 2005; Silvera et al. 2009; Silvera et al. 2010).

Estimating the prevalence of CAM

CAM species are widely distributed throughout semi-arid tropical and subtropical environments, including epiphytes in the humid tropics that must endure intermittent or seasonal water availability. Excluding the Orchidaceae, approximately 9,000 species are estimated to perform CAM (Winter and Smith 1996). However, the Orchidaceae alone may contribute an additional 10,400 species assuming that of the estimated 26,000 orchid species (Pfahl et al. 2008) approximately 10% engage in strong CAM and 30% engaging in weak CAM (Silvera et al. 2005). Thus, about 19,400 species or about 6.5% of vascular 14 plant species might engage in CAM to varying degrees. The extent of CAM expression generally correlates with the degree of adaptation to more xeric ecological niches (Kluge et al. 2001; Pierce et al. 2002; Zotz 2004). A recent survey of 1,022 orchid species of

Panama and Costa Rica using stable isotopic measurements documented that the number of CAM species increases with decreasing precipitation with the majority of CAM species at sites between sea level and 500 m and no CAM species above 2,400 m (Silvera et al. 2009). Similarly, while 19% of epiphytic species in a wet lowland forest (Zotz

2004) and 26% of epiphytic orchid species in the humid tropical forest of Papua New

Guinea (Earnshaw 1987) displayed CAM isotopic values, these percentages increased to

40% in a lowland forest in Panama (Zotz and Ziegler 1997) and 62% in an Australian forest (Winter et al. 1983). Also within a single site, the percentage of CAM epiphytes tended to increase from shaded understory sites to exposed canopy sites. For example, in a moist tropical forest in Panama, CAM is more prevalent in emergent layers and exposed tree canopies when compared to understory sites (Zotz and Ziegler 1997).

13 The differential enzyme-mediated discrimination against CO2 during photosynthetic carbon assimilation between C3 photosynthesis and CAM results in different whole-tissue carbon ratios (!13C) (Ehleringer and Osmond 1989). CAM species

13 13 ! C values typically range from –22 to –12‰, whereas for C3 plants, ! C values may range from –33 to –22‰ (Ehleringer and Osmond 1989). Thus, !13C values have become widely used as a rapid and relatively inexpensive screening method for determining CAM activity. A recent study of how closely !13C values reflect the proportion of day and night

CO2 fixed revealed that “the typical CAM plant” gains about 71% to 77% of its carbon through nocturnal fixation (Winter and Holtum 2002). However, surveys using only !13C 15 values to determine the number of CAM-equipped species can underestimate the number of CAM species that obtain less than one-third or less of its carbon in the dark (Winter and Holtum 2002). Recent surveys that include measurements of nocturnal tissue acidification have identified a greater number of CAM species than surveys using isotopic composition measurements alone (Pierce et al. 2002; Silvera et al. 2005).

Furthermore, surveys conducted during the rainy season might not reveal the presence of facultative CAM species that exhibit CAM only under water deficit stress conditions.

Thus, current estimates of the taxonomic distribution of CAM using stable isotopes are likely to underestimate the prevalence of CAM.

Integrative studies that attempt to map carbon-isotopic ratio surveys with molecular phylogenies remain limited. Although a well-resolved and comprehensively sampled molecular phylogeny of the Aizoaceae exists (Klak et al. 2003a; Klak et al.

2003b), the occurrence of CAM has not been mapped on the available phylogenetic tree.

Similarly, an incomplete molecular phylogeny has been established for about two-thirds of species within the Agavaceae (Good-Avila et al. 2006), however, most species are expected to perform CAM. A detailed molecular phylogenetic reconstruction of the

Vanilloideae with emphasis on the genus Vanilla, which surveyed 47 of the 110 different species has also been constructed using four plastid genes, however, no attempt was made to map the occurrence of CAM (Bouetard et al. 2010). DNA-based molecular phylogenies are well established for the Bromeliaceae (e.g. Crayn et al. 2004; Barfuss et al. 2005; Jabaily and Sytsma 2010). Carbon-isotopic ratios collected from 1,873 of 2,885 bromeliad species revealed that CAM photosynthesis and the epiphytic habit evolved a minimum of three times in this family (Crayn et al. 2004). Molecular phylogenies have 16 also been established for 31 of the estimated 350 species within the Clusiaceae with

CAM arising independently within two of the three major groups of Clusia species as estimated by carbon-isotope ratio analysis (Gehrig et al. 2003; Vaasen et al. 2002).

Orchids as a model for the study of CAM evolution

The Orchidaceae is the largest family of flowering plants with >800 genera and about

26,000 species worldwide, of which about three-fourths are estimated to be epiphytic

(Atwood 1986; Dressler 1993). Orchids exhibit a large number of morphological, anatomical, ecological and physiological characteristics that allow them to exist within diverse ecosystems and ecological niches with the greatest diversity existing in mountainous regions of the tropics (Cribb and Govaerts 2005). One such characteristic is the expression of CAM. Orchids are among the largest and most taxonomically well- studied family of CAM plants. For example, from 1994-2004 DNA sequences from

4,262 orchids have been deposited in GenBank (Cameron 2005). Chase et al. (2003) proposed an updated classification for the family based upon many recent and ongoing molecular phylogenetic studies. Orchids contain a useful distribution of C3 and CAM species (Silvera et al. 2010). In contrast, within other families, such as the Agavaceae,

Cactaceae, and the Didiereaceae, nearly all species are CAM and thus do not permit the evaluation of CAM evolutionary progression. Of families that display a mixture of both

C3 and CAM species (e.g., Aizoaceae, Bromeliaceae, Crassulaceae), the Orchidaceae has a well-resolved molecular phylogeny at an advanced stage as summarized by in five volumes of the Genera Orchidacearum (Pridgeon et al. 1999-2009). 17

The diversity expressed by orchids is crucial in linking CAM expression to vegetative morphology such as leaf thickness, habitat specialization such as epiphytism, and adaptive radiation spanning moisture gradients (Dressler 1993; Silvera et al. 2009;

Williams et al. 2001a). Silvera et al. (2005) used a combination of !13C isotopic ratios and titratable acidity measurements to survey the presence of CAM in 200 Panamanian native orchids species. The survey produced a bimodal distribution of !13C values with peaks around –15‰ (signifying strong CAM) and –28‰ (signifying C3 photosynthesis), comparable to other broad surveys employing !13C value measurements. Within the peak

13 of C3 photosynthesis ! C values, titratable acidity measurements revealed a second peak indicative of species with low capacities for nocturnal CO2 fixation (weak CAM). Taking into account both !13C values and titratable acidity measurements suggest that CAM appears to be more widespread among tropical epiphytic orchids than previously suggested. Up to 50% of species may exhibit some level of CAM photosynthesis (Silvera et al. 2005). However, fewer than 4% of all known orchid species have been sampled to date (Silvera et al. 2005).

Mapping the occurrence of CAM within the Orchidaceae

A key prerequisite for the phylogenetic reconstruction of the evolutionary origins of

CAM is a sufficiently robust and densely sampled phylogeny for the family based on molecular and morphological characters. The subtribe Oncidiinae is one of the most highly derived clades of orchids of the New World, with a great variety of diversity in chromosome number, vegetative features and floral characteristics (Chase et al. 2005).

Oncidiinae is the second largest orchid subtribe and comprises about 69 genera and 18 around 1600 species, the vast majority of which are epiphytic (Williams et al. 2001a;

Williams et al. 2001b). This subtribe is one of the most intensively sampled clades within Orchidaceae. About 600 of 1600 (37%) of species have been sampled using data for both nuclear and plastid DNA sequences as well as morphological characters

(Williams et al. 2001a; Williams et al. 2001b; Chase 2009), providing an excellent basis from which to study CAM evolution. The monophyly of the Oncidiinae and phylogenetic relationships of related genera have been evaluated by combined data from the internal transcribed spacer of nuclear ribosomal (nrITS DNA) and three plastid regions (matK, trnL intron, and the trnL-F intergenic spacer) producing highly resolved cladograms (Williams et al. 2001a; Williams et al. 2001b). Members of the Oncidiinae occupy a wide variety of epiphytic sites, from large limbs that are exposed in the canopy of tropical forests, to dense shades composed of small axes of lianas, shrubs or twigs occurring at the understory level (Chase 1988; Chase et al. 2005). Leaf morphology is also highly variable, with species showing a gradient from thick-succulent terete or conduplicate leaves to species showing thin conduplicate leaves. Species within

Oncidiinae also show a gradient of CAM expression, from C3 photosynthesis to strong

CAM (Silvera et al. 2005). Ancestral state reconstruction of the occurrence of CAM onto a phylogeny of orchids shows multiple independent origins of CAM with several reversal events (Fig. 3) and correlated divergence of photosynthetic pathways and epiphytism explained by the prevalence of CAM in the low-elevation epiphytes and increased diversification of high-elevation epiphytes (Silvera et al. 2009). Similarly, ancestral state reconstruction of CAM onto a phylogeny of Oncidiinae species suggests at least eight independent origins of CAM and weak CAM within the clade (Fig. 4). 19

Molecular Evolution of CAM

The progression of photosynthetic pathways has consistently been shown to be from C3 ancestors to CAM photosynthesis. However, the genetic changes required for this progression (and reversion) remain unclear. The multiple independent evolutionary origins of CAM and the observation that all of the enzymatic requirements to perform

CAM already exist in most plant cells, particularly stomatal guard cells, might suggest that CAM evolution involves relatively few genetic changes. The available molecular data from C4 cycle enzymes support this view in that none of the C4 or CAM cycle enzymes or corresponding genes are unique to these plants (Westhoff and Gowik 2004).

However, given the large number of anatomical and biochemical requirements for CAM

(Fig. 2) and the complexity of the regulatory changes associated with modulation of stomatal behavior and gene expression patterns associated with CAM (Cushman et al.

2008), we suggest that number of genetic changes are likely to be many.

Molecular markers for studying CAM evolution

The cytosolic enzyme phosphoenolpyruvate carboxylase (EC 4.1.1.31; PEPC) catalyzes the ß-carboxylation of phosphoenolpyruvate, with oxaloacetate and inorganic phosphate as products, and serves various functions in plants (Chollet et al. 1996; Nimmo 2000). In addition to anaplerotic roles in leaves and nonphotosynthetic tissues, PEPC catalyzes the initial fixation of atmospheric CO2 into C4-dicarboxylic acids in CAM and C4 photosynthesis. For the PEPC gene family, non-photosynthetic and photosynthetic isoforms are present in C3, C4 and CAM species. These non-photosynthetic ‘C3 isoforms’ 20

might have served as the starting point for the evolution of the C4 and CAM isogenes. In

C4 plants, key determinants for the evolution of the C4 cycle include duplication of ancestral non-photosynthetic or C3 isogenes, followed by the acquisition of increased mRNA and protein expression, with organ- and cell-type specific expression patterns of the C4 photosynthetic isogenes largely due to transcriptional changes in gene expression

(Furumoto et al. 2000; Westhoff and Gowik 2004). C4 and CAM-specific isoforms of

PEPC are distinguished by their elevated mRNA and protein expression in leaf tissues of

CAM plants. Evidence from comparative analysis of C3, C3-C4 intermediates, and C4

Flaveria species suggests that C4 photosynthetic PEPC isoforms have evolved from ancestral non-photosynthetic or C3 isoforms by gene duplication and acquired distinct kinetic and regulatory properties mediated by discrete amino acid changes (Bläsing et al.

2000; Bläsing et al. 2002; Engelmann et al. 2002, 2003; Westhoff and Gowik 2004).

In CAM plants, an evolutionary progression of gene family changes similar to that described for C4 plants is thought to have occurred from detailed studies of PEPC gene families from C3 photosynthesis, weak CAM and strong CAM species (Fig. 5).

Among CAM species, multiple PEPC isogenes have been described for a given plant, with CAM specific PEPC isoforms exhibiting enhanced mRNA expression relative to the expression of C3 PEPC isoforms (Cushman et al. 1989; Gehrig et al. 1995; Gehrig et al.

2001; Gehrig et al. 2005). A comparison of four Clusia species: a C3 species, two C3-

CAM intermediates with water-deficit stress-inducible CAM, and one strong, constitutive

CAM species, revealed that the ability to conduct nocturnal CO2 fixation was well correlated with PEPC amount and activity (Borland et al. 1998) and PEPC mRNA and 21 protein expression was a major factor underpinning the genotypic capacity for CAM

(Taybi et al. 2004).

Detailed comparisons of PEPC isoforms from C3 photosynthesis, weak CAM and strong CAM species within the Orchidaceae are underway. As in C4 plants, distinct kinetic and regulatory properties are expected to be conferred by discrete amino acid changes in CAM-specific isoforms of PEPC. In addition, CAM isoform genes for PEPC and PEPC kinase appear to have evolved unique expression patterns that are under circadian clock control with expression patterns that are distinct from those expressed in

C3 plants (Taybi et al. 2004; Boxall et al. 2005; Cushman et al. 2008). However, the evolutionary recruitment of gene family members must extend beyond those involved in

C4 acid metabolism and include those genes that control the large reciprocating pools of storage carbohydrates, which can account for up to 20% of the total leaf dry weight within a cell (Winter and Smith 1996; Dodd et al. 2002; Dodd et al. 2003). Some CAM plants can accumulate soluble sugars (e.g., sucrose, glucose, fructose) and polysaccharides (e.g., fructan, galactomannan) in extra-chloroplastic compartments, while other species store both starch within the plastid and glucose (Christopher and

Holtum 1996). Detailed studies have revealed at least eight distinct combinations of malate decarboxylation and carbohydrate storage strategies in CAM plants (Christopher and Holtum 1996). These various carbohydrate accumulation patterns likely reflect the evolutionary history of the species, rather than by carbon flow constraints of the pathway

(Winter and Smith 1996; Winter and Holtum 2002).

Partial nucleotide sequences of PEPC have provided a valuable molecular phylogenetic marker for understanding the evolution of metabolic pathways in which 22

PEPC is involved. PEPC sequences are useful, not only because the gene is ubiquitous in prokaryotes and plants, but also because the marker can provide information about the tissue-specific expression patterns and metabolic roles of specific gene family members

(Gehrig et al. 2001). For example, Gehrig et al. (2001) used expression changes during leaf development to infer potential CAM-related isogenes of PEPC relative to non-CAM isoforms expressed predominantly in non-photosynthetic roots (Gehrig et al. 2001).

However, tissue-specific expression alone is inadequate to infer a CAM-related function.

The relative abundance of each isoform in CAM-performing tissue must be confirmed in order to designate the most abundantly expressed isoform as CAM-specific (Cushman et al. 1989; Gehrig et al. 1995; Gehrig et al. 2001; Gehrig et al. 2005). Interestingly, three leafless orchid species with chloroplast containing CAM-performing aerial roots (Winter et al. 1985), also expressed PEPC isoforms that clustered with PEPC isoforms recovered from CAM-performing leaves of other species, but not with PEPC isoforms from nonphotosynthetic aerial roots (Gehrig et al. 2001). This observation suggests that such

“shootless” species do not make use of the root-inherent isoform for photosynthetic carbon assimilation, but rather express an additional PEPC isoform that conducts the initial fixation of CO2 needed for CAM.

Characterization of PEPC isogenes from Kalanchoë pinnata revealed seven distinct PEPC isogenes: four in leaves and three in roots. Sequence similarity comparisons and distance neighbor-joining calculations separate the seven PEPC isoforms into two clades, one of which contains the three PEPCs found in roots (Gehrig et al. 2005). The second clade contains the four isoforms found in leaves and is divided into two branches, one of which contains two PEPCs most similar to previously described 23

CAM isoforms. Of these two isoforms, however, only one exhibited abundant expression in CAM-performing leaves, but not in very young leaves, which do not exhibit CAM, suggesting this isoform encodes a CAM-specific PEPC. Protein sequence calculations suggest that all isogenes are likely derived from a common ancestor gene, presumably by serial gene duplication events. However, higher plant genomes also encode a gene for a non-phosphorylable type of PEPC that is more closely related to bacterial and algal types of PEPC that serves as a likely candidate for such an ancestral progenitor gene (Sanchez and Cejudo 2003).

Comprehensive examination of PEPC gene families along with phylogenetic tree construction from C3 photosynthesis, weak CAM and strong CAM orchids, indicates that paralogous PEPC genes evolved independently, probably from one or more ancient duplication events, as evidenced by the clustering of closely-related Oncidiinae PEPC sequences (K. Silvera, unpublished data). Abundantly expressed CAM-Ppc genes from

CAM species cluster together and belong to a sister group of weak CAM and C3 most abundant Ppc-genes. The least abundant isoforms from C3, weak CAM and CAM species cluster separately, and presumably belong to Ppc genes involved in anaplerotic function

(Fig. 5).

Circadian clock regulated markers

In addition to PEPC, two other genes have been characterized recently and are likely to be excellent markers for tracing CAM evolution within the context of circadian clock evolution: PEPC kinase and the glucose-6-phosphate/Pi translocator (Gpt2). In CAM plants, PEPC is activated at night via phosphorylation of a Ser residue near the N- 24 terminus rendering the enzyme more sensitive to PEP and the positive effectors glucose-

6-phosphate (Glc-6-P) and triose-P and less sensitive to the allosteric inhibitor, malate

(Chollet et al. 1996; Nimmo 2000). This phosphorylation is carried out by PEPC kinase

(PPCK), a dedicated, calcium-independent Ser/Thr protein kinase, the expression of which is controlled by the circadian clock (Hartwell et al. 1999; Taybi et al. 2000).

However, the circadian regulation of PPCK mRNA abundance can also be regulated by metabolic signals, such as malate accumulation (Borland et al. 1999). Thus, both circadian and metabolic signals appear to modulate PPCK transcript abundance, which in turn regulates PPCK activity, the phosphorylation/activation state of PEPC, and the degree of nocturnal carbon assimilation (Nimmo 2000). In a comparison of four Clusia species: a C3 species, two C3-CAM intermediates, and a strong, constitutive CAM species, the circadian modulation of PPCK mRNA abundance correlated with the performance of CAM and appeared linked to day/night changes in malate and soluble sugar content. However, circadian fluctuations in PPCK mRNA abundance were not evident in the C3, species and one of the C3-CAM intermediates (Taybi et al. 2004).

In a more recent study in the C3-CAM Mesembryanthemum crystallinum, the expression of the glucose-6-phosphate/Pi translocator gene (Gpt2) was undetectable in plants performing C3, but was preferentially expressed in leaves of CAM-induced plants under circadian clock control (Kore-eda et al. 2005). In summary, three major changes appear to have occurred during the evolution of progenitor genes in order to function in

CAM (Fig. 2). First, CAM isoform genes are highly expressed in plants performing

CAM. Secondly, CAM isoform genes appear to have evolved leaf-specific or leaf- 25 preferential expression patterns. Thirdly, CAM isoform genes appear to have evolved expression patterns that are under circadian clock control.

Circadian clock specialization during CAM evolution

In addition to being an important ecophysiological adaptation to limiting water supply,

CAM is also one of the best-characterized physiological rhythms in plants and represents an interesting example of circadian clock specialization (Wilkins 1992; Lüttge 2003;

Wyka et al. 2004). Because all of the enzymatic machinery for CAM is present within a single cell, competing carboxylation reactions by PEPC is regulated, in part, by reversible protein phosphorylation catalyzed by a PEPC kinase under circadian clock control (Hartwell et al. 1999; Taybi et al. 2000). RUBISCO activity is also modulated, with peak activity apparent during mid to late part of the light period (Maxwell et al.

1999; Griffiths et al. 2002). PEPC kinase transcript abundance is inversely correlated with cytosolic malate concentrations suggesting that malate exerts a negative effect on

PEPC kinase gene expression or mRNA stability and appears to override its circadian control (Borland et al. 1999). Circadian control of the large, reciprocating pools of carbohydrates (Dodd et al. 2003) and associated transport activities (Häusler et al. 2000;

Kore-eda et al. 2005) and the distinction between different classes of carbon pools are also critical for the performance of CAM (Borland and Dodd 2002).

Comparison of the steady-state mRNA abundance patterns of seven circadian clock components in the facultative CAM plant M. crystallinum operating in either C3 or

CAM mode, suggest that the central clock is very similar to that in Arabidopsis and it is not perturbed by development or salinity stress (Boxall et al. 2005). However, different 26 clock components can be used in different ways to alter clock outputs. Evidence for this comes from the observations that ZEITLUPE (ZTL), a gene that does not exhibit an oscillating expression pattern of mRNA abundance in Arabidopsis, does in M. crystallinum. Furthermore, the circadian abundance profile of McZTL exhibits a more prolonged period of expression and McTOC1 (TIMING OF CAB EXPRESSION 1) exhibits increased mRNA abundance following CAM induction. Additional support for alterations in the circadian clock control outputs in M. cystallinum comes from oligonucleotide-based microarray experiments that document that the shift from C3 to

CAM is accompanied by shifts in the phase at which peak expression occurs (Cushman et al. 2008). A large proportion (70%) of Arabidopsis genes that exhibit circadian fluctuations in transcript abundance also respond to environmental stress (i.e. low temperature, salt, and drought; Kreps et al. 2002). Such rhythmic expression of stress- adaptive genes may prepare the plant to better withstand a stress or exploit a limiting resource (Eriksson and Millar 2003). Given that water deficit stress is likely to be one of the driving forces behind CAM evolution (Raven and Spicer 1996), ancestral C3 progenitors of CAM plants may have evolved clocks which exerted pervasive control over metabolism as a means of maintaining metabolic homeostasis under stressful environments (Borland and Taybi 2004).

ACKNOWLEDGMENTS

This work was supported by funding from the U.S. Environmental Protection Agency under the Greater Research Opportunities Graduate Program (Agreement no. MA

91685201 to K.S.); National Science Foundation NSF IOB-0543659 (to J.C.C.); and 27

Smithsonian Tropical Research Institute (to K.W.). We are indebted to Kirah Van Sickle

(Animania, LLC) for assistance with figure preparation and to J. Andrew C. Smith

(Oxford University) for his contributions to Table 2. EPA has not formally reviewed this publication. The views expressed in this publication are solely those of the authors and the EPA does not endorse any products or commercial services mentioned in this publication. This publication was also made possible by NIH Grant Number P20 RR-

016464 from the INBRE Program of the National Center for Research Resources through its support of the Nevada Genomics, Proteomics and Bioinformatics Centers.

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Table 1. !13C values, nocturnal fluctuation in titratable acidity, and leaf traits from 173 orchid species. Titratable acidity and leaf traits are represented by the average ± standard deviation from 86 C3 species, 45 CAM species and 42 weak CAM species. SLA signifies Specific Leaf Area (area per unit dry mass); FM/DM signifies the ratio of fresh mass to dry mass. Values in parentheses represent the fold-change in titratable acidity and corresponding leaf traits in C3 photosynthesis compared with weak CAM or weak CAM compared with strong CAM.

C3 Weak CAM Strong CAM !13C range (‰) –32.0 to –22.0 –32.0 to –22.0 –21.9 to –10.0 Titratable Acidity 2.0±3.2 12.2±9.5 (+6.1) 75.8±60.2 (+6.21) (!H+) Leaf thickness (mm) 0.5±0.4 0.5±0.3 (0) 2.03±2.01 (+4.06) SLA (cm2 g-1) 167±78 145±59 (–0.8) 72±31 (–2.01) FM/DM 6.1±2.1 6±1.8 (0) 10.5±4.1 (+1.75)

Table 2. Taxonomic distribution of CAM plants, including family, order and exemplar species. The list has been updated upon that of Smith and Winter (1996), and Holtum et al. 2007. Family nomenclature follows those provided by the Angiosperm Phylogeny website (Stevens 2006). Major groups are represented in capital letters.

Family Order Examples LYCOPODIOPHYTA Isoetaceae Isoetales Isoetes (quillworts)

PTERIDOPHYTA Polypodiaceae Polypodiales Pyrrosia Vittariaceae Polypodiales Vittaria

MAGNOLIOPHYTA Agavaceae=Asparagaceae Asparagales Agave Aizoaceae Caryophyllales Mesembryanthemum Alismataceae Alismatales Sagittaria Anacampserotaceae Caryophyllales Grahamia Araceae Alismatales Zamioculcas Asphodelaceae=Xanthorrhoeaceae Asparagales Aloe, Haworthia Apiaceae Apiales aquatic Lilaeopsis Apocynaceae Gentianales Pachypodium Asteraceae Asterales Senecio Bromeliaceae Poales Aechmea, Tillandsia Commelinaceae Commelinales Callisia Cactaceae Caryophyllales Cactus Clusiaceae Malpighiales Clusia Crassulaceae Saxifragales Crassula, Kalanchoë Cucurbitaceae Cucurbitales Xerosicyos Didiereaceae Caryophyllales Didierea Euphorbiaceae Malpighiales Euphorbia, Monadenium Geraniaceae Geraniales Sarcocaulon Gesneriaceae Lamiales Codonanthe Hydrocharitaceae Alismatales Vallisneria Lamiaceae Lamiales Plectranthus Montiaceae Caryophyllales Calandrinia Orchidaceae Asparagales Oncidium, Phalaenopsis Oxalidaceae Oxalidales Oxalis Passifloraceae Malpighiales Adenia Piperaceae Piperales Peperomia Plantaginaceae Lamiales Littorella Portulacaceae Caryophyllales Portulaca, Portulacaria Rubiaceae Gentianales Myrmecodia Ruscaceae=Asparagaceae Asparagales Sansevieria, Dracaena Vitaceae Vitales Cissus Welwitschiaceae Gnetales Welwitschia Zamiaceae Cycadales Dioon

Figure 1. CAM metabolite fluxes during nighttime (left panel) and daytime (right panel). During nighttime (CAM phase I), while stomata are open, atmospheric CO2 enters the cell and is fixed as bicarbonate by phosphoenolpyruvate carboxylase (PEP carboxylase) leading to the formation of oxaloacetate and malate, which is accumulated and stored as malic acid in the vacuole. During daytime (CAM phases II-IV), while stomata are closed, malic acid exits the vacuole and malate is decarboxylated. CO2 released is then refixed by RUBISCO through the Calvin cycle within the chloroplast. Pyruvate serves as a substrate of the gluconeogenic pathway where carbohydrates are regenerated and stored. Figure modified from Taiz and Ziegler (2003) Plant Physiology. 3rd Edition.

Figure 2. Major requirements hypothesized for the evolution of CAM along the evolutionary progression from C3 photosynthesis to weak CAM and strong CAM indicated by color transitions from green to orange and red, respectively. The relative contribution of nocturnal CO2 uptake increases as indicated by increased dark shading and height of triangle. Fluctuations in titratable acidity increase due to diel fluctuations in organic acid accumulation and reciprocal fluctuations in storage carbohydrates such as starch, glucans, or soluble hexoses as indicated by white (daytime) and black (nighttime) arrows demonstrating increasing diel flux dynamics. These fluctuations are accompanied by associated increases in transport activities across the tonoplast (e.g., vacuolar H+-ATPase), mitochondrial, and chloroplast envelope membranes (not shown). Putative expansion in the complexity of gene families due to gene duplication events accompanied by selective recruitment of CAM- specific isogenes is indicated by orange and red lines. Diagnostic indicators of gene recruitment include a progressive increase in leaf-specific or leaf-preferential mRNA and protein expression and overall increased expression of PEPC, decarboxylating (e.g., PEPCK or NADP+-/NAD+-ME) enzymes and enzymes of both the glycolytic and gluconeogenic pathways indicated by orange and red colored leaves and bars, respectively. CAM evolution is well correlated with increased leaf succulence characterized by mesophyll cells with increased cell size, increased mesophyll tissue and leaf thickness due to large storage vacuoles as indicated by the increased thickness of leave models drawn in side view. Finally, circadian clock control is thought to progress by both shifts in the phase of circadian clock output and increased magnitude of circadian expression patterns, particularly at the level of mRNA expression. Green shaded box, daytime, black shaded box, nighttime. 45

Figure 3. Summary tree for the classification of the family Orchidaceae. Names on the clades represent the five orchid subfamilies where pink shading highlight large speciation events. Circles represents the number of subtribes identified within each subfamily. Red circles represent subtribes in which CAM is present. White circles represent subtribes in which CAM is absent. The percentage of subtribes with CAM is given in each subfamilies. Tree is redrawn from Silvera et al. (2009).

Figure 4. Summary tree for the appearance of weak and strong CAM among Oncidiinae species. Major clades are represented by capital letters and clade relationships are derived from Williams et al. (2001). Photosynthetic pathway based on !13C and titratable acidity was mapped onto this cladogram using MCCLADE V.4.08 with ACCTRAN optimization using Pescatorea (Zygopetalinae) as outgroup. Those lineages with C3 photosynthesis are represented by white bars; those with CAM photosynthesis are shown in black; those with weak CAM are shown in gray, and those with equivocal results are represented by striped bars. 47

Figure 5. Evolution of PEPC genes (Ppc) in the Orchidaceae. CAM-Ppc genes from CAM species cluster together, and belong to a sister group of weak CAM and C3 most abundant Ppc-genes. The least abundant isoforms from C3, weak CAM and CAM species cluster separately, and presumably represent Ppc genes having anaplerotic as opposed to photosynthetic functions (K. Silvera, unpublished data).

Chapter II

Crassulacean acid metabolism and epiphytism linked to adaptive

radiations in the Orchidaceae

Katia Silvera*, Louis S. Santiago, John C. Cushman, and Klaus Winter

Department of Biochemistry & Molecular Biology MS 200, University of Nevada, Reno,

NV 89557-0200 (KS, JCC); 2Smithsonian Tropical Research Institute, P.O. Box 0843-

03092, Balboa, Ancón, Republic of Panama (KS, KW); and Department of Botany &

Plant Sciences, University of California, 2150 Batchelor Hall, Riverside, CA 92521

(LSS).

Corresponding author;

Katia Silvera

E-mail address: [email protected]

Published in April 2009, Plant Physiology Vol 149: 1838-1847

ABSTRACT

Species of the large family Orchidaceae display a spectacular array of adaptations and rapid speciations that are linked to several innovative features including specialized pollination syndromes, colonization of epiphytic habitats, and the presence of crassulacean acid metabolism (CAM), a water conserving photosynthetic pathway. To better understand the role of CAM and epiphytism in the evolutionary expansion of tropical orchids, we sampled leaf carbon isotopic composition of 1,103 species native to

Panama and Costa Rica, performed character state reconstruction and phylogenetic trait analysis of CAM and epiphytism, and related strong CAM, present in 10% of species surveyed, with climatic variables and the evolution of epiphytism in tropical regions.

Altitude was the most important predictor of photosynthetic pathway when all environmental variables were taken into account, with CAM being most prevalent at low altitudes. By creating integrated orchid trees to reconstruct ancestral character states, we found that C3 photosynthesis is the ancestral state and that CAM has evolved at least 10 independent times with several reversals. A large CAM radiation event within the

Epidendroideae, the most species-rich epiphytic clade of any known plant group, is linked to a Tertiary species radiation that originated 65 million years ago. Our study shows that parallel evolution of CAM is present among subfamilies of orchids, and correlated divergence between photosynthetic pathways and epiphytism can be explained by the prevalence of CAM in low elevation epiphytes and rapid speciation of high- elevation epiphytes in the Neotropics, contributing to the astounding diversity in the

Orchidaceae. 50

INTRODUCTION

Crassulacean acid metabolism (CAM) is a taxonomically widespread photosynthetic pathway that has evolved in plants of CO2- and water-limited environments, including tropical forest canopies with intermittent or seasonal water availability, hot semi-arid regions, and some aquatic environments. The CAM pathway is characterized by the temporal separation of carbon fixation between nocturnal CO2 fixation by phosphoenolpyruvate carboxylase (PEPC) in the cytosol and daytime decarboxylation of organic acids to release CO2 that is then refixed by ribulose-1,5-biphosphate carboxylase/oxygenase (Rubisco) in the chloroplast (Ting, 1985). CAM photosynthesis is found in approximately 7% of vascular plant species from 36 families (Smith and Winter,

1996; Holtum et al., 2007). About 10% of all vascular plant species are estimated to be epiphytes (Benzing, 1989), many of which exhibit CAM (Lüttge, 2004). CAM vascular epiphytes (mostly orchids and bromeliads) are an important component of the biomass and species richness of tropical forest canopies (Benzing, 1987; Lüttge, 2004; Zotz,

2004). Bromeliads, aroids and orchids, are three of the very few flowering plant lineages that were able to successfully colonize epiphytic niches (Gentry and Dodson, 1987). Yet, orchids are particularly species-rich relative to these other epiphytic groups (Gravendeel et al., 2004), making Orchidaceae a prime subject for understanding mechanisms of evolutionary radiation and diversification. About 72% of orchid species are estimated to be epiphytic (Benzing, 1989; Gravendeel et al., 2004) with the majority of these being restricted to tropical regions. Tropical forest canopies are rich in epiphytic CAM plant diversity (Benzing, 1987; Winter and Smith, 1996; Lüttge, 2004). CAM has been found in 62% and 26% of epiphytic orchid species in Australian and New Guinean rainforests, 51 respectively (Winter et al., 1983; Earnshaw et al., 1987), 42% of orchid species in a moist lowland forest site in Panama (Zotz and Ziegler, 1997), and up to 100% of the epiphytic flora in a Mexican dry forest (Mooney et al., 1989). The abundance of CAM species in such habitats is related to limited water availability. Within a single site, the percentage of CAM epiphytes increases with canopy height, from 7% in the forest understory, to

25% at intermediate heights, and to 50% in exposed canopy sites (Zotz and Ziegler,

1997). CAM species are also found in contrasting habitats such as very arid and very moist sites, which provides evidence of the biochemical flexibility of this photosynthetic adaptation (Dodd et al., 2002). Several researchers have postulated that in addition to

CAM, mechanisms such as production of dust-like seeds capable of long-distance dispersal, germination associations with mycorrhizae, absorptive velamentous photosynthetic root tissue capable of rapid water uptake, and reproductive features that promote specialized pollination syndromes, have contributed to diversification of epiphytic orchids (Benzing, 1987; Gravendeel et al., 2004; Peakall, 2007; Mondragon-

Palomino and Theissen, 2008). Whether CAM is linked to epiphytic diversification and species radiations throughout evolutionary time remains poorly understood.

Orchid systematics is now at an advanced stage allowing ancestral state reconstruction and correlated evolution analysis of key adaptive traits within the context of a highly resolved phylogeny (Chase et al., 2003). We used whole leaf tissue carbon

13 isotopic composition (! C) as a rapid screening method to establish whether CO2 assimilation occurs predominantly by strong CAM or C3 photosynthesis. Strong CAM

13 and C3 photosynthetic tissues exhibit distinct ! C values because CAM plants are

13 enriched in C relative to C3 plants (Rundel et al., 1979; Winter, 1979; Winter et al., 52

1983; Kluge et al., 1991; Zotz and Ziegler, 1997; Crayn et al., 2001; Crayn et al., 2004;

Zotz, 2004; Silvera et al., 2005), whereas weak CAM species show overlapping !13C values with C3 species, and measurable changes in day/night titratable acidity (Silvera et al., 2005). We present data on the prevalence of CAM in the largest family of angiosperms using the regional, highly diversified, and well-described tropical orchid flora of Panama and Costa Rica (Dressler, 1993; Hammel et al., 2003), and evaluate its relationship with climate and its role in the mega-diversification of epiphytes. We used available phylogenetic information to determine the distribution of photosynthetic pathways among orchid species and relate these evolutionary patterns to climate-driven habitat preferences and epiphytism. We also used phylogenetic comparative methods to demonstrate statistically the evolutionary association between epiphytism and photosynthetic pathways and evaluate the role of CAM in species radiations in the

Orchidaceae.

MATERIAL AND METHODS

Site description

Panama and Costa Rica are equatorial tropical countries located between 7-11º N and 77-

80º W. The Panamanian Isthmus serves as a land bridge between North and South

America, and fosters a rich intermixture of plant and animal life that has migrated between the continents. The two principal mountain ranges, the Tabasará Mountains

(Cordillera Central) in the west, and the Cordillera de San Blas in the east, divide the country into Atlantic and Pacific facing slopes. Costa Rica is similarly divided into

Caribbean and Pacific slopes by the Cordillera Central and the Cordillera de Talamanca. 53

Two distinct regional seasons driven by latitudinal movement of the inter-tropical convergence zone produce a dry season from December-May with shorter, less intense dry seasons in sites with greater annual precipitation (Dressler, 1993). Annual precipitation is generally greater on the Caribbean coast (1,500-5,550 mm per year) than on the Pacific coast (1,140-2,290 mm per year). We present data from sites ranging in mean annual precipitation between 1,652-5,204 mm per year, mean annual temperatures ranging from 14.9-27.6 ºC, mean relative humidity ranging from 79.7-89.2%, and altitude from sea level to 3,290 m.

Carbon isotope analysis

Small fragments (2-5 mg) of leaf tissue were collected from a combination of 12 live specimens from Selby Botanical Gardens and 1,091 species from five Herbaria: Missouri

Botanical Gardens Herbarium (MOBOT), Marie Selby Botanical Gardens Herbarium

(SEL), University of Florida Herbarium (FLAS), University of Panama Herbarium

(Universidad de Panama), and the Smithsonian Tropical Research Institute Herbarium

(SCZ). Leaf samples were analyzed for carbon stable isotopic composition (!13C) at the

Center for Stable Isotope Biogeochemistry at the University of California, Berkeley, with an isotope ratio mass spectrometer (Finnigan-MAT Delta Plus XT). 13C/12C ratios were calculated relative to the Pee Dee belemnite standard using the relationship:

!13C (‰) = [(13C/12C in sample)/(13C/12C in standard) – 1] X 1000.

13 We categorized species into C3 or CAM based on leaf ! C values commonly observed

13 for C3 photosynthesis plants ranging from –33 to –22‰, and with ! C values typical of

CAM plants ranging from –22 to –12‰ (Ehleringer and Osmond, 1989). 54

Climatic Data

We used altitude and geographic coordinates to determine climate variables for each herbarium specimen sampled for carbon isotope analysis. For each specimen we recorded species name, herbarium code and collection number, growth form (epiphytic, terrestrial or both), collection location, elevation, and coordinates. We used average altitude for species in which a range was given. For species with missing entries, verbal descriptions of location were converted to altitude and coordinates using the online Global Gazette

Version 2.0 (http://www.fallingrain.com/world/) for Panama and Costa Rica. Coordinate information for each specimen, was then used to generate mean annual temperature, mean relative humidity, mean annual precipitation, and mean annual daily temperature range (DTR), using the Climatic Research Unit application CRU_CL_2.0 (New et al.

2002) (http://www.cru.uea.ac.uk/cru/data/tmc.htm). From DTR we calculated mean monthly minimum (Tmin) and maximum temperature (Tmax). Simple and multiple regression analyses were used to determine relationships between climatic variables and

!13C using SAS statistics software (version 9.1.3).

Orchid phylogeny and species nomenclature

We constructed two tree hypotheses for the Orchidaceae (genera and subtribes) using

Phylomatic 2 (Webb et al., 2008) and Mesquite v. 2.5 (Maddison and Maddison, 2008).

The relationship among genera and overall tree topology followed the strict consensus tree resulting from a combined analysis of psaB + rbcL gene sequences (Cameron, 2004).

The relationship among subtribes and overall topology followed the summary tree of the revised classification of Orchidaceae (Chase et al., 2003). Resulting subtribe and genera 55 trees retained existing polytomies. The overall tree topology differs between these two trees because of different taxon and regions sampled within each publication. Because of the large size of the Orchidaceae, relationships among genera were further expanded in the genera tree using recent publications and nomenclatural changes to the Oncidiinae

(Williams et al., 2001; Williams et al., 2001; Dressler and Williams, 2003), Laeliinae

(Higgins, 1997; van den Berg et al., 2000; Dressler, 2002; Dressler and Higgins, 2003),

Pleurothallidinae (Pridgeon and Chase, 2001; Pridgeon et al., 2001; Luer, 2004),

Cranichideae (Salazar et al., 2003), and Maxillarieae (Whitten et al., 2000; Ojeda et al.,

2005; Whitten et al., 2007). Overall, our nomenclature is consistent with recent publications on nomenclatural changes presented above, the Field Guide to the Orchids of Costa Rica and Panama (Dressler, 1993), and the Royal Botanic Gardens Kew World

Checklist of Monocotyledons nomenclatural database and associated authority files

(http://apps.kew.org/wcsp/home.do).

Phylogenetic and statistical analysis

Presence and absence of CAM were traced onto the genera tree based on !13C (this study) and titratable acidity measurements (Silvera et al., 2005). Genera were labeled as CAM if at least one species within the genus was known to conduct CAM. Character state reconstruction was performed by maximum likelihood in Mesquite v. 2.5 (Maddison and

Maddison, 2008), using a marginal probability reconstruction with Asymmetrical parameter Markov-k model of evolution with an estimated forward rate of 0.1020 and backward rate of 0.2007 corresponding to gain or loss of CAM, respectively. Root state frequency in this analysis was set to be the same as equilibrium, and we specified a bias 56 of less than 1 indicating that backward changes (loss of CAM) are more probable than forward changes. This model was chosen over parsimony or Mk1 model (gain and loss of

CAM are equally weighted), based on the prior assumption that the gain of CAM is less probable because it requires the appearance and coordination of multiple genes, whereas the loss of CAM could result from a single loss-of-function mutation. This prior assumption would likely avoid the false rejection of Dollo’s law when reconstructing a character (Goldberg and Igic, 2008). Because both trees were the result of compilation of different phylogenies, branch lengths were not available and analyses were performed with branch lengths set as equal to 1. Presence and absence of CAM based on !13C, was traced onto the orchid subtribe tree using the same reconstruction method as the orchid genera tree. Subtribes were labeled as CAM if at least one genus within the subtribe was known to conduct CAM based on the information presented in this paper and previously published information (Smith and Winter, 1996; Silvera et al., 2005). The results of analyses using two different assumptions about the probability of each character state at the root (state frequency same as equilibrium and state frequency equal) gave nearly identical outputs for both genera and subtribe trees. The results were displayed as likelihood states reported as the proportion of total likelihood and represented as pie diagrams for the subtribe tree, and as proportional to weights in each branch for the genera tree.

To analyze the relationship between photosynthetic pathway and epiphytism, leaf

!13C was coded for each genus as the mean for all species of that genus. Epiphytism was coded for each species as 0 for terrestrial and 1 for epiphytic, and for the genus as the mean for all species of that genus. For simplicity, both epiphytes and lithophytes were 57 considered as epiphytes. Trait correlation between leaf !13C and epiphytism was evaluated with Pearson product-moment correlation coefficients across orchid genera.

The same genus-level coding was used to perform correlated divergence analysis between !13C and epiphytism in the Analysis of Traits (AOT) module in PHYLOCOM

(Felsenstein, 1985; Webb et al., 2008) using the genera tree for Orchidaceae. Branch lengths were estimated by calibrating the tree for orchid genera at a single point (first bifurcation, 76 MYA) using age estimate for the Orchidaceae from Ramirez et al.

(Ramirez et al., 2007). We then used the branch length adjustment utility in Phylocom

(BLADJ) to perform age interpolations for undated nodes and produce phylogenetic distances. Node ages were treated as approximations. AOT calculates standardized divergence of extant taxa and handles polytomies by ranking species based on the value of the independent variable where the median is used to create two groups (Pagel, 1992).

Correlated divergence analysis was performed by constructing a 0-intercept linear regression between divergence in trait 1 (!13C) and divergence in trait 2 (epiphytism) among sister groups throughout the genera tree, so that divergence width was based on differences in traits between descendent lines (Garland et al., 1993; Garland and Díaz-

Uriarte, 1999; Webb et al., 2008). We used divergence width instead of independent contrasts as a measure of absolute trait radiation because the standard deviation

(divergence width) can be used when polytomies are present in the phylogeny (Moles et al., 2005; Beaulieu et al., 2007). We also used contribution index as a measure of the amount of present-day variation within a trait that is attributed to a focal clade (Moles et al., 2005).

RESULTS

A total of 1,022 Panamanian and Costa Rican orchid species from 147 genera covering

802 sites, and a total of 1,103 species-site combinations were analyzed. Our study covered 4% of the total number of orchids (24,910 species) as described by Chase et al.

(2003), and about 68% of the total estimated number of species from Panama and Costa

Rica (ca. 1,500 species) as described by Dressler (1993). Variation in whole-tissue !13C values ranged from -37.1‰ to -11.4‰, with an overall mean of -27.7‰. The isotopic values among orchid species showed a bimodal distribution with the majority of species showing values near -28‰ indicative of the C3 photosynthetic pathway, and a smaller mode near -16‰ indicative of the CAM pathway. We found that 924 species (90%)

13 belong to the cluster of mainly C3 photosynthesis and had ! C values more negative than

-22‰, whereas 98 species (10%) belong to the cluster of mainly CAM and had !13C values less negative than -22‰.

The percentage of CAM species showed a steady decline with increasing altitude with the largest number of CAM species at sites from 0-500 m altitude, with no strong

CAM species being observed above 2400 m (Fig. 1A). The number of orchid species was lowest at high elevations (3,000 to 3,500 m, Fig. 1B), with the greatest number of species at mid-elevation (1000-1500 m Fig. 1B). Based on bivariate regression, !13C was most strongly related to daily temperature range (DTR, R2 = 0.176, P # 0.0001, Fig. 2), mean annual rainfall (R2 = 0.07, P = 0.0392), altitude (R2 = 0.05, P # 0.0001), mean maximum

2 2 temperature (Tmax, R = 0.05, P # 0.0001), and mean minimum temperature (Tmin, R =

0.05, P # 0.0001), but was not significantly related to mean monthly relative humidity

(RH, R2 = 0.0004, P = 0.8549). Multiple regression analysis showed that when all 59 variables were taken into account, !13C was most strongly related to altitude (P =

0.0013), followed by RH (P = 0.0065), Tmax (P = 0.0157), Tmin (P = 0.0156), and DTR (P

= 0.0164), but was not significantly related to mean annual rainfall. The best fit multiple regression model for the effects of environmental variables on !13C included altitude,

2 RH, Tmax, Tmin, and DTR (F = 14.47, R = 0.0709, P # 0.0001, df = 882).

Our phylogenetic analyses support multiple origins of CAM within the orchid family (Fig. 3, Fig. 4). CAM is present in 43% of orchid subtribes worldwide (23 out of

53 subtribes show CAM, Fig. 4), and has evolved at least 10 independent times among

Neotropical orchid genera, with two large CAM radiation events, one within the subtribe

Oncidiinae, and a second one within the subtribe Laeliinae, both belonging to the large epiphytic subfamily Epidendroideae (Fig. 3). Across the entire Orchidaceae family, maximum-likelihood estimates suggest that C3 photosynthesis is the ancestral state and that CAM has evolved multiple times, with a large radiation event in the Epidendroid group (20 out of 33 subtribes show CAM, Fig. 4). Our data also indicate the possibility of several reversal events within the subtribes Oncidiinae and Laeliinae (Fig. 3). However, more carbon isotope sampling in these clades is required in order to confirm the presence or absence of CAM in species within respective genera. The majority of species in this study are epiphytic (88%) compared to terrestrial (10.5%), with a low percentage showing both growth forms (1.5%). When using maximum likelihood to trace epiphytism as a character state across the orchid phylogeny, we found that the terrestrial habit is the ancestral state within tropical orchids and similar to CAM, the epiphytic habit is derived

(data not shown). 60

Across all genera a positive relationship between the presence of epiphytism and

!13C was observed (Fig. 5A, r = 0.30; p < 0.0001), explained by the increased use of

CAM among epiphytes. Similar to the intergeneric relationship, we found a significant positive relationship between epiphytism and !13C using divergence analysis (Fig. 5B, r =

0.34; p < 0.0001), indicating correlated evolution of photosynthetic pathway properties

13 associated with less discrimination against CO2 during photosynthesis, and the epiphytic habit. Correlated divergence analysis indicates that the positive cross-genera relationship between epiphytism and !13C is driven by radiation events that occurred deep within the phylogenetic genera tree. Node 68 (Fig. 3), belonging to the subtribe

Oncidiinae, showed the highest contribution index value due to a prominent split between

CAM genera and C3 genera within closely-related species. All species of 10 out of 16 genera within this node are CAM and epiphytes (Macroclinium, Notylia, Ionopsis,

Scelochilus, Comparettia, Rodriguezia, Leochilus, Plectophora, Goniochilus, Trizeus).

Similarly, clade Oeceoclades-Eulophia and clade Vanilla-Cleistes (Fig. 3) showed significantly high divergence width and contribution index values. Each of these radiations contributed to the observed positive significant cross-genera divergence.

DISCUSSION

The challenge presented by the Orchidaceae is to understand how this group evolved with the large array of adaptive characteristics that has allowed a multitude of species-rich radiations and colonization of diverse terrestrial ecosystems worldwide. Our data demonstrate patterns of CAM evolution across the Orchidaceae, including multiple independent origins of CAM, several reversals indicating the evolutionary flexibility of 61

CAM (Fig. 3 and Fig. 4), and parallel evolution of CAM across subfamilies (Fig. 4). Our data also indicate that divergences in photosynthetic pathway and epiphytism have been consistently correlated through evolutionary time (Fig. 5), related to the prevalence of

CAM epiphytic species in lower elevations and abundant species diversification of high elevation epiphytes.

The !13C bimodal distribution found in this study is consistent with other studies

(Griffiths and Smith, 1983; Pierce et al., 2002; Holtum et al., 2004; Silvera et al., 2005;

Motomura et al., 2008), showing disruptive selection on C3 photosynthesis and strong

CAM !13C values with very few intermediate !13C values. Strong CAM is present in about 10% of orchids species studied and the largest proportion is distributed at lower altitude. We found a weak relationship between climatic variables and photosynthetic pathway in this study, partly because Panama and Costa Rica are concentrated in a high precipitation region, and partly because the coarse climate variables analyzed do not fully describe the microclimate of epiphytic habitats. Nonetheless, our data demonstrate that

CAM has contributed to exploitation of epiphytic habitats through mid-elevation tropical montane environments. Orchid species richness in our study, with the largest peak around

1000-1500 m of altitude, is consistent with the rule of the mid altitudinal bulge. The mechanism creating this bulge has been explained by the interaction between temperature and water gradients caused by increasing altitude (Whittaker and Niering, 1975; Peet,

1978; Zhao et al., 2005). Higher moisture availability from elevations around 1,000 m are likely to provide a more suitable environment for epiphytic lineages (Gentry and Dodson,

1987; Cardelús et al., 2006), whereas sites above 2,000 m might have reduced canopy height with less habitat for epiphyte colonization. Increases in !13C with increasing 62

altitude among C3 photosynthesis groups (Fig. 2), due to the reduced photosynthetic carbon isotope discrimination at high elevation, independent of CAM (Körner et al.,

1988), can contribute to the !13C – epiphytism relationship. However, previous studies have shown that the presence of weak CAM can also contribute to a second peak of abundance within the C3 photosynthesis isotopic cluster (Winter and Holtum, 2002;

Silvera et al., 2005). Evaluation of the evolutionary role of weak CAM will require extensive in situ measurements of dawn/dusk titratable acidity, an endeavor limited by the accessibility of live specimens in the field. Nevertheless, the contribution of CAM- independent factors in determining the !13C of high elevation epiphytes and the role of weak CAM as an evolutionary reservoir for further CAM radiations in changing environments deserve further study.

We provide evidence that C3 photosynthesis is the ancestral state in the

Orchidaceae, and that CAM has evolved multiple independent times (Fig. 3 and Fig. 4).

There is strong evidence that evolutionary progression of photosynthesis in plants has been from C3 photosynthetic ancestors to derived weak CAM to strong CAM modes

(Pilon-Smits et al., 1996; Crayn et al., 2004), paralleled by progression from terrestrial ancestors to epiphytic growth forms. Phylogenetic tree construction using PEPC sequences have shown that sequences from CAM species within the Oncidiinae clade cluster together, and differ from CAM sequences of species in the Vanillineae clade, providing evidence that these paralogous genes probably arose from one or more ancient duplications, thus adding an additional line of evidence for the multiple origins of CAM

(Silvera et al., unpublished data). A recent study using the genus Cymbidium concluded that weak CAM was the ancestral state and that C3 and strong CAM were derived 63

(Motomura et al., 2008). However, the results were restricted to species within one genus and limit the conclusions that can be applied at the family level. The change from terrestrial to epiphytic habitat is paralleled by a change from monopodial to sympodial growth form, a common feature in epiphytic orchids often associated with pseudobulbs, which provide tolerance to drought stress. Our study also demonstrates parallel evolution of CAM among four of the five subfamilies of orchids (Epidendroideae, Orchidoideae,

Cypripedioideae, and Vanilloideae). The two largest orchid subfamilies, Orchidoideae and Epidendroideae, are considered to have diversified early in the Tertiary (Ramirez et al., 2007). The progressive aridification and declining CO2 concentrations during the

Tertiary (Pearson and Palmer, 2000) are likely factors that have contributed to the large

CAM radiation in Epidendroideae, which has produced the most species-rich epiphytic clades of any known plant group. CAM and epiphytism in orchids are likely to have originated about 65 MYA. The CAM radiation events within the subfamily

Epidendroideae (Fig. 4), and especially within the Neotropical subtribes Oncidiinae and

Laeliinae (Fig. 3), are thus in line with paleo-environmental conditions that favored the evolution of CAM (Monson, 1989; Ehleringer and Monson, 1993; Raven and Spicer,

1996).

Epiphytism in orchids is a pantropical phenomenon and our study shows significant correlation between photosynthetic pathways and epiphytism (Fig. 5), indicating that throughout evolutionary time, divergence in !13C is consistently accompanied by divergence in epiphytism and demonstrating a functional relationship between these traits. Correlated divergence between photosynthetic pathway and epiphytism is likely an important factor contributing to the burst of speciation that 64 occurred in diverse epiphytic orchid clades (subtribe Oncidiinae and Laeliinae, Fig. 3).

These clades are species-rich due to colonization of epiphytic habitats, and species within them were able to colonize wider habitat ranges compared with other plant species, due to the development of CAM as a water-conserving mode of photosynthesis, ultimately contributing to the large floristic diversity found in the Neotropics. Correlated divergence between photosynthetic pathways also indicates that epiphytic orchid species developed

CAM convergently to other epiphytic or terrestrial plants species under the selectional pressure arising from the search for light in rainforest habitats. Our study shows that strong CAM species are restricted to epiphytic habitats, except for Oeceoclades maculata, a terrestrial invasive orchid species native to Africa with recent distribution in the Neotropics from Argentina to Florida (Stern, 1988; Cohen and Ackerman, 2008). The presence of strong CAM in this genus is intriguing and whether or not the presence of

CAM contributes to invasiveness in this species is unknown.

CONCLUSION

This study demonstrates several patterns of CAM evolution across the Orchidaceae, including multiple independent origins of CAM, several reversal events indicating the evolutionary flexibility of CAM, and parallel evolution of CAM across subfamilies.

Divergences in photosynthetic pathway and epiphytism have been consistently correlated through evolutionary time and are related to the prevalence of CAM epiphytic species in lower elevations and abundant species diversification of high elevation epiphytes.

Overall, our study reveals biochemical underpinnings and evolutionary interactions between CAM as a water-saving mode of photosynthesis and colonization of epiphytic 65 habitats that have contributed to some of the most substantial plant speciations known to exist.

ACKNOWLEDGEMENTS

Special thanks go to Dr. Bruce Holst (Selby Botanical Gardens); Kent Perkins, Dr. Mark

Whitten and Dr. Norris Williams (FLAS Herbarium); Dr. Jim Solomon (MOBOT

Herbarium); and Mireya Correa (Univ of Panama Herbarium) for assisting with herbarium collections. We gratefully acknowledge M Whitten, K Neubig, L Endara, the

Santiago Lab, and two anonymous reviewers for comments to improve this paper; Dr.

Todd Dawson and Dr. Stefania Mambelli (UC Berkeley) for assisting with isotopic analysis; Dr. Karen Schlauch (UNR) for statistical advice, Dr. Doug Altshuler for

Mesquite advice; Cristina Milsner and Michael O’Leary for assisting with database entry; and Vanessa Boukili (UC Berkeley) and Becky Albion (UNR) for assistance in the lab.

This work is supported, in part, by funding from the Environmental Protection Agency

(EPA) under the Greater Research Opportunities Graduate Program Assistance

Agreement No. MA 91685201 (to KS); National Science Foundation NSF IOB-0543659

(to JCC); NSF DEB-0706813 (to LSS), NIH Grant P20 RR-016464 from the INBRE

Program of the National Center for Research Resources supporting the Nevada

Genomics, Proteomics and Bioinformatics Center, the Andrew W. Mellon Foundation through the Smithsonian Tropical Research Institute (to KW), and the Nevada

Agricultural Experiment Station as publication NAES 03087114. EPA has not formally reviewed this publication. The views expressed in this publication are solely those of the 66 authors and the EPA does not endorse any products or commercial services mentioned in this publication. 67

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Figure 1. A, Percentage of CAM species as a function of altitude. B, Total number of species as a function of altitude. Black bars represent C3 photosynthesis species, and red bars represent CAM species. 73

Figure 2. Relationship between !13C and climatic variables. Each point represents a species-site combination. Black points represent C3 photosynthesis species, and red points represent CAM species. 74

Figure 3. Orchidaceae tree showing the relationship among 147 tropical genera. Tribes (in boldface) and subtribes are shown. Photosynthetic pathways based !13C (this study) and titratable acidity derived from Silvera et al. (2005) were mapped onto the cladogram using Mesquite version 2.5. Those lineages that show C3 photosynthesis are represented by black lines, and those with CAM are highlighted in red. The arrow indicates node 68, which shows a large contribution index to divergence correlation. These data support the multiple, independent evolutionary origins of CAM. 75

Figure 4. Orchidaceae tree showing the relationship among 53 subtribes. Black vertical lines represent five major lineages recognized as subfamilies. Presence and absence of CAM based on this study and published information derived from Silvera et al. (2005), and Smith and Winter (1996) were mapped onto the cladogram using Mesquite version 2.5. Those lineages that show CAM are depicted in red. The red area within each pie chart indicates the relative support for different ancestor states. These data support the multiple, independent evolutionary origins of CAM. 76

Figure 5. Pairwise correlation between !13C and epiphytism (A), and divergence in !13C and epiphytism among radiations in the tree for orchid genera (B). Epiphytism is represented as a gradient from which 0 represents terrestrial forms (no epiphytism) and 1 represents epiphytic forms. Values in between 0 and 1 represent genera with species that have both growth forms. Black points represent C3 photosynthesis genera, and red represent points represent CAM genera.

Chapter III

Incidence of crassulacean acid metabolism in the Orchidaceae

derived from carbon isotope ratios: a checklist of the flora of

Panama and Costa Rica

Katia Silvera1,2,*, Louis S. Santiago3, John C. Cushman1, and Klaus Winter2

1Biochemistry & Molecular Biology MS 200, University of Nevada, Reno, NV 89557-

0200; 2Smithsonian Tropical Research Institute, P.O. Box 0843-03092, Balboa, Ancón,

Republic of Panama; and 3Botany & Plant Sciences, University of California, 2150

Batchelor Hall, Riverside, CA 92521.

*Corresponding author;

Katia Silvera

E-mail address: [email protected]

Accepted for publication, Botanical Journal of the Linnean Society 78

ABSTRACT

Leaf carbon stable isotopic composition of 1,002 orchid species representing 61% of the total number of orchid species described for Panama and Costa Rica were obtained from herbarium specimens to survey the occurrence of crassulacean acid metabolism (CAM).

Carbon isotopic composition of leaf material showed a bimodal distribution with modes at -28 ‰ indicating C3 photosynthesis, and at -15 ‰, indicating pronounced CAM photosynthesis. Strong CAM was present in 9.5% of species and in 33 of 163 genera studied. Twelve of these genera were not previously known to contain species exhibiting

CAM. A checklist of orchids of Panama and Costa Rica and their !13C values is included, as well as an updated list of all known orchid genera that possess species with the ability to perform CAM.

Keywords: carbon isotopes- Central America- epiphytes- herbarium- orchids- photosynthetic pathway

INTRODUCTION

Crassulacean acid metabolism (CAM) is one of three photosynthetic pathways found in vascular plants for the assimilation of atmospheric CO2. In contrast to C3 and C4 plants,

CAM plants can perform substantial net CO2 uptake from the atmosphere at night, reducing the amount of water lost per unit carbon assimilated (e.g., greater water-use efficiency) (Cushman, 2001; Winter, Aranda & Holtum, 2005; Winter & Smith, 1996).

CAM species are widely distributed throughout semi-arid tropical and subtropical 79 environments, including epiphytic habitats in the humid tropics. Thus far, CAM has been reported for 343 genera in 36 families (Holtum, Winter, Weeks & Sexton, 2007), and about 7% of all vascular plant species are estimated to possess CAM (Holtum, Winter,

Weeks & Sexton, 2007; Smith & Winter, 1996). Improving this estimate requires detailed work in species-rich families with large expected numbers of CAM species, such as the

Orchidaceae (Chase, Cameron, Barrett & Freudenstein, 2003).

Several studies indicate that CAM photosynthesis may be widespread among tropical epiphytic orchids (Earnshaw, Winter, Ziegler, Stichler, Cruttwell, Kerenga,

Cribb, Wood, Croft, Carver & Gunn, 1987; Silvera, Santiago, Cushman & Winter, 2009;

Silvera, Santiago & Winter, 2005; Winter, Wallace, Stocker & Roksandic, 1983). Winter and Smith (2006) anticipated that 50% of tropical epiphytic orchids might perform CAM.

Indeed, a survey of over 200 tropical orchid species showed strong and weak CAM in approximately 100 species (Silvera et al., 2005). In the current study, we targeted the rich orchid flora of Panama and Costa Rica, the 1,640 species of which are well described by

Dressler (1993). Carbon isotopic composition (!13C) of bulk leaf tissue was used as a method for determining the photosynthetic pathway in 1,002 of these species, thus allowing us to distinguish between the presence of strong CAM or C3 photosynthesis.

13 13 This method takes advantage of differences in ! C of CAM and C3 species because ! C values of leaf carbon reflect the proportion of CO2 gained during daytime via C3 photosynthesis and nighttime via the CAM pathway (Ehleringer & Osmond, 1989;

Winter et al., 2005; Winter & Holtum, 2002). However, because several species that have

13 a C3-type ! C can also perform low level CAM activity (i.e., weak CAM), using only stable isotopic measurements to determine photosynthetic pathways can underestimate 80 the number of species capable of performing CAM when used in large surveys (Pierce,

Winter & Griffiths, 2002; Silvera et al., 2005; Winter, Garcia & Holtum, 2008; Winter &

Holtum, 2002). Analyses that combine nocturnal acidification measurements and diel-gas patterns with isotopic composition are often employed to further categorize whether species are C3, weak CAM or strong CAM (Pierce et al., 2002; Silvera et al., 2005). In this study we used stable isotope measurements as a rapid screening method to categorize

13 species as either C3 or strong CAM based on whether ! C values ranged from -33‰ to -

22‰, which is typical of C3 photosynthesis, or from -22‰ to 12‰, which is typical for

CAM plants (Ehleringer & Osmond, 1989). The aims of this study were to update the number of orchid species performing CAM worldwide, provide a framework for further studies, and provide a checklist of orchid genera in which the presence of CAM has been reported.

MATERIAL AND METHODS

SITE DESCRIPTION

Panama and Costa Rica are tropical, Central American countries located between 7º and

11º N, and 77 º to 85º W, and form the narrowest part of the Mesoamerican Isthmus, thus serving as a land bridge between North and South America, and, therefore, fostering a rich intermixture of plant and animal life that has migrated between the continents. This region contains all of the tropical life zones described by Holdridge (Holdridge, 1967), with climatic ranges of mean annual precipitation from approximately 1,140-5,500 mm and mean annual temperature from 14-28 ºC. Much of the regional climatic variation is caused by an elevation range from sea level to 3,820 m, which results in an orographic 81 distribution of precipitation. This diversity of habitats, therefore, covers the complete range of conditions in which tropical orchids are known to occur (Dressler, 1993), including native, urban, agricultural, and disturbed habitats.

CARBON ISOTOPE ANALYSIS

Small fragments (2-5 mg) of leaf tissue were collected from herbarium specimens of

1,002 orchid species encompassing 163 genera at five herbaria: Missouri Botanical

Gardens Herbarium (MO), Marie Selby Botanical Gardens Herbarium (SEL), University of Florida Herbarium (FLAS), University of Panama Herbarium (PMA), and the

Smithsonian Tropical Research Institute Herbarium (SCZ). This sampling includes 61% of the orchid flora of Panama and Costa Rica described by Dressler (1993). Orchid nomenclature followed a combination of Dressler (1993) and the Royal Botanic Gardens

Kew World Checklist of Monocotyledons nomenclatural database and associated authority files (http://apps.kew.org/wcsp/home.do). Leaf samples were analyzed for carbon stable isotopic composition (!13C) at the Center for Stable Isotope

Biogeochemistry, University of California Berkeley, using a continuous flow isotope ratio mass spectrometer (Finnigan-MAT Delta Plus XT). Ratios of 13C/12C were calculated relative to the Pee Dee belemnite standard (Belemnitella americana) using the relationship:

!13C (‰) = [(13C/12C sample)/(13C/12C standard) – 1] · 1000.

Long-term external precision for !13C analyses is ± 0.22‰ when compared to standards. Based on the differential enzyme-mediated discrimination patterns against

13 CO2 during photosynthetic carbon assimilation, CAM and C3 species have different, but 82

13 overlapping values of whole leaf ! C values, such that values observed for C3 plants range from –33 to –22‰, whereas species with !13C values characteristic of CAM plants range from –22 to –12‰ (Ehleringer & Osmond, 1989; Osmond, Allaway, Sutton,

Troughton, Queiroz, Lüttge & Winter, 1973; Pierce et al., 2002; Santiago, Silvera,

Andrade & Dawson, 2005; Silvera et al., 2005).

Because the !13C values of samples can be influenced by leaf development, and

13 because non-photosynthetic tissues in C3 species tend to be enriched in C compared to leaves (Cernusak, Tcherkez, Keitel, Cornwell, Santiago, Knohl, Barbour, Williams,

Reich, Ellsworth, Dawson, Griffiths, Farquhar & Wright, 2009), only mature healthy leaves were sampled and non-photosynthetic tissues were avoided. Similarly, specimens collected from the field were preferred over those grown in greenhouses or artificial

13 conditions, to avoid effects of variable ! C of source CO2 in closed environments.

Variation in !13C values from herbarium specimens of 2-3 individuals of the same species measured in 80 orchid species showed a standard deviation from ± 0.01 to 3.2 for both C3 and strong CAM species (K. Silvera, unpubl. data). Variation within leaves of the sample species can also occur, however, these differences are small relative to !13C variation between C3 and strong CAM leaf tissue. For example, a recent study of the variation in !13C values of multiple leaves from individual trees collected at different times of the year showed standard deviation of ± 0.1 to 0.5 in C3 species (Holtum &

Winter, 2005), and a study of the variation in !13C values of mature tissue of plants cultivated side-by-side showed standard deviation values of ± 0.1 to 0.3 in CAM species

(Winter et al., 2005). 83

We also performed a literature survey to update a previous list (Smith & Winter,

1996) of the total number of orchid genera worldwide in which the presence of CAM has been reported.

RESULTS

Variation of bulk leaf !13C values of orchids from Panama and Costa Rica ranged from -

36.5‰ to -11.4‰, with an average of -27.6‰. The frequency distribution of isotopic values among study species showed a bimodal distribution with the majority of species at values around -28 ‰ typical of C3 photosynthesis, and a second mode at -15 ‰, typical of strong CAM photosynthesis (Fig. 1). Out of 1,002 species, 907 (90.5%) belong to the

13 C3 photosynthesis cluster, whereas 95 (9.5%) were strong CAM species based on ! C measurements.

Table 1 provides detailed voucher information and !13C values of all species studied. Strong CAM was present in 33 of 163 (20%) genera surveyed in this study. All species surveyed within genera Barkeria, Brassavola, Campylocentrum, Cattleya,

Caularthron, Chelyorchis, Comparettia, Encyclia, Goniochilus, Guarianthe, Ionopsis,

Laelia, Leochilus, Macroclinium, Myrmecophila, Notylia, Oeceoclades, Ornithocephalus,

Plectophora, Rodriguezia, Scelochilus, Trichocentrum and Trizeuxis showed strong

CAM. In addition, strong CAM was present in at least one species of the following genera: Elleanthus, Epidendrum, Heterotaxis, Lockhartia, Maxillaria, Myoxanthus,

Oncidium, Pleurothallis, and Vanilla.

The large neotropical genus Epidendrum with approximately 1,200 species worldwide was the most sampled in this study (143 species), and contained the largest 84 proportion of CAM species (24, Table 1). The second most sampled genus, Maxillaria

(110 species), with approximately 550 species worldwide, had only one species with strong CAM. Only 2 of 50 species of another large genus, Pleurothallis, which is currently under extensive revision, showed strong CAM. Scaphyglottis and Sobralia, the fourth and fifth most sampled genera in this study, did not show any strong CAM species.

All species that belonged to the genera Cohniella and Lophiaris, which were previously part of Oncidium, but have now been merged into the genus Trichocentrum (Williams,

Chase, Fulcher & Whitten, 2001), showed strong CAM.

Our checklist represents the first known report of the presence of CAM in 12 genera, bringing the total number of orchid genera known to contain CAM species to 90, considering previous literature reports and recent nomenclatural changes (Table 2).

DISCUSSION

Our finding that the distribution of C3 and CAM photosynthetic pathways among orchids of Panama and Costa Rica shows a bimodal distribution along the complete isotopic range of study species is consistent with previous isotope-based screening campaigns of

CAM taxa (Crayn, Winter & Smith, 2004; Griffiths & Smith, 1983; Holtum, Aranda,

Virgo, Gehrig & Winter, 2004; Neales & Hew, 1975; Pierce et al., 2002; Silvera et al.,

2005; Winter et al., 1983). That the majority of species are clustered around the C3 isotopic range, with a smaller cluster around the CAM isotopic range and very few species with !13C values in between (Fig 1), is indicative of specialization in carbon assimilation pathways, and likely biochemical and/or anatomical limitations associated with extremes within these isotopic ranges (Silvera et al., 2005). This bimodal 85

distribution suggests that strong CAM or C3 photosynthesis is favored over intermediate metabolisms or mixed systems. Species rely on one pathway or the other likely based on available ecological niches. Additionally, intermediate species are likely to exhibit anatomical and physiological limitations associated with the assimilation of equal amounts of carbon through both pathways (Silvera et al., 2005). Whether these intermediate species show mixed strategies or population variation in the field remains unknown. Strong CAM is present in 9.5% of the study species, but this number does not include species in which CAM is weakly expressed. Because !13C was used as a

13 screening method, we cannot account for those species that have ! C characteristic of C3 species, but obtain up to one-third of their carbon through nocturnal CO2 fixation, as is the case of weak CAM and facultative CAM species (Silvera et al., 2005; Winter et al.,

2008; Winter & Holtum, 2002; Winter & Holtum, 2007). Previous work in the

Orchidaceae has shown that roughly 30% of species with isotopic values characteristic of

C3 photosynthesis can show weakly expressed CAM (Silvera et al., 2005). For example, a number of species in the genera Brassia, Sobralia, Cischweinfia, Coryanthes,

Dimerandra, Sobralia and Eriopsis, Mormodes, Peristeria, Scaphyglottis, Trichopilia, and Trigonodium exhibited nocturnal increases in leaf tissue acidity indicative of CAM

13 although ! C values were in the C3 range. The detection of weak CAM requires the study of live specimens in which either titratable acidity and/or net CO2 exchange is measured, and thus, requires more elaborate experimental sampling methods than !13C screening of herbarium specimens (Sinclair, 1983; Winter & Holtum, 2002).

The previous report of 20% strong CAM species among 214 mainly Central

American species studied by Silvera et al. (2005), compared to 9.5% strong CAM species 86 in the current much larger survey, may be explained by the fact that the former study had a bias towards commercially valuable lowland orchid species adapted to drier sites

(Silvera et al., 2009). In contrast, the current, more complete study considered orchids from a much broader range of habitats including high elevation sites where strong CAM species occur less frequently. Nonetheless, if the predicted number of weak CAM species were added to the estimates of strong CAM species in the current study, then the total number of species exhibiting an ability to perform CAM in the Panama-Costa Rica region is expected to increase markedly above the 9.5% established by strong CAM species alone.

In a previous extensive survey of CAM in the neotropical family Bromeliaceae,

Crayn et al. (2004) found strong CAM in 826 of 1,873 bromeliad species (44%), which surpasses the proportion of strong CAM species for orchids of Panama and Costa Rica. In

Panama and Costa Rica most collection sites for orchids receive well over 1,000 mm of annual precipitation and there is little coverage of tropical and subtropical dry forests, in which CAM epiphytes are favored.

Our study represents the largest !13C survey of orchids ever performed and provides a fairly representative picture of the occurrence of strong CAM among the local flora of Panama and Costa Rica. Even so, this survey covers only 4% of the total number of orchid species known worldwide. Further extensive !13C sampling of orchid species from other regions is needed to improve our knowledge of the incidence and functional significance of CAM in this large family of vascular plants.

ACKNOWLEDGMENTS

We are extremely grateful to Dr. Bruce Holst from Marie Selby Botanical Gardens; Kent

Perkins, Dr. Mark Whitten and Dr. Norris Williams from University of Florida

Herbarium; Dr. Jim Solomon from Missouri Botanical Gardens Herbarium; Mireya

Correa from University of Panama Herbarium; and Carmen Galdames from the

Smithsonian Tropical Research Institute Herbarium for access to herbarium collections.

We are indepted to Dr. Todd Dawson and Dr. Stefania Mambelli (UC Berkeley) for assistance with isotopic analysis. This work was supported by funding from the U.S.

Environmental Protection Agency under the Greater Research Opportunities Graduate

Program (Agreement no. MA 91685201 to K.S.); National Science Foundation NSF IOB-

0543659 (to J.C.C.); NSF DEB-0706813 (to L.S.S.); and the Andrew W. Mellon

Foundation through the Smithsonian Tropical Research Institute (to K.W.). EPA has not formally reviewed this publication. The views expressed in this publication are solely those of the authors and the EPA does not endorse any products or commercial services mentioned in this publication.

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Pierce S, Winter K, Griffiths H. 2002. Carbon isotope ratio and the extent of daily CAM use by Bromeliaceae. New Phytologist 156: 75-83. Santiago LS, Silvera K, Andrade JL, Dawson TE. 2005. Use of stable isotopes in tropical biology. Interciencia 30: 536-542. Silvera K, Santiago LS, Cushman JC, Winter K. 2009. Crassulacean Acid Metabolism and Epiphytism Linked to Adaptive Radiations in the Orchidaceae. Plant Physiology 149: 1838-1847. Silvera K, Santiago LS, Winter K. 2005. Distribution of crassulacean acid metabolism in orchids of Panama: evidence of selection for weak and strong modes. Functional Plant Biology 32: 397-407. Sinclair R. 1983. Water relations of tropical epiphytes. Evidence for Crassulacean Acid Metabolism. Journal of Experimental Botany 35: 1-7. Smith JAC, Winter K. 1996. Taxonomic distribution of crassulacean acid metabolism. In: Winter K and Smith JAC, eds. Crassulacean Acid Metabolism: Biochemistry, Ecophysiology and Evolution. Berlin Heidelberg: Springer-Verlag. 427-436. Williams NH, Chase MW, Fulcher T, Whitten WM. 2001. Molecular systematics of the Oncidiinae based on evidence from four DNA sequence regions: expanded circumscriptions of Cyrtochilum, Erycina, Otoglossum, and Trichocentrum and a new genus (Orchidaceae). Lindleyana 162: 113-139. Winter K, Aranda J, Holtum JAM. 2005. Carbon isotope composition and water-use efficiency in plants with Crassulacean acid metabolism. Functional Plant Biology 32: 381-388. Winter K, Garcia M, Holtum JAM. 2008. On the nature of facultative and constitutive CAM: environmental and developmental control of CAM expression during early growth of Clusia, Kalanchoë, and Opuntia. Journal of Experimental Botany 59: 1829-1840. Winter K, Holtum JAM. 2002. How closely do the !13C values of crassulacean acid metabolism plants reflect the proportion of CO2 fixed during day and night? Plant Physiology 129: 1843-1851. Winter K, Holtum JAM. 2007. Environment or development? Lifetime net CO2 exchange and control of the expression of crassulacean acid metabolism in Mesembryanthemum crystallinum. Plant Physiology 143: 98-107. Winter K, Smith JAC. 1996. Crassulacean acid metabolism: current status and perspectives. In: Winter K and Smith JAC, eds. Crassulacean Acid Metabolism: Biochemistry, Ecophysiology and Evolution. Berlin Heidelberg: Springer-Verlag. 389-426. Winter K, Wallace BJ, Stocker GC, Roksandic Z. 1983. Crassulacean acid metabolism in Australian vascular epiphytes and some related species. Oecologia 57: 129- 141.

Figure 1. Frequency of leaf !13C values for 1,002 Panamanian and Costa Rican orchid species. Each bar represents a 1‰ range of !13C values. Samples were collected at the Marie Selby Botanical Gardens Herbarium (SEL), Missouri Botanical Gardens Herbarium (MO), University of Florida Herbarium (FLAS), University of Panama Herbarium (PMA), and Smithsonian Tropical Research Institute Herbarium (SCZ).

SUBFAMILY Tribe Species Herbariu Accession d13C (‰) m number SUBFAMILY VANILLOIDEAE Tribe Pogoniieae Cleistes rosea Lindl. MO 3042034 -28.8 Tribe Vanilloideae Vanilla inodora Schiede (Vanilla pfaviana FLAS 205833 -29.0 Rchb.f.) *Vanilla planifolia Jacks. ex Andrews MO 1986318 -15.3 *Vanilla pompona Schiede MO 2481055 -15.9 *Vanilla trigonocarpa Hoehne (Vanilla pauciflora Dressler) FLAS 168044 -21.7 SUBFAMILY CYPRIPEDIOIDEAE Phragmipedium humboldtii (Warsz. ex Rchb.f.) J.T.Atwood & MO 4272449 -26.6 Dressler Phragmipedium longifolium (Warsz. & Rchb.f.) Rolfe FLAS 202849 -31.7 Phragmipedium warszewiczianum (Rchb.f.) Christenson FLAS 149690 -26.8 Selenipedium chica Rchb.f. & Warsz. SEL 45447 -31.8 SUBFAMILY ORCHIDOIDEAE Tribe Cranichideae Subtribe Goodyerinae Aspidogyne roseoalba (Dressler) Ormerod SEL 67280 -34.6 Aspidogyne tuerckheimii (Schltr.) Garay MO 2601082 -34.8 Erythrodes sp. Blume SEL 66795 -28.3 Goodyera erosa (Ames & C.Schweinf.) Ames SEL 68616 -30.7 Goodyera fimbrilabia Ormerod SEL 68461 -33.8 Goodyera micrantha Schltr. SEL 66719 -36.5 Kreodanthus sarcochilus E.A.Christ. sp. nov. ined. MO 2908235 -31.5 Microchilus calophylla (Rchb.f.) Ormerod [Erythrodes PMA 44668 -34.9 calophyllas (Rchb.f.) Ames] Microchilus nigrescens (Schltr.) Ormerod SEL 68600 -35.6 Microchilus tridax (Rchb.f.) Ormerod FLAS 205754 -34.4 Microchilus vesicifer (Rchb.f.) Ormerod SEL 64831 -35.3 Microchilus whitefoordiae Ormerod SEL 56638 -33.5 Platythelys epidendroides ined. MO 3224443 -33.1 Platythelys maculata (Hook.) Garay MO 2241194 -34.2 Platythelys querceticola (Lindl.) Garay SEL 71119 -34.4 Subtribe Spiranthinae Beloglottis costaricence (Rchb.f.) Schltr. SEL 57543 -33.7 92

Coccineorchis bracteosa (Ames & C.Schweinf.) Garay SEL 75623 -32.3 Coccineorchis cernua (Lindl.) Garay SEL 61512 -31.0 Coccineorchis cristata Szlach. MO 3479970 -29.9 Coccineorchis navarrensis (Ames) Garay MO 3311782 -31.1 Coccineorchis standleyi (Ames) Garay (Stenorrhynchos MO 2606908 -30.9 standleyi Ames) Coccineorchis warszewicziana Szlach.Rutk. & Mytnik MO 3659052 -26.8 Cyclopogon elatus (Sw.) Schltr. MO 4272041 -32.8 Cyclopogon miradorensis Schltr. MO 2938757 -31.5 Cyclopogon plantagineus Schltr. MO 3502879 -30.0 Eurystyles standleyi Ames FLAS 185783 -31.8 Pelexia funckiana (A.Rich. & Galeotti) Schltr. MO 2928674 -34.1 Pelexia smithii (Rchb.f.) Garay SEL Live 1991 -27.1 -0131 Sacoila lanceolata (Aubl.) Garay MO 2012389 -28.1 Sarcoglottis sceptrodes (Rchb.f.) Schltr. SEL 60172 -30.9 (Sarcoglottis hunteriana Schltr.) Sarcoglottis sceptrodes (Rchb.f.) Schltr. SEL Live 2003 -28.1 -0136A Sarcoglottis smithii (Rchb.f.) Schltr. SEL 69065 -29.8 Sarcoglottis woodsonii (L.O.Williams) Garay MO 1172202 -27.0 Stenorrhynchos speciosum (Jacq.) Rich. ex Spreng. MO 3032040 -32.6 Subtribe Cranichidinae Baskervilla colombiana Garay MO 4273395 -34.5 Cranichis reticulata Rchb.f. MO 3271517 -31.7 Cranichis saccata Ames SEL 56635 -30.3 Cranichis wageneri Rchb.f. SEL 68643 -29.6 Gomphichis adnata (Ridl.) Schltr. [Gomphichis costaricensis FLAS 202846 -30.4 (Schltr.) Ames] Gomphichis hetaerioides Schltr. SEL 52169 -30.6 Ponthieva brenesii Schltr. MO 2628583 -30.1 Ponthieva ephippium Rchb.f. MO 1171744 -30.1 Ponthieva formosa Schltr. MO 5345769 -27.9 Ponthieva inaudita Rchb.f. MO 5587748 -31.7 Ponthieva racemosa (Walter) C.Mohr SEL 61047 -29.2 Ponthieva tuerckheimii Schltr. FLAS 180035 -34.2 Prescottia stachyodes (Sw.) Lindl. SEL 61437 -30.5 Pseudocentrum hoffmannii Rchb.f. MO 3878256 -28.3 Pterichis galeata Lindl. SEL 65456 -28.3 Pterichis habenarioides (F.Lehm. & Kraenzl.) Schltr. SEL 1585 -27.9 Solenocentrum costaricense Schltr. MO 2481258 -31.9 Tribe Orchideae Subtribe Orchidinae Habenaria alata Hook. MO 2353007 -29.0 93

Habenaria avicula Schltr. MO 2323571 -34.1 Habenaria clypeata Lindl. SCZ 2195 -28.6 Habenaria dentifera C.Schweinf. MO 5052280 -30.5 Habenaria distans Griseb. SEL 68284 -35.2 Habenaria eustachya Rchb.f. MO 2061019 -31.7 Habenaria lactiflora A.Rich. & Galeotti MO 3532267 -30.1 Habenaria lankesteri Ames SEL 66890 -30.6 Habenaria mediocris Dressler MO 5345789 -27.9 Habenaria monorrhiza (Sw.) Rchb.f. SEL 71389 -33.1 Habenaria petalodes Lindl. MO 1785355 -28.9 Habenaria repens Nutt. SEL 68294 -31.1 Habenaria rodeiensis Barb.Rodr. MO 2937360 -30.7 Habenaria strictissima Rchb.f. PMA 17532 -33.9 Habenaria trifida Kunth SEL 14012 -27.1 Habenaria wercklei Schltr. MO 2323000 -28.4 SUBFAMILY EPIDENDROIDEAE Tribe Neottieae Palmorchis powellii (Ames) C.Schweinf. & Correll MO 5345774 -34.5 Palmorchis silvicola L.O.Williams PMA 17796 -32.3 Palmorchis trilobulata L.O.Williams FLAS 205794 -35.1 Palmorchis trinotata Dressler SEL 15670 -33.6 Tribe Sobralieae Elleanthus aurantiacus (Lindl.) Rchb.f. FLAS no number -27.5 Elleanthus bradeorum Schltr. MO 4302326 -25.6 *Elleanthus capitatus (Poepp. & Endl.) Rchb.f. MO 5325352 -21.7 Elleanthus caricoides Nash FLAS 178179 -29.7 Elleanthus cynarocephalus (Rchb.f.) Rchb.f. MO 5345761 -31.4 Elleanthus fractiflexus Schltr. MO 5345780 -31.2 Elleanthus glaucophyllus Schltr. FLAS 188734 -27.5 Elleanthus graminifolius (Barb.Rodr.) Lojtnant SCZ 2127 -26.4 Elleanthus hymenophorus (Rchb.f.) Rchb.f. FLAS 205748 -30.1 Elleanthus jimenezii (Schltr.) C.Schweinf. MO 5345770 -28.3 Elleanthus lancifolius C.Presl FLAS 178181 -30.5 Elleanthus laxus Schltr. MO 4964566 -26.2 Elleanthus lentii Barringer MO 2481114 -29.5 Elleanthus longibracteatus (Lindl. ex Griseb) Fawc. MO 5161467 -31.2 Elleanthus muscicola Schltr. MO 5345752 -31.9 Elleanthus poiformis Schltr. MO 5345800 -29.8 Elleanthus robustus (Rchb.f.) Rchb.f. MO 5605116 -27.3 Elleanthus scopula Schltr. MO 5324713 -27.5 Elleanthus stolonifer Barringer FLAS no number -29.5 Elleanthus tillandsiodes Barringer PMA 46870 -29.5 94

Elleanthus tonduzii Schltr. FLAS 178176 -25.5 Elleanthus tricallosus Ames & C.Schweinf. FLAS no number -30.5 Elleanthus wercklei Schltr. PMA 32657 -29.8 Sobralia albolutea Dressler FLAS 205812 -24.8 Sobralia allenii L.O.Williams SEL 1714 -26.1 Sobralia amabilis (Rchb.f.) L.O.Williams MO 4304243 -28.6 Sobralia atropubescens Ames & C.Schweinf. SEL 71279 -24.6 Sobralia bletiae Rchb.f. (Sobralia suaveolens Rchb.f.) MO 2072598 -27.0 Sobralia callosa L.O.Williams MO 3594106 -29.2 Sobralia candida (Poepp. & Endl.) Rchb.f. MO 5345810 -30.6 Sobralia carazoi Lank. & Ames SEL 66695 -27.2 Sobralia chrysostoma Dressler MO 5170269 -29.3 Sobralia corazoi Lank. & Ames MO 4649925 -29.2 Sobralia decora Bateman MO 1939722 -31.7 Sobralia doremiliae Dressler MO 4952726 -26.5 Sobralia fragrans Lindl. MO 4952724 -25.2 Sobralia helleri A.D.Hawkes FLAS no number -28.7 Sobralia kerryae Dressler MO 5345793 -29.0 Sobralia labiata Warsz. Rchb.f. SEL 10885 -28.1 Sobralia lancea Garay FLAS 178187 -28.9 Sobralia leucoxantha Rchb.f. MO 5079696 -31.3 Sobralia lindleyana Rchb.f. MO 4270746 -28.4 Sobralia luteola Rolfe FLAS 205819 -31.0 Sobralia macra Schltr. FLAS 178188 -28.1 Sobralia macrophylla Rchb.f. MO 2057755 -28.6 Sobralia mucronata Ames & C.Schweinf. FLAS 205822 -26.1 Sobralia nutans Dressler MO 4270747 -25.9 Sobralia powellii Schltr. FLAS 205823 -30.8 Sobralia quinata Dressler SEL 66722 -30.0 Sobralia undatocarinata C.Schweinf. MO 4951499 -31.6 Sobralia valida Rolfe MO 5345758 -29.1 Sobralia warszewiczii Rchb.f. (Sobralia bradeorum Schltr.) FLAS 205814 -26.1 Sobralia wilsoniana Rolfe MO 1980717 -26.2 Tribe Tropidieae Corymborkis flava (Sw.) Kuntze FLAS 205741 -32.0 Corymborkis forcipigera (Rchb.f. & Warsz.) L.O.Williams SEL 75632 -33.6 Tribe Triphoreae Monophyllorchis microstyloides (Rchb.f.) Garay SEL 68889 -30.7 Psilochilus macrophyllus (Lindl.) Ames MO 3608753 -28.7 Tribe Calypsoeae Govenia ciliilabia Ames & C.Schweinf. MO 3500547 -26.9 Govenia quadriplicata Rchb.f. MO 4242855 -31.1 95

Govenia utriculata (Sw.) Lindl. (Govenia powellii Schltr.) MO 1943608 -27.5 Tribe Epidendreae Chysis violacea Dressler SEL Live 2007 -25.7 -0058A Subtribe Ponerinae Isochilus carnosiflorus Lindl. SEL 71569 -28.8 Isochilus linearis (Jacq.) R.Br. SEL 68053 -27.9 Isochilus major Cham. & Schltdl. SEL 68059 -27.4 Subtribe Bletiinae Bletia campanulata La Llave & Lex. MO 2614940 -23.0 Bletia purpurea (Lam.) DC. SEL 79845 -28.9 Subtribe Pleurothallidinae *Acianthera decipiens (Ames & C.Schweinf.) Pridgeon & SEL 13493 -15.1 M.W.Chase (Pleurothallis decipiens Ames & C.Schweinf.) Acianthera glumacea (Lindl.) Pridgeon & M.W.Chase MO 4273643 -30.0 (Pleurothallis alexandrae Schltr.) Acianthera johnsonii (Ames) Pridgeon & M.W.Chase SEL 13488 -29.1 (Pleurothallis johnsonii Ames) *Acianthera lepidota (L.O.Williams) Pridgeon & M.W.Chase SEL 15513 -14.6 (Pleurothallis lepidota L.O.Williams) *Acianthera lojae (Schltr.) Luer (Pleurothallis citrophila Luer) SEL 15714 -17.1 *Acianthera oscitans (Ames) Pridgeon & M.W.Chase SEL 13524 -17.6 (Pleurothallis oscitans Ames) Acianthera pantasmi (Rchb.f.) Pridgeon & M.W.Chase SEL 57542 -23.9 (Pleurothallis pantasmi Rchb.f.) *Acianthera pubescens (Lindl.) Pridgeon & M.W.Chase MO 4658643 -16.5 (Pleurothallis pubescens Lindl.) *Acianthera sicaria (Lindl.) Pridgeon & M.W.Chase MO 2662908 -20.4 (Pleurothallis alpina Ames) *Acianthera verecunda (Schltr.) Pridgeon & M.W.Chase MO 2049780 -11.7 (Pleurothallis verecunda Schltr.) Acostaea bicornis Luer SEL 52031 -29.7 Acostaea costaricensis Schltr. MO 2338552 -31.0 Acostaea pleurothalloides Schltr. MO 2896003 -29.8 Anathallis barbulata (Lindl.) Pridgeon & M.W.Chase MO 2060988 -28.9 (Pleurothallis abjecta Ames) Anathallis cuspidata (Luer) Pridgeon & M.W.Chase MO 3326498 -28.6 (Pleurothallis cuspidata Luer) Anathallis dolichopus (Schltr.) Pridgeon & M.W.Chase MO 3498723 -33.0 (Pleurothallis dolichopus Schltr.) Anathallis lewisiae (Ames) Solano & Soto Arenas SEL 3040 -27.6 (Pleurothallis lewisae Ames) Anathallis polygonoides (Griseb.) Pridgeon & M.W.Chase SEL 14030 -25.2 (Pleurothallis polygonoides Griseb.) Anathallis sclerophylla (Lindl.) Pridgeon & M.W.Chase SEL 55642 -28.4 (Pleurothallis sclerophylla Lindl.) Barbosella circinata Luer SCZ 13291 -30.0 Barbosella dolichoriza Schltr. MO 5463884 -25.8 Barbosella geminata Luer SEL 67906 -31.3 96

Barbosella orbicularis Luer FLAS no number -30.0 Barbosella prorepens (Rchb.f.) Schltr. MO 5463907 -26.9 Brachionidium calypso Luer MO 3872241 -31.2 Brachionidium cruzae L.O.Williams MO 3220275 -30.7 Brachionidium dressleri Luer MO 2481263 -32.2 Brachionidium filamentosum Luer & Hirtz MO 2928662 -32.1 Brachionidium folsomii Dressler MO 4026942 -32.0 Brachionidium haberi Luer MO 3708380 -31.5 Brachionidium polypodium Luer MO 3493256 -30.9 Brachionidium pusillum Ames & C.Schweinf. MO 4390476 -31.8 Diodonopsis erinacea (Rchb.f.) Pridgeon & M.W.Chase MO 3311771 -31.0 (Masdevallia erinacea Rchb.f.) Diodonopsis pygmaea (Kraenzl.) Pridgeon & M.W.Chase SEL 66552 -33.5 (Masdevallia pygmaea Kraenzl.) Dracula astuta (Rchb.f.) Luer SEL 38195 -28.5 Dracula erythrochaete (Rchb.f.) Luer MO 3152269 -31.7 Dracula gaskelliana (Rchb.f.) Luer SEL 51038 -24.8 Dresslerella hispida (L.O.Williams) Luer PMA 4176 -29.3 (Pleurothallis hispida L.O.Williams) Dresslerella pertusa (Dressler) Luer MO 2107591 -29.3 Dryadella butcheri Luer MO 2136946 -29.6 Dryadella dressleri Luer SEL 52038 -28.1 Dryadella gnoma (Luer) Luer SEL 52054 -31.7 Dryadella guatemalensis (Schltr.) Luer SEL 77210 -29.6 Dryadella odontostele Luer MO 3515525 -31.4 Echinosepala lappiformis (A.H.Heller & L.O.Williams) SEL 13221 -27.1 Pridgeon & M.W.Chase [Myoxanthus lappiformis (A.H.Heller & L.O.Williams) Luer] Echinosepala sempergemmata (Luer) Pridgeon & M.W.Chase MO 2623825 -28.3 [Myoxanthus sempergemmatus (Luer) Luer] Echinosepala uncinata (Fawc.) Pridgeon & M.W.Chase MO 3659007 -31.3 [Myoxanthus uncinatus (Fawc.) Luer] Lepanthes antilocapra Luer & Dressler PMA 54852 -27.7 Lepanthes blepharistes Rchb.f. SEL 38136 -30.3 Lepanthes brunnescens Luer MO 3716824 -31.9 Lepanthes chameleon Ames SEL 50565 -27.8 Lepanthes ciliisepala Schltr. SEL 50547 -31.5 Lepanthes coeloglossa Luer SEL 66774 -31.1 Lepanthes crossota Luer MO 3432797 -29.2 Lepanthes eciliata Schltr. (Lepanthes cascajalensis Ames) SEL 66485 -34.8 Lepanthes elata Rchb.f. PMA 26726 -29.4 Lepanthes eximia Ames SEL 50533 -31.6 Lepanthes grandiflora Ames & C.Schweinf. (Lepanthes PMA 54864 -26.5 mulderae Luer) Lepanthes helleri A.D.Hawkes (Lepanthes comet-halleyi Luer) SEL 65298 -26.8 97

Lepanthes horichii Luer SEL 66481 -32.4 Lepanthes horrida Rchb.f. SEL 38152 -23.2 Lepanthes jimenezii Schltr. SEL 66396 -29.3 Lepanthes lindleyana Oerst. & Rchb.f. SEL 66398 -27.5 Lepanthes maxonii (Schltr.) PMA 54866 -33.0 Lepanthes monteverdensis Luer & R.Escobar SEL 65261 -28.4 Lepanthes myiophora Luer SEL 65257 -30.9 Lepanthes mystax Luer & R.Escobar SEL 51984 -28.3 Lepanthes psyche Luer SEL 52006 -25.7 Lepanthes pygmaea Luer SEL 54355 -28.9 Lepanthes tipulifera Rchb.f. SEL 50519 -29.1 Lepanthes turialvae Rchb.f. SEL 38208 -24.4 Lepanthes wendlandii Rchb.f. SCZ 12438 -31.4 Masdevallia attenuata Rchb.f. SEL 51987 -29.3 Masdevallia calura Rchb.f. SEL 28499 -32.4 Masdevallia chasei Luer SEL 67910 -25.6 Masdevallia chontalensis Rchb.f. MO 2480972 -30.3 Masdevallia collina L.O.Williams SEL 13514 -25.4 Masdevallia cupularis Rchb.f. SEL 28543 -26.8 Masdevallia lata Rchb.f. SEL 58981 -30.8 Masdevallia laucheana J.Fraser SEL 18301 -26.9 Masdevallia livingstoneana Roezl & Rchb.f. MO 1982949 -30.6 Masdevallia molossoides Kraenzl. SEL 71080 -29.6 Masdevallia morenoi Luer SEL 24765 -26.9 Masdevallia nidifica Rchb.f. MO 3201961 -30.5 Masdevallia olmosii Koniger & Sijm PMA 54880 -31.1 Masdevallia picturata Rchb.f. SEL 38156 -28.2 Masdevallia rafaeliana Luer SEL 28521 -31.3 Masdevallia reichenbachiana Endres ex Rchb.f. SEL 18321 -27.3 Masdevallia rolfeana Kraenzl. SEL 80335 -28.0 Masdevallia scabrilinguis Luer SEL 51989 -28.3 Masdevallia schizopetala Kraenzl. SEL 68839 -31.0 Masdevallia striatella Rchb.f. SEL 67909 -30.8 Masdevallia tonduzii Woolward SEL 41162 -24.6 Masdevallia tubuliflora Ames SEL 28381 -32.8 *Myoxanthus colothrix (Luer) Luer MO 2242082 -15.8 Myoxanthus exasperatus (Lindl.) Luer (Pleurothallis SEL 13454 -27.5 peduncularis Lindl.) Myoxanthus hirsuticaulis (Ames & C.Schweinf.) Luer MO 2199648 -29.2 Myoxanthus octomeriae (Schltr.) Luer MO 1208672 -23.5 Myoxanthus scandens (Ames) Luer (Pleurothallis scandens PMA 4273 -27.6 Ames) Myoxanthus speciosus (Luer) Luer MO 2107232 -27.6 98

Myoxanthus trachyclamys (Schltr.) Luer SEL 93277 -28.3 Octomeria costaricensis Schltr. MO 2241388 -31.6 Octomeria graminifolia (L.) R.Br. MO 3586975 -26.1 Phloeophila pelecaniceps (Luer) Pridgeon & M.W.Chase MO 2136945 -24.2 [Luerella pelecaniceps (Luer) Braas] Phloeophila peperomioides (Ames) Garay MO 2601868 -27.2 (Pleurothallis peperomioides Ames) Platystele brenneri Luer MO 5463927 -29.3 Platystele calymma Luer MO 5463931 -30.0 Platystele caudatisepala (C.Schweinf.) Garay MO 5463937 -32.5 Platystele compacta (Ames) Ames MO 2481019 -28.8 Platystele jungermannioides (Schltr.) Garay SEL 66549 -29.6 Platystele lancilabris (Rchb.f.) Schltr. SEL 73330 -32.1 Platystele microtatantha (Schltr.) Garay SEL 66545 -30.3 Platystele ovalifolia (H.Focke) Garay & Dunst. MO 2937293 -33.3 Platystele oxyglossa (Schltr.) Garay MO 2623594 -31.0 Platystele pedicellaris (Schltr.) Garay SEL 66548 -29.2 Platystele stenostachya (Rchb.f.) Garay SEL 79273 -29.3 Platystele taylorii Luer MO 5464003 -29.9 Pleurothallis allenii L.O.Williams MO 3311759 -28.3 Pleurothallis annectens Luer SEL Live 1977 -30.8 -1746A Pleurothallis archicolonae Luer SEL 51977 -30.0 Pleurothallis bivalvis Lindl. (Pleurothallis antonensis MO 2937577 -30.1 L.O.Williams) Pleurothallis bothros Luer SEL 68904 -24.7 Pleurothallis cardiochila L.O.Williams MO 2167250 -28.8 Pleurothallis cardiothallis Rchb.f. SEL 68854 -29.0 Pleurothallis chloroleuca Lindl. (Pleurothallis ventricosa MO 4273588 -28.0 Lindl.) Pleurothallis colossus Kraenzl. ex Kerch MO 3138467 -30.8 Pleurothallis coriacardia Rchb.f. SEL 93279 -30.9 Pleurothallis crescentilabia Ames SEL 70789 -28.0 Pleurothallis crocodiliceps Rchb.f. MO 2606987 -26.5 Pleurothallis cucumeris Luer SEL 29856 -29.0 Pleurothallis dentipetala Rolfe ex Ames MO 4273633 -31.7 Pleurothallis discoidea Lindl. MO 4273433 -26.0 Pleurothallis divaricans Schltr. SEL 15633 -27.6 Pleurothallis dorotheae Luer SEL 51094 -26.8 Pleurothallis dressleri Luer MO 2205136 -29.2 *Pleurothallis ellipsophylla L.O.Williams MO 1227011 -12.2 Pleurothallis emecocaulon Schltr. MO 4658642 -28.0 Pleurothallis eumecocaulon Schltr. MO 2740035 -28.6 Pleurothallis excavata Schltr. (Pleurothallis concaviflora MO 2060987 -28.2 C.Schweinf.) 99

Pleurothallis hemileuca Luer MO 2941700 -25.1 Pleurothallis homalantha Schltr. PMA 25398 -31.7 Pleurothallis isthmica Luer MO 2167251 -29.4 Pleurothallis longipedicellata Ames & C.Schweinf. MO 3432760 -32.7 Pleurothallis loranthophylla Rchb.f. MO 2167252 -27.8 Pleurothallis luctuosa Rchb.f. SEL 51082 -24.9 Pleurothallis mammillata Luer SEL 18360 -27.8 *Pleurothallis nuda (Klotzsch) Rchb.f. (Pleurothallis SEL Live 2002 -21.1 hemirhoda Lindl.) -0146A Pleurothallis oncoglossa Luer SEL 28556 -26.4 Pleurothallis pallida Luer MO 2787483 -30.5 Pleurothallis palliolata Ames MO 3224437 -28.9 Pleurothallis peculiaris Luer MO 2908220 -29.1 Pleurothallis phyllocardia Rchb.f. MO 1934966 -23.4 Pleurothallis phyllocardioides Schltr. MO 4274961 -29.5 Pleurothallis picta Hook MO 4658641 -30.0 Pleurothallis pleurothalloides (Cogn.) Handro MO 3772357 -32.3 Pleurothallis pruinosa Lindl. MO 2999579 -27.2 Pleurothallis radula Luer SEL 66569 -30.6 Pleurothallis rectipetala Ames & C.Schweinf. (Pleurothallis MO 2937272 -29.4 scitula Luer) Pleurothallis rhodoglossa Schltr. MO 2937585 -29.7 Pleurothallis rowleei Ames MO 4273624 -32.4 Pleurothallis rubella Luer MO 4273626 -30.2 Pleurothallis ruscifolia (Jacq.) R.Br. MO 4272349 -29.0 Pleurothallis sanchoi Ames SEL 70961 -28.0 Pleurothallis titan Luer MO 4273442 -26.3 Pleurothallis tonduzii Schltr. SEL 70974 -30.0 Pleurothallis uncinata Fawc. MO 4273417 -30.0 Pleurothallis volcanica Luer MO 2937253 -31.6 Pleurothallopsis tubulosa Lindl. [Restrepiopsis tubulosa SEL 66734 -30.9 (Lindl.) Luer] Pleurothallopsis ujarensis (Rchb.f.) Lindl. [Restrepiopsis SEL 15411 -25.5 ujarensis (Rchb.f.) Luer] Restrepia muscifera (Lindl) Rchb.f. ex Lindl. MO 2241438 -24.1 Restrepia trichoglossa F.Lehm. ex Sander MO 3520961 -31.7 (Restrepia subserrata Schltr.) Restrepiella ophiocephala (Lindl.) Garay & Dunst. SEL Live 2002 -28.6 -0132 Scaphosepalum clavellatum Luer MO 2481134 -28.1 Scaphosepalum microdactylum Rolfe MO 2353021 -29.9 Scaphosepalum pittieri Schltr. PMA 4498 -29.0 Specklinia acrisepala Ames & C.Schweinf. (Pleurothallis MO 4273437 -28.6 acrisepala Ames & C.Schweinf.) Specklinia amparoana (Schltr.) Luer (Pleurothallis amparoana MO 2928621 -27.0 100

Schltr.) Specklinia aristata (Hook.) Pridgeon & M.W.Chase SEL 28484 -29.7 (Pleurothallis aristata Hook.) Specklinia barbae (Schltr.) Luer (Pleurothallis barbae Schltr.) SEL 16875 -31.2 Specklinia brighamii (S.Watson) Pridgeon & M.W.Chase MO 2937332 -31.7 (Pleurothallis brighamii S.Watson) Specklinia cactantha (Luer) Pridgeon & M.W.Chase SEL 3005 -29.9 (Pleurothallis cactantha Luer) Specklinia calyptrostele (Schltr.) Pridgeon & M.W.Chase MO 2060999 -29.8 (Pleurothallis calyptrostele Schltr.) Specklinia condylata (Luer) Pridgeon & M.W.Chase SEL 52013 -30.8 (Pleurothallis condylata Luer) Specklinia convallaria (Schltr.) Luer (Pleurothallis convallaria SEL 13505 -27.4 Schltr.) Specklinia corniculata (Sw.) Steud. [Pleurothallis corniculata MO 2928703 -28.7 (Sw.) Lindl.] Specklinia costaricensis (Rolfe) Pridgeon & M.W.Chase MO 2249598 -27.1 (Pleurothallis costaricensis Rolfe) Specklinia endotrachys (Rchb.f.) Pridgeon & M.W.Chase MO 2635149 -27.8 (Pleurothallis endotrachys Rchb.f.) Specklinia fimbriata Ames & C.Schweinf. (Pleurothallis setosa SEL 70908 -30.2 C.Schweinf.) Specklinia fuegi (Rchb.f.) Solano & Soto Arenas (Pleurothallis SEL 51089 -28.1 fuegi Rchb.f.) Specklinia fulgens (Rchb.f.) Pridgeon & M.W.Chase SEL 66744 -31.7 (Pleurothallis fulgens Rchb.f.) Specklinia glandulosa (Ames) Pridgeon & M.W.Chase MO 2937304 -29.7 (Pleurothallis glandulosa Ames) Specklinia grobyi (Bateman ex Lindl.) F.Barros (Pleurothallis MO 1941061 -27.4 grobyi Bateman ex Lindl.) Specklinia guanacastensis (Ames & C.Schweinf.) Pridgeon & SEL 56628 -30.7 M.W.Chase (Pleurothallis guanacastensis Ames & C.Schweinf.) Specklinia herpestes (Luer) Pridgeon & M.W.Chase SEL 28483 -28.0 (Pleurothallis herpestes Luer) Specklinia lanceola (Sw.) Lindl. (Pleurothallis lateritia SEL 31467 -28.8 Rchb.f.) Specklinia microphylla (A.Rich. & Galeotti) Pridgeon & MO 4273430 -31.2 M.W.Chase (Pleurothallis microphylla A.Rich. & Galeotti) Specklinia recula (Luer) Luer (Pleurothallis recula Luer) SEL 3147 -28.6 Specklinia segregatifolia (Ames & C.Schweinf.) Solano & Soto MO 2999583 -29.4 Arenas (Pleurothallis segregatifolia Ames & C.Schweinf.) Specklinia simmleriana (Rendle) Luer (Pleurothallis periodica SEL 67698 -32.6 Ames) Specklinia strumosa (Ames) Pridgeon & M.W.Chase SEL 71129 -29.0 (Pleurothallis strumosa Ames) Specklinia tribuloides (Sw.) Pridgeon & M.W.Chase SCZ 13265 -27.7 [Pleurothallis tribuloides (Sw.) Lindl.] Specklinia tripterantha (Rchb.f.) Luer (Pleurothallis MO 2914918 -25.7 tripterantha Rchb.f.) Specklinia turrialbae (Luer) Luer (Pleurothallis turrialbae SEL 66885 -32.4 Luer) Specklinia uniflora (Lindl.) Pridgeon & M.W.Chase SEL 24898 -28.1 101

(Pleurothallis uniflora Lindl.) Stelis alajuelensis Pridgeon & M.W.Chase SEL 51171 -28.5 (Pleurothallis ramonensis Schltr.) Stelis alta Pridgeon & M.W.Chase (Pleurothallis grandis SEL 51085 -29.1 Rolfe) Stelis aprica Lindl. SEL 57538 -27.8 Stelis argentata Lindl. MO 5751821 -30.4 Stelis atrorubens L.O.Williams MO 4274955 -32.5 Stelis brunnea (Dressler) Pridgeon & M.W.Chase (Salpistele MO 3106368 -27.3 brunnea Dressler) Stelis butcheri Luer SEL 13718 -28.2 Stelis canae (Ames) Pridgeon & M.W.Chase (Pleurothallis MO 2482054 -32.9 canae Ames) Stelis carnosilabia (A.H.Heller & A.D.Hawkes) Pridgeon & SEL 68353 -26.7 M.W.Chase (Pleurothallis cf. carnosilabia A.H.Heller & A.D.Hawkes) Stelis carpinterae (Schltr.) Pridgeon & M.W.Chase MO 3138486 -29.5 (Pleurothallis carpinterae Schltr.) Stelis ciliaris Lindl. (Stelis fimbriata R.K.Baker) MO 2302031 -30.9 Stelis cresentiicola Schltr. MO 2057880 -31.4 Stelis cylindrata Pridgeon & M.W.Chase (Pleurothallis MO 3432761 -29.2 macrantha L.O.Williams) Stelis deregularis Barb.Rodr. (Pleurothallis deregularis SEL 73592 -26.9 Barb.Rodr.) Stelis despectans Schltr. MO 2623608 -29.7 Stelis dracontea (Luer) Pridgeon & M.W.Chase (Pleurothallis SEL 66575 -28.0 dracontea Luer) Stelis fortunae (Luer & Dressler) Pridgeon & M.W.Chase MO 2937588 -31.8 (Pleurothallis fortunae Luer & Dressler) Stelis gelida (Lindl.) Pridgeon & M.W.Chase (Pleurothallis SEL 57539 -30.8 gelida Lindl.) Stelis gigantea Pridgeon & M.W.Chase MO 3432755 -25.7 (Pleurothallis powellii Schltr.) Stelis gracilis Ames (Stelis panamensis Schltr.) SEL 38792 -28.2 Stelis guttata (Luer) Pridgeon & M.W.Chase (Pleurothallis SEL 2000 -29.2 guttata Luer) Stelis hymenantha Schltr. MO 2241176 -27.2 Stelis immersa (Linden & Rchb.f.) Pridgeon & M.W.Chase SEL 71244 -26.5 (Pleurothallis immersa Linden & Rchb.f.) Stelis imraei (Lindl.) Pridgeon & M.W.Chase (Pleurothallis MO 4658647 -29.5 vaginata Schltr.) Stelis janetiae (Luer) Pridgeon & M.W.Chase (Pleurothallis SEL 51084 -29.5 janetiae Luer) Stelis jimenezii Schltr. (Stelis gratiosa Luer) SEL 41265 -31.8 Stelis lankesteri Ames MO 2999589 -26.9 Stelis macrophylla Kunth (Pleurothallis pittieri Schltr.) SEL 10721 -24.8 Stelis maculata Pridgeon & M.W.Chase (Salpistele lutea SEL 3029 -29.1 Dressler) Stelis maxima Lindl. MO 3595552 -31.8 Stelis megachlamys (Schltr.) Pupulin (Pleurothallis MO 4069555 -25.6 102 tuerckheimii Schltr.) Stelis microchila Schltr. MO 4274940 -28.8 Stelis montana L.O.Williams MO 2144451 -31.4 Stelis morganii (Stelis dressleri Luer) SEL 28514 -28.0 Stelis multirostris (Rchb.f.) Pridgeon & M.W.Chase SEL 70902 -26.0 [Pleurothallis racemiflora (Sw.) Lindl. ex Hook] Stelis mystax (Luer) Pridgeon & M.W.Chase (Pleurothallis MO 2983915 -32.5 mystax Luer) Stelis papillifera (Rolfe) Pridgeon & M.W.Chase (Pleurothallis SEL 71236 -31.8 papillifera Rolfe) Stelis pardipes Rchb.f. MO 5751820 -29.4 Stelis parvula Lindl. SEL 68156 -30.7 Stelis pompalis (Ames) Pridgeon & M.W.Chase (Pleurothallis MO 5309890 -27.8 pompalis Ames) Stelis powellii Schltr. MO 2923133 -30.6 Stelis purpurascens A.Rich. & Galeotti SEL 16788 -26.7 Stelis purpurea (Ruiz & Pav) Willd. SEL 56497 -30.5 Stelis segoviensis (Rchb.f.) Pridgeon & M.W.Chase SEL 33107 -27.2 (Pleurothallis wercklei Schltr.) Stelis semperflorens Luer SEL 38813 -30.5 Stelis simplex (Ames & C.Schweinf.) Pridgeon & M.W.Chase SEL 71233 -27.4 (Pleurothallis simplex Ames & C.Schweinf.) Stelis spathulata Poepp. & Endl. MO 3593455 -31.9 Stelis storkii Ames MO 4274953 -29.0 Stelis superbiens Lindl. MO 2941697 -29.4 Stelis thymochila (Luer) Pridgeon & M.W.Chase (Pleurothallis MO 3432763 -30.3 thymochila Luer) Stelis transversalis Ames SEL 38814 -31.1 Stelis umbelliformis Hespenh. & Dressler SEL 35739 -29.7 Stelis vestita Ames MO 2481246 -27.1 Stelis williamsii Ames SEL 11171 -27.4 Trichosalpinx arbuscula (Lindl.) Luer MO 2637405 -27.3 Trichosalpinx blaisdellii (S.Watson) Luer SEL 54595 -29.0 Trichosalpinx carinilabia (Luer) Luer MO 3432787 -28.4 Trichosalpinx cedralensis (Ames) Luer MO 2149613 -27.5 Trichosalpinx ciliaris (Lindl.) Luer SEL 18356 -28.5 Trichosalpinx dura (Lindl.) Luer (Pleurothallis foliata Griseb.) MO 3131353 -28.2 Trichosalpinx memor (Rchb.f.) Luer MO 2926925 -23.8 Trichosalpinx orbicularis (Lindl.) Luer MO 2635146 -25.7 Trichosalpinx pergrata (Ames) Luer MO 2623595 -33.5 Trichosalpinx rotundata (C.Schweinf.) Dressler MO 4593874 -27.2 Trichosalpinx tantilla (Luer) Luer MO 2937349 -28.8 Trisetella dressleri (Luer) Luer MO 2060985 -31.0 Trisetella triaristella (Rchb.f.) Luer SEL 50944 -28.6 Trisetella triglochin (Rchb.f.) Luer SEL 66543 -33.9 103

Zootrophion dayanum (Rchb.f.) Luer SEL Live 2002 -25.5 -0130A Zootrophion endresianum (Kraenzl.) Luer SEL 29843 -29.5 Subtribe Laeliinae Acrorchis roseola Dressler MO 3479948 -27.8 Arpophyllum giganteum Hartw. ex Lindl. FLAS 205734 -25.1 *Barkeria lindleyana Bateman ex Lindl. SEL 68061 -18.3 *Brassavola nodosa (L.) Lindl. FLAS 205736 -14.2 *Cattleya dowiana Bateman SEL 12473 -12.7 *Caularthron bilamellatum (Rchb.f.) R.E.Schult. SCZ 2142 -14.3 Dimerandra emarginata (G.Mey) Hoehne SEL 65609 -27.5 Dimerandra isthmii Schltr. MO 2146310 -27.0 *Encyclia amanda (Ames) Dressler MO 5345792 -16.2 *Encyclia ceratistes (Lindl.) Schltr. SEL 68224 -15.0 *Encyclia cordigera (Kunth) Dressler MO 3398419 -15.0 *Encyclia gravida (Lindl.) Schltr. SCZ 2134 -16.5 *Encyclia stellata (Lindl.) Schltr. SEL 74287 -18.2 Epidendrum acrostigma Hágsater & García-Cruz MO 5046291 -27.1 Epidendrum adnatum Ames & C.Schweinf. SEL 56620 -31.3 Epidendrum alfaroi Ames & C.Schweinf. SEL 65934 -28.8 Epidendrum allenii L.O.Williams MO 3201936 -28.7 *Epidendrum amparoana Schltr. MO 2928619 -17.0 Epidendrum anceps Jacq. SEL Live 2004 -23.3 -0032A Epidendrum anoglossoides Ames & C.Schweinf. SEL 53592 -28.9 Epidendrum anoglossum Schltr. SEL 64569 -26.3 Epidendrum antonense Hágsater MO 2940045 -29.2 Epidendrum barbae Rchb.f. SEL 66926 -28.9 Epidendrum barbeyanum Kraenzl. PMA 27026 -24.1 *Epidendrum baumannianum Schltr. MO 3273717 -17.9 Epidendrum bilobatum Ames SEL 80007 -23.8 Epidendrum bisulcatum Ames MO 2604919 -29.5 *Epidendrum cardiophorum Schltr. PMA 8578 -12.6 Epidendrum carpophorum Barb.Rodr. MO 2903062 -28.6 Epidendrum centropetalum Rchb.f. [Oerstedella centropetala SEL 64014 -28.2 (Rchb.f.) Rchb.f.] *Epidendrum chogoncolonchence Hágsater & Dodson SEL Live 2006 -16.3 -0062 *Epidendrum ciliare L. MO 3877024 -12.4 Epidendrum cirrhochilum F.Lehm. & Kraenzl. MO 2896001 -30.2 Epidendrum cocleense Ames, F.T.Hubb & C.Schwienf. MO 5779467 -24.0 Epidendrum cordiforme C.Schweinf. SEL 51207 -27.7 *Epidendrum coriifolium Lindl. (Epidendrum fuscopurpureum FLAS 140579 -20.4 Schltr.) 104

*Epidendrum coronatum Ruiz & Pav. MO 2782358 -18.7 Epidendrum corrifolium Lindl. PMA 17936 -22.1 Epidendrum cresentiloba Ames [Oerstedella cresentiloba MO 3887635 -30.5 (Ames) Hágsater] Epidendrum criniferum Rchb.f. SEL Live 1974 -27.4 -0030-054B Epidendrum cryptanthum L.O.Williams SEL 61978 -31.8 Epidendrum dentilobum Ames PMA 54879 -26.5 Epidendrum difforme Jacq. PMA 26356 -25.9 Epidendrum eburneum Rchb.f. MO 3492205 -26.5 Epidendrum ellipsophyllum L.O.Williams MO 3289595 -34.0 Epidendrum endresii Rchb.f. [Oerstedella endresii (Rchb.f.) MO 4251244 -30.2 Hágsater] Epidendrum exasperatum Rchb.f. MO 5781131 -29.2 Epidendrum exile Ames MO 2937322 -28.2 Epidendrum firmum Rchb.f. SEL 69032 -25.3 Epidendrum flexicaule Schltr. SEL 66961 -33.0 Epidendrum flexuosissimum C.Schweinf. MO 3271515 -28.6 Epidendrum folsomii Hágsater & E.Santiago MO 2937618 -26.8 Epidendrum fortunae Hágsater & Dressler MO 2928625 -30.4 Epidendrum fuscinun (Dressler) Hágsater (Oerstedella fuscina MO 3493362 -28.4 Dressler) *Epidendrum cf. galeochilum Hágsater & Dressler MO 4302604 -21.5 Epidendrum gibbosum L.O.Williams SEL 61540 -29.3 *Epidendrum glumibracteum Rchb.f. SEL 68265 -18.8 Epidendrum goniorhachis Schltr. SEL 51179 -28.8 Epidendrum hunterianum Schltr. MO 4302609 -24.8 *Epidendrum ibaguense Kunth in F.W.H. von Humboldt, PMA 19147 -15.6 A.J.A.Bonpland & C.S.Kunth (Epidendrum decipiens Lindl.) *Epidendrum imatophyllum Lindl. SCZ 2171 -15.7 Epidendrum incomptum Rchb.f. MO 5779473 -29.5 Epidendrum insolatum Barringer PMA 26749 -26.0 Epidendrum insulanum Schltr. SEL 16873 -23.8 Epidendrum intermixtum Ames & C.Schweinf. SEL 56434 -27.4 Epidendrum isomerum Schltr. FLAS 83823 -27.4 Epidendrum isthmii Schltr. MO 3496953 -29.6 Epidendrum jefeallenii Hágsater & García-Cruz MO 5779463 -28.8 Epidendrum jefestigma Hágsater & García-Cruz MO 4302495 -27.2 Epidendrum lacustre Lindl. MO 3431736 -27.8 Epidendrum lancilabium Schltr. SEL 38433 -28.3 Epidendrum lankesteri Ames SEL 66929 -32.2 Epidendrum laucheanum Bonhof ex Rolfe SEL 64616 -30.2 Epidendrum lechleri Rchb.f. SEL 66962 -27.0 *Epidendrum lockhartioides Schltr. MO 4305160 -17.4 105

Epidendrum longibracteatum Hágsater & García-Cruz MO 5779481 -28.8 *Epidendrum macroclinium Hook. SEL 54601 -20.5 Epidendrum maduroi Hágsater & García-Cruz MO 4304250 -29.2 Epidendrum magnibracteatum Ames (Epidendrum palmense SEL 64833 -27.4 Ames) Epidendrum mantisreligiosae Hágsater MO 2937236 -25.4 Epidendrum microdendron Rchb.f. SEL 66951 -28.5 Epidendrum miserrimum Rchb.f. MO 2937377 -30.0 Epidendrum mora-retanae Hágsater SEL Live 2003 -27.0 -0298A Epidendrum muscicola Schltr. FLAS 185780 -28.4 Epidendrum myodes Rchb.f. MO 5781126 -29.8 Epidendrum nocturnum Jacq. MO 4298834 -24.0 Epidendrum notabile Schltr. MO 3431746 -31.6 Epidendrum nutantirhachis Ames & C.Schweinf. MO 5781122 -26.0 Epidendrum octomerioides Schltr. SEL 57544 -25.8 Epidendrum odontochilum Hágsater MO 4305124 -25.3 *Epidendrum oerstedii Rchb.f. MO 4336238 -16.0 Epidendrum oxyglossum Schltr. MO 5779461 -25.0 Epidendrum pajitense C.Schweinf. PMA 454 -25.4 Epidendrum pallens Rchb.f. MO 2481241 -32.4 Epidendrum palmidium Hágsater MO 3289619 -25.9 Epidendrum panamense Schltr. MO 5779456 -33.4 Epidendrum paniculatum Ruiz & Pav. FLAS 107426 -28.4 Epidendrum pansamalae Schltr. [Oerstedella pansamalae SEL 17468 -30.8 (Schltr.) Hágsater] Epidendrum paraguastigma Hágsater & García-Cruz MO 4298846 -27.9 Epidendrum paranthicum Rchb.f. [Epidanthus paranthicus SCZ 2155 -30.1 (Rchb.f.) L.O.Williams] *Epidendrum parkinsonianum Hook. MO 2896587 -16.5 Epidendrum parviexasperatum Hágsater [Oerstedella SEL 70558 -28.8 parviexasperata Hágsater] Epidendrum paucifolium Schltr. MO 3203974 -29.5 *Epidendrum peperomia Rchb.f. (Epidendrum porpax Rchb.f.) SEL 9709 -19.4 Epidendrum pergameneum (Rchb.f.) MO 5779477 -25.6 Epidendrum phragmites A.H.Heller & L.O.Williams FLAS no number -27.9 Epidendrum phyllocharis Rchb.f. MO 2928742 -30.2 *Epidendrum physodes Rchb.f. SEL 19445 -16.6 [Physinga physodes (Rchb.f) Brieger & Bicalho] Epidendrum piliferum Rchb.f. MO 5779493 -30.0 Epidendrum plagiophyllum Hágsater SEL 79970 -25.7 Epidendrum platystigma Rchb.f. MO 4298832 -29.6 Epidendrum pleurothalloides Hágsater MO 5781117 -29.2 Epidendrum polyanthum Lindl. SEL 76135 -32.0 Epidendrum polychlamys Schltr. PMA 54853 -27.7 106

Epidendrum powellii Schltr. MO 3431747 -28.6 Epidendrum pseudoramosum Schltr. SEL 66960 -30.2 Epidendrum pseudoschumannianum Fowlie [Oerstedella SEL 9175 -24.7 pseudoschumannianum (Fowlie) Hágsater] Epidendrum pseudo-wallisii Schltr. [Oerstedella pseudo- MO 3131340 -30.5 wallisii (Schltr.) Hágsater] Epidendrum pumila Rolfe [Oerstedella pumila (Rolfe) MO 2937253 -27.4 Hágsater] Epidendrum puteum Standl. & L.O.Williams FLAS 83240 -31.1 *Epidendrum radicans Pav. ex Lindl. SEL 66697 -17.0 Epidendrum ramonianum Schltr. SEL 68149 -28.2 Epidendrum ramosum Jacq. MO 5779489 -25.6 Epidendrum repens Cogn. SEL 56837 -28.3 Epidendrum rigidiflorum Schltr. MO 5781118 -27.0 *Epidendrum rigidum Jacq. MO 5161584 -20.2 Epidendrum rugosum Ames MO 5779484 -29.7 Epidendrum sanchoi Ames SCZ 2185 -28.3 Epidendrum sancti-ramoni Kraenzl. MO 2710890 -26.8 Epidendrum santaclarense Ames MO 5779474 -27.9 *Epidendrum scalpelligerum Rchb.f. SEL 12315 -18.8 *Epidendrum schlecterianum Ames MO 3273716 -16.3 Epidendrum schumannianum Schltr. [Oerstedella MO 3131341 -30.4 schumanniana (Schltr.) Hágsater] *Epidendrum sculptum Rchb.f. MO 2937362 -16.8 Epidendrum selaginella Schltr. MO 3138485 -28.9 *Epidendrum stamfordianum Bateman SCZ 2187 -16.3 Epidendrum stevensii Hágsater SEL 51187 -27.1 Epidendrum stolidium Hágsater (Oerstedella ornata Dressler) MO 3111760 -28.3 Epidendrum storkii Ames SEL 65543 -23.6 *Epidendrum strobiliferum Rchb.f. SEL 56811 -21.9 Epidendrum strobiliodes Garay & Dunst. SEL Live 2003 -25.0 -0243A Epidendrum subnutans Ames & C.Schweinf. MO 5779492 -30.5 Epidendrum summerhayesii Hágsater MO 2937511 -27.1 Epidendrum suturatum Hágsater & Dressler MO 2908228 -30.5 Epidendrum talamancanum (J.T.Atwood) Mora-Ret. & García MO 2623817 -28.4 Castro Epidendrum tenuisulcata (Dressler) Hágsater (Neowilliamsia PMA 22869 -28.8 tenuisulcata Dressler) Epidendrum tetraceros Rchb.f. [Oerstedella tetraceros MO 3857308 -29.2 (Rchb.f.) Hágsater] Epidendrum thurstonorum Dodson & Hágsater MO 3432783 -22.2 Epidendrum trachythece Schltr. SEL 66932 -29.4 Epidendrum trialatum Hágsater MO 5779466 -29.5 Epidendrum turialvae Rchb.f. MO 5779468 -30.5 107

Epidendrum veraguasense Hágsater MO 2928579 -30.4 Epidendrum veroscriptum Hágsater SEL 57336 -28.9 (Epidendrum scriptum L.) Epidendrum vincentinum Lindl. SEL 67057 -31.0 Epidendrum wallisii Rchb.f. MO 5779488 -25.9 Epidendrum wercklei Schltr. SEL 64859 -28.3 *Guarianthe patinii (Cogn.) Dressler & W.E.Higgins SEL Live 1991 -18.8 -0251A *Guarianthe skinneri (Bateman) Dressler & W.E.Higgins MO 4964599 -16.6 Homalopetalum pumilio (Rchb.f.) Schltr. MO 4904348 -26.2 Jacquiniella equitantifolia (Ames) Dressler SEL 74233 -26.3 Jacquiniella globosa (Jacq.) Schltr. FLAS 185785 -27.3 Jacquiniella pedunculata Dressler MO 2061010 -27.7 Jacquiniella teretifolia (Sw.) Britton & P.Wilson SEL 13948 -23.5 *Laelia rubescens Lindl. SEL Live 1990 -16.6 -0497 *Myrmecophila tibicinis (Bateman ex Lindl.) Rolfe MO 4956371 -12.9 Nidema boothii (Lindl.) Schltr. SEL 56408 -28.8 Prosthechea abbreviata (Schltr.) W.E.Higgins FLAS 186628 -25.0 Prosthechea aemula (Lindl.) W.E.Higgins SEL 39359 -27.2 Prosthechea brassavolae (Rchb.f.) W.E.Higgins MO 3111765 -28.5 Prosthechea campylostalix (Rchb.f.) W.E.Higgins MO 2481059 -28.7 Prosthechea chacaoensis (Rchb.f.) W.E.Higgins FLAS 205798 -27.3 Prosthechea chimborazoensis (Schltr.) W.E.Higgins MO 4304248 -26.6 Prosthechea cochleata (L.) W.E.Higgins [Encyclia cochleata SEL 75613 -28.5 (L.) Dressler] Prosthechea crassilabia (Poepp. & Endl.) Carnevali & FLAS no number -29.2 I.Ramírez Prosthechea fragrans (Sw.) W.E.Higgins SEL 74286 -27.7 Prosthechea ionocentra (Rchb.f.) W.E.Higgins MO 2897374 -23.9 Prosthechea ionophlebia (Rchb.f.) W.E.Higgins MO 3878255 -26.7 Prosthechea livida (Lindl.) W.E.Higgins MO 3421921 -29.6 Prosthechea ochracea (Lindl.) W.E.Higgins SEL 72340 -24.7 Prosthechea prismatocarpa (Rchb.f.) W.E.Higgins MO 2007425 -24.0 Prosthechea pseudopygmaea (Finet) W.E.Higgins SEL 51185 -26.7 Prosthechea pygmaea (Hook.) W.E.Higgins MO 2938765 -31.0 Prosthechea racemifera (Dressler) W.E.Higgins (Encyclia SEL 56835 -29.2 racemifera Dressler) Prosthechea racemifera (Dressler) W.E.Higgins MO 2584335 -29.0 Prosthechea sima (Dressler) W.E.Higgins MO 2011426 -26.7 Prosthechea spondiada (Rchb.f.) W.E.Higgins SEL 9989 -23.0 Prosthechea vagans (Ames) W.E.Higgins SEL 73347 -24.8 Prosthechea vespa (Vell.) W.E.Higgins MO 2066184 -28.6 Reichenbachanthus cuniculatus (Schltr.) Pabst MO 4904326 -25.1 Reichenbachanthus reflexus (Lindl.) Porto & Brade MO 4893571 -23.5 108

Reichenbachanthus subulatus (Schltr.) Dressler SEL 1592 -25.9 Scaphyglottis acostaei (Schltr.) C.Schweinf. FLAS 205802 -30.1 Scaphyglottis amparoana (Schltr.) Dressler SEL 72167 -27.9 Scaphyglottis arctata (Dressler) B.R.Adams MO 3311767 -27.3 Scaphyglottis behrii (Rchb.f.) Benth. & Hook.f. ex Hemsl. MO 1986378 -28.4 Scaphyglottis bidentata (Lindl.) Dressler (Hexisea bidentata SCZ 12459 -28.9 Lindl.) Scaphyglottis bifida (Rchb.f.) C.Schweinf. SEL 72234 -28.4 Scaphyglottis bilineata (Rchb.f.) Schltr. FLAS 187198 -33.3 Scaphyglottis boliviensis (Rolfe) B.R.Adams FLAS 205803 -27.5 Scaphyglottis chlorantha B.R.Adams MO 2480921 -28.5 Scaphyglottis clavata Dressler MO 2199651 -28.7 Scaphyglottis corallorrhiza (Ames) Ames, F.T.Hubb. & FLAS 187202 -28.0 C.Schweinf. Scaphyglottis coriacea (L.O.Williams) Dressler (Platyglottis MO 2166595 -30.1 coriacea L.O.Williams) Scaphyglottis crurigera (Bateman ex Lindl.) Ames & Correll MO 3152084 -28.9 Scaphyglottis densa (Schltr.) B.R.Adams FLAS 205807 -29.0 Scaphyglottis fusiformis (Griseb.) R.E.Schult. MO 2118428 -26.5 Scaphyglottis gigantea Dressler MO 3772331 -24.9 Scaphyglottis imbricata (Lindl.) Dressler [Hexisea imbricata SEL 61820 -27.3 (Lindl.) Rchb.f.] Scaphyglottis jimenezii Schltr. FLAS 187201 -28.5 Scaphyglottis laevilabium Ames MO 5752973 -30.2 Scaphyglottis lindeniana (A.Rich. & Galeotti) L.O.Williams FLAS 187203 -26.5 Scaphyglottis longicaulis S.Watson MO 2107216 -29.0 Scaphyglottis mesocopis (Endres & Rchb.f.) Benth. & Hook.f. FLAS 205808 -30.3 ex Hemsl. Scaphyglottis micrantha (Lindl.) Ames & Correll SEL 72179 -29.5 Scaphyglottis minutiflora Ames & Correll FLAS 205809 -31.3 Scaphyglottis modesta (Rchb.f.) Schltr. FLAS 187204 -29.7 Scaphyglottis pachybulbon (Schltr.) Dressler SEL 64835 -25.7 Scaphyglottis panamensis B.R.Adams MO 3152092 -28.3 Scaphyglottis prolifera (R.Br.) Cogn. (Scaphyglottis cuneata PMA 4275 -27.8 Schltr.) Scaphyglottis prolifera (R.Br.) Cogn. MO 3479950 -27.1 Scaphyglottis pulchella (Schltr.) L.O.Williams PMA 42863 -28.0 Scaphyglottis punctulata (Rchb.f.) C.Schweinf. MO 3152106 -28.5 Scaphyglottis reflexa Lindl. MO 3152077 -26.8 Scaphyglottis robusta B.R.Adams MO 3152076 -30.5 Scaphyglottis sessiliflora B.R.Adams MO 3303748 -31.7 Scaphyglottis sigmoidea (Ames & C.Schweinf.) B.R.Adams MO 5313064 -32.1 Scaphyglottis spathulata C.Schweinf. MO 2353012 -30.9 Scaphyglottis stellata Lodd. ex Lindl. MO 1882542 -28.1 Tribe Arethuseae 109

Subtribe Arethusinae Arundina graminifolia (D.Don) Hochr. MO 2172184 -26.7 Tribe Malaxideae Crossoglossa blephariglottis (Schltr.) Dressler ex Dodson MO 4901759 -29.2 Crossoglossa elliptica Dressler MO 4273409 -33.7 Crossoglossa eustachys (Schltr.) Dressler ex Dodson SEL 70526 -30.3 Crossoglossa fratrum (Schltr.) Dressler ex Dodson SEL 68232 -32.9 Crossoglossa tenuis Dressler & Dodson MO 3586973 -32.2 Crossoglossa tipuloides (Lindl.) Dodson MO 2481253 -31.0 Liparis elata Lindl. FLAS 205769 -32.8 Liparis nervosa (Thunb.) Lindl. PMA 36657 -31.1 Malaxis blephariglottis (Schltr.) Ames SEL 68647 -32.0 Malaxis brachyrrhynchos (Rchb.f.) Ames MO 2397418 -28.8 Malaxis excavata (Lindl.) Kuntze [Malaxis hastilabia (Rchb.f.) MO 3432768 -30.9 Kuntze] Malaxis pandurata (Schltr.) Ames MO 3311774 -33.3 Malaxis simillima (Rchb.f.) Kuntze MO 2908231 -28.3 Tribe Cymbidieae Subtribe Catasetinae Catasetum bicolor Klotzsch MO 2611986 -26.3 Catasetum viridiflavum Hook. MO 2049501 -27.7 Clowesia warczewitzii (Lindl. & Paxton) Dodson MO 955919 -27.6 Cycnoches egertonianum Bateman PMA 45143 -31.4 Cycnoches warscewiczii Rchb.f. FLAS 181632 -26.3 Cyrtopodium punctatum (L.) Lindl. MO 5310331 -26.8 Dressleria allenii H.G.Hills SEL Live 1976 -25.5 -0056-019A Dressleria suavis (Ames & C.Schweinf.) Dodson SEL Live 1990 -25.8 -0738A Dressleria dilecta (Rchb.f.) Dodson SEL 65792 -25.4 Dressleria eburnea (Rolfe) Dodson SEL 77420 -25.3 Dressleria helleri Dodson FLAS 181504 -27.3 Subtribe Eulophiinae Eulophia alta (L.) Fawc. & Rendle MO 2048564 -29.6 *Oeceoclades maculata (Lindl.) Lindl. SCZ 12455 -16.7 Subtribe Eriopsidinae Eriopsis biloba Lindl. MO 3332444 -28.5 Subtribe Oncidiinae Ada allenii (L.O.Williams ex C.Schweinf.) N.H.Williams FLAS 218961 -28.8 Ada chlorops (Endres & Rchb.f.) N.H.Williams MO 4272326 -26.8 Aspasia epidendroides Lindl. SEL 60171 -26.8 Aspasia principissa Rchb.f. MO 2105212 -30.0 Brassia arcuigera Rchb.f. SEL 32535 -28.0 110

Brassia caudata (L.) Lindl. SEL 56655 -24.2 Brassia gireoudiana Rchb.f. & Warsz. SEL 12427 -26.1 Brassia verrucosa Bateman ex Lindl. subsp. verrucosa SEL 53583 -26.1 *Chelyorchis ampliata (Lindl.) Dressler & N.H.Williams MO 2999590 -16.0 Cischweinfia dasyandra (Rchb.f.) Dressler & N.H.Williams MO 4901772 -28.0 Cischweinfia pusilla (C.Schweinf.) Dressler & N.H.Williams FLAS 218932 -30.2 *Comparettia falcata Poepp. & Endl. SEL 13957 -14.3 Cuitlauzina convallarioides (Schltr.) Dressler & N.H.Williams MO 2914922 -30.4 Cuitlauzina egertonii (Lindl.) Dressler & N.H.Williams MO 2167465 -23.2 Cyrtochiloides ochmatochila (Rchb.f.) N.H.Williams & MO 3714801 -24.6 M.W.Chase (Oncidium ochmatochilum Rchb.f.) Cyrtochiloides panduriformis (Ames & C.Schweinf.) SEL 64717 -28.8 N.H.Williams & M.W.Chase (Oncidium panduriforme Kunth) Cyrtochilum maduroi (Dressler) Senghas FLAS 219011 -27.3 (Oncidium maduroi Dressler) Erycina crista-galli (Rchb.f.) N.H.Williams & M.W.Chase MO 3399696 -27.0 [Psygmorchis crista-galli (Rchb.f.) Dodson] Erycina pumilio (Rchb.f.) N.H.Williams & M.W.Chase MO 5175516 -23.2 [Psygmorchis pumilio (Rchb.f.) Dodson & Dressler] Erycina pusilla (L.) N.H.Williams & M.W.Chase [Psygmorchis MO 2928717 -22.8 pusilla (L.) Dodson & Dressler] *Goniochilus leochilinus (Rchb.f.) M.W.Chase SEL 68262 -13.6 *Ionopsis satyrioides (Sw.) Rchb.f. SEL 56626 -15.5 *Ionopsis utricularioides (Sw.) Lindl. SCZ 2197 -12.6 *Leochilus labiatus (Sw.) Kuntze SEL 62746 -15.2 *Leochilus leochilinus (Rchb.f.) Dressler & Williams PMA 46935 -13.3 *Leochilus scriptus (Scheidw.) Rchb.f. SEL 62866 -13.1 *Lockhartia acuta (Lindl.) Rchb.f. SCZ 2200 -21.4 Lockhartia amoena Endres & Rchb.f. SEL 880 -26.1 Lockhartia hercodonta Rchb.f. ex Kraenzl. SEL 9177 -29.4 Lockhartia micrantha Rchb.f. SCZ 2201 -24.9 Lockhartia oerstedii Rchb.f. SEL 57666 -25.0 Lockhartia pittieri Schltr. SCZ 12973 -24.6 *Macroclinium lineare (Ames & C.Schweinf.) Dodson MO 2286436 -14.0 Mesospinidium horichii I.Bock MO 4971080 -29.3 Mesospinidium panamense Garay MO 4971083 -30.1 Mesospinidium warscewiczii Rchb.f. SEL 73416 -31.4 Miltonioides carinifera (Rchb.f.) Senghas & Lückel [Oncidium MO 2941768 -27.7 cariniferum (Rchb.f.) Beer] Miltoniopsis roezlii (Rchb.f.) God.-Leb. MO 2937288 -31.2 Miltoniopsis warszewiczii (Rchb.f.) Garay & Dunst. MO 4272346 -27.8 *Notylia albida Klotzsch FLAS 218934 -17.8 *Notylia pentachne Rchb.f. MO 1979166 -14.7 *Notylia pittieri Schltr. MO 5175874 -15.8 111

Oncidium allenii Dressler MO 2937527 -29.0 Oncidium ansiferum Rchb.f. MO 5325844 -28.7 Oncidium anthocrene Rchb.f. (Oncidium powellii Schltr.) MO 2937237 -27.9 Oncidium bracteatum Warsz. & Rchb.f. SEL 68181 -32.9 Oncidium bryolophotum Rchb.f. MO 3201963 -30.2 Oncidium cheirophorum Rchb.f. MO 2627293 -26.8 Oncidium dichromaticum Rchb.f. (Oncidium cabagrae Schltr.) PMA 49320 -29.7 Oncidium ensatum Lindl. MO 3399703 -27.4 Oncidium exalatum Hágsater MO 2937525 -28.8 Oncidium fuscatum Rchb.f. MO 3398418 -32.3 Oncidium imitans Dressler FLAS 219162 -28.8 Oncidium isthmi Schltr. SEL Live 1996 -24.5 -0169B Oncidium lineoligerum Rchb.f. & Warsz. (Oncidium stenotis MO 5175517 -28.3 Rchb.f.) Oncidium luteum Rolfe MO 3393412 -27.5 Oncidium nebulosum Lindl. MO 3382709 -27.7 (Oncidium klotzschianum Rchb.f.) Oncidium obryzatoides Kraenzl. MO 4364574 -28.6 Oncidium panamense Schltr. FLAS 219026 -28.1 Oncidium parviflorum L.O.Williams MO 4273711 -31.9 *Oncidium polycladium Rchb.f. ex Lindl. MO 3776494 -17.8 Oncidium punctulatum Dressler SEL 60039 -27.9 Oncidium schroederianum (O’Brien) Garay & Stacy MO 3399704 -28.1 Oncidium stenobulbon Kraenzl. SEL Live 1990 -28.4 -0803A Oncidium warszewiczii Rchb.f. MO 4272328 -27.0 Oncidium zelenkoanum Dressler & Pupulin FLAS 218812 -26.8 *Ornithocephalus bicornis Lindl. FLAS 174569 -11.4 *Ornithocephalus cochleariformis C.Schweinf. FLAS 174570 -14.1 *Ornithocephalus cryptantha (C.Schweinf. & P.H.Allen) SEL 6480 -16.0 Toscano & Dressler [Sphyrastylis cryptantha (C.Schweinf. & P.H.Allen) Garay] *Ornithocephalus dressleri (Toscano) Toscano & Dressler FLAS 213134 -19.0 *Ornithocephalus inflexus Lindl. FLAS 174567 -11.8 *Ornithocephalus lankesteri Ames SEL 68075 -12.5 *Ornithocephalus powellii Schltr. FLAS 174573 -12.9 Otoglossum brevifolium (Lindl.) Garay & Dunst. [Otoglossum MO 4273416 -27.8 chiriquense (Rchb.f.) Garay & Dunst.] Otoglossum globuliferum (Kunth) N.H.Williams & M.W.Chase MO 3877029 -27.1 Pachyphyllum crystallinum Lindl. MO 4971074 -30.6 Pachyphyllum hispidulum (Rchb.f.) Garay & Dunst. MO 3872739 -26.9 *Plectrophora alata (Rolfe) Garay FLAS 219042 -17.7 Rhynchostele bictoniensis (Bateman) Soto Arenas & Salazar MO 3393413 -24.8 *Rodriguezia compacta Schltr. MO 1169468 -14.6 112

*Rodriguezia lanceolata Ruiz & Pav. (Rodriguezia secunda MO 1823279 -15.0 Kunth) Rossioglossum schlieperianum (Rchb.f.) Garay & G.C.Kenn SEL 68350 -26.9 *Scelochilus tuerckheimii Schltr. SEL 51193 -13.1 Sigmatostalix abortiva L.O.Williams MO 3244136 -27.4 Sigmatostalix hymenantha Schltr. PMA 49367 -28.2 Sigmatostalix integrilabris Pupulin MO 3716786 -27.2 Sigmatostalix macrobulbon Kraenzl. MO 2894583 -28.8 Sigmatostalix picta Rchb.f. (Sigmatostalix guatemalensis MO 2999573 -28.1 Schltr.) Sigmatostalix picturatissima Kraenzl. MO 1934885 -29.7 Systeloglossum acuminatum Ames & C.Schweinf. SEL 66799 -30.7 Systeloglossum costaricense Schltr. SEL 16892 -28.4 Systeloglossum panamense Dressler & N.H.Williams MO 4971081 -28.1 Telipogon biolleyi Schltr. SEL 11325 -27.3 Telipogon boylei (J.T.Atwood) N.H.Williams & Dressler SEL 68264 -29.6 (Stellilabium boylei J.T.Atwood) Telipogon costaricensis Schltr. FLAS 205827 -25.3 Telipogon storkii Ames & C.Schweinf. SEL 72276 -29.2 Ticoglossum krameri (Rchb.f.) Halb. SEL 1240 -26.6 Ticoglossum oerstedii (Rchb.f.) Halb. FLAS 187206 -32.1 *Trichocentrum ascendens (Lindl.) M.W.Chase & MO 3399702 -16.3 N.H.Williams [Cohniella ascendens (Lindl.) Christenson] *Trichocentrum caloceras Endres & Rchb.f. SEL 15424 -14.4 *Trichocentrum candidum Lindl. [Hybochilus inconspicuous PMA 47037 -13.5 (Kraenzl.) Schltr.] *Trichocentrum capistratum Linden & Rchb.f. SEL 9184 -12.0 *Trichocentrum carthagenense (Jacq.) M.W.Chase & MO 2928704 -17.2 N.H.Williams [Oncidium carthagenense (Jacq.) Sw.] *Trichocentrum cebolleta (Jacq.) M.W.Chase & N.H.Williams SEL 10497 -11.7 [Cohniella cebolleta (Jacq.) Christenson] *Trichocentrum costaricensis Mora-Ret. & Pupulin SEL Live 2003 -14.6 -0184A *Trichocentrum lacerum (Lindl.) ined. [Cohniella nuda SEL 10539 -11.5 (Bateman ex Lindl.) Christenson subsp. stipitatum] *Trichocentrum nuda (Bateman ex Lindl.) M.W.Chase & MO 2937239 -13.8 N.H.Williams [Cohniella nuda (Bateman ex Lindl.) Christenson subsp. nuda] *Trichocentrum pfavii Rchb.f. SEL 52142 -13.6 Trichopilia punicea Dressler & Pupulin SEL Live 2004 -24.3 -0039A Trichopilia similis Dressler FLAS 178506 -29.3 Trichopilia suavis Lindl. & Paxton FLAS 205832 -30.2 Trichopilia turialbae Rchb.f. FLAS 182083 -29.2 *Trizeuxis falcata Lindl. SEL 11352 -12.8 Xylobium colleyi (Bateman ex Lindl.) Rolfe MO 4901770 -29.5 Xylobium elongatum (Lindl. & Paxton) Hemsl. SEL 68043 -32.7 113

Xylobium foveatum (Lindl.) G.Nicholson FLAS 178752 -29.7 Xylobium sulfurinum (Lem.) Schltr. MO 2107251 -27.0 Subtribe Maxillariinae Chrysocycnis tigrina (C.Schweinf.) J.T.Atwood (Maxillaria SEL 69030 -30.8 tigrina C.Schweinf.) Cryptocentrum calcaratum (Schltr.) Schltr. MO 2941738 -22.9 Cryptocentrum gracilipes Schltr. MO 4964577 -28.2 Cryptocentrum flavum Schltr. MO 1934875 -28.1 Cryptocentrum gracillimum Ames & C.Schweinf. SEL 66794 -29.1 Cryptocentrum latifolium Schltr. SEL 196 -25.6 Cryptocentrum lehmannii (Rchb.f.) Garay SEL 53591 -24.2 Cryptocentrum standleyi Ames SEL 14003 -32.6 *Heterotaxis crassifolia Lindl. SEL 59784 -14.7 Heterotaxis discolor (Lodd. ex Lindl.) Ojeda & Carnevali PMA 46822 -29.1 Heterotaxis maleolens (Schltr.) Ojeda & Carnevali SEL 67303 -23.9 *Heterotaxis valenzuelana (A.Rich.) Ojeda & Carnevali SEL 10315 -21.8 [Maxillaria valenzuelana (A.Rich.) Nash] Lycaste campbellii C.Schweinf. SEL Live 1990 -24.6 -0560A Lycaste leucantha (Klotzsch) Lindl. MO 3494248 -29.1 Lycaste macrophylla (Poepp. & Endl.) Lindl. PMA 4386 -27.4 Lycaste powellii Schltr. MO 4305418 -29.1 Lycaste schilleriana Rchb.f. MO 2628639 -30.3 Lycaste tricolor Rchb.f. SEL 9488 -26.1 Maxillaria amplifora C.Schweinf. SEL 84849 -31.1 Maxillaria acervata Rchb.f. MO 4336352 -29.9 Maxillaria aciantha Rchb.f. MO 2117309 -23.2 Maxillaria acostae Schltr. SEL 56446 -28.7 Maxillaria acuminata Lindl. FLAS 212827 -23.7 Maxillaria acutifolia Lindl. MO 2166550 -31.1 Maxillaria adendrobium (Rchb.f.) Dressler SEL 85549 -29.9 Maxillaria adolphi (Schltr.) Ames & Correll SEL 80019 -24.8 Maxillaria alba (Hook.) Lindl. SEL 80303 -29.7 Maxillaria allenii L.O.Williams SEL 56397 -29.5 Maxillaria amesiana hort. FLAS 212837 -28.3 Maxillaria anceps Ames & C.Schweinf. SEL 68148 -24.9 Maxillaria angustisegmenta Ames & C.Schweinf. SEL 56355 -27.3 Maxillaria angustissima Ames, F.T.Hubb. & C.Schweinf. SEL 81901 -29.2 Maxillaria arachnitiflora Ames & C.Schweinf. SEL 56656 -28.8 Maxillaria aurea (Poepp. & Endl.) L.O.Williams MO 2929223 -27.9 Maxillaria bicallosa (Rchb.f.) Garay FLAS 212860 -28.6 Maxillaria biolleyi (Schltr.) L.O.Williams SEL 84898 -26.1 Maxillaria brachybulbon Schltr. MO 2241365 -26.6 114

Maxillaria bracteata (Schltr.) Ames & Correll SEL 65933 -28.6 Maxillaria bradeorum (Schltr.) L.O.Williams SEL 51191 -32.5 Maxillaria brevilabia Ames & Correll SEL 85591 -32.0 Maxillaria burgeri J.T.Atwood SEL 53523 -27.4 Maxillaria caespitifica Rchb.f. SEL 85592 -31.5 Maxillaria calantha Schltr. FLAS 212874 -27.8 Maxillaria camaridii Rchb.f. MO 1941309 -25.4 Maxillaria campanulata C.Schweinf. SEL 79661 -25.7 Maxillaria chartacifolia Ames & C.Schweinf. SEL 67341 -30.0 Maxillaria chionantha J.T.Atwood FLAS 212884 -30.0 Maxillaria concavilabia Ames & Correll SEL 85542 -32.1 Maxillaria conduplicata (Ames & C.Schweinf.) L.O.Williams SEL 85589 -27.3 Maxillaria confusa Ames & C.Schweinf. SEL 73648 -26.0 Maxillaria costaricensis Schltr. SEL 72570 -29.7 Maxillaria cryptobulbon Carnevali & J.T.Atwood SEL 56367 -28.8 *Maxillaria ctenostachys Rchb.f. FLAS 212900 -21.7 Maxillaria cucullata Lindl. (Maxillaria punctostriata Rchb.f.) MO 4909652 -25.7 Maxillaria dendrobioides (Schltr.) L.O.Williams FLAS 212911 -28.2 Maxillaria dichotoma (Schltr.) L.O.Williams SEL 72812 -30.4 Maxillaria diuturna Ames & C.Schweinf. SEL 45578 -28.9 Maxillaria dressleriana Carnevali & J.T.Atwood SEL 79814 -28.6 Maxillaria endresii Rchb.f. SEL 57560 -29.0 Maxillaria exaltata (Kraenzl.) C.Schweinf. SEL 71467 -26.8 Maxillaria falcata Ames & Correll SEL 80324 -28.1 Maxillaria flava Ames, F.T.Hubb & C.Schweinf. SEL 84863 -29.7 Maxillaria fragans J.T.Atwood SEL 84868 -27.7 Maxillaria friedrichsthalii Rchb.f. SEL 66963 -26.0 Maxillaria fuerstenbergiana Schltr. MO 2117310 -26.9 Maxillaria fulgens (Rchb.f.) L.O.Williams SCZ 2208 -26.0 Maxillaria galantha J.T.Atwood & Carnevali MO 3303806 -27.2 Maxillaria gomeziana J.T.Atwood SEL 56309 -30.3 Maxillaria grandiflora (Kunth) Lindl. FLAS 212947 -24.8 Maxillaria hedwigiae Hamer & Dodson SEL 57540 -30.2 Maxillaria hennisiana Schltr. FLAS 212957 -28.7 Maxillaria horichii Senghas SEL 80309 -24.8 Maxillaria inaudita Rchb.f. SEL 70973 -25.8 Maxillaria lankesteri Ames SEL 84858 -28.5 Maxillaria lawrenceana (Rolfe) Garay & Dunst. FLAS 212972 -27.3 Maxillaria lepidota Lindl. FLAS 212976 -29.1 Maxillaria linearifolia Ames & C.Schweinf. SEL 71084 -29.6 Maxillaria longicolumna J.T.Atwood MO 2928592 -28.7 Maxillaria longiloba (Ames & C.Schweinf.) J.T.Atwood SEL 71303 -28.2 115

Maxillaria longipes Lindl. FLAS 212982 -29.7 Maxillaria longipetiolata Ames & C.Schweinf. SEL 66540 -31.0 Maxillaria lutheri J.T.Atwood SEL 84855 -28.4 Maxillaria meridensis Lindl. SEL 67335 -26.0 Maxillaria microphyton Schltr. SEL 84853 -28.7 Maxillaria minor (Schltr.) L.O.Williams SEL 71060 -28.7 Maxillaria monteverdensis J.T.Atwood & Barboza SEL 73752 -29.6 Maxillaria moralesii Carnevali & J.T.Atwood SEL 88715 -32.3 Maxillaria nasuta Rchb.f. SEL 56322 -32.3 Maxillaria neglecta (Schltr.) L.O.Williams MO 1980691 -28.8 Maxillaria nicaraguensis (Hamer & Garay) J.T.Atwood SEL 67919 -25.9 Maxillaria nutans Lindl. FLAS 213012 -28.0 Maxillaria obscura Linden & Rchb.f. SEL 85539 -26.6 Maxillaria oreocharis Schltr. SEL 61433 -27.4 Maxillaria paleata (Rchb.f.) Ames & Correll SEL 56395 -31.1 Maxillaria parviflora (Poepp. & Endl.) Garay SEL 85537 -31.7 Maxillaria pittieri (Ames) L.O.Williams SEL 56318 -29.3 Maxillaria ponerantha Rchb.f. FLAS no number -29.1 Maxillaria porrecta Lindl. SEL 959 -30.3 Maxillaria pseudoneglecta J.T.Atwood SEL 67950 -31.0 Maxillaria quadrata Ames & Correll SEL 84896 -24.2 Maxillaria ramonensis Schltr. SEL 76268 -31.4 Maxillaria ramosa Ruiz & Pav. (Maxillaria repens SEL 67916 -28.2 L.O.Williams) Maxillaria reichenheimiana Endres & Rchb.f. FLAS 213049 -31.0 Maxillaria ringens Rchb.f. SEL 52172 -28.1 Maxillaria rodrigueziana J.T.Atwood MO 2940838 -28.7 Maxillaria rubioi Dodson FLAS 213058 -31.5 Maxillaria rufescens Lindl. MO 2481151 -27.2 Maxillaria sanderiana Rchb.f. ex Sander FLAS 213065 -28.0 Maxillaria sanguinea Rolfe MO 1227035 -27.0 Maxillaria scalariformis J.T.Atwood SEL 83980 -29.0 Maxillaria schlechteriana (C.Schweinf.) J.T.Atwood SEL 71982 -30.8 Maxillaria scorpioidea Kraenzl. SEL 15510 -25.7 Maxillaria setigera Lindl. (Maxillaria callichroma Rchb.f.) FLAS 212875 -28.1 Maxillaria sigmoidea (C.Schweinf.) Ames & Correll SEL 71978 -26.7 Maxillaria suaveolens Barringer FLAS 213078 -26.8 Maxillaria tenuifolia Lindl. FLAS 213082 -25.3 Maxillaria tonduzii (Schltr.) Ames & Correll SEL 79809 -30.2 Maxillaria trilobata Ames & Correll SEL 71468 -30.1 Maxillaria tubercularis J.T.Atwood SEL 79818 -33.2 Maxillaria turkeliae Christenson FLAS 213085 -27.9 116

Maxillaria tutae J.T.Atwood SEL 93558 -28.1 Maxillaria umbratilis L.O.Williams SEL 67690 -32.1 Maxillaria uncata Lindl. SEL 54571 -29.7 Maxillaria vaginalis Rchb.f. SEL 72771 -33.3 Maxillaria valerioi Ames & C.Schweinf. SEL 56324 -27.7 Maxillaria variabilis Bateman ex Lindl. SEL 61459 -29.6 Maxillaria vittariifolia L.O.Williams SEL 1114 -29.0 Maxillaria wercklei (Schltr.) L.O.Williams SEL 52183 -27.9 Mormolyca ringens (Lindl.) Gentil MO 3587134 -24.1 Trigonidium egertonianum Bateman ex Lindl. MO 1976848 -29.0 Trigonidium insigne Rchb.f. ex Benth. & Hook.f. SEL 1918 -26.0 Trigonidium lankesteri Ames FLAS 212813 -25.4 Trigonidium riopalenquense Dodson MO 2937380 -31.2 Subtribe Stanhopeinae Coryanthes maculata Hook. MO 4302493 -27.8 Gongora armeniaca (Lindl.) Rchb.f. SEL Live 2003 -27.2 -0159A Gongora claviodora Dressler FLAS 205756 -29.8 Gongora fulva Lindl. MO 955901 -28.9 Gongora gibba Dressler MO 1934888 -30.0 Gongora horichiana Fowlie FLAS 178784 -30.4 Gongora quinquenervis Ruiz & Pav. FLAS 165503 -25.1 Houlletia odoratissima Linden ex Lindl. & Paxton PMA 4054 -28.1 Houlletia tigrina Linden ex Lindl. SEL Live 2002 -29.9 -0133A Kegeliella atropilosa L.O.Williams & A.H.Heller SEL Live 1994 -27.3 -0371C Polycycnis gratiosa Endres & Rchb.f. MO 2481104 -30.5 Polycycnis muscifera (Lindl. & Paxton) Rchb.f. MO 2926914 -32.5 Polycycnis ornata Garay MO 1941570 -30.8 Stanhopea avicula Dressler FLAS 178208 -28.0 Stanhopea cirrhata Lindl. SEL Live 1975 -25.1 -0023-041B Stanhopea costaricensis Rchb.f. MO 1782468 -28.9 Stanhopea ecornuta Lem. MO 4336209 -30.2 Stanhopea panamensis N.H.Williams & W.M.Whitten SEL Live 1976 -28.0 -0056-016A Stanhopea pulla Rchb.f. FLAS 181688 -23.1 Stanhopea wardii Lodd. ex Lindl. SEL Live 1975 -29.3 -0023-031A Subtribe Coeliopsidinae Coeliopsis hyacinthosma Rchb.f. MO 4273419 -30.2 Peristeria elata Hook. MO 3432826 -27.8 Subtribe Zygopetalinae Euryblema anatonum (Dressler) Dressler [Chondrorhyncha MO 2937369 -33.8 117 anatona (Dressler) Senghas] Chondrorhyncha bicolor Rolfe MO 3714803 -28.9 Chondrorhyncha crassa Dressler MO 4622799 -32.4 Chondrorhyncha reichenbachiana Schtr. FLAS 152213 -26.4 Chondroscaphe atrilinguis Dressler PMA 54869 -31.7 Cochleanthes aromatica (Rchb.f.) R.E.Schult. & Garay SEL 68352 -29.8 Cryptarrhena guatemalensis Schltr. MO 2622834 -29.1 Cryptarrhena lunata R.Br. MO 3496948 -29.0 Dichaea ciliolata Rolfe MO 3484233 -27.2 Dichaea costaricensis Schltr. MO 4893570 -28.8 Dichaea cryptarrhena Rchb.f. ex Kraenzl. SEL 64574 -26.8 Dichaea dammeriana Kraenzl. MO 3114545 -32.4 Dichaea dressleri Folsom MO 2928645 -31.3 Dichaea elliptica Dressler & Folsom MO 3595561 -31.9 Dichaea fragrantissima Folsom MO 2814449 -28.4 Dichaea globosa Dressler & Pupulin FLAS 212758 -25.0 Dichaea hystricina Rchb.f. MO 4893574 -28.3 Dichaea lankesteri Ames FLAS 185771 -28.4 Dichaea morrisii Fawc. & Rendle SEL 68107 -28.9 Dichaea muricatoides Hamer & Garay FLAS 205742 -26.1 Dichaea neglecta Schltr. MO 4964297 -31.0 Dichaea oxyglossa Schltr. SEL 68896 -29.7 Dichaea panamensis Lindl. SEL 56506 -32.9 Dichaea poicillantha Schltr. SEL 63725 -23.2 Dichaea standleyi Ames FLAS 185774 -30.3 Dichaea tenuifolia Schltr. MO 4963040 -32.4 Dichaea trichocarpa (Sw.) Lindl. FLAS 202842 -27.4 Dichaea trulla Rchb.f. MO 2937231 -30.4 Dichaea tuerckheimii Kraenzl. MO 2814172 -29.7 Dichaea violacea Folsom MO 2923116 -29.8 Galeottia grandiflora A.Rich. MO 1947355 -29.4 Huntleya burtii (Endres & Rchb.f.) Rolfe MO 2240379 -27.7 Huntleya fasciata Fowlie SCZ 2193 -25.6 Huntleya lucida (Rolfe) Rolfe MO 3595549 -28.7 Kefersteinia auriculata Dressler MO 4901757 -31.5 Kefersteinia costaricensis Schltr. MO 4893555 -27.7 Kefersteinia excentrica Dressler & Mora-Ret. MO 3716832 -26.5 Kefersteinia lactea (Rchb.f.) Schltr. MO 4893548 -29.1 Koellensteinia kellneriana Rchb.f. MO 1780929 -26.2 Pescatoria cerina (Lindl. & Paxton) Rchb.f. FLAS 152210 -31.9 Pescatoria dayana Rchb.f. MO 2926912 -30.9 Stenotyla picta (Rchb.f.) Dressler [Chondrorhyncha picta SEL 67925 -32.0 118

(Rchb.f.) Senghas] Warczewiczella discolor (Lindl.) Rchb.f. [Cochleanthes FLAS 181609 -22.6 discolor (Lindl.) R.E.Schult. & Garay] Warczewiczella lipscombiae (Rolfe) Fowlie [Cochleanthes FLAS 181757 -26.1 lipscombiae (Rolfe) Garay] Warrea costaricensis Schltr. MO 4893556 -30.2 Tribe Vandeae Subtribe Polystachyinae Polystachya foliosa (Hook.) Rchb.f. MO 3887632 -28.4 Subtribe Angraecinae *Campylocentrum brenesii Schltr. SEL 70631 -19.2 *Campylocentrum micranthum (Lindl.) Rolfe SCZ 2117 -15.3 *Campylocentrum panamense Ames SEL 59739 -13.6 *Campylocentrum schiedei (Rchb.f.) Benth. ex Hemsl. SEL 70629 -13.5 Subtribe Dendrobiinae Bulbophyllum aristatum (Rchb.f.) Hemsl. SEL 56821 -28.1 Subtribe Collabiinae Calanthe calanthoides (A.Rich. & Galeotti) Hamer & Garay SEL 68932 -32.0 Calanthe mexicana Rchb.f. FLAS 82929 -29.0 Spathoglottis plicata Blume MO 5345753 -29.9 Herbarium is listed as followed: Missouri Botanical Gardens Herbarium (MO), Marie Selby Botanical Gardens Herbarium (SEL), University of Florida Herbarium (FLAS), University of Panama Herbarium (PMA), and the Smithsonian Tropical Research Institute Herbarium (SCZ). Live specimens and reference number correspond to species available at the Selby Botanical Gardens live collection. Taxonomic names in brackets and parentheses are those recorded on the herbarium sheets, which have been replaced by new names.

119

Table 2. List of orchid genera containing species with CAM. This list has been updated upon that of Smith & Winter 1996, using a literature search and the results of this study.

Orchid Genus Reference Acianthera This study Aerangis Smith & Winter 1996 Aeranthes Smith & Winter 1996 Aerides Smith & Winter 1996 Angraecum Smith & Winter 1996 Arachnis Smith & Winter 1996 Ascocentrum Smith & Winter 1996 Aspasia Silvera et al. 2005 Barkeria This study Brassavola Smith & Winter 1996, Zotz & Ziegler 1997, Silvera et al. 2005, This study Brassia Silvera et al. 2005 Bulbophyllum Smith & Winter 1996, Silvera et al. 2005 Cadetia Smith & Winter 1996 Calanthe Smith & Winter 1996 Campylocentrum Zotz & Ziegler 1997, Zotz 2004, This study Cattleya Smith & Winter 1996, Zotz & Ziegler 1997, Silvera et al. 2005, This study Caularthron Smith & Winter 1996, Zotz & Ziegler 1997, Zotz 2004, This study Chelyorchis (=Oncidium) Silvera et al. 2005, This study Chiloschista Smith & Winter 1996 Cischweinfia Silvera et al. 2005 Coelogyne Smith & Winter 1996, Silvera et al. 2005 Comparettia This study Coryanthes Silvera et al. 2005 Cymbidium Smith & Winter 1996 Cyrtopodium Smith & Winter 1996 Dendrobium Smith & Winter 1996 Dendrophylax (=Polyradicion) Smith & Winter 1996 Dimerandra Smith & Winter 1996, Zotz & Ziegler 1999, Silvera et al. 2005 Elleanthus This study Encyclia Smith & Winter 1996, Silvera et al. 2005, This study Epidendrum Smith & Winter 1996, Zotz & Ziegler 1997, Zotz 2004, Silvera et al. 2005, This study Eria Smith & Winter 1996 Eriopsis Silvera et al. 2005 Eulophia (=Lissochilus) Smith & Winter 1996 Flickingeria Smith & Winter 1996 Goniochilus This study Guarianthe Silvera et al. 2005, This study Heterotaxis Silvera et al. 2005, This study Ionopsis Silvera et al. 2005, This study Jacquiniella Zotz 2004 Jumellea Smith & Winter 1996 120

Laelia Smith & Winter 1996, This study Leochilus This study Lockhartia Zotz & Ziegler 1997, Zotz 2004, Silvera et al. 2005, This study Luisia Smith & Winter 1996 Macroclinium Silvera et al. 2005, This study Maxillaria Smith & Winter 1996, Zotz & Ziegler 1997, Zotz 2004, Silvera et al. 2005, This study Microcoelia Smith & Winter 1996 Micropera Smith & Winter 1996 Mobilabium Smith & Winter 1996 Mormodes Silvera et al. 2005 Myoxanthus This study Myrmecophila This study Notylia Zotz & Ziegler 1997, Zotz 2004, Silvera et al. 2005, This study Oeceoclades This study Oeonia Smith & Winter 1996 Oncidium Smith & Winter 1996, Silvera et al. 2005, This study Ornithocephalus Zotz & Ziegler 1997, Zotz 2004, Silvera et al. 2005, This study Paphiopedilum Smith & Winter 1996 Peristeria Silvera et al. 2005 Phalaenopsis Smith & Winter 1996 Pholidota Smith & Winter 1996 Plectorrhiza Smith & Winter 1996 Plectophora This study Pleurothallis Smith & Winter 1996, Zotz & Ziegler 1997, Zotz 2004, Silvera et al. 2005, This study Prosthechea Silvera et al. 2005 Rhinerrhiza Smith & Winter 1996 Robiquetia Smith & Winter 1996 Rodriguezia Zotz & Ziegler 1997, Silvera et al. 2005, This study Saccolabiopsis Smith & Winter 1996 Saccolabium Smith & Winter 1996 Sarcochilus Smith & Winter 1996 Scaphyglottis Silvera et al. 2005 Scelochilus This study Schoenorchis Smith & Winter 1996 Sobralia Silvera et al. 2005 Solenangis Smith & Winter 1996 Sophronitis Smith & Winter 1996 Specklinia Silvera et al. 2005 Taeniophyllum Smith & Winter 1996 Thrixspermum Smith & Winter 1996 Thunia Smith & Winter 1996 Trichocentrum Zotz 2004, Silvera et al. 2005, This study Trichoglottis Smith & Winter 1996 Trichopilia Silvera et al. 2005 121

Trigonidium Silvera et al. 2005 Trizeuxis This study Tuberolabium Smith & Winter 1996 Vanda Smith & Winter 1996 Vanilla Smith & Winter 1996, Zotz & Ziegler 1997, Silvera et al. 2005, This study

122

Chapter IV

Dynamics of the phosphoenolpyruvate carboxylase (Ppc) gene family in

the Orchidaceae (Subtribe Oncidiinea) during the evolution of

Crassulacean Acid Metabolism

ABSTRACT

In plants performing Crassulacean acid metabolism (CAM), phosphoenolpyruvate carboxylase (PEPC) catalyzes the initial fixation of atmospheric CO2 into C4-dicarboxylic acids forming oxaloacetate and inorganic phosphate as a product. PEPC is a ubiquitous enzyme that belongs to a multigene family with each gene encoding a function- and tissue-specific isoform of the enzyme. CAM-specific PEPC isoforms are thought to have evolved in response to water deficit from ancestral non-photosynthetic C3 isoforms by gene duplication and acquired transcriptional control sequences that mediate increased mRNA expression and leaf-specific or leaf-preferential expression patterns.

In this study, we characterize the diversity of PEPC genes families in ten closely- related orchid species from the Subtribe Oncidiinae with a range of photosynthetic pathways from C3-photosynthesis (Oncidium maduroi, Ticoglossum krameri, and

Oncidium sotoanum) to weak CAM (Oncidium panamense, Oncidium sphacelatum,

Gomesa flexuosa and Rossioglossum insleayi) to strong CAM (Rossioglossum ampliatum, Trichocentrum nanum, and Trichocentrum carthaginense) in order to 123 understand patterns of PEPC family diversification over evolutionary time. Genotypic differences in PEPC gene family structure were correlated with the capacity to perform

CAM measured by 24 h gas exchange, titratable acidity and isotopic measurements. We found that up to five PEPC isoforms are present in orchids, with one putative CAM- specific PEPC isogene with discrete amino acid changes identified in CAM species based on cDNA clone sampling, and an evident shift in PEPC isoform abundance from 2-3 isoforms in C3 species to 3-4 isoforms in weak CAM species to 4-5 isoforms in strong

CAM species. This apparent expansion of gene family size was likely the result of gene duplication events followed by sequence divergence. Because current interpretations are based solely on cDNA sampling frequencies, expression patterns will be confirmed by real-time (RT)-PCR from samples collected over a 24 h period. Validation of the isotopic analysis and the molecular genetic analysis of PEPC gene family using 24-hour gas

exchange showed that weak CAM species exhibit limited amounts of nocturnal CO2 uptake and fixation when compared to strong CAM species. The results highlight the physiological role of PEPC in CAM function, and serve as an example of how evolution by gene duplication is explained by adaptive divergence between duplicated genes.

INTRODUCTION

Phosphoenolpyruvate carboxylase (PEPC; EC 4.1.1.31) is a ubiquitous cytosolic enzyme with a wide distribution and is found in photosynthetic organisms such as higher plants and green algae, cyanobacteria and photosynthetic bacteria, and also in non- photosynthetic bacteria and protozoa (Chollet et al., 1996; Izui et al., 2004). PEPC is involved in C4-dicarboxylic acid metabolism and catalyzes the irreversible ß- 124

– 2+ carboxylation of phosphoenolpyruvate (PEP) in the presence of HCO3 and Mg yielding oxaloacetate (OAA) and inorganic phosphate (Pi). In CAM species, OAA is then converted to C4 organic acids (usually malic acid), which are stored in the vacuole at night. During the subsequent day, organic acids from the vacuole are decarboxylated resulting in CO2 release and refixation by ribulose-1,5-bisphosphate carboxylase/oxygenase (RUBISCO). This CO2 concentrating mechanism of “CO2 pump” effectively suppresses photorespiration and improves water use efficiency (WUE) relative to C3 and C4 plant species.

PEPC genes (Ppc) have important roles in plants performing C4 photosynthesis and CAM, because they catalyze the initial fixation of atmospheric CO2 into C4- dicarboxylic acids. In addition, PEPC has many other physiological roles in plants, which include an anaplerotic role in the cell (replenishing biosynthetic precursors to the Krebs

Cycle), maintaining cellular pH, supplying carbon to N2-fixing legume root nodules, absorption and transport of cations in roots, stomata movements, fruit maturation, and seed germination and maturation (Chollet et al., 1996; Echevarria and Vidal, 2003; Izui et al., 2004).

PEPC in vascular plants belongs to a multigene family with each gene encoding a function and tissue-specific isoforms of the enzyme (Gehrig et al., 1995; Chollet et al.,

1996; Gehrig et al., 1998; Gehrig et al., 2001; Izui et al., 2004). For example, in all plants a ‘housekeeping isoform’ or C3 isoform of PEPC catalyses anaplerotic reactions. In contrast, a CAM-specific isoform is responsible for the primary CO2 fixation in CAM leaf tissues. Photosynthetic isoforms of PEPC can be distinguished by their elevated mRNA and protein expression in leaf tissues (Cushman et al., 1989, Gehrig et al., 2005). 125

Vascular plant PEPCs undergo reversible phosphorylation at a conserved Ser near the N terminus of the gene catalyzed by a dedicated PEPC kinase (Hartwell et al., 1999; Taybi et al., 2000). Presently, full-length PEPC genes have been characterized from bacteria, vascular plants, cyanobacteria, and protozoa (Izui et al., 2004). Phylogenetic analyses suggest that plant PEPC had evolved from a common ancestral bacterial origin (Chollet et al., 1996; Izui et al., 2004; Sanchez and Cejudo, 2003; Westhoff and Gowik, 2004).

The currently available molecular data support the view that none of the C4 or

CAM enzymes or genes are unique to C4 or CAM plants (Westhoff and Gowik, 2004).

For example, Ppc has non-photosynthetic gene family isoforms present in all C3 plants, which are also present in non-photosynthetic tissues of C4 and CAM species, suggesting that these ubiquitous isoforms served as the starting point for the evolution of the C4 and

CAM genes (Cushman and Bohnert, 1999; Monson, 1999). Also, evidence from Ppc and comparative analysis of the C3, C3- C4 intermediates, and C4 species in the genus

Flaveria, suggests the acquisition of distinct kinetic and regulatory properties of C4 cycle enzymes is mediated by discrete amino acid changes (Blasing et al., 2000; Blasing et al.,

2002; Westhoff and Gowik, 2004).

Gene duplication events are documented as an important step in gene diversification. Duplication followed by recombination of genes facilitates neofunctionalization. However, the manner in which duplicated genes diverge over evolutionary times is not fully understood (West-Eberhard, 2003). The phenomenon by which genes belonging to a multigene family differentiate in terms of function and how multiple duplications of one divergent gene cause secondary duplication events, is part of 126 concerted evolution, a series of changes in nucleotide sequences within family members that ultimately causes the formation of divergent set of duplicates (West-Eberhard, 2003).

CAM-specific Ppc isoforms are thought to have evolved in response to water deficit from ancestral non-photosynthetic C3 isoforms by gene duplication and acquired transcriptional control sequences that mediate increased mRNA expression and leaf- specific or leaf-preferential expression patterns (Gehrig et al., 2001; Taybi et al., 2004;

Gehrig et al., 2005). For example, seven distinct Ppc isogenes have been recently recovered in the CAM species Kalanchoë pinnata, which contains four isogenes in leaves and three in roots (Gehrig et al., 2005). In Mesembryanthemum crystallinum a CAM- specific isoform is expressed during the induction of CAM, in addition to an uninduced housekeeping isoform (Cushman and Bohnert 1989). In orchids performing CAM, at least three isoforms are expected in leaves with at least one isoform predicted to be

CAM-specific, and two or more predicted to be C3-specific based on relative abundance

(Gehrig et al. 2001).

In this study, we characterize the diversity of PEPC genes in closely-related orchid species (Oncidiinae) with photosynthetic pathways ranging from C3 to weak CAM to CAM in order to understand PEPC gene family structure dynamics throughout evolutionary time, and to determine whether or not differences in PEPC gene family structure and recruitment correlate with the expression of CAM as measured by 24 h gas exchange, titratable acidity and isotopic measurements. The results highlight the evolutionary diversification of PEPC gene families in closely-related species with C3- photosynthesis, weak CAM and CAM, and the role of gene duplication events in neofunctionalization for CAM. 127

MATERIAL AND METHODS

Plant Material

Leaf tissues were obtained from a selected subset of closely-related species within the

Subtribe Oncidiinae (Orchidaceae), based on !13C values and titratable acidity measurements (Table 1). The phylogenetic relationship among species is depicted in

Figure 1. Orchids were grown in a closed greenhouse at the University of Nevada, Reno and the University of California, Riverside. Daily temperature within the greenhouse varied from 17˚C to 35˚C with RH varying from 40% to 80%, and an annual mean photon flux density (PFD) of 200 "mol m-2 sec-1. Plants were watered daily and nutrients were supplied once a week with a combination of slow release fertilizer (Osmocote 19-6-

12 formula) and commercial fertilizer solution (Schultz 19-31-17 formula). Samples for molecular analysis were collected from mature healthy leaves from each species at 2 PM and 2 AM to account for putative circadian differences in the expression of PEPC. Root samples were also collected at the same times from mature plants and stored separately.

All samples were flash frozen in liquid nitrogen immediately after harvesting, and stored at –80ºC for isolation of total RNA.

Gas exchange measurements

CO2 gas exchange was measured in a mature orchid leaf for each of ten species studied, by sealing the leaf inside a Plexiglass cuvette in a controlled environmental chamber. The hole between the cuvette and the leaf was sealed with a nonporous synthetic rubber sealant, thus delimiting the leaf from the rest of the plant. The plants were well-watered and maintained in growth media while conducting measurements to avoid stresses. The 128 only exception was for Trichocentrum nanum for which the whole plant (including roots) was placed inside the cuvette. Net CO2 exchange was measured continuously using a flow-through-gas-exchange system (Walz, Effeltrich, Germany) operating at 2 L air min-1, and monitored with a LI-6252 infrared gas analyzer (LI-COR, Inc., Lincoln, NE,

USA) operating in the absolute mode without interruption for up to 5 days. The temperature inside the cuvette/chamber was 25°C during the day, and 22°C during the night under ambient CO2 concentration, dew-point of the air was 18°C, and light intensity was 300 "mol m-2s-1.

Extraction of RNA

Total RNA was extracted using the RNeasy Midi Kit (Qiagen, Inc., Valencia, CA) with a modified PEG-RNA extraction method that utilizes high-molecular weight polyethylene glycol (Gehrig et al., 2000), that has proven successful with succulent and non-succulent orchid tissues. RNA integrity was examined by 1% gel electrophoresis, and the quality and concentration was examined using a NanoDrop ND-1000 UV-V spectrophotometer

(NanoDrop Technologies Inc., Rockland, DE).

RT-PCR amplification and cloning

An 1,100-bp fragment was amplified by reverse transcription (RT)-PCR using two degenerate primers as previously defined for Ppc (Taybi et al., 2004; Gehrig et al., 2005;

Kore-eda et al., 2005) and slightly modified for orchid specificity. The degenerate primers were: Forward 5’-TC(ACGT) GA(CT) TC(ACGT) GG(ACG) AA(AG) GA(CT)

GC-3’ and Reverse 5’-GC(AGT) GC(AG) AT(AG) CC(CT) TTC AT(GT) G-3’. Using 129 the OneStep RT-PCR kit (Qiagen, Inc., Valencia, CA), 500 ng of total RNA were reversed transcribed and amplified following manufacturer’s instructions. Final concentrations of the reaction components were 400 "M for each dNTP, 1x 5X Qiagen

OneStep RT-PCR Buffer containing 12.5 mM MgCl2, and 1"M PEPC Forward and

Reverse primers. The following conditions were used: Reverse transcription at 50ºC for

30 minutes, initial PCR activation at 95ºC for 15 minutes, amplification of 39 cycles at

94ºC for 1 minute, 40ºC for 1 minute, 72ºC for 1 minute, and final extension of 72ºC for

10 minutes.

RT-PCR products were purified by agarose gel electrophoresis, recovered using

QIAquick Gel Extraction Kit (Qiagen, Inc., Valencia, CA), and cloned into the TA-

TOPO cloning vector pCR2.1 vector system (InvitrogenTM Life Technologies, Carlsbad,

CA), and transformed into XL1-Blue competent cells or TOP10 competent cells following manufacturer’s instructions. Bacterial cells containing plasmids from 100 randomly selected clones per species were grown in terrific broth (TB) liquid media for

16 hours at 37ºC with vigorous shaking. Bacterial cells were then harvested by centrifugation at 13,000 x g and plasmids were purified using the Qiagen Plasmid Mini-

Kit (Qiagen, Inc., Valencia, CA) following manufacturer’s instructions. cDNA clones were then analyzed by EcoR1 digestion and electrophoresis on 1% agarose gels stained with 0.5 "g/ml ethidium bromide. Selected plasmids were sequenced (Genomics Center,

University of Nevada Reno) by using the ABI BigDye™ Terminator Cycle sequencing

Ready Reaction Kit v3.1 and an ABI 3730 Sequence Analyzer (Applied Biosystems,

Foster City, CA), using the M13 forward (5’-TGTAAAACGACGGCCAGT-3’) and reverse (5’-GAGCGGATAACAATTTCACACAG-3’) primer sets. 130

Sequence analysis

Raw sequences were edited manually by removing vector sequences using MacVector v11.1 software (MacVector, Inc. Cary, NC). PEPC fragments were verified by identifying conserved amino acid sequences using the Basic Local Alignment Search

Tool (BLAST) database search tool at the National Center for Biotechnology Information

(NCBI). Alignment matrices for each species were then obtained using the ClustalW tool in MacVector and analyzed by constructing trees using phylogenetic reconstructions with the distance method Neighbor-Joining (NJ, as a tool in MacVector), which makes no assumptions of constant divergence among sequences. Sites containing gaps were ignored. Consensus sequences for each isoform were generated in each species separately. Trees were generated by including bootstrap analyses using 1,000 replicates performed by the absolute number of difference distance method that estimates the total number of differences between sequences using the isoform sequences for all species.

The tree of nucleotides and protein sequences generated by NJ were then rooted using the

E. coli PEPC sequence as an outgroup.

RACE amplification

Amplification and sequencing of the 3’ end of the cDNA fragment for PEPC for three species (Oncidium maduroi, Rossioglossum ampliatum, and Oncidium panamense) were recovered by 3’ RACE system (SMARTTM RACE cDNA Amplification, BD Bioscience

Clontech, Mountain View, USA) following manufacturer’s instructions and using gene- specific primers designed from the sequencing results obtained from the initial degenerate RT-PCR (Table 2). RACE products were generated using the SMART™ 131

RACE cDNA amplification kit (BD Biosciences), and cDNA sequence analysis were performed using a BigDye™ Terminator Sequencing kit and an ABI 3730 Sequence

Analyzer (Applied Biosystems) at the Genomics Center of the University of Nevada

Reno. Raw sequences were edited manually by removing vector sequences using

MacVector v11.1 software (MacVector, Inc., Cary, NC). Sequences were then assembled to the 1,100 bp isoform fragments for each species using the assembly project function in

MacVector.

RESULTS AND DISCUSSION

Oncidiinae species and characterization

The subtribe Oncidiinae represents one of the most highly derived clades of orchids of the neotropics, with a great variety of diversity in chromosome number, vegetative features and floral characteristics (Chase et al. 2005). In this study, ten closely-related species within the Oncidiinae with a range of photosynthetic pathways from C3- photosynthesis (Oncidium maduroi, Ticoglossum krameri, and Oncidium sotoanum) to weak CAM (Oncidium panamense, Oncidium sphacelatum, Gomesa flexuosa and

Rossioglossum insleayi) to strong CAM (Rossioglossum ampliatum, Trichocentrum nanum, and Trichocentrum carthaginense) were selected based on !13C values and titratable acidity measurements (Table 1) and used to characterize the diversity of PEPC genes families. The relationship among these species is depicted in Figure 1.

Photosynthetic pattern of Oncidiinae species

An intrinsic characteristic of plants performing CAM is inverted stomatal rhythms, 132

wherein stomata are open during the night thus allowing net CO2 uptake to occur, catalyzed by PEPC during Phase I. Leaves performing CAM show higher net CO2 uptake at night when compared with daytime CO2 uptake. In our study, well-watered C3 species showed no such nocturnal uptake (Fig. 2), whereas weak CAM species showed fluctuations in net CO2 uptake between day and night with the majority of the CO2 taken up during the day and a small non-significant net CO2 loss during the beginning of the night (Fig. 3), and strong CAM species showed pronounced CO2 uptake during the night

(Fig. 4).

Oncidium maduroi, Oncidium sotoanum, Oncidium cheirophorum and

Rossioglossum krameri (all C3 species, Fig. 2) showed net CO2 uptake exclusively during the daytime with a slight CO2 loss during the nighttime. This loss of CO2 was insignificant when compared to the large gain of CO2 taken during the day. Interestingly,

Oncidium sphacelatum, Oncidum panamense, Gomesa flexuosa and Rossioglossum

13 insleayi, were species that showed ! C values characteristic of C3 species and significant titratable acidity fluctuation between day and night typical of plants performing CAM

(Table 1). Based on these two characteristics, these species were categorized as weak

CAM (Table 1). Winter and Holtum (2002) demonstrated that species that have !13C values characteristic of C3 species could obtain up to one third of their carbon via the

CAM pathway. Such a small nocturnal carbon gain can only be detected by the combination of carbon-isotope-ratio analysis, titratable acidity and 24 h gas exchange measurements. Indeed, we found that these weak CAM species showed patterns of CO2 uptake during nighttime typical of CAM species but at a greatly reduced magnitude (Fig.

3). Both Oncidium sphacelatum and Oncidium panamense showed very similar 133

fluctuations in net CO2 uptake between day and night with the majority of the CO2 taken up during the day and a small non-significant net CO2 loss during the beginning of the night period (Fig. 3, top two panels). In Gomesa flexuosa, net CO2 loss occurred toward the end of the night, probably due to depletion of starch needed for PEP production (Fig.

3, middle panel). Rossioglossum insleayi showed a small decrease in net CO2 loss at around midday probably linked to stomatal closure needed to reduce transpiration (Fig. 3, bottom panel). In all cases, the majority of net CO2 uptake occurred during the daytime, with a second smaller peak of net CO2 uptake during the night, meaning that these species are capable of opening their stomata at night and fixing CO2 through the CAM pathway under well-watered conditions (Fig. 3).

Trichocentrum carthaginense (a strong CAM species) maintained closed stomata during the day as evidenced by the lack of net CO2 uptake during daytime (Fig. 4, top panel). There was a slight CO2 loss during midday and right before nighttime, which was accompanied by a decrease in transpiration. Stomata gradually close as daytime approaches, reducing water loss (Fig. 4). In contrast, Rossioglossum ampliatum (a strong

CAM species, Fig. 4 middle panel) and Trichocentrum nanum (a strong CAM species,

Fig. 4 bottom panel) showed daytime net CO2 uptake during the afternoon hours, associated with C3 photosynthesis during the day. During this time, stomata are open and low transpiration rates should occur. The majority of net CO2 uptake, however, occurs during the middle of the nighttime when stomata are closed and transpiration rates are minima. A gradual decrease in CO2 uptake was evident as daytime approached, which was likely linked to a depletion of starch within the cells, which was needed to produce

PEP as the CO2 acceptor (Cushman et al., 2008). The degree of CAM is likely limited by 134 vacuole size, with a minimum volume being needed to store large quantities of malic acid during the night, which are decarboxylated during the daytime to ultimately produce PEP from carbohydrates during the night (Winter 1985). Root respiration for Trichocentrum nanum was included because the whole plant was measured within the cuvette due to the small size of the plant. Root respiration represented the larger contribution of CO2 loss during nighttime and daytime when compared to the other CAM species (Fig. 4).

Net CO2 uptake by CAM plants over 24 h has been reported to be optimal for day and night temperatures of 25ºC and 15ºC, respectively, with optimal daily net CO2 uptake determined by temperature at night when the majority of the CO2 is fixed (Nobel, 1991).

During drought, orchid species rely mainly on their leaf and pseudobulb water storage reserves. Such water storage combined with a higher water use efficiency can contribute to a larger period of net CO2 uptake compared to C3 species. In terms of emery cost,

CAM plants utilize more ATP compared to C3 species and can be as much as 20% less efficient than C3 species and 10% less efficient than C4 plants. CAM and C4 plants have additional energy requirements due to additional enzymatic steps. CAM plants, for example, have the extra expense of transporting malate at night into the vacuole. During the fixation of CO2 into carbohydrates, C3 species require 3 ATP and 2 NADPH, C4 species require 5 ATP and 2 NADPH, and CAM species require 5.5-6.5 ATP and 2

NADPH (Nobel, 1991). However CAM plants have an advantage in environments with intermittent water supply due to the increased water conserving nature of the CAM pathway. Further work should investigate outstanding questions such as: 1) Do weak

CAM orchid species switch to full CAM mode due to environmental stress as in the case of facultative CAM? 2) Is there fluctuation of net CO2 uptake at night due to seasonal 135 changes? 3) Are the 24 h gas exchange patterns observed in this study maintained regardless of environmental variation?

Identification of multiple PEPC isogenes in Oncidiinae species

We sampled 1,200 cDNA and sequenced 1,000 cDNA clones coding a partial sequence of the key CAM enzyme Phosphoenolpyruvate carboxylase (PEPC) after transcription of mRNA using well-watered closely-related Oncidiinae species (Table 1). We used degenerate RT-PCR to sample a fragment of 1,100 bp, which encompasses the C- terminal third of the PEPC coding region (Fig. 5). By using this partial sequence, we were able to distinguish distinct isogenes without the need to isolate the full 3,000 bp sequence, because this fragment comprises the active center of the enzyme, which is conserved in all plant species (Fig. 6). Also, the remaining portions of the fragment were variable enough to allow differentiation among isoforms (Gehrig et al., 2001; Izui et al.,

2004). In this regard, the use of partial length PEPC sequences, instead of full-length sequences has proven to be useful in comparing sequences across species in the context of molecular phylogeny and taxonomy, thus saving time and financial resources for researchers interested in using PEPC as a molecular marker (Gehrig et al., 2001). We found 35 PEPC isogenes from ten Oncidiinae species, which are summarized in Table 3.

From the continuum of photosynthetic pathways from C3 to weak CAM to strong CAM, we found evidence of gene duplication events. Species performing C3 photosynthesis encode 2-3 isogenes, whereas species with weak CAM encode 3-4 isogenes, and species with strong CAM encode 4-5 isogenes (Table 3). PEPC multiple nucleotide sequence alignments of 1,100 bp fragment showed 76-99% similarity score values with 430 136 invariant nt positions across all isogenes. PEPC multiple protein sequence alignments of

363 amino acids showed 84-99% similarity score values with 271 invariant amino acid positions across all isogenes. Comparisons of the Oncidiinae 1,100 bp PEPC fragment sequences with 223-875 bp fragments of PEPC contigs resulting from 454 pyrosequencing of a strong CAM species Rossioglossum ampliatum (contigs 36974,

10159 and 35735, see Chapter 5 for 454 pyrosequencing details) showed an overall 74-

99% similarity score with 164-612 invariant sites (Table 4).

Phylogenetic analysis was performed using Neighbor joining in order to distinguish the different PEPC lineages. PEPC sequences of ten Oncidiinae species formed two main clusters (both with 100 bootstrap support), with one branch comprising the least abundant isoforms for all species (branch A, Fig. 7), and the second cluster comprising the most abundant isoforms for all species (branch B, Fig. 7). This pattern suggests that these two lineages evolved in parallel. Genes within one lineage (branch B

Fig. 7), presumably derived from gene duplication events, evolved a role in photosynthesis in CAM (represented by arrow in Fig. 7), whereas genes within the other separate lineage (branch A) likely retained an anaplerotic ‘housekeeping’ role.

Interestingly, we did not find a distinct PEPC isogene that could be associated exclusively with root tissue. Gehrig et al. (2005) found three root PEPC that contained an insertion of 8 amino acids towards the C-terminus of the enzyme. PEPC from root tissues in all of the orchid species studied here showed isoforms found in leaves and no insertion was evident. Because aerial roots in epiphytic orchids engage in photosynthesis, the

PEPC genes found in roots of epiphytic orchids are most likely the same as those found in leaves (Kwok-Ki et al., 1983). 137

Sequence analysis showed that the most abundant isoform for all strong CAM species studied cluster together (Fig. 7). The trees generated, both at the protein and nucleotide level (Figs. 7-8), showed that all isogenes were derived from a common ancestor. The clustering of the highly abundant isoforms present in photosynthetic tissues of CAM species is evidence that these paralog genes are likely recruited for CAM function. The observed sequence relatedness among them indicates convergent evolution for assuming the enzymatic functionality associated with CAM. A differentiation among paralogous genes occurs within branch B, such that PEPC sequences separated into distinct groups: the first clade represented by an arrow in Fig. 7 (bootstrap support of

97%), contained PEPC sequences from the most abundant isoform from strong CAM species, presumably with a role in CAM that evolved from gene duplication events.

Although a polytomy was present in Branch B, the most abundant isoforms for weak

CAM and C3 species clustered together (82% bootstrap support). The remaining groups within branch B were poorly resolved (Fig. 7). The most abundant isoform in weak

CAM species are likely involved in CAM photosynthesis, whereas the most abundant isoforms in C3 species are likely involved in leaf anaplerotic functions. Detailed analysis of the mRNA abundance patterns of these isogenes is needed to resolve their putative functional contributions to CAM. Interestingly, Trichocentrum nanum Ppc5 was quite distinct from all other isoforms contained in branch B, as evidenced by the separate lineage (Fig. 7). This very low abundant isoform from a strong CAM species might be related to a non-photosynthetic PEPC lineage. Although it might be related to the ancestral bacterial type PEPC lineage, more detailed studies are needed in order to confirm the putative function of this very rare isoform. This clade also contained low 138

abundance isoforms from a weak CAM species and a C3 species (Oncidium sphacelatum

Ppc3, Oncidium sotoanum Ppc3). These two low abundance genes share a high similarity with the highly abundant PEPC from CAM species, but are not likely to function in CAM photosynthesis. This is likely due to functional diversification of paralogous genes, or possibly due to past reversal events from CAM back to C3 photosynthesis.

The reversion of characters (reversal events) or the independent loss of a character would require convergent evolution by independent loss of identical genetic sites, an event that is difficult to trace, especially when reversals occur in widely separate clades

(See Chapter 2 for examples of reversals within the Orchidaceae). Reversals in CAM can be defined as the secondary presence of the primitive character state (C3 photosynthesis) not homologous with the ancestral or pleisomorphic state found in the common ancestor.

Some reversions are induced by environmental conditions, and others are the product of gradual loss of complex traits (West-Eberhard, 2003). Recurrent reversion within closely- related clades suggests the involvement of multiple pathways that permits the re- expression of different regulatory mechanisms. Also, the independent origin of CAM in distantly related clades within plants is evidence that CAM likely involves the amplification of genes that are already present in the most distant common C3 ancestor.

The association of CAM with drier environments and stressful conditions suggests a role for environmental influences in the origin and evolution of CAM in distant lineages. This suggestion is also supported by the observation that the extent of CAM expression generally correlates with the degree of adaptation to more xeric ecological niches (Kluge et al. 2001, Pierce et al. 2002, Zotz 2004). Water limitation and the resulting limitation of

CO2 brought about by stomatal closure and the reductions in atmospheric CO2 139 concentrations during the late Tertiary period (Pearson and Palmer, 2000) might have provided the selective pressures for the evolution of CAM over the last 40-100 million years (Ehleringer and Monson, 1993; Monson 1989; Raven and Spicer, 1996).

Within the Orchidaceae, the presence of CAM is evolutionary labile, and prone to parallel evolution and reversals (see Chapter 2 for examples of parallel evolution of CAM within the Orchidaceae) especially within clades that contain large number of epiphytic species (Subfamily Epidendroideae, Silvera et al. 2009). The frequency of reversal and parallel evolution of CAM reflects a strong local selective pressure for conditions where

CAM is an advantage (such as the epiphytic habitat of orchids). For example, an epiphytic population colonizing a new niche where drought is present, such as deciduous forests with a long dry season, would be under selective pressure to evolve a set of photosynthetic characteristics, among other adaptive traits, that might confer an advantage over populations that would be competing for the same sites. Response to these selective pressures would include biochemical shifts and molecular genetic re- arrangements that can promote the shift from C3 to CAM, and imply a direct role of environmental cues in promoting changes in genotypes. After CAM is established within a lineage, the expression of CAM can be plastic, as illustrated by the existence of

‘facultative’, ‘inducible’ or ‘optional’ CAM or C3-CAM intermediate species that engage in CAM in response to environmental stimuli such as drought stress (Griffith 1998;

Winter 1985; Winter et al., 2008). This flexibility combined with the fact that the enzymes present in CAM are ubiquitous in all plants might explain why multiple independent origins of CAM, as well as reversals, have been observed within the

Orchidaceae (Silvera et al., 2009). 140

The PEPC protein tree for Oncidiinae species showed less resolution when compared to the nucleotide tree (Fig. 8). Nonetheless, the overall clustering of PEPC lineages between the nucleotide and protein trees was similar. The PEPC protein tree for

Oncidiinae species and various other orchid CAM species showed that PEPC sequences tended to arrange phylogenetically according to taxa. For example, all Dendrobium PEPC clustered together (Fig. 9). Within taxa, PEPC sequences tended to arrange according to function. This pattern is likely due to the different functional relationships among paralog and ortholog genes within the evolution of PEPC in orchid lineages. Nonetheless, we found that PEPC sequences within orchids cluster into two main groups (branch A and B,

Fig. 9), one related to CAM function and the other related to housekeeping functions.

Using 3’RACE-PCR technique, we were able to isolate the last six amino acid residues followed by the 3’ untranslated region (UTR) and the poly(A) tail, for two isoforms of each of three Oncidiinae species (Fig. 10). As we continue to isolate the

UTRs of the remaining isoforms, we might find unique isoforms not found by the random sampling of cDNA clones produced by degenerate RT-PCR. Figures 11-13 show the multiple nucleotide sequence alignments between each of three PEPC contigs (35735,

10159, and 36974) obtained from 454 pyrosequencing of a strong CAM species

(Rossioglossum ampliatum) within the corresponding 1,100 bp fragment of the PEPC isoforms of three Oncidiinae species (Oncidium maduroi, Oncidium panamense, and

Rossioglossum ampliatum) obtained from cDNA cloning sampling. The comparisons using these two lines of evidence validate the identity of the PEPC fragments found by cDNA cloning sampling. The multiple sequence alignments for contig 10159 showed single nucleotide polymorphic sites with 133 invariant nt sites out of the 220 bp 141 fragment, 164 invariant nt sites out of the 223 bp fragment for contig 36974, and 612 invariant nt sites out of the 885 bp fragment for contig 35735 (Fig. 11-13). Nucleotide similarity scores ranged from 71.5% between Rossioglossum ampliatum Ppc4 and contig

10159, to up to 99.7% between Rossioglossum ampliatum Ppc1 and contig 35735 (Table

4). Because this study was based on over 1,000 plasmids, we are confident that we have recovered nearly all PEPC isogenes expressed in the leaf and roots of the ten species studied covering the range of photosynthetic pathways from C3 to weak CAM to strong

CAM.

CONCLUSIONS

The present study shows that the PEPC gene family in orchids has likely undergone gene family expansion during the transition from C3 to weak CAM to strong

CAM. The multiple independent evolutionary origins of CAM in orchids suggest that evolution from C3 to weak and strong CAM might involve relatively few genetic changes.

Evidence from C4 Flaveria species suggests that C4 photosynthesis PEPC isoforms evolved from ancestral non-photosynthetic C3 isoforms by gene duplication events and further acquisition of distinct kinetic and regulatory properties (Blasing et al., 2002;

Gehrig et al., 2005). Here, we provide evidence that a similar progression of events likely occurred during CAM evolution as demonstrated by an increase in the number of PEPC isoforms in strong CAM species. Our analysis of the PEPC gene family structure and mRNA relative abundance, using closely-related species exhibiting C3, weak CAM and strong CAM, found that as many as five isoforms were present in orchids performing

CAM, with one putative CAM-specific PEPC isogene with discrete amino acid changes 142 identified within each CAM species based on cDNA clone sampling. Validation of the isotopic analysis and the molecular genetic analysis of PEPC gene family using 24-hour gas exchange showed that weak CAM species exhibit a limited degree of nocturnal CO2 uptake when compared to strong CAM species. This study demonstrated several patterns of CAM evolution in orchids, thus improving our understanding of the functional significance and evolutionary origins of CAM.

ACKNOWLEDGMENTS

We would like to thank Dr. Klaus Winter at STRI in Panama for the use of the 24 h gas exchange system and Dr. Gaspar Silvera for supplying the orchid plant species.

Additionally, we want to thank Becky Albion for technical assistance in the lab, Letty

Rodriguez and Cristina Milsner for assistance with PEPC cloning sampling, and the

Cushman Lab for support. We want to also thank Dr. Mark Whitten, Dr. Norris Williams, and Kurt Neubig for providing Oncidiinae DNA matrices and trees. This work was supported by funding from the U.S. Environmental Protection Agency under the Greater

Research Opportunities Graduate Program Agreement no. MA 91685201 (to K.S.) and

National Science Foundation NSF IOB-0543659 (to J.C.C.). This publication was also made possible by NIH Grant Number P20 RR-016464 from the INBRE Program of the

National Center for Research Resources through its support of the Nevada Genomics,

Proteomics and Bioinformatics Centers. The views expressed in this publication are solely those of the authors and the EPA does not endorse any products or commercial services mentioned in this publication. 143

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146

Table 1. !13C values, leaf thickness and titratable acidity for ten Oncidiinae species. Titratable acidity is represented by the mean±SD of 3 replicates at morning and evening (Silvera et al. 2005 and new measurements from this study). *Denotes significance between means of the morning and evening at P<0.05 as determined by a Student’s t-test. NS, not significant.

Leaf Leaf H+ H+ $H+ Photosynthetic !13C thickness (evening) (morning) pathway Species name (‰) (mm) ("mol H+g–1 FW) Rossioglossum ampliatum – 15.3 1.59 5.5±1.3 153.5±3.6 148.0* Strong CAM Trichocentrum nanum –17.2 3.40 18.8±4.1 57.3±5.2 38.50* Strong CAM Trichocentrum –12.2 2.32 12.5±0.4 77.3±3.4 64.8* Strong CAM carthaginense Rossioglossum insleayi –22.5 1.10 16.3±0.5 34.9±12 18.60* Weak CAM Oncidium panamense –26.2 0.54 11.5±0.7 33.2±0.3 21.7* Weak CAM Oncidium sphacelatum –27.9 0.53 8.3±6.3 31.2±2.1 22.9* Weak CAM Gomesa flexuosa –24.4 0.26 37.9±9.4 74.0±10.4 36.1* Weak CAM Rossioglossum krameri –31.7 0.35 14.4±2.5 14.8±2.4 0.4 NS C3 Oncidium sotoanum –25.2 0.25 2.9±0.4 2.72±1.6 –0.2 NS C3 Oncidium maduroi –24.7 0.24 17.3±1.9 19.5±0.8 2.2 NS C3

147

Table 2. Gene specific primer sets (GSP 5’– 3’ position) used for 3’RACE amplification of the cDNA fragment coding for Ppc isoform for ten Oncidiinae species and 36 isoforms. GSP1-1 to 1-23 were used for the first round of amplification, and GSP2-1 to GSP2-23 were used in the second round of amplification to reduce non-specific products.

GSP1-1 AACTCTGAACGTCTGCCAAGCCTACGCG GSP1-2 AACTCTGAACGTTTGCCAAGCCTACACA GSP1-3 GACCCTGAATGTGTGCCAAGCCTTCACC GSP1-4 GACGCTGAATGTGTGCCAAGCCTTCACC GSP1-5 AACTCTGAACGTTTGCCAAGCCTACACG GSP1-6 TACTCTGAACGTTTGCCAAGCCTGCACA GSP1-7 GACCCTGAATGTGTGCCAAGCCTTCACT GSP1-8 GACCCTGAATGTGTGCCAAGCTTTCACC GSP1-21 CGACTCAGGCTCCGCTACCCG GSP1-22 TTGGCAGTTGTACAAGGCTCAGGAG GSP1-23 GGCAGCTATATAAGACTCAAGG GSP2-1 GCGAATCAGAGATCCAAGCGATCATGG GSP2-2 GCGGATCCGAGATCCAAGCAATGGTCA GSP2-3 GCGAATCAGAGATCCAAGCGATCATCG GSP2-4 GCGAATCAGAGATCCGAGCGATCATCA GSP2-5 GCGAATCAGAGATCCGAGCGATCATCC GSP2-6 GCGAATCAGAGATCCAAGCGATCATGG GSP2-7 AAGGATCAGAGACCCAAATTTTCTTGT GSP2-8 AAGGATCAGAGACCCAAAATTTCTTGT GSP2-9 AAGGATCAGAGATCCAAATTTTCTTGT GSP2-10 AAGAATCAGAGACCCAAATTTTCTTGTG GSP2-12 GCGGATCAGAGATCCAAGCGATCGTCA GSP2-13 GCGGATCAGAGATCCAAGCGATCATCG GSP2-14 GCGGATCAGAGATCCAAGCAATGGTCA GSP2-15 GCGGATCCGAGATCCAAGCGACGGTCA GSP2-16 AAGGATCAGAGACCCAAATTTTCATGT GSP2-21 AACGTTTGCCAAGCCTACACGC GSP2-22 GGCTCAGGAGGATCTCATAAAAGTG GSP2-23 GAAAGTAGCCAAGGAATTTGGA

148

Table 3. Different functional PEPC isoforms from ten Oncidiinae species based on relative abundance of clone sampling.

Species name Isoforms Relative Functional abundance (%) Designation Gomesa flexuosa (Weak CAM) Ppc1 66 CAM Ppc2 31 C3 Ppc3 2 C3 Ppc4 1 C3 Oncidium maduroi (C3) Ppc1 88 C3 Ppc2 11 C3 Ppc3 1 C3 Oncidium panamense (Weak CAM) Ppc1 73 CAM Ppc2 19 C3 Ppc3 7.5 C3 Oncidium sotoanum (C3) Ppc1 86 C3 Ppc2 10 C3 Ppc3 4 C3 Oncidium sphacelatum (Weak CAM) Ppc1 96 CAM Ppc2 2.5 C3 Ppc3 1 C3 Rossioglossum ampliatum (CAM) Ppc1 70 CAM Ppc2 20 C3 Ppc3 5 C3 Ppc4 5 C3 Rossioglossum insleayi (Weak CAM) Ppc1 76 CAM Ppc2 11 C3 Ppc3 10 C3 Ppc4 2 C3 Rossioglossum krameri (C3) Ppc1 92 C3 Ppc2 8 C3 Trichocentrum carthaginense (CAM) Ppc1 84 CAM Ppc2 9 C3 Ppc3 4 C3 Ppc4 3 C3 Trichocentrum nanum (CAM) Ppc1 59 CAM Ppc2 20 C3 Ppc3 16 C3 Ppc4 4 C3 Ppc5 1 C3

149

OM1 OM3 RA1 OP1 RA2 RA3 OP2 OM2 OP3 RA4 CON O_maduroi_Ppc1 (OM1) 100.0 94.6 95.1 95.5 93.7 79.4 96.0 78.5 79.4 79.4 78.9 O_maduroi_Ppc3 (OM3) 94.6 100.0 94.2 94.6 93.3 78.0 95.1 76.2 77.1 76.7 77.6 R_ampliatum_Ppc1 (RA1) 95.1 94.2 100.0 96.0 93.3 79.8 95.1 78.9 78.9 78.9 79.4 O_panamense_Ppc1 (OP1) 95.5 94.6 96.0 100.0 95.1 80.7 97.3 80.3 80.3 79.8 80.3 R_ampliatum_Ppc2 (RA2) 93.7 93.3 93.3 95.1 100.0 79.8 93.7 78.8 77.6 78.5 78.9 R_ampliatum_Ppc3 (RA3) 79.4 78.0 79.8 80.7 79.8 100.0 80.3 96.0 96.4 96.4 96.9 O_panamense_Ppc2 (OP2) 96.0 95.1 95.1 97.3 93.7 80.3 100.0 79.4 80.3 79.4 79.4 O_maduroi_Ppc2 (OM2) 78.5 76.2 78.9 80.3 78.8 96.0 79.4 100.0 97.3 96.0 96.9 O_panamense_Ppc3 (OP3) 79.4 77.1 78.9 80.3 77.6 96.4 80.3 97.3 100.0 96.0 97.3 R_ampliatum_Ppc4 (RA4) 79.4 76.7 78.9 79.8 78.5 96.4 79.4 96.0 96.0 100.0 97.8 Contig 36974 (CON) 78.9 77.6 79.4 80.3 78.9 96.9 79.4 96.9 97.3 97.8 100.0

CON RA1 OP1 OM1 OP2 OM3 OM2 OP3 RA4 RA3 RA2 Contig 35735 (CON) 100.0 99.7 93.9 94.4 93.7 94.5 76.8 76.9 76.5 76.7 94.1 R_ampliatum_Ppc1(RA1) 99.7 100.0 94.3 94.7 93.9 94.6 77.0 77.1 76.6 76.9 94.1 O_panamense_Ppc1(OP1) 93.9 94.3 100.0 97.7 97.1 97.4 78.2 78.2 77.7 78.1 93.9 O_maduroi_Ppc1 (OM1) 94.4 94.7 97.7 100.0 98.4 97.7 78.5 78.5 78.2 78.3 94.1 O_panamense_Ppc2(OP2) 93.7 93.9 97.1 98.4 100.0 97.0 78.3 78.5 77.9 77.8 93.0 O_maduroi_Ppc3 (OM3) 94.5 94.6 97.4 97.7 97.0 100.0 78.6 78.6 77.9 78.4 93.6 O_maduroi_Ppc2 (OM2) 76.8 77.0 78.2 78.5 78.3 78.6 100.0 98.6 97.1 96.9 75.9 O_panamense_Ppc3(OP3) 76.9 77.1 78.2 78.5 78.5 78.6 98.6 100.0 96.7 96.6 76.0 R_ampliatum_Ppc4(RA4) 76.5 76.6 77.7 78.2 77.9 77.9 97.1 96.7 100.0 97.0 75.8 R_ampliatum_Ppc3(RA3) 76.7 76.9 78.1 78.3 77.8 78.4 96.9 96.6 97.0 100.0 76.0 R_ampliatum_Ppc2(RA2) 94.1 94.1 93.9 94.1 93.0 93.6 75.9 76.0 75.8 76.0 100.0

CON RA2 OM1 OP2 OM3 OP1 RA1 OM2 OM3 RA4 RA3 Contig 10159 (CON) 100.0 79.1 84.1 83.6 83.2 83.2 82.3 75.5 75.0 74.5 75.5 R_ampliatum_Ppc2 (RA2) 79.1 100.0 95.0 95.5 93.2 95.0 96.4 71.4 70.9 71.4 71.4 O_maduroi_Ppc1 (OM1) 84.1 95.0 100.0 99.5 97.7 98.2 96.8 74.5 74.1 73.6 74.5 O_panamense_Ppc2 (OP2) 83.6 95.5 99.5 100.0 97.7 98.6 97.3 74.5 74.1 73.6 74.5 O_maduroi_Pp3 (OM3) 83.2 93.2 97.7 97.7 100.0 98.2 95.0 74.1 73.6 72.7 74.1 O_panamense_Ppc1 (OP1) 83.2 95.0 98.2 98.6 98.2 100.0 95.9 74.1 73.6 73.2 74.1 R_ampliatum_Ppc1 (RA1) 82.3 96.4 96.8 97.3 95.0 95.9 100.0 74.1 73.6 73.2 74.1 O_maduroi_Ppc2_(OM2) 75.5 71.4 74.5 74.5 74.1 74.1 74.1 100.0 99.1 95.0 95.9 O_panamense_Ppc3_(OM3) 75.0 70.9 74.1 74.1 73.6 73.6 73.6 99.1 100.0 94.1 95.0 R_ampliatum_Ppc4_(RA4) 74.5 71.4 73.6 73.6 72.7 73.2 73.2 95.0 94.1 100.0 96.4 R_ampliatum_Ppc3_(RA3) 75.5 71.4 74.5 74.5 74.1 74.1 74.1 95.9 95.0 96.4 100.0

150

Figure 1. Phylogeny for ten Oncidiinae species. The matrix includes nrITS 1&2, plastid regions ycf1 (a ca. 1,200 bp portion from 5' end, and ca. 1,500 bp portion from 3' end), matK, and the trnH-psbA intergenic spacer using Eulophia graminea as the outgroup (data unpublished, matrix to generate the tree was provided by N.H. Williams and M.W. Whitten). Values on the branches represent bootstrap support. Red represents species with strong CAM, orange species with weak CAM, and green species with C3 photosynthesis. Representative images of corresponding floral morphology are shown to the right of each species designator. 151

Figure 2. Continuous net CO2 uptake by C3 orchid species during a 12 h light (white bar) and 12 h dark period (black bar). 152

Figure 3. Continuous net CO2 uptake by weak CAM orchid species during a 12 h light (white bar) and 12 h dark period (black bar). 153

Figure 4. Continuous net CO2 uptake by strong CAM orchid species during a 12 h light (white bar) and 12 h dark period (black bar). Root respiration for Trichocentrum nanum is included. 154

155

156 157 158 159 160

161 162 163 164 165 166 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185 186 187 188 189

Figure 5. Aligned partial nucleotide sequence of 1,100 base pairs from 35 PEPC isoforms from ten Oncidiinae species: Trichocentrum nanum (CAM), Rossioglossum ampliatum (CAM), Trichocentrum carthaginense (CAM), Gomesa flexuosa (Weak CAM), Oncidium sphacelatum (Weak CAM), Oncidium panamense (Weak CAM), Rossioglossum insleayi (Weak CAM), Oncidium sotoanum (C3), Oncidium maduroi (C3), Rossioglossum krameri (C3). Dark shading represents identical residues, and light shading represents conserved residues. 190

191 192 193 194 195

Figure 6. Aligned partial protein sequence of 363 aminoacid from 35 PEPC isoforms from ten Oncidiinae species: Trichocentrum nanum (CAM), Rossioglossum ampliatum (CAM), Trichocentrum carthaginense (CAM), Gomesa flexuosa (Weak CAM), Oncidium sphacelatum (Weak CAM), Oncidium panamense (Weak CAM), Rossioglossum insleayi (Weak CAM), Oncidium sotoanum (C3), Oncidium maduroi (C3), Rossioglossum krameri (C3). Dark shading represents identical residues, and light shading represents conserved residues. Solid line below the sequence from positions 37-55 represent a highly conserved region of - the catalytic subdomain that participates in PEP/HCO3 binding. 196

Figure 7. Phylogenetic tree derived from neighbor joining analysis of 1,100-bp alignment of Ppc nucleotides sequences from ten closely-related Oncidiinae species. Isoforms were assigned based on relative abundance (depicted as percentage) with Ppc1 being always the most abundant isoform followed by Ppc2-Ppc5. Bootstrap values (1000 replicates) are shown on the branches. (A) Cluster with the least abundant isoforms for all species (non-photosynthetic PEPC). (B) Cluster with the most abundant isoforms for all species including the PEPC genes likely recruited for CAM (indicated by arrow). Distance was calculated using the absolute number of differences between sequences (gap sites were ignored). The tree was rooted with the bacterial type PEPC (Ppc4) from Arabidopsis thaliana. 197

Figure 8. Phylogenetic tree derived from neighbor joining analysis of 363 amino acid alignments of Ppc from ten closely-related Oncidiinae species. Isoforms were assigned based on relative abundance (depicted as percentage) with Ppc1 being always the most abundant followed by Ppc2-Ppc5. Bootstrap values (1,000 replicates) are shown on the branches. (A) Cluster with the least abundant isoforms for all species (non-photosynthetic PEPC). (B) Cluster with the most abundant isoforms for all species including the PEPC genes likely recruited for CAM. Distance was calculated using the absolute number of differences between sequences (gap sites were ignored). The tree was rooted with the bacterial type PEPC from Arabidopsis thaliana. 198

Figure 9. Phylogenetic tree derived from neighbor joining analysis of 363 amino acid alignment of Ppc from ten closely-related Oncidiinae species and various CAM orchid species. Isoforms were assigned based on relative abundance (depicted as percentage) with Ppc1 being always the most abundant followed by Ppc2-Ppc5. Bootstrap values (1000 replicates) are shown on the branches. (A) Cluster with the least abundant isoforms for all species (non-photosynthetic PEPC). (B) Cluster with the most abundant isoforms for all species including the PEPC genes likely recruited for CAM. Distance was calculated using the absolute number of differences between sequences (gap sites were ignored). The tree was rooted with the bacterial type PEPC (Ppc4) from Arabidopsis thaliana. 199

Figure 10. Protein alignment for six isoforms from three Oncidiinae species, including the 3’UTR region recovered using 3’RACE. Dark shading represents identical residues and light shading represents conserved residues.

200 201 202 203 204 205

Figure 11. Aligned partial nucleotide sequence of 876 base pairs between PEPC contig 35735 obtained from 454 pyrosequencing of a strong CAM species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (Weak CAM), and Oncidium maduroi (C3). Dark shading represents identical residues, and light shading represents conserved residues.

206 207

Figure 12. Aligned partial nucleotide sequence of 221 base pairs between PEPC contig 10159 obtained from 454 pyrosequencing of a strong CAM species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (Weak CAM), and Oncidium maduroi (C3). Dark shading represents identical residues, and light shading represents conserved residues. 208

209

Figure 13. Aligned partial nucleotide sequence of 224 base pairs between PEPC contig 36974 obtained from 454 pyrosequencing of a strong CAM species (Rossioglossum ampliatum) and ten PEPC isoforms obtained from cDNA cloning sampling from three Oncidiinae species: Rossioglossum ampliatum (CAM), Oncidium panamense (Weak CAM), and Oncidium maduroi (C3). Dark shading represents identical residues, and light shading represents conserved residues. 210

Chapter V

Large-scale microarray gene expression profiling in Oncidiinae orchids

along the C3 to CAM evolutionary continuum

ABSTRACT

We conducted 454 sequencing of cDNA prepared from RNA sampling from leaf, flower, pseudobulb and root tissue of the strong CAM species Rossioglossum ampliatum, which resulted in 189 Mb of DNA sequence, 41,115 contigs, and 100,889 singletons. To understand the molecular mechanisms responsible for the recruitment of CAM-specific genes, a custom NimbleGen microarray was designed and fabricated, and contained 60 mer probesets. mRNA expression profiling using a C3 species (Oncidium maduroi), a weak CAM species (Oncidium panamense), and a strong CAM species (Rossioglossum ampliatum), showed that C3 and weak CAM species had average hybridization intensities that diverged from the strong CAM species by 2 and 3 percent, respectively. We used the log2 ratio (R) of relative transcript abundance between contrasting photosynthetic pathways as an index of variation in genetic regulation needed to perform different pathways. From 13,566 genes that showed a significant 4.6-fold difference in expression levels from the comparisons between CAM, C3 and weak CAM, 4,520 genes showed a greater than 4.6-fold increase in the ratio of CAM/C3 (RCAM/C3) relative transcript abundance, whereas 3,745 genes showed a greater than 4.6-fold decrease in RCAM/C3 211 transcript abundance. Similarly, 4,514 genes showed a greater than 4.6-fold increase in the ratio of weak CAM/C3 (RWCAM/CAM) relative transcript abundance, whereas 3,528 genes showed a greater than 4.6-fold decrease in RWCAM/CAM transcript abundance. Also,

3,378 genes showed a greater than 4.6-fold increase in the ratio of weak CAM/C3

(RWCAM/C3) compared to a 3,561 genes showing a greater than 4.6-fold decrease in

RWCAM/C3 transcript abundance. A maximal increase or decrease in relative transcript abundance of more than 1,000- and 500-fold, respectively, was observed. The results of this study provide an extensive catalogue of gene expression patterns that lay the foundation for future work aimed at understanding CAM-specific expression patterns and the functional requirements needed for CAM. Furthermore, these data can be used to improve our understanding of molecular genetic underpinnings required for CAM by setting the stage for in-depth analysis of those transcriptional control regions responsible for altered gene expression patterns associated with CAM evolution.

INTRODUCTION

CAM is one of the best-understood examples of an ecophysiological metabolic adaptation for plants in CO2 and water-limited environments, including tropical epiphytes that must thrive in epiphytic habitats with intermittent or seasonal water availability

(Silvera et al. 2005). However, relatively little is known about its evolutionary origin.

The expression of CAM is highly plastic and largely controlled by environmental conditions. For example, the C3-CAM species Mesembryanthemum crystallinum can complete its life cycle in the C3 mode if grown under non-stressed conditions (Winter and

Holtum 2007) or it can switch from C3 to CAM if grown under various stress treatments 212 such as drought, high light, or high salinity concentrations (Winter and Holtum 2005).

During the evolution of CAM from C3 photosynthesis, to weak CAM, to strong CAM modes, many CAM-specific genes are likely to be differentially expressed. Also, selective recruitment of genes is expected to be more strongly expressed and circadianly regulated, as a diagnostic indicator of their recruitment, to serve CAM-related functions.

For example, many genes with CAM function show shifts in expression patterns with as much as 28% of genes (out of 8,455 genes) showing a significant change in mRNA expression patterns following induction from C3 to CAM in M. crystallinum (Cushman et al. 2008). In Arabidopsis, up to 11% of genes (out of 11,521 genes), showed significant diurnal expression patterns with as much as 2% of those being associated with circadian rhythms (Schaffer et al. 2001).

Studies using microarray analysis can be used to identify underlying changes in phases due to environmental stresses or physiological cues, gain knowledge of potential gene function and coordination of gene expression, and are particularly helpful in the discovery of new putative genes involved in biochemical pathways. For example, changes in gene expression between vegetative phases or developmental transition between juvenile to adult phases assessed in maize using microarray analysis, showed that the largest class of genes induced were those involved in photosynthesis, with 221 expressed sequence tags (EST) showing increased abundance in juveniles and 28 ESTs showing increased abundance in adults (Strable et al. 2008). Another study using microarray analysis of an Arabidopsis mutant (CHL27) deficient in the chlorophyll (Chl- pigments that trap light energy in the antenna system) biosynthetic pathway, showed that several nuclear genes involved in photosynthesis were repressed, including genes 213 involved in the light-harvesting complex I and II and Photosystem I and II (Bang et al.

2008). Similarly, a study using microarray analysis to compare the photosynthetic acclimation of plants growing in different environment, found that the transfer of plants from low to high light was accompanied by a substantial, but transient, increase in expression of a gene encoding a glucose-6-phosphate/phosphate translocator (GPT2) with a key role in determining the fitness of plants (Athanasiou et al. 2010). However, understanding the mechanisms that coordinate large number of genes through the progression from C3 to weak CAM to strong CAM remains a challenge.

In this study we used functional genomics tools in the form of a cDNA library constructed using 454 pyrosequencing of cDNA from the strong CAM orchid species

(Rossioglossum ampliatum), and the use of custom oligonucleotide microarray to survey mRNA expression patterns diagnostic of CAM evolution. Estimates based on recent published reports in maize and Arabidopsis using GS20 instrumentation (~100 bp read length, Emrich et al. 2007), suggest that tags obtained for approximately 70-80% of expressed genes are likely to be represented and allow for the assembly of full-length sequences for oligonucleotide-based microarray design. With the introduction of the GS

FLX instrument, the average read length has increased to ~200 bp resulting in more robust assembly of cDNA contigs.

The orchid cDNA library produced can be used to conduct a cross-species comparative analyses of key pathways, regulatory and transporter components, and genes involved in signaling, with other CAM model species such as Mesembryanthemum crystallinum, Kalanchoë daigremontiana and Clusia species (Cushman, 2001). Using the

454-pyrosequencing library as a template for the presence of CAM in other species 214

within the subtribe Oncidiinae, we can determine expression profiles diagnostic for C3 photosynthesis, C3-CAM intermediates (weak CAM), and strong CAM to gauge the degree of gene expression changes that have occurred during the course of evolution. mRNA expression profiling was conducted using a C3 species (Oncidium maduroi), a weak CAM species (Oncidium panamense), and a strong CAM species (Rossioglossum ampliatum) by comparison of average hybridization intensities across the C3 to weak

CAM to strong CAM diversion.

MATERIAL AND METHODS

Plant material

Oncidiinae orchid plants were grown in a closed greenhouse at the University of Nevada

Reno. Daily temperature within the greenhouse varied from 17˚C to 35˚C with RH varying from 40% to 80%, and photon flux density (PFD) of 200 !mol m-2 sec-1. Plants were watered daily and nutrients were supplied once a week with a combination of slow release fertilizer (Osmocote 19-6-12 formula) and commercial fertilizer solution (Schultz

19-31-17 formula). Samples for 454 pyrosequencing were collected from combined tissues from pseudobulb, leaves at various developmental stages, healthy roots, flowers, inflorescence stems and buds, using a strong CAM orchid species (Rossioglossum ampliatum). Samples for microarray analysis were collected from combined leaf and root tissues collected at 02:00 h from 4 biological replicates for each of a C3 species

(Oncidium maduroi), a weak CAM species (Oncidium panamense), and a strong CAM species (Rossioglossum ampliatum). All samples were flash frozen in liquid nitrogen immediately after harvesting, and stored at –80ºC for isolation of RNA. 215

RNA isolation

Tissue was ground in liquid nitrogen and total RNA was isolated using the RNeasy Midi

Kit (Qiagen, Inc., Valencia, CA) with a modified PEG-RNA extraction method that utilizes high-molecular weight polyethylene glycol (Gehrig et al. 2000). RNA produced was cleaned from proteins with phenol:chloroform extractions and further purified using the RNeasy Mini Kit (Qiagen, Inc.) following manufacturer’s instructions. RNA quality was examined by gel electrophoresis, and the integrity and concentration was examined using a NanoDrop ND-1000 UV-Vis spectrophotometer (Nanodrop Technologies Inc.,

Rockland, DE).

454 Life Sciences (Roche) Sequencing Technology

A high-throughput, ultra-deep EST sequencing (454 Life Sciences, a Roche company,

Branford, CT) was constructed from cDNA library of a strong CAM species

(Rossioglossum ampliatum). Massively parallel 454 sequencing-by-synthesis was completed using the Roche GS-FLX Genome Sequencer. EST sequencing using 454- pyrosequencing technologies allows for a more cost-effective and comprehensive analysis of the transcriptome (20-fold increase in number of reads). A directional cDNA library was constructed using UNI-ZAP directional cloning vector (Stratagene, Inc. La

Jolla, CA) and normalized (Bonaldo et al. 1996) using the W.M. Keck Biotechnology

Center’s cDNA library normalization and subtraction service (University of Illinois at

Urbana-Champaign; http://www.biotec.uiuc.edu). The use of normalized libraries allowed the identification and characterization of low abundance transcripts. Quality control procedures to validate the normalized status of the cDNA library included pilot 216 sequencing of 192 clones to survey the frequency of abundant transcripts such as those encoding the RUBISCO small subunit.

NimbleGen microarry fabrication and analysis

A Roche NimbleGen custom microarray was designed using the fabrication and design services (http://www.NimbleGen.com), and the mRNA expression profiling hybridizations produced was processed as Affymetrix data (Davletova et al. 2005). The array contained 41,115 genes and controls, with each gene model represented by 10 x 60 mer probes. Heterologous hybridization was used to target RNA from a weak CAM species (Oncidium panamense) and a C3 species (Oncidium maduroi) to the array constructed from a strong CAM species (Rossioglossum ampliatum), and homologous hybridization was used to target RNA from the strong CAM species Rossioglossum ampliatum. RMA (Robust Multi-Array Average; Bioconductor rel. 1.2) was used to normalize the microarray data (Irizarry et al. 2003). Principal component analysis (PCA) and hierarchical clustering were used to visualize structure in the data. Expression values were computed from the raw Pairs files by first applying the RMA model of probe- specific background correction of PM (perfect match) probes. These corrected probe values were then normalized via quantile normalization, and a median polish was applied to compute one expression measure from all probe values (Cushman et al. 2008).

Resulting expression values were log2-transformed. Data quality was monitored by digestion curves describing trends in RNA degradation between the 5’ end and the 3’ end in each probeset. Pearson correlation coefficients and Spearman rank coefficients were computed on the RMA expression values (log2-transformed) for each set of biological 217 replicates. Clustering was performed using the statistical programming language R, with a modified version of the hclust procedure to enable clustering with correlation. A

Bonferroni correction was applied for multiple testing with a specified cutoff, to generate a list of genes with significant differential expression. Concordance in gene expression profiles across species comparisons was quantified as fold-changes.

RESULTS AND DISCUSSION

Massively parallel 454 sequencing-by-synthesis using Roche GS-FLX Genome

Sequencer of two large plates yielded 1,028,667 cleansed and assembled reads, a total of

41,655 contigs and 100,899 singletons. A total of 39,058 probes were converted to a custom designed Rossioglossum ampliatum microarray and used to survey mRNA expression patterns between closely-related Oncidiinae species expressing a range of photosynthetic pathways from C3 to weak CAM to strong CAM modes. Four duplicate cDNA samples per species were hybridized to the custom NimbleGen microarray. The data were cleansed by removing inconsistently hybridizing elements.

After performing an ANOVA and a Bonferroni correction, about 31,000 probe sets were retained from the original 39,058 probe sets. From these, 13,566 genes showed significant 4.6-fold changes from the comparisons between CAM, weak CAM and C3. A subset of 4,520 genes showed a greater than 4.6-fold increase in the ratio of CAM/C3

(RCAM/C3) transcript abundance, whereas 3,735 genes showed a greater than 4.6-fold decrease in RCAM/C3 transcript abundance. Similarly, 4,514 genes showed a greater than

4.6-fold increase in the ratio of weak CAM/C3 (RWCAM/CAM) relative transcript abundance, whereas 3,528 genes showed a greater than 4.6-fold decrease in RWCAM/CAM 218 transcript abundance. Also, 3,378 genes showed a greater than 4.6-fold increase in the ratio of weak CAM/C3 (RWCAM/C3) compared to 3,561 genes showing a greater than 4.6- fold decrease in RWCAM/C3 transcript abundance. A maximal increase or decrease in relative transcript abundance of more than 1,000- and 500-fold, respectively, was observed.

Principal component analysis (PCA) was used to partition the transcriptome variability into coordinates and visualize the variance among the different species sampled (Fig. 1). The two principal components generated explained 97.8% of the overall transcriptome variability (87.9% for axis 1 and 9.9% for axis 2) and partitioned the data into two distinct groups, one represented by weak CAM species and C3 species combined, and a second group represented by strong CAM species alone (Fig. 1), indicating pronounced differences in expression patterns between strong CAM and C3 or weak CAM. These results were consistent with the comparison between mRNA expression changes between the weak CAM and CAM (Fig. 2A), C3 vs. CAM (Fig. 2B), and weak CAM vs. C3 (Fig. 2C) performed by computing the fold-changes (log2-scale)

st nd for the 1 and 2 standard deviation. The scatterplots showed that CAM and C3, and weak CAM and CAM were very different (Fig. 2), whereas C3 and weak CAM were more similar than other comparisons, based on mRNA expression patterns. The two horizontal lines representing fold-changes correspond to a ratio of 1 and -1 on the log2

(ratio) scale.

Expression levels were also determined for the RMA-normalized probeset values of the microarrays and a 4.6-fold cutoff filter was used to identify genes with 4.6-fold differences in expression for comparison of RWCAM/CAM and RCAM/C3 (Fig. 3). P-value was 219 calculated and corrected by Benjiamani-Hochberg multiple test correction and only those genes with one or more 4.6-fold difference and adj. P<0.05 were considered (Fig. 3).

When looking at the log2-transformed intensity ratio RWCAM/CAM and RCAM/C3, the larger number of genes that showed increased mRNA abundance was found between CAM and

C3 (Red group, Fig. 3) followed by genes with increased mRNA abundance between weak CAM and CAM (Yellow group, Fig. 3). Similarly, the larger number of genes with decreased mRNA abundance was found between weak CAM and CAM (Dark purple group, Fig. 3) followed by genes with decreased mRNA abundance between CAM and

C3. Of interest from an evolutionary point of view, will be the study of the circadian clock control of those genes that show significant increased mRNA abundance between

CAM and C3, and also show minimal changes between weak CAM and CAM (Yellow group, Fig. 3).

Categorization of CAM-related genes

A list of 169 genes that exhibited significant changes in transcript abundance and are related to the CAM pathway are shown in Table 1. From these, genes involved in starch degradation and synthesis, cellular signal and communication, and glycolysis and gluconeogenesis were the most abundant (Table 1). Genes with a role in starch degradation and mobilization have a central role in CAM function. For example, 40- 60% of the starch degraded at night is used to supply the PEP and exporting of soluble sugars

(Borland and Dodd, 2002). Also, expression of genes encoding starch degradation enzymes is under circadian clock control and regulated by day length (Lu et al. 2005)

Genes related to the glycolysis and gluconeogenesis pathways are needed for anaplerotic 220

functions including the fueling of nocturnal CO2 fixation by PEPC, the flow of carbon through gluconeogenesis and starch biosynthesis (Cushman et al. 2008).

Genes that showed the most decrease in transcript abundance between C3 and weak CAM species (RC3/WCAM, Table 2) included those encoding functions associated with dehydration-responsiveness, oxygen binding, transcription factors and cytochromes.

From these, dehydration responsive enzymes showed up to 64-fold change between weak

CAM and C3 (Table 2). Many known dehydration-responsive genes are involved in drought responses and include late embryogenesis abundant (LEA) protein, and are involved in the prevention of water loss and dehydration responses.

Genes that showed the most decrease in transcript abundance between CAM and

C3 species (RCAM/C3, Table 3) included those encoding functions associated with the synthesis of starch, transferases, oxygen binding, transaminases, proteases, and hydrolases. From these genes, starch synthase showed a 256-fold change between CAM and C3. This result highlights the role of starch mobilization in CAM function.

Genes that showed the most decrease in transcript abundance between CAM and weak CAM species (RCAM/WCAM, Table 4) include those encoding functions associated with oxygen and calmodulin binding, helicases, transcription factors, transaminases, transferases, transporters and synthesis of starch. These genes showed from 32 to 128- fold changes between CAM and weak CAM, and are predicted to have roles in metabolite mobilization and carbon flux, and are important components of the self- sustained rhythms of nocturnal CO2 fixation needed for CAM function.

Genes that showed the most increase in transcript abundance between C3 and weak CAM species (RC3/WCAM, Table 5) included those encoding functions associated 221 with hydrolases, lipoxygenases, protein folding, transporters, transferases, calmodulin binding, and hydrolases. These genes showed 16 to 64-fold changes between C3 and weak CAM species, and are involved in carbohydrate synthesis and degradation. Genes that showed the most increase in transcript abundance between RCAM/C3, and RCAM/WCAM

(Table 6-7) included a majority of unidentified transcripts with 200- to 1000-fold changes in transcript abundance. From those identified genes, lipoxygenase, hydrolyase/oxygen binding protein, and a cyclin-dependent kinase also showed changes from 200- to 1000- fold.

Many of the most induced genes with important roles in CAM function, included phosphoenolpyruvate carboxylase, phosphoenolpyruvate carboxylase kinase, glucose-6- phosphate translocator, glyceraldehyde-3-phosphate dehydrogenase, pyruvate orthophosphate dikinase, carbonic anhydrase, alpha- and beta-amylase, fructose 1,6 biphosphate, fructose biphosphate aldolase, peroxisomal NAD-malate dehydrogenase, malate dehydrogenase NADP, cytosolic triose phosphate isomerase, and the blue light receptor 2 (Fig. 4).

Genes that also have an important role in CAM function or transposable elements constitute an important portion of the genome and are maintained by DNA methylations, under the control of environmental factors that can activate or repress them by DNA demethylation (Chinnusamy and Zhu, 2009). Epigenetic stress memory or the ability of plants to cope with stress by phenotypic plasticity adjustment is an important mechanism of stress resistance. In CAM plants, DNA hypermethylation is involved in stress-induced switches from C3 to CAM (Dyachenko et al. 2006). DNA methylation and nucleosome histone post-translational modification, therefore, play a key role in gene expression and 222 plant development under environmental stresses associated with mRNA changes between

C3 and CAM. Many stress-regulated and stress tolerance genes are expected to be involved in changes in histone modifications and DNA methylation. MicroRNAs are also widely known to regulate genes involved in plant growth and development, in addition to having a role in plant abiotic stress by targeting Cu/Zn superoxide dismutases (SODs) and transcription factors. SOD convert toxic reactive oxygen species (ROS) to less toxic hydrogen peroxide (Shukla et al. 2008). More recently, stress inducible microRNAs from

Arabidopsis have been identified and found to be unregulated during salt, osmotic and cold stresses (Liu et al. 2008). Transcription factors also positively regulate photosynthesis related genes or plastid differentiation (Nakamura et al. 2009). Future studies that include comparison between day and night cycles coupled with environmental drought stresses in these orchid species are needed to distinguish those genes that are involved in epigenetic regulation and stress regulation, and those genes that are under circadian clock control, to ultimately tease apart the metabolite control that are needed to maintain CAM rhythms in plants.

Conclusions

In conclusion, we report the results of the first large-scale mRNA expression profiling performed in orchid species. From about 30,000 probe sets, more than 4,000 genes showed a greater than 4.6-fold increase in relative transcript abundance in RCAM/C3, and more than 3,000 genes showed a greater than 4.6-fold decrease in relative transcript abundance in RCAM/C3. The results provided here will serve as a platform for future studies, and will provide a catalogue of gene expression patterns available for work 223 aimed at understanding CAM specific expression patterns that can be used to further understand gene regulation by in-depth analysis of the transcriptional control regions responsible for altered gene expression patterns associated with CAM evolution. Future studies that include expression profiling over the day/night cycle of these orchid species, studies including drought stress, and those coupled with 24 h gas exchange, and metabolite analysis will aid in understanding the circadian rhythm of genes involved in

CAM.

ACKNOWLEDGMENTS

We would like to thank Dr. Karen Schlauch and Richard Tillett for microarray analysis and graphics preparation, Becky Albion for technical assistance in the lab, Dr. Gaspar

Silvera for supplying the orchid plant species, and the Cushman Lab for support. This work was supported by funding from the U.S. Environmental Protection Agency under the Greater Research Opportunities Graduate Program Agreement no. MA 91685201 (to

K.S.) and National Science Foundation NSF IOB-0543659 (to JCC). This publication was also made possible by NIH Grant Number P20 RR-016464 from the INBRE

Program of the National Center for Research Resources through its support of the Nevada

Genomics, Proteomics and Bioinformatics Centers. The views expressed in this publication are solely those of the authors and the EPA does not endorse any products or commercial services mentioned in this publication. 224

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Table 1. List of genes with CAM-related function derived from microarray data. Values represent normalized data expressed as hybridization intensity values. Leaves were collected at 2 AM from four biological replicates from a C3 species Oncidium maduroi, a weak CAM species Oncidium panamense and a strong CAM species Rossioglossum insleayi.

Weak Adjusted Feature CAM C3 CAM CAM/ ANOVA Description C3 p-value CONTIG10801 2397.68 135.45 169.11 17.70 0.0000 2-phosphoglycerate kinase-related CONTIG38691 25.12 31.86 33.99 0.79 0.0580 2-phosphoglycerate kinase-related CONTIG23911 45.25 160.64 99.47 0.28 0.0019 2-phosphoglycerate kinase-related 2,3-biphosphoglycerate-independent phosphoglycerate mutase family protein / CONTIG38089 5567.33 3525.03 2842.90 1.58 0.0006 phosphoglyceromutase family protein 2,3-biphosphoglycerate-independent phosphoglycerate mutase-related / phosphoglyceromutase- CONTIG01760 2323.45 3050.20 3584.73 0.76 0.0001 related 2,3-biphosphoglycerate-independent phosphoglycerate mutase, putative / phosphoglyceromutase, CONTIG00841 21088.77 9860.72 9026.33 2.14 0.0000 putative CONTIG06407 1121.27 1147.42 852.93 0.98 0.0735 ADP-glucose pyrophosphorylase family protein CONTIG00677 3303.78 4256.78 4769.06 0.78 0.0008 ADP-glucose pyrophosphorylase family protein CONTIG13134 1100.64 2126.85 2069.45 0.52 0.0000 ADP-glucose pyrophosphorylase family protein CONTIG30886 734.03 349.78 234.08 2.10 0.0013 ATAMY1, AMY1 | AMY1/ATAMY1 (alpha-amylase-like); alpha-amylase CONTIG04273 44.33 95.96 83.95 0.46 0.0059 ATAMY1, AMY1 | AMY1/ATAMY1 (alpha-amylase-like); alpha-amylase CONTIG01213 537.27 3202.76 2918.21 0.17 0.0000 ATAMY2, AMY2 | AMY2/ATAMY2 (alpha-amylase-like 2); alpha-amylase CONTIG11505 7643.00 3560.19 6525.96 2.15 0.0004 ATAMY3, AMY3 | AMY3/ATAMY3 (alpha-amylase-like 3); alpha-amylase CONTIG12749 2946.15 1567.99 3197.61 1.88 0.0001 ATAMY3, AMY3 | AMY3/ATAMY3 (alpha-amylase-like 3); alpha-amylase CONTIG39452 507.57 446.43 426.69 1.14 0.5484 ATAMY3, AMY3 | AMY3/ATAMY3 (alpha-amylase-like 3); alpha-amylase ATGNA1 | ATGNA1 (Arabidopsis thaliana glucose-6-phosphate acetyltransferase 1); N- CONTIG27111 27.94 97.43 53.48 0.29 0.0234 acetyltransferase/ glucosamine 6-phosphate N-acetyltransferase CONTIG29814 8964.62 5485.85 5608.63 1.63 0.0020 ATGWD3, OK1, PWD | PWD (phosphoglucan water dikinase); catalytic CONTIG23316 1071.51 2340.09 1448.42 0.46 0.0376 ATGWD3, OK1, PWD | PWD (phosphoglucan water dikinase); catalytic CONTIG11968 24.97 26.73 132.08 0.93 0.0625 ATISA3, ISA3 | ATISA3/ISA3 (isoamylase 3); alpha-amylase CONTIG38932 4539.81 5982.18 11125.55 0.76 0.0002 ATISA3, ISA3 | ATISA3/ISA3 (isoamylase 3); alpha-amylase CONTIG32115 794.06 5664.08 2811.56 0.14 0.0000 ATISA3, ISA3 | ATISA3/ISA3 (isoamylase 3); alpha-amylase CONTIG03146 914.65 8247.33 5941.34 0.11 0.0000 ATISA3, ISA3 | ATISA3/ISA3 (isoamylase 3); alpha-amylase ATNADP-ME1 | ATNADP-ME1 (NADP-malic enzyme 1); malate dehydrogenase (oxaloacetate-decarboxylating) (NADP+)/ malic enzyme/ oxidoreductase, acting on NADH or CONTIG26988 266.08 7614.48 9460.11 0.03 0.0000 NADPH, NAD or NADP as acceptor ATNADP-ME3 | ATNADP-ME3 (NADP-malic enzyme 3); malate dehydrogenase (oxaloacetate-decarboxylating) (NADP+)/ malic enzyme/ oxidoreductase, acting on NADH or CONTIG33907 19008.27 19161.35 20135.84 0.99 0.7667 NADPH, NAD or NADP as acceptor ATNADP-ME4 | ATNADP-ME4 (NADP-malic enzyme 4); malate dehydrogenase (oxaloacetate-decarboxylating) (NADP+)/ malic enzyme/ oxidoreductase, acting on NADH or CONTIG01284 6144.51 7921.50 2786.45 0.78 0.0006 NADPH, NAD or NADP as acceptor CONTIG01021 13692.00 22971.48 11861.02 0.60 0.0002 ATNADP-ME4 | ATNADP-ME4 (NADP-malic enzyme 4); malate dehydrogenase 227

(oxaloacetate-decarboxylating) (NADP+)/ malic enzyme/ oxidoreductase, acting on NADH or NADPH, NAD or NADP as acceptor ATNADP-ME4 | ATNADP-ME4 (NADP-malic enzyme 4); malate dehydrogenase (oxaloacetate-decarboxylating) (NADP+)/ malic enzyme/ oxidoreductase, acting on NADH or CONTIG40840 574.03 5288.28 5553.41 0.11 0.0000 NADPH, NAD or NADP as acceptor ATPHS2, PHS2 | ATPHS2/PHS2 (alpha-glucan phosphorylase 2); phosphorylase/ transferase, CONTIG38596 5881.74 4072.39 3032.66 1.44 0.0004 transferring glycosyl groups ATPHS2, PHS2 | ATPHS2/PHS2 (alpha-glucan phosphorylase 2); phosphorylase/ transferase, CONTIG00014 3660.11 4298.36 2823.23 0.85 0.1468 transferring glycosyl groups CONTIG35735 35531.89 11887.00 16190.48 2.99 0.0002 ATPPC1 | ATPPC1 (Phosphoenolpyruvate carboxylase 1); phosphoenolpyruvate carboxylase CONTIG38637 6953.46 2628.34 1598.22 2.65 0.0019 ATPPC1 | ATPPC1 (Phosphoenolpyruvate carboxylase 1); phosphoenolpyruvate carboxylase CONTIG29985 19086.43 7539.23 9085.66 2.53 0.0009 ATPPC1 | ATPPC1 (Phosphoenolpyruvate carboxylase 1); phosphoenolpyruvate carboxylase CONTIG29413 3003.63 5526.18 3630.70 0.54 0.0008 ATPPC1 | ATPPC1 (Phosphoenolpyruvate carboxylase 1); phosphoenolpyruvate carboxylase CONTIG36974 9465.97 21498.77 15267.53 0.44 0.0000 ATPPC1 | ATPPC1 (Phosphoenolpyruvate carboxylase 1); phosphoenolpyruvate carboxylase CONTIG10159 2235.47 6251.72 10590.51 0.36 0.0000 ATPPC1 | ATPPC1 (Phosphoenolpyruvate carboxylase 1); phosphoenolpyruvate carboxylase CONTIG16824 13669.09 35285.08 14065.68 0.39 0.0000 ATPPC2 | ATPPC2 (Phosphoenolpyruvate carboxylase 2); phosphoenolpyruvate carboxylase CONTIG30557 12532.71 4229.42 2521.97 2.96 0.0000 ATPPC3 | ATPPC3 (Phosphoenolpyruvate carboxylase 3); phosphoenolpyruvate carboxylase CONTIG09191 19979.71 9986.10 8546.99 2.00 0.0016 ATPPC3 | ATPPC3 (Phosphoenolpyruvate carboxylase 3); phosphoenolpyruvate carboxylase CONTIG09192 8406.49 5501.00 5860.81 1.53 0.1921 ATPPC3 | ATPPC3 (Phosphoenolpyruvate carboxylase 3); phosphoenolpyruvate carboxylase CONTIG02012 10159.00 7242.90 7755.01 1.40 0.0221 ATPPC3 | ATPPC3 (Phosphoenolpyruvate carboxylase 3); phosphoenolpyruvate carboxylase CONTIG12507 2793.76 4692.14 4380.56 0.60 0.0015 ATPPC3 | ATPPC3 (Phosphoenolpyruvate carboxylase 3); phosphoenolpyruvate carboxylase CONTIG18296 439.09 5099.74 7013.16 0.09 0.0000 ATPPC3 | ATPPC3 (Phosphoenolpyruvate carboxylase 3); phosphoenolpyruvate carboxylase CONTIG09267 876.87 661.91 1146.35 1.32 0.0003 ATPU1, ATLDA | ATLDA/ATPU1 (Pullulanase 1); alpha-amylase/ limit dextrinase CONTIG26670 497.77 1058.37 1633.86 0.47 0.0009 ATPU1, ATLDA | ATLDA/ATPU1 (Pullulanase 1); alpha-amylase/ limit dextrinase ATTDT, ATSDAT | ATSDAT/ATTDT (Tonoplast dicarboxylate transporter); malate CONTIG33417 1060.91 424.53 305.21 2.50 0.0699 transmembrane transporter/ sodium:dicarboxylate symporter ATTDT, ATSDAT | ATSDAT/ATTDT (Tonoplast dicarboxylate transporter); malate CONTIG33456 993.58 550.76 869.29 1.80 0.0114 transmembrane transporter/ sodium:dicarboxylate symporter ATTDT, ATSDAT | ATSDAT/ATTDT (Tonoplast dicarboxylate transporter); malate CONTIG37346 2769.36 2030.50 761.37 1.36 0.0080 transmembrane transporter/ sodium:dicarboxylate symporter CONTIG12283 17672.13 6422.81 5666.65 2.75 0.0000 BE3, SBE2.1 | SBE2.1 (Starch branching enzyme 2.1); 1,4-alpha-glucan branching enzyme CONTIG11371 14607.54 5738.39 4970.85 2.55 0.0000 BE3, SBE2.1 | SBE2.1 (Starch branching enzyme 2.1); 1,4-alpha-glucan branching enzyme CONTIG19797 313.04 606.90 311.74 0.52 0.0017 BMY4, BAM7 | BAM7/BMY4 (beta-amylase 7); beta-amylase CONTIG08791 332.10 1107.29 964.80 0.30 0.0001 BMY4, BAM7 | BAM7/BMY4 (beta-amylase 7); beta-amylase CONTIG05263 984.99 3826.32 2575.11 0.26 0.0000 BMY4, BAM7 | BAM7/BMY4 (beta-amylase 7); beta-amylase CONTIG15864 202.93 1262.24 858.18 0.16 0.0000 BMY7, TR-BAMY, BAM1 | BAM1/BMY7/TR-BAMY (beta-amylase 1); beta-amylase CONTIG31776 23247.94 8037.72 822.41 2.89 0.0000 BMY8, BAM3, CT-BMY | CT-BMY (beta-amylase 3, Beta-amylase 8); beta-amylase CONTIG08828 2085.21 4459.90 3487.86 0.47 0.0001 BMY8, BAM3, CT-BMY | CT-BMY (beta-amylase 3, Beta-amylase 8); beta-amylase CONTIG36019 451.67 1245.37 1450.67 0.36 0.0000 BMY9, BAM2 | BAM2/BMY9 (beta-amylase 2); beta-amylase CONTIG32032 187.68 1177.23 493.10 0.16 0.0000 CA1 | CA1 (carbonic anhydrase 1); carbonate dehydratase/ zinc ion binding CA18, BETA CA2, CA2 | CA2 (beta carbonic anhydrase 2); carbonate dehydratase/ zinc ion CONTIG28739 2764.98 287.54 1085.08 9.62 0.0000 binding CA18, BETA CA2, CA2 | CA2 (beta carbonic anhydrase 2); carbonate dehydratase/ zinc ion CONTIG36917 22741.87 4051.22 2553.97 5.61 0.0001 binding CONTIG31438 558.18 638.64 594.35 0.87 0.4746 carbonic anhydrase family protein / carbonate dehydratase family protein CONTIG08712 161.71 169.81 1051.13 0.95 0.0000 carbonic anhydrase family protein 228

CONTIG06263 631.96 892.77 1338.27 0.71 0.1337 carbonic anhydrase family protein CONTIG14687 129.99 427.75 1111.49 0.30 0.0000 carbonic anhydrase family protein CONTIG02202 6611.50 14924.55 5357.76 0.44 0.0000 dicarboxylate/tricarboxylate carrier (DTC) CONTIG31063 3097.30 2382.04 2333.81 1.30 0.1542 DIT1 | DIT1 (dicarboxylate transporter 1); oxoglutarate:malate antiporter CONTIG32333 17150.00 27247.89 21404.63 0.63 0.0003 DIT1 | DIT1 (dicarboxylate transporter 1); oxoglutarate:malate antiporter | CONTIG03747 7965.17 5321.66 8060.44 1.50 0.0004 EMB2729, BE1 | BE1/EMB2729 (branching enzyme 1); alpha-amylase CONTIG15929 2223.47 5916.49 4490.24 0.38 0.0039 EMB2729, BE1 | BE1/EMB2729 (branching enzyme 1); alpha-amylase fructose-1,6-bisphosphatase, putative / D-fructose-1,6-bisphosphate 1-phosphohydrolase, CONTIG05658 3927.43 33.32 55.83 117.86 0.0000 putative / FBPase, putative fructose-1,6-bisphosphatase, putative / D-fructose-1,6-bisphosphate 1-phosphohydrolase, CONTIG05661 9663.41 11489.83 7930.91 0.84 0.1755 putative / FBPase, putative fructose-1,6-bisphosphatase, putative / D-fructose-1,6-bisphosphate 1-phosphohydrolase, CONTIG01989 21515.18 4521.25 5170.67 4.76 0.0000 putative / FBPase, putative fructose-1,6-bisphosphatase, putative / D-fructose-1,6-bisphosphate 1-phosphohydrolase, CONTIG09454 26.60 54.57 48.36 0.49 0.0059 putative / FBPase, putative CONTIG28489 3472.20 63.76 76.18 54.46 0.0000 fructose-bisphosphate aldolase, putative CONTIG10012 29567.37 13551.31 23100.36 2.18 0.0000 fructose-bisphosphate aldolase, putative CONTIG00459 38670.49 27013.63 42058.72 1.43 0.0042 fructose-bisphosphate aldolase, putative CONTIG23954 58.31 126.73 85.54 0.46 0.1281 fructose-bisphosphate aldolase, putative CONTIG27919 11183.09 14342.84 9926.27 0.78 0.0035 fructose-bisphosphate aldolase, putative CONTIG11435 27.30 50.52 53.92 0.54 0.0973 fructose-bisphosphate aldolase, putative CONTIG02426 843.54 50.51 41.99 16.70 0.0000 fructose-bisphosphate aldolase, putative CONTIG36650 49537.35 51270.82 46360.26 0.97 0.0900 fructose-bisphosphate aldolase, putative CONTIG32937 470.11 99.79 68.66 4.71 0.0002 fructose-bisphosphate aldolase, putative CONTIG26775 22842.76 15597.67 30809.25 1.46 0.0000 fructose-bisphosphate aldolase, putative CONTIG34400 7693.27 2933.58 5050.46 2.62 0.0005 G6PD1 | G6PD1 (glucose-6-phosphate dehydrogenase 1); glucose-6-phosphate dehydrogenase CONTIG32823 1131.74 1434.86 2108.72 0.79 0.0000 G6PD2 | G6PD2 (glucose-6-phosphate dehydrogenase 2); glucose-6-phosphate dehydrogenase CONTIG05709 17673.83 11609.04 6033.66 1.52 0.0000 G6PD5 | G6PD5 (glucose-6-phosphate dehydrogenase 5); glucose-6-phosphate dehydrogenase CONTIG15589 520.38 328.50 333.24 1.58 0.1693 G6PD6 | G6PD6 (glucose-6-phosphate dehydrogenase 6); glucose-6-phosphate dehydrogenase CONTIG00729 3924.57 7477.55 7840.40 0.52 0.0000 GAMMA CA1 | GAMMA CA1 (Gamma carbonic anhydrase 1); carbonate dehydratase CONTIG17724 31.30 88.02 64.62 0.36 0.0001 GAMMA CA2, APFI | APFI; carbonate dehydratase GAMMA CAL2 | GAMMA CAL2 (Gamma carbonic anhydrase-like 2); acyltransferase/ CONTIG35220 691.83 37.80 175.10 18.30 0.0000 transferase GAMMA CAL2 | GAMMA CAL2 (Gamma carbonic anhydrase-like 2); acyltransferase/ CONTIG00828 1591.18 1121.06 962.79 1.42 0.0002 transferase CONTIG28163 21889.07 524.94 1472.14 41.70 0.0000 GAPA-2 | GAPA-2; glyceraldehyde-3-phosphate dehydrogenase CONTIG39782 51117.90 43732.44 37987.19 1.17 0.0020 GAPA-2 | GAPA-2; glyceraldehyde-3-phosphate dehydrogenase GAPB | GAPB (glyceraldehyde-3-phosphate dehydrogenase B subunit); glyceraldehyde-3- CONTIG04525 13906.49 3427.25 4639.00 4.06 0.0001 phosphate dehydrogenase GAPC-1, GAPC | GAPC (glyceraldehyde-3-phosphate dehydrogenase C subunit); CONTIG39176 28257.75 9578.11 13905.19 2.95 0.0000 glyceraldehyde-3-phosphate dehydrogenase GAPC-1, GAPC | GAPC (glyceraldehyde-3-phosphate dehydrogenase C subunit); CONTIG29435 25474.36 26804.19 30378.72 0.95 0.0295 glyceraldehyde-3-phosphate dehydrogenase CONTIG36407 22934.76 7505.38 14232.44 3.06 0.0001 GAPC-2 | GAPC-2; glyceraldehyde-3-phosphate dehydrogenase CONTIG39404 25613.63 9160.66 9473.36 2.80 0.0000 GAPC-2 | GAPC-2; glyceraldehyde-3-phosphate dehydrogenase CONTIG20135 392.35 680.56 1304.42 0.58 0.0009 GAPCP-2 | GAPCP-2; glyceraldehyde-3-phosphate dehydrogenase 229

CONTIG36724 13834.73 2567.44 4450.49 5.39 0.0000 glucan phosphorylase, putative CONTIG00844 2659.21 942.02 1075.83 2.82 0.0111 glucan phosphorylase, putative CONTIG36781 7054.05 7971.36 10081.55 0.88 0.0143 glucose-6-phosphate isomerase, cytosolic (PGIC) CONTIG05952 2461.74 3919.50 5018.59 0.63 0.0000 glucose-6-phosphate/phosphate translocator-related CONTIG01676 49.55 65.88 47.13 0.75 0.1227 glucose-6-phosphate/phosphate translocator, putative CONTIG27489 622.22 577.13 911.23 1.08 0.0005 glycerol-3-phosphate dehydrogenase, putative CONTIG06305 2536.98 4256.01 2303.52 0.60 0.0001 glycerol-3-phosphate transporter, putative / glycerol 3-phosphate permease, putative CONTIG05914 6681.80 1035.22 1115.85 6.45 0.0000 glycerol-3-phosphate transporter, putative / glycerol 3-phosphate permease, putative CONTIG18769 10650.45 24226.11 26814.68 0.44 0.0000 glycerol-3-phosphate transporter, putative / glycerol 3-phosphate permease, putative GPT1 | GPT1 (glucose-6-phosphate transporter 1); antiporter/ glucose-6-phosphate CONTIG06250 974.72 129.66 89.41 7.52 0.0000 transmembrane transporter GPT1 | GPT1 (glucose-6-phosphate transporter 1); antiporter/ glucose-6-phosphate CONTIG02056 1146.98 3874.28 3364.16 0.30 0.0000 transmembrane transporter GPT2 | GPT2 (glucose-6-phosphate/phosphate translocator 2); antiporter/ glucose-6-phosphate CONTIG33315 1101.20 400.27 905.01 2.75 0.0015 transmembrane transporter GPT2 | GPT2 (glucose-6-phosphate/phosphate translocator 2); antiporter/ glucose-6-phosphate CONTIG02442 158.73 117.39 454.81 1.35 0.0009 transmembrane transporter GPT2 | GPT2 (glucose-6-phosphate/phosphate translocator 2); antiporter/ glucose-6-phosphate CONTIG37173 370.81 2918.03 1663.30 0.13 0.0000 transmembrane transporter CONTIG17069 397.56 145.23 69.51 2.74 0.0055 malate dehydrogenase (NAD), mitochondrial CONTIG36954 10375.41 957.54 1858.33 10.84 0.0000 malate dehydrogenase (NAD), mitochondrial, putative CONTIG19913 25.12 36.33 38.24 0.69 0.0462 malate dehydrogenase (NAD), mitochondrial, putative CONTIG31652 6873.77 8091.34 7726.41 0.85 0.5831 malate dehydrogenase (NADP), chloroplast, putative CONTIG09276 21920.87 39282.54 39709.40 0.56 0.0000 malate dehydrogenase (NADP), chloroplast, putative CONTIG33747 32891.26 15940.76 22814.31 2.06 0.0000 malate dehydrogenase, cytosolic, putative CONTIG01838 3979.52 4727.15 10788.26 0.84 0.0000 malate dehydrogenase, cytosolic, putative CONTIG01839 2547.64 3329.49 4365.50 0.77 0.0001 malate dehydrogenase, cytosolic, putative CONTIG24450 27.18 39.56 53.93 0.69 0.0009 malate dehydrogenase, cytosolic, putative CONTIG06243 1693.92 2672.88 2395.40 0.63 0.0015 MDH | MDH (malate dehydrogenase); malate dehydrogenase PCK1, PEPCK | PCK1/PEPCK (phosphoenolpyruvate carboxykinase 1); ATP binding / CONTIG22281 2643.88 1632.47 1575.51 1.62 0.1658 phosphoenolpyruvate carboxykinase (ATP) PCK1, PEPCK | PCK1/PEPCK (phosphoenolpyruvate carboxykinase 1); ATP binding / CONTIG00902 521.14 386.93 399.41 1.35 0.4143 phosphoenolpyruvate carboxykinase (ATP) CONTIG35920 219.65 196.54 1012.54 1.12 0.0000 PEPKR1 | PEPKR1 (Phosphoenolpyruvate carboxylase-related kinase 1); kinase CONTIG17416 29.30 29.93 28.74 0.98 0.8141 PEPKR1 | PEPKR1 (Phosphoenolpyruvate carboxylase-related kinase 1); kinase CONTIG06839 21245.19 5770.37 8798.55 3.68 0.0000 PGK | PGK (phosphoglycerate kinase) CONTIG32724 29914.77 9252.55 8133.83 3.23 0.0000 PGK | PGK (phosphoglycerate kinase) CONTIG32647 28825.42 23398.38 25557.85 1.23 0.0767 PGK1 | PGK1 (phosphoglycerate kinase 1); phosphoglycerate kinase CONTIG04784 44.64 27.73 29.28 1.61 0.0005 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG01420 95.63 739.60 769.13 0.13 0.0000 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG06492 139.52 94.00 153.05 1.48 0.0243 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG06913 261.67 416.07 204.02 0.63 0.0037 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG17397 138.89 513.30 431.47 0.27 0.0000 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG08782 213.59 162.56 122.97 1.31 0.0347 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG30207 276.24 6068.08 5114.06 0.05 0.0000 phosphoglycerate/bisphosphoglycerate mutase family protein CONTIG33657 9012.06 166.74 208.50 54.05 0.0000 PHR2 | PHR2 (Photolyase/blue-light receptor 2) 230

CONTIG29423 13624.90 447.15 465.85 30.47 0.0000 PMDH2 | PMDH2 (peroxisomal NAD-malate dehydrogenase 2); malate dehydrogenase CONTIG13382 5585.27 12328.52 9478.10 0.45 0.0000 PMDH2 | PMDH2 (peroxisomal NAD-malate dehydrogenase 2); malate dehydrogenase CONTIG01940 1732.84 11154.43 16935.82 0.16 0.0000 PMDH2 | PMDH2 (peroxisomal NAD-malate dehydrogenase 2); malate dehydrogenase CONTIG40060 15895.98 1710.41 2982.19 9.29 0.0000 PPCK1 | PPCK1 (Phosphoenolpyruvate carboxylase kinase); kinase CONTIG33835 7570.05 1900.61 1402.75 3.98 0.0000 PPCK1 | PPCK1 (Phosphoenolpyruvate carboxylase kinase); kinase CONTIG39720 9572.55 5132.36 7588.77 1.87 0.0571 PPCK1 | PPCK1 (Phosphoenolpyruvate carboxylase kinase); kinase CONTIG21405 46.56 27.68 28.32 1.68 0.0079 PPCK1 | PPCK1 (Phosphoenolpyruvate carboxylase kinase); kinase CONTIG03970 491.05 1491.93 1130.66 0.33 0.0001 PPCK1 | PPCK1 (Phosphoenolpyruvate carboxylase kinase); kinase CONTIG34766 16404.73 2834.65 4860.97 5.79 0.0000 PPDK | PPDK (pyruvate orthophosphate dikinase); kinase CONTIG26315 49879.12 11826.10 13426.23 4.22 0.0000 PPDK | PPDK (pyruvate orthophosphate dikinase); kinase CONTIG40725 43092.24 22830.44 27961.65 1.89 0.0000 PPDK | PPDK (pyruvate orthophosphate dikinase); kinase CONTIG36433 5729.34 3195.82 5080.17 1.79 0.0289 PPDK | PPDK (pyruvate orthophosphate dikinase); kinase CONTIG35974 4291.75 225.91 269.99 19.00 0.0000 pyruvate decarboxylase family protein CONTIG04286 11708.87 4929.08 4510.78 2.38 0.0004 pyruvate decarboxylase family protein CONTIG09417 617.76 333.06 85.54 1.85 0.0000 pyruvate decarboxylase, putative CONTIG23045 6525.79 1640.08 578.05 3.98 0.0000 pyruvate decarboxylase, putative CONTIG28795 10083.56 5990.54 2144.80 1.68 0.0000 pyruvate decarboxylase, putative CONTIG06491 15954.89 7075.15 8673.97 2.26 0.0006 SBE2.2 | SBE2.2 (starch branching enzyme 2.2); 1,4-alpha-glucan branching enzyme CONTIG12481 4933.51 2279.84 1446.44 2.16 0.0007 SBE2.2 | SBE2.2 (starch branching enzyme 2.2); 1,4-alpha-glucan branching enzyme CONTIG06106 4102.06 4100.92 2574.05 1.00 0.0000 SSI, ATSS1 | ATSS1/SSI (starch synthase I); transferase, transferring glycosyl groups CONTIG11657 7181.40 7368.37 4365.99 0.97 0.0004 SSI, ATSS1 | ATSS1/SSI (starch synthase I); transferase, transferring glycosyl groups CONTIG15914 2629.69 5946.14 3909.92 0.44 0.0011 SSI, ATSS1 | ATSS1/SSI (starch synthase I); transferase, transferring glycosyl groups CONTIG32001 3749.11 3320.66 6112.88 1.13 0.0032 TIM | TIM (triosephosphate isomerase); triose-phosphate isomerase CONTIG25995 25.74 46.02 37.55 0.56 0.2624 TIM | TIM (triosephosphate isomerase); triose-phosphate isomerase CONTIG21735 26.93 63.07 58.39 0.43 0.0123 TIM | TIM (triosephosphate isomerase); triose-phosphate isomerase CONTIG40118 12473.20 38.23 33.94 326.24 0.0000 TPI, ATCTIMC | ATCTIMC (cytosolic triose phosphate isomerase); triose-phosphate isomerase CONTIG00243 30941.35 12631.31 20693.12 2.45 0.0000 TPI, ATCTIMC | ATCTIMC (cytosolic triose phosphate isomerase); triose-phosphate isomerase CONTIG28914 49432.70 31008.65 36964.05 1.59 0.0001 TPI, ATCTIMC | ATCTIMC (cytosolic triose phosphate isomerase); triose-phosphate isomerase CONTIG36583 546.32 950.41 956.06 0.57 0.0004 TPI, ATCTIMC | ATCTIMC (cytosolic triose phosphate isomerase); triose-phosphate isomerase CONTIG17356 1819.47 7376.16 4418.28 0.25 0.0001 UPF1, LBA1 | LBA1/UPF1 (low-level beta-amylase 1); RNA helicase CONTIG19343 37.86 814.61 228.92 0.05 0.0000 UPF1, LBA1 | LBA1/UPF1 (low-level beta-amylase 1); RNA helicase

231

Table 2. List of the 30 genes exhibiting the most decrease in transcript abundance between C3 and weak CAM species. Fold decrease (indicated by one standard deviation representing a 4.6-fold change) is expressed as the maximal log2 ratio (C3/weak CAM) at 2:00 AM.

log2 Adjusted Feature ratio p-value Hit_ID Description CONTIG12751 -6.76 6.88E-09 — Dehydration-responsive protein- CONTIG13669 -6.57 2.60E-08 AT4G14360.1 related CONTIG25546 -6.05 2.26E-08 — — CYP72A14 | CYP72A14 (cytochrome P450, family 72, subfamily A, CONTIG40689 -5.94 4.02E-09 AT3G14680.1 polypeptide 14); oxygen binding CONTIG07565 -5.92 2.87E-09 — — NDHF | Chloroplast encoded NADH CONTIG33581 -5.56 2.50E-08 ATCG01010.1 dehydrogenase unit Pentatricopeptide (PPR) repeat- CONTIG28156 -5.32 7.07E-08 AT5G60960.1 containing protein CONTIG29281 -5.31 1.18E-07 — — CONTIG19462 -5.28 1.41E-08 — — CYP704A1 | CYP704A1 (cytochrome P450, family 704, subfamily A, CONTIG14810 -5.26 3.67E-09 AT2G44890.1 polypeptide 1) CONTIG27667 -5.24 9.36E-07 — — CONTIG05442 -5.21 2.50E-07 — — Pyridine nucleotide-disulphide CONTIG40571 -5.16 1.61E-08 AT3G44190.1 oxidoreductase family protein Similar to hypothetical protein MtrDRAFT_AC148817g12v2 CONTIG32616 -5.16 3.73E-07 AT4G00585.1 [Medicago truncatula] ATSRP30, ATSRP30.1, ATSRP30.2 | ATSRP30.1 Arabidopsis thaliana Serine/Arginine protein 30.1); CONTIG02172 -5.15 1.04E-09 AT1G09140.1 RNA binding Mitotic phosphoprotein Nprime end CONTIG17040 -5.13 7.56E-08 AT3G16310.1 (MPPN) family protein Invertase/pectin methylesterase CONTIG06872 -5.02 1.65E-06 AT4G25260.1 inhibitor family protein lipid-binding serum glycoprotein CONTIG00298 -4.98 5.05E-08 AT3G20270.1 family protein CONTIG03134 -4.97 7.41E-08 — — SOC1, AGL20 | AGL20 AGAMOUS- CONTIG38190 -4.96 2.56E-08 AT2G45660.1 LIKE 20; transcription factor Transducin family protein / WD-40 CONTIG12702 -4.93 7.27E-08 AT3G49660.1 repeat family protein CONTIG31968 -4.93 1.14E-07 — — CONTIG32308 -4.91 9.37E-08 — — CONTIG31858 -4.90 3.50E-07 AT5G59320.1 LTP3 | LTP3 (Lipid transfer protein 232

3); lipid binding IDH1 | IDH1 Isocitrate Dehydrogenase 1); isocitrate CONTIG38233 -4.90 1.06E-08 AT4G35260.1 dehydrogenase (NAD+) Dehydration-responsive protein- CONTIG02405 -4.86 2.97E-08 AT5G14430.1 related CONTIG31725 -4.84 8.06E-08 AT2G02710.3 PAC motif-containing protein CONTIG33411 -4.84 2.70E-07 — — CONTIG14324 -4.78 5.00E-07 — — CONTIG22581 -4.78 8.16E-08 — —

233

Table 3. List of the 30 genes exhibiting the most decrease in transcript abundance between CAM and C3 species. Fold decrease (indicated by one standard deviation representing a 4.6-fold change) is expressed as the maximal log2 ratio (CAM/C3) at 2:00 AM.

log2 Adjusted Feature ratio p-value Hit_ID Description CONTIG26762 -8.53 4.73E-10 AT1G32900.1 Starch synthase, putative CONTIG27362 -8.36 3.82E-09 AT1G32900.1 Starch synthase, putative CONTIG33083 -7.54 7.05E-09 AT1G32900.1 Starch synthase, putative ATGSTF13 | ATGSTF13 (Arabidopsis thaliana Glutathione S-transferase CONTIG37978 -7.37 1.15E-07 AT3G62760.1 (class phi) 13); glutathione transferase SCPL35 | SCPL35 (serine carboxypeptidase-like 35); serine CONTIG21626 -7.32 8.79E-09 AT5G08260.1 carboxypeptidase CONTIG23566 -7.23 5.94E-09 AT5G01720.1 F-box family protein (FBL3) ATHCX1, CAX1-LIKE, ATCAX3, CAX3 | CAX3 (cation exchanger 3); CONTIG30112 -7.20 1.65E-07 AT3G51860.1 cation:cation antiporter CONTIG25390 -7.19 5.75E-10 AT5G11330.1 Monooxygenase family protein CYP89A6 | CYP89A6 (cytochrome P450, family 87, subfamily A, CONTIG17576 -7.12 7.33E-09 AT1G64940.1 polypeptide 6); oxygen binding CONTIG14038 -7.09 7.15E-09 — — ATBCAT-3, BCAT3 | ATBCAT- 3/BCAT3 (Branched-chain aminotransferase 3); Branched-chain- CONTIG37300 -6.97 1.85E-09 AT3G49680.1 amino-acid transaminase/ catalytic PDR6, ATPDR6 | ATPDR6/PDR6 (Pleiotropic drug resistance 6); ATPase, coupled to transmembrane CONTIG12760 -6.96 1.93E-08 AT2G36380.1 movement of substances CONTIG32868 -6.86 2.54E-09 AT4G27830.1 glycosyl hydrolase family 1 protein HIS1-3 | HIS1-3 (HISTONE H1-3); CONTIG29231 -6.81 1.10E-08 AT2G18050.1 DNA binding CONTIG31600 -6.72 7.33E-09 — — HDG2 | homeobox-leucine zipper family protein /lipid-binding START CONTIG17630 -6.68 1.73E-08 AT1G05230.1 domain-containing protein CONTIG18907 -6.53 1.89E-09 AT5G11700.1 glycine-rich protein CONTIG04893 -6.53 9.07E-08 AT5G60190.1 Ulp1 protease family protein CONTIG21098 -6.51 1.49E-07 AT1G45191.2 Glycosyl hydrolase family 1 protein CONTIG21535 -6.48 1.40E-09 — — Similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G73390.2); similar to unknown protein [Arabidopsis CONTIG32300 -6.44 3.60E-08 AT1G17940.1 thaliana] (TAIR:AT1G73390.1); 234

similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G73390.3); similar to unnamed protein product [Vitis vinifera] (GB:CAO62047.1); contains InterPro domain BRO1 (InterPro:IPR004328) CONTIG33505 -6.30 7.82E-07 — — Similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G17940.1); similar to unknown [Populus trichocarpa] (GB:ABK95976.1); similar to unnamed protein product [Vitis vinifera] (GB:CAO62047.1); contains InterPro domain BRO1 CONTIG40390 -6.26 1.54E-06 AT1G73390.1 (InterPro:IPR004328) CONTIG31039 -6.23 2.11E-08 — — Endonuclease/exonuclease/phosphatase CONTIG17104 -6.21 3.83E-07 AT2G32010.1 family protein CONTIG04853 -6.03 6.29E-09 — — Proton-dependent oligopeptide CONTIG33762 -6.02 3.50E-08 AT5G01180.1 transport (POT) family protein CONTIG30625 -5.99 7.13E-09 — — Similar to unknown protein [Arabidopsis thaliana] (TAIR:AT5G65440.2); similar to unknown protein [Arabidopsis thaliana] (TAIR:AT5G65440.1); similar to unknown protein [Arabidopsis thaliana] (TAIR:AT5G48310.1); similar to unnamed protein product [Vitis vinifera] (GB:CAO61624.1); similar to unnamed protein product [Vitis vinifera] (GB:CAO22682.1); similar to hypothetical protein OsJ_008178 [Oryza sativa (japonica cultivar- group)] (GB:EAZ24695.1); contains InterPro domain Globin-like CONTIG09409 -5.99 1.13E-08 AT4G24610.1 (InterPro:IPR009050) ATNUDT12 | ATNUDT12 (Arabidopsis thaliana nudix hydrolase CONTIG05361 -5.98 2.66E-08 AT1G12880.1 homolog 12); hydrolase

235

Table 4. List of the 30 genes exhibiting the most decrease in transcript abundance between CAM and weak CAM species. Fold decrease (indicated by one standard deviation representing a 4.6-fold change) is expressed as the maximal log2 ratio (CAM/Weak CAM) at 2:00 AM.

log2 Adjusted Feature ratio p-value Hit_ID Description CYP89A6 | CYP89A6 (Cytochrome P450, family 87, subfamily A, CONTIG17576 -7.11 7.33E-09 AT1G64940.1 polypeptide 6); oxygen binding CONTIG39994 -7.00 8.47E-09 — — CONTIG39875 -6.96 2.96E-09 — — CONTIG18907 -6.93 1.89E-09 AT5G11700.1 Glycine-rich protein CONTIG04785 -6.68 1.40E-09 AT3G26780.1 Catalytic CONTIG30625 -6.67 7.13E-09 — — CONTIG38595 -6.66 4.16E-07 — — CAF, SUS1, SIN1, ASU1, EMB76, EMB60, DCL1 | DCL1 (Dicer-Like1); ATP-dependent helicase/ ribonuclease CONTIG21222 -6.62 1.89E-09 AT1G01040.1 III IAA26, PAP1 | PAP1 (Phytochrome Associated Protein 1); Transcription CONTIG09447 -6.62 1.62E-09 AT3G16500.1 factor CYP704A1 | CYP704A1 (cytochrome P450, family 704, subfamily A, CONTIG14810 -6.60 3.67E-09 AT2G44890.1 polypeptide 1); oxygen binding CONTIG00001 -6.54 6.02E-08 AT1G51990.1 O-methyltransferase family 2 protein Similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G25370.1); similar to unnamed protein product [Vitis vinifera] (GB:CAO41620.1); contains InterPro domain Protein of unknown function DUF1639 CONTIG14789 -6.52 4.22E-10 AT3G60410.1 (InterPro:IPR012438) ATBCAT-3, BCAT3 | ATBCAT- 3/BCAT3 (Branched-chain aminotransferase 3); Branched-chain- CONTIG37300 -6.49 1.85E-09 AT3G49680.1 amino-acid transaminase/ catalytic CSLA03, ATCSLA3, ATCSLA03 | ATCSLA03 (Cellulose synthase-like A3); transferase, transferring glycosyl CONTIG09864 -6.38 4.73E-10 AT1G23480.1 groups ATKT1P, ATKUP1, KUP1, ATKT1 | ATKT1 (Arabidopsis thaliana K+ uptake 1); Potassium ion CONTIG17894 -6.35 1.57E-08 AT2G30070.1 transmembrane transporter CONTIG27362 -6.25 3.82E-09 AT1G32900.1 Starch synthase, putative CONTIG35101 -6.24 7.33E-09 AT1G27450.1 APT1, ATAPT1, APRT | APT1; 236

adenine phosphoribosyltransferase HDG2 | Homeobox-leucine zipper family protein / lipid-binding START CONTIG17630 -6.22 1.73E-08 AT1G05230.1 domain-containing protein ATKT1P, ATKUP1, KUP1, ATKT1 | ATKT1 (Arabidopsis thaliana K+ uptake 1); Potassium ion CONTIG11119 -6.19 1.00E-07 AT2G30070.1 transmembrane transporter RLK1 | RLK1 (RECEPTOR-LIKE PROTEIN KINASE 1); carbohydrate CONTIG21835 -6.15 4.70E-09 AT5G60900.1 binding / kinase ATHCX1, CAX1-LIKE, ATCAX3, CAX3 | CAX3 (cation exchanger 3); CONTIG30112 -6.07 1.65E-07 AT3G51860.1 cation:cation antiporter Similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G73390.2); similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G73390.1); similar to unknown protein [Arabidopsis thaliana] (TAIR:AT1G73390.3); similar to unnamed protein product [Vitis vinifera] (GB:CAO62047.1); contains InterPro domain BRO1 CONTIG32300 -6.04 3.60E-08 AT1G17940.1 (InterPro:IPR004328) IQD32 | IQD32 (IQ-domain 32); CONTIG04703 -6.03 6.70E-09 AT1G19870.1 calmodulin binding CES101 | CES101 (CALLUS EXPRESSION OF RBCS 101); CONTIG34197 -6.01 1.98E-08 AT3G16030.1 carbohydrate binding / kinase CONTIG09443 -6.00 2.89E-10 AT5G54630.1 Zinc finger protein-related CONTIG14004 -6.00 3.97E-09 — — CONTIG31600 -5.93 7.33E-09 — — CONTIG27579 -5.90 5.68E-08 — — CONTIG12327 -5.89 1.64E-07 AT5G64700.1 Nodulin MtN21 family protein CONTIG26762 -5.86 4.73E-10 AT1G32900.1 Starch synthase, putative

237

Table 5. List of the 30 genes exhibiting the most increase in transcript abundance between C3 and weak CAM species. Fold increase (indicated by one standard deviation representing a 4.6-fold change) is expressed as the maximal log2 ratio (C3/weak CAM) at 2:00 AM. log2 Adjusted Feature ratio p-value Hit_ID Description CONTIG12953 6.55 6.43E-10 AT1G02850.1 Glycosyl hydrolase family 1 protein CONTIG21098 6.53 1.49E-07 AT1G45191.2 Glycosyl hydrolase family 1 protein ATLOX2, LOX2 | LOX2 CONTIG39427 6.18 2.12E-08 AT3G45140.1 (Lipoxygenase 2) ATLOX2, LOX2 | LOX2 CONTIG22507 5.94 1.44E-08 AT3G45140.1 (Lipoxygenase 2) CONTIG30663 5.84 3.67E-10 — — CONTIG05028 5.70 1.08E-08 — — AGL22, SVP | SVP (Short Vegetative CONTIG32097 5.66 2.96E-09 AT2G22540.1 phase); transcription factor MUB6 | MUB6 (Membrane- anchored-Ubiquitin-fold protein 6 CONTIG33318 5.63 1.07E-08 AT1G22050.1 precursor) CONTIG32777 5.58 1.46E-08 — — CONTIG40399 5.55 4.74E-08 — — CONTIG12425 5.48 2.27E-09 — — CONTIG06196 5.43 4.78E-09 AT4G35890.1 La domain-containing protein Integral membrane transporter family CONTIG03830 5.41 3.22E-08 AT1G79710.1 protein CONTIG11114 5.24 8.43E-07 — — CONTIG05606 5.23 1.47E-08 — — CSLB04, ATCSLB4, ATCSLB04 | ATCSLB04 (Cellulose synthase-like B4); transferase/ transferase, CONTIG22568 5.17 2.46E-07 AT2G32540.1 transferring glycosyl groups CONTIG36204 5.11 1.22E-07 — — CONTIG15961 5.00 4.75E-08 — — CONTIG05413 4.96 2.13E-07 — — CONTIG33505 4.96 7.82E-07 — — CONTIG12954 4.93 1.22E-06 AT1G02850.2 Glycosyl hydrolase family 1 protein CONTIG22859 4.92 1.59E-06 — — CONTIG15744 4.90 3.01E-06 — — CONTIG35747 4.89 7.70E-08 AT5G57580.1 Calmodulin-binding protein CONTIG23157 4.87 5.75E-10 — — CONTIG06598 4.86 9.44E-08 AT3G05990.1 Leucine-rich repeat family protein CONTIG12864 4.84 4.08E-09 — — CONTIG14107 4.83 7.69E-08 AT1G02850.1 Glycosyl hydrolase family 1 protein Similar to unknown protein [Arabidopsis thaliana] (TAIR:AT3G04040.1); similar to unnamed protein product [Vitis CONTIG07822 4.79 3.08E-08 AT2G41945.1 vinifera] (GB:CAO23431.1) CONTIG31720 4.73 1.73E-07 — — 238

Table 6. List of the 30 genes exhibiting the most increase in transcript abundance between CAM and C3 species. Fold increase (indicated by one standard deviation representing a 4.6-fold change) is expressed as the maximal log2 ratio (CAM/C3) at 2:00 AM.

log2 Adjusted Feature ratio p-value Hit_ID Description CONTIG40221 10.12 8.04E-10 — — CONTIG37111 10.10 1.19E-09 — — CONTIG33920 10.04 1.40E-09 — — CONTIG37008 10.03 1.12E-09 — — CONTIG20390 10.00 9.07E-09 — — CONTIG36853 9.94 1.61E-09 — — CONTIG27941 9.76 3.55E-11 — — CONTIG40148 9.71 4.58E-10 — — CONTIG27442 9.41 3.21E-09 — — CONTIG23958 9.31 1.25E-09 — — CONTIG36158 9.29 1.51E-10 — — CONTIG06153 9.26 1.40E-10 — — CONTIG37672 9.26 4.73E-10 — — CONTIG39394 9.25 3.55E-11 — — CONTIG02035 9.25 2.54E-09 — — CONTIG14400 9.19 6.75E-11 — — CONTIG05740 9.13 3.55E-10 — — CONTIG40318 9.12 1.11E-09 — — CONTIG35583 9.09 3.11E-08 — — CONTIG37201 9.06 2.45E-09 — — CONTIG27716 9.06 9.18E-09 — — CONTIG38059 9.03 3.08E-08 — — CONTIG40774 8.98 1.25E-08 — — CONTIG37470 8.93 1.10E-09 — — CONTIG39805 8.91 2.51E-09 — — CDKB2;1 | CDKB2;1 (Cyclin- CONTIG01003 8.86 4.67E-10 AT1G76540.1 dependent kinase B2;1); kinase CONTIG40914 8.85 4.33E-09 — — CONTIG15515 8.85 1.44E-09 — — CONTIG35534 8.83 2.90E-10 — — CONTIG32511 8.83 1.13E-08 — —

239

Table 7. List of the 30 genes exhibiting the most increase in transcript abundance between CAM and weak CAM species. Fold increase (indicated by one standard deviation representing a 4.6-fold change) is expressed as the maximal log2 ratio (CAM/weak CAM) at 2:00 AM.

log2 Adjusted Feature ratio p-value Hit_ID Description CONTIG37008 10.58 1.12E-09 — — CONTIG20390 10.31 9.07E-09 — — ATLOX2, LOX2 | LOX2 CONTIG22507 10.27 1.44E-08 AT3G45140.1 (Lipoxygenase 2) CONTIG33920 10.22 1.40E-09 — — CONTIG37111 10.07 1.19E-09 — — CONTIG27941 9.95 3.55E-11 — — CONTIG36853 9.89 1.61E-09 — — CONTIG37201 9.66 2.45E-09 — — CONTIG40148 9.61 4.58E-10 — — CONTIG36158 9.51 1.51E-10 — — CONTIG06153 9.37 1.40E-10 — — CONTIG02035 9.37 2.54E-09 — — CONTIG39805 9.34 2.51E-09 — — CONTIG05740 9.32 3.55E-10 — — CONTIG40318 9.29 1.11E-09 — — CONTIG37672 9.29 4.73E-10 — — CONTIG39394 9.26 3.55E-11 — — CONTIG40763 9.24 3.50E-10 — — CONTIG27716 9.24 9.18E-09 — — CONTIG36618 9.22 6.06E-12 — — CONTIG35583 9.05 3.11E-08 — — CONTIG01333 9.05 4.02E-08 — — CONTIG39821 9.01 4.03E-08 — — CONTIG14400 9.00 6.75E-11 — — CONTIG23958 8.93 1.25E-09 — — CYP74A, AOS | AOS (Allene oxide CONTIG28868 8.92 1.77E-10 AT5G42650.1 synthase); hydrolyase/oxygen binding CONTIG36985 8.86 1.15E-08 — — CDKB2;1 | CDKB2;1 (Cyclin- CONTIG01003 8.84 4.67E-10 AT1G76540.1 dependent kinase B2;1); kinase PSAF | PSAF (photosystem I subunit CONTIG40858 8.84 2.61E-09 AT1G31330.1 F) CONTIG40914 8.81 4.33E-09 — —

240

Figure 1. Principal component analysis (PCA) of mRNA expression profiles of three Oncidiinae species and four biological replicates per species. Red circles represent the strong CAM species (Rossioglossum ampliatum), blue squares represent the C3 species (Oncidium maduroi), and green diamonds represent the weak CAM species (Oncidium panamense).

241

(A)

(B)

242

(C)

Figure 2. Normalized mRNA expression for the comparison between weak CAM and CAM (A); C3 and CAM (B) and weak CAM and C3 (C). The diagonal lines represent the one-to-one correspondence line bracketed by one and two standard deviation, representing 4.6-fold and 21-fold differences in mRNA expression, respectively. 243

Figure 3. Plot of log2-transformed intensity ratios showing increased or decreased mRNA abundance values of probe sets between weak CAM and CAM, and between

CAM and C3. Colors represent categories of gene expression: Red represents genes with increased mRNA abundance in CAM relative to C3 with minimal changes between weak CAM and CAM. Yellow represents genes with increased mRNA abundance in weak

CAM relative to CAM with minimal changes between CAM and C3. Dark purple represent genes with decreased mRNA abundance in weak CAM relative to CAM with minimal changes between CAM to C3. Green represents genes with decreased mRNA abundance in CAM relative to C3 with minimal changes between CAM and C3 progression. Orange, light green, blue and light purple represent genes that show intermediate mRNA abundance changes among weak CAM, CAM, and C3. Only points that show one or more 4.6-fold difference and an adjusted P <0.05 were considered (see text for details). 244

20 Phosphoenolpyruvate n

o carboxylase 1 i s

s (Contig 35735) e r 15 p x e

A N R m

10 d e m r o f s n

a 5 r t - 2 g o l

0 CAM Weak C3 CAM

Phosphoenolpyruvate n o i carboxylase 2 s s

e (Contig 16824) r

p 15 x e

A N R m

10 d e m r o f s n 5 a r t - 2 g o l

CAM Weak C3 CAM

245

20 n o

i Phosphoenolpyruvate s

s carboxylase 3 e r

p (Contig 02012) x 15 e

A N R m

d 10 e m r o f s n

a 5 r t - 2 g o l

CAM Weak C3 CAM

n 20

o Pyruvate orthophosphate i s

s dikinase PPDK e r (Contig 26315) p x 15 e

A N R m

d 10 e m r o f s n 5 a r t - 2 g o l 0 CAM Weak C3 CAM 246

20 n

o Carbonic anhydrase 1 i s s (Contig 32032) e r p 15 x e

A N R m 10 d e m r o f s n a 5 r t - 2 g o l

0 CAM Weak C3 CAM

n Carbonic anhydrase 2 o i

s (Contig 28739) s e r

p 15 x e

A N R m

10 d e m r o f s

n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

247

n o i Phosphoenolpyruvate s s

e carboxylase kinase 1 r

p (Contig 33835) x 15 e

A N R m 10 d e m r o f s n

a 5 r t - 2 g o l

CAM Weak C3 CAM

n 20 o i

s Malate dehydrogenase (NADP) s e (Contig 09276) r p x 15 e

A N R m

d 10 e m r o f s n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

248

20 n

o Blue light receptor 2 i s s (Contig 33657) e r p 15 x e

A N R m

10 d e m r o f s

n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

n Glyceraldehyde-3-phosphate o i

s dehydrogenase s e

r (Contig 28163) p

x 15 e

A N R m 10 d e m r o f s

n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

249

20 n o

i Glucose-6-phosphate s s translocator 1 e r

p (Contig 02056) x 15 e

A N R m

d 10 e m r o f s n a 5 r t - 2 g o l

0 CAM Weak C3 CAM

20 n o i Glucose-6-phosphate s s

e translocator 2 r

p (Contig 02442) x 15 e

A N R m 10 d e m r o f s

n 5 a r t - 2 g o l 0 CAM Weak C3 CAM

250

n 20 o i

s Fructose-biphosphate s

e aldolase r p

x 15 (Contig 28489) e

A N R m

10 d e m r o f

s 5 n a r t - 2 g o l 0

CAM Weak C3 CAM

20 n o i

s Fructose-1,6-biphosphate s e (Contig 05658) r p

x 15 e

A N R

m 10 d e m r o f s 5 n a r t - 2 g o l 0

CAM Weak C3 CAM

251

n 20 o i Gamma carbonic s s e anhydrase-like 2 r p x 15 (Contig 35220) e

A N R m 10 d e m r o f s

n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

n 20 o i

s Pyruvate decarboxylase s e

r family protein p

x 15 (Contig 35974) e

A N R m

10 d e m r o f

s 5 n a r t - 2 g o l 0

CAM Weak C3 CAM

252

n 20 o i Peroxisomal NAD-malate s s

e dehydrogenase 2 r p x 15 (Contig 29423) e

A N R m

10 d e m r o f s 5 n a r t - 2 g o l 0

CAM Weak C3 CAM

20 n

o Cytosolic triose i s

s phosphate isomerase e r

p (Contig 40118)

x 15 e

A N R

m 10

d e m r o f

s 5 n a r t - 2 g o l 0

CAM Weak C3 CAM

253

20 n o i

s alpha-amylase s e

r (Contig 09267) p

x 15 e

A N R m 10 d e m r o f s

n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

n 20 o i s

s beta-amylase e r (Contig 19797) p x

e 15

A N R m

d 10 e m r o f s

n 5 a r t - 2 g o l 0

CAM Weak C3 CAM

Figure 4. Average mRNA expression (±SD) of selected genes with CAM-related functions that exhibit shifts in expression patterns between CAM, weak CAM and C3 species. Red represents CAM species, orange represents weak CAM species, and green represents C3 species. 254

Chapter VI

Concluding Remarks

The varying degrees to which CAM is expressed can be explained by a continuum of photosynthetic pathways from C3 photosynthesis to weakly expressed CAM to fully expressed CAM arising from the evolutionary history of a particular species. The plasticity of CAM is governed by the evolutionary disposition of each species, whether under development control in a constitutive CAM species or under environmental control in a facultative CAM species CAM. In neotropical orchids, the CAM pathway can be found in up to 50% of species (Silvera et al. 2005). The distribution of photosynthetic

pathways shows that C3 photosynthesis is the ancestral state and that CAM has evolved independently several times within the Orchidaceae. Using phylogenetic trait analysis, we found that divergences in photosynthetic pathway and epiphytism are consistently correlated through evolutionary time and are related to the prevalence of CAM epiphytes in lower elevations and abundant species diversification of high elevation epiphytes. The multiple independent evolutionary origins of CAM in orchids suggest that evolution from

C3 to weak and strong CAM might involve relatively few genetic changes.

In plants performing CAM, phosphoenolpyruvate carboxylase (PEPC) catalyzes the initial fixation of atmospheric CO2 into C4-dicarboxylic acids forming oxaloacetate and inorganic phosphate as a product. PEPC is a ubiquitous enzyme that belongs to a multigene family with each gene encoding a function- and tissue-specific isoform of the enzyme. We found that up to five PEPC isoforms are present closely-related orchids from 255 the Subtribe Oncidiinae, with one putative CAM-specific PEPC isogene with discrete amino acid changes identified in CAM species based on cDNA clone sampling, and an evident shift in PEPC isoform abundance from 2-3 isoforms in C3 species to 3-4 isoforms in weak CAM species to 4-5 isoforms in strong CAM species. Validation of the isotopic analysis and the molecular genetic analysis of PEPC gene family using 24-hour gas

exchange showed that weak CAM species exhibit limited amounts of nocturnal CO2 uptake and fixation when compared to strong CAM species.

Our understanding of the CAM photosynthetic pathway is advancing, especially at the molecular genetic level. Gene sequence information has proliferated quickly and will provide a solid foundation for future research into CAM evolution. Transcriptome sequencing has been performed in a strong CAM orchid species (Rossioglossum ampliatum) along with the fabrication of a custom oligonucleotide microarray, which will permit mRNA expression patterns to be compared within closely-related C3 photosynthesis and weak CAM species as a way to define gene expression changes associated with CAM evolution. From 13,566 genes that showed a significant 4.6-fold difference in expression levels from the comparisons between CAM, C3 and weak CAM,

4,520 genes showed a greater than 4.6-fold increase in the ratio of CAM/C3 relative transcript abundance, whereas 3,745 genes showed a greater than 4.6-fold decrease in the ratio of CAM/C3.

Future projects aimed at testing whether the presence or absence of cis-regulatory elements responsible for circadian clock controlled expression patterns would also allow to determine whether or not the regulation of the CAM-specific expression patterns are governed by evolutionary changes within the 5’ flanking regions. Weak CAM species 256 are of particular interest because a reservoir of duplicated genes that have undergone neofunctionalization from C3 ancestral genes are expected to be present, in addition to C3 genes. The molecular evolution of CAM can be expected to occur as postulated for the evolution of C4 photosynthesis. Gene duplication events followed by neofunctionalization and subfunctionalization is likely to occur through differentiation of the cis-regulatory elements that control tissue-specific patterns of expression (Monson 2003). The major distinction for CAM evolution is that temporal regulation of gene expression patterns will have greater importance than cell-specific expression patterns. In both cases, differentiation within the coding region can be expected to produce alternative functional domains within proteins.

Undoubtedly, the presence of CAM in evolutionary lineages must be defined at the molecular level, in order to understand which changes in genetic material allowed the evolutionary progression from C3 to strong CAM, and possible reversal events linked to a changing environment, and to further determine whether low level CAM in certain lineages might serve as a genetic reservoir for adaptive radiations. The use of phylogenetic comparative methods is particularly useful in the testing of correlated evolutionary changes of multiple CAM traits together (e.g., molecular, physiological, anatomical and environmental traits). Larger carbon-isotope ratio surveys should be attempted and these should be performed in conjunction with titratable acidity measurements from live specimens. Future research is also needed to verify gene expression in species across a variety of taxa under well watered and watered-deficit conditions to discover facultative CAM species that might be missed by carbon-isotope ratio surveys alone. Proteomic studies targeting temporal changes in protein abundance or 257 posttranslational modifications are also expected to improve our understanding of the circadian regulation, especially when coupled with mRNA expression profiling of both coding mRNAs and non-coding miRNAs in selected CAM species. Ultimately, an integrated approach that combines molecular strategies, genetic approaches, phylogenetic analysis, ecophysiology, and bioinformatics will aid in our understanding of the molecular evolution of CAM photosynthesis.

258

LITERATURE CITED

Monson RK (2003) Gene duplication, neofunctionalization, and the evolution of C4 photosynthesis. International Journal of Plant Sciences 164, S43-S54.

Silvera K, Santiago LS, Winter K (2005) Distribution of crassulacean acid metabolism in orchids of Panama: evidence of selection of weak and strong modes. Functional Plant Biology 32, 397-407.