Transcriptional Regulation of Select Light-Harvesting Genes during Photoacclimation in

Lympha mucosa gen. et sp. prov. (, Rhodophyta)

A thesis presented to

the faculty of

the College of Arts and Sciences of Ohio University

In partial fulfillment

of the requirements for the degree

Master of Science

Joshua R. Evans

August 2017

© 2017 Joshua R. Evans. All Rights Reserved. 2 This thesis titled

Transcriptional Regulation of Select Light-Harvesting Genes during Photoacclimation in

Lympha mucosa gen. et sp. prov. (Batrachospermales, Rhodophyta)

by

JOSHUA R. EVANS

has been approved for

the Department of Environmental and Plant Biology

and the College of Arts and Sciences by

Morgan L. Vis

Professor of Environmental and Plant Biology

Robert Frank

Dean, College of Arts and Sciences 3 ABSTRACT

EVANS, JOSHUA R., M.S., August 2017, Environmental and Plant Biology

Transcriptional Regulation of Select Light-Harvesting Genes during Photoacclimation in

Lympha mucosa gen. et sp. prov. (Batrachospermales, Rhodophyta)

Director of Thesis: Morgan L. Vis

The strictly freshwater red algal order Batrachospermales has undergone

numerous taxonomic rearrangements in the recent past to rectify the paraphyly of its

largest genus Batrachospermum. These systematic investigations have led to the

description of new genera and species as well as re-circumscription of some taxa.

Specimens collected from two locations in southeastern USA were initially identified as

being allied to Batrachospermum sensu lato, but could not be assigned to any previously

described species. Comparison of DNA sequence data for two gene regions and

morphology with other batrachospermalean taxa resulted in the proposal of a new

monospecific genus Lympha mucosa gen. et sp. prov. to accommodate these specimens.

A phylogeny of L. mucosa showed it is sister to the genus Volatus, but has morphological

similarities with Batrachospermum sections Turfosa and Virescentia. This new taxon

adds to the freshwater red algal diversity of southeastern USA, a region already known

for biodiversity and high endemism of the aquatic flora and fauna.

Lympha mucosa occurs in open and shaded sites of temperate streams and is

abundant during summer months. Although most freshwater red algal taxa are considered

shade-adapted, many species exhibit differences in photosynthetic rates and

characteristics that indicate they have a much greater ability to acclimate to higher

irradiances. Specimens of L. mucosa were collected from open (sun-acclimated) or 4 shaded (shade-acclimated) sites and were exposed to low (<20 µmol photon m-2 s-1) or high (220 µmol photon m-2 s-1) for 72h in controlled conditions to examine

photoacclimation. To observe regulation for this process at the transcriptional level, the

L. mucosa plastid genome was assembled to provide sequence data for photosynthetic

genes involved with light harvesting machinery. Of the six light-harvesting genes

selected, two involved with photosystem I and one involved with phycoerythrin synthesis

were downregulated at high light. This is the first evidence of transcriptional regulation

as a potential mechanism for acclimation to varying irradiances in a freshwater red alga. 5 DEDICATION

This work is dedicated to my parents, Karen and Harry, without whom I would not be on

this journey, and to my nephews.

“No man ever steps in the same river twice, for it’s not the same river and he’s not the

same man” - Heraclitus

6 ACKNOWLEDGMENTS

A number of people have helped me shape and conduct this project and I would still be at the starting line without them. I would first like to acknowledge my advisor, Dr.

Morgan Vis. During my short time in her lab, my ability to conduct critical research and be independent has become stronger because of her knowledge and support, and I have gained many new skills that I will take with me on the next step of my career. I am honored to have been her student, and in the future, I hope to be even half the mentor she was for me. Many thanks go to members of my thesis committee, Drs. Sarah Wyatt and

Harvey Ballard, who have taken the time to critically evaluate my work and provide constructive feedback. I am indebted to Dr. Wyatt, who generously provided me with space and encouragement to learn something new and expand my skills base, and for shaping some of my thoughts on education. I thank Dr. Ballard for helpful comments, specifically with the second chapter of this thesis, and for graciously inviting us to his field accommodations. Colin Kruse is thanked for taking the time to troubleshoot RNA extractions with me, even if it meant not working with Arabidopsis. I appreciate his collaborative spirit and desire to help others. Dr. Erin Murphy is thanked for generously allowing me to use her qPCR machine. Additional acknowledgements go to past and present members of the Vis lab (Emily Keil-Loudner, Danny Wolf, Nathan St. Amour,

Lexie Redmond) for their assistance and support. Dr. Bill Broach, Al Meyers, and Anne

Sternberger are thanked for helpful discussions. I would like to acknowledge the support and expertise provided by the Ohio University Genomics Facility in completion of NGS sequencing. This research was partially funded through Original Works Grants provided by the Ohio University Graduate Student Senate. 7 TABLE OF CONTENTS

Page

Abstract ...... 3 Dedication ...... 5 Acknowledgments ...... 6 List of Tables ...... 9 List of Figures ...... 10 Chapter 1: Introduction ...... 11 References ...... 13 Chapter 2: Adding to the Freshwater Red Algal Diversity in North America: Lympha mucosa gen. et sp. prov. (Batrachospermales, Rhodophyta)...... 17 Introduction ...... 17 Methods...... 19 Sample Collection ...... 19 DNA Extraction and Amplification ...... 19 Phylogenetic Analyses ...... 20 Morphological Measurements ...... 21 Results ...... 22 Molecular Results ...... 22 Morphological Results ...... 23 ...... 23 Discussion ...... 26 References ...... 28 Chapter 3: Plastid Genomics and Relative Expression Analysis of Light-Harvesting Genes in Lympha mucosa Exposed to Low and High Irradiances ...... 37 Introduction ...... 37 Methods...... 42 Sample Collection and Experimental Design ...... 42 DNA Extraction and Sequencing ...... 44 Plastid Assembly and Annotation ...... 45 RNA Extraction ...... 46 Gene Selection and RT-qPCR Primer Design ...... 47 cDNA Synthesis and RT-qPCR Conditions ...... 49 8 Gene Expression Analysis ...... 50 Results ...... 51 Plastid Genome ...... 51 Gene Expression Analyses ...... 53 Discussion ...... 55 Freshwater Nemaliophycidae Plastid Genomes ...... 55 Differentially Expressed Genes in Light Treatments ...... 57 Implications of this Research ...... 63 References ...... 65 Chapter 4: Conclusions ...... 84 References ...... 85 Appendix A: Genbank Accession Numbers for COI-5P and rbcL Sequence Data of Batrachospermales and Thoreales Used for Phylogenetic Analyses: Chapter 2 ...... 88 Appendix B: Experimental Design for Lympha mucosa Samples Exposed to Low or High Light: Chapter 3 ...... 90 Appendix C: Total Quantity, RNA Integrity Number (RIN), and Absorbance Ratio (260/280) for each Lympha mucosa RNA Sample: Chapter 3 ...... 91 Appendix D: Two-Way Analysis of Variance Tables Using Data Normalized to rps3: Chapter 3 ...... 92 Appendix E: Two-Way Analysis of Variance Tables Using Data Normalized to eRF3: Chapter 3 ...... 94 Appendix F: List of Plastid Genomes for Taxa in the Nemaliophycidae: Chapter 3 ...... 96 Appendix G: RT-qPCR Optimization Data: Chapter 3 ...... 98

9 LIST OF TABLES

Page

Table 2-1 Morphometrics of vegetative and reproductive characters for Lympha mucosa gen. et sp. prov...... 33

Table 3-1 Site conditions for the Kinniconick Creek, KY ...... 74

Table 3-2 Primer sets that were used in RT-qPCR for each target and reference gene in

Lympha mucosa ...... 74

10 LIST OF FIGURES

Page

Figure 2-1 Maximum likelihood (ML) phylogeny of Lympha mucosa gen. et sp. prov. with other genera of Batrachospermales ...... 34

Figure 2-2 Morphological characteristics of Lympha mucosa gen et sp. prov ...... 35

Figure 3-1 Map of potential target and reference genes for expression analysis of

Lympha mucosa photoacclimation...... 76

Figure 3-2 The ³ 189,825 bp Lympha mucosa plastid genome...... 77

Figure 3-3 Comparison of the Lympha mucosa plastid genome to all other sequenced plastid genomes of freshwater Nemaliophycidae (Rhodophyta) ...... 78

Figure 3-4 Log2 fold change of psaA relative expression normalized to rps3 ...... 79

Figure 3-5 Log2 fold change of petF relative expression normalized to rps3 ...... 80

Figure 3-6 Log2 fold change of psbA relative expression normalized to rps3 ...... 81

Figure 3-7 Log2 fold change of HV60 relative expression normalized to eRF3 ...... 82

Figure 3-8 Log2 fold change in relative expression of genes encoding the main pigment- binding proteins in the phycobilisome, normalized to rps3...... 83

11 CHAPTER 1: INTRODUCTION

The red (Rhodophyta) are an early-diverging lineage of ~7,160 currently described species (Guiry and Guiry 2017) that are distributed in marine, freshwater, terrestrial, and hot spring environments. Combined with the distributional diversity, a number of divergent traits can be found in the , including a range of life histories, morphologies, and physiologies. For example, the heteromorphic and sporophyte generations of a single species have frequently been described as two distinct genera until the life history was known (see Hawkes 1990), and lifestyle strategies ranging from free-living to epiphytic/zoic and parasitic forms have been described (e.g., Clayden and Saunders 2010, Blouin and Lane 2012). These highly diverse forms and strategies have allowed the red algae to radiate and adapt to many environments. Nevertheless, this group requires more systematic and ecological study to truly understand these adaptations.

Freshwater lineages of the red algae occur throughout the red algal tree of life and are found in a diverse array of freshwater habitats (Sheath 1984). Unicellular members of the Cyanidiales, an early-diverging lineage within the Rhodophyta, predominantly occur as extremophiles in sulfuric and hot spring habitats (Ciniglia et al.

2004). However, the majority of freshwater taxa are mesophilic with a worldwide distribution in lentic and lotic freshwater (Sheath 1984). Although only encompassing

~5% of red algal species, freshwater taxa are highly diverse, with the majority occurring in three strictly freshwater orders of the subclass Nemaliophycidae (Lam et al. 2016).

Interestingly, these three orders are no more related to one another than they are to marine orders in the subclass (Lam et al. 2016), suggesting the transition to freshwater 12 environments and subsequent evolutionary radiation has evolved independently more than once in the red algae. Of these orders, the Batrachospermales is the most species rich, but the systematics has been updated in recent years with new genera and species continually being described (e.g., Entwisle et al. 2009, Vis et al. 2012, Salomaki et al.

2014, Chapuis et al. 2017). Moreover, these species differ considerably in their distribution, phenology, and photosynthetic characteristics.

The red algae commonly occur and thrive in aquatic habitats receiving low irradiance, and in the case of freshwater taxa, these features tend to have seasonality in temperate habitats with deciduous tree canopy in summer. Traditionally, these organisms have been considered shade-adapted given their distribution and ecology. These adaptations are likely achieved by mechanisms in the red algal plastid. Red and green plastids () are hypothesized to have arisen following an endosymbiotic event between a photosynthesizing cyanobacterium and an ancestral eukaryote (Cavalier-

Smith 2000, McFadden 2001). However, red plastids do not undergo significant differentiation during development, and there is evidence that they are the progenitor to photosynthesis in many other protist groups, such as the heterokonts and apicomplexans

(Moreira et al. 2000, Cavalier-Smith 2013, Keeling 2013).

Red plastids are highly divergent from other plastids. All plastids use chl. a as a photopigment for light harvesting, but red algae utilize unique accessory pigments in place of other chlorophyll molecules that are found in other eukaryotic plastids (Gantt

1990). The absorption spectra of these accessory pigments differ from other chlorophylls and allow red algae to exploit different wavelengths of light for photosynthetic processes.

In addition, red plastid genomes can have a larger gene repertoire than green algae, 13 including several photosynthetic gene families, and still retain other genomic features including group II introns and some transcription factors (Minoda et al. 2005, Muñoz-

Gómez et al. 2017). Therefore, studies of red algal plastids are critical to understand the evolution of this organelle, and for photosynthetic studies within the phylum.

The purpose of this thesis was to continue systematic revision within the

Batrachospermales and provide plastid gene expression data, as it relates to photoacclimation for a freshwater red alga. In the following chapters, Lympha mucosa gen. et sp. prov. is distinguished from other batrachospermalean lineages using morphological and phylogenetic evidence and is described as a new regionally endemic and monospecific genus of freshwater red algae (Chapter 2). Lympha mucosa is a taxon that occurs in a variety of irradiances, making it an excellent candidate for studying photoacclimation with gene expression analyses. The third chapter incorporates sequencing of the L. mucosa plastid genome as a tool to study select light-harvesting genes that are encoded in the red algal plastid. A controlled light experiment using L. mucosa was conducted to examine transcriptional regulation as a mechanism for photoacclimation and provide insights into this highly diverse and adaptive group of photosynthetic organisms.

References

Blouin, N.A. & Lane, C.E. 2012. Red algal parasites: models for a life history evolution

that leaves photosynthesis behind again and again. BioEssays 34:226-235.

Cavalier-Smith, T. 2000. Membrane heredity and early evolution. Trends

Plant Sci. 5:174-182. 14 Cavalier-Smith, T. 2013. Symbiogenesis: mechanisms, evolutionary consequences, and

systematic implications. Annu. Rev. Ecol. Evol. Syst. 44:145-172.

Chapuis, I.S., Necchi Jr., O., Zuccarello, G.C., Xie, S.L., Aboal, M., Sánchez Castillo,

P.M. & Vis, M.L. 2017. A new genus, Volatus and four new species of

Batrachospermum sensu stricto (Batrachospermales, Rhodophyta). Phycologia

56:454-468.

Ciniglia, C., Yoon, H.S., Pollio, A., Pinto, G. & Bhattacharya, D. 2004. Hidden

biodiversity of the extremophilic Cyanidiales red algae. Mol. Ecol. 13:1827-1838.

Clayden, S.L. & Saunders, G.W. 2010. Recognition of Rubrointrusa membranacea gen et

comb. nov. Rhodonematella subimmersa gen. et comb. nov. (with a

reinterpretation of the life history) and the Meiodiscaceae fam. nov. within the

Palmariales (Rhodophyta). Phycologia 49:283-300.

Entwisle, T.J., Vis, M.L., Chiasson, W.B., Necchi, O. & Sherwood, A.R. 2009.

Systematics of the Batrachospermales (Rhodophyta) – a synthesis. J. Phycol.

45:704-715.

Gantt, E. 1990. Pigmentation and photoacclimation. In Biology of Red Algae [Eds] Cole,

K.M. & Sheath, R.G. Cambridge University Press, Cambridge, UK. pp203-219.

Guiry, M.D. & Guiry, G.M. 2017. Algaebase. World-wide electronic publication,

National University of Ireland, Galway. http://www.algaebase.org; searched on 25

June 2017.

Hawkes, M.W. 1990. Reproductive strategies. In Biology of Red Algae [Eds] Cole, K.M.

& Sheath, R.G. Cambridge University Press, Cambridge, UK, pp455-476. 15 Keeling, P.J. 2013. The number, speed, and impact of plastid endosymbiosis in

eukaryotic evolution. Ann. Rev. Plant Biol. 64:583-607.

Lam, D.W., Verbruggen, H., Saunders, G.W. & Vis, M.L. 2016. Multigene phylogeny of

the red algal subclass Nemaliophycidae. Mol. Phyl. Evol. 94:730-736.

McFadden, G.I. 2001. Primary and secondary endosymbiosis and the origin of plastids. J.

Phycol. 37:951-959.

Minoda, A., Nagasawa, K., Hanaoka, M., Horiuchi, M., Takahashi, H. & Tanka, K. 2005.

Microarray profiling of plastid gene expression in a unicellular red alga,

Cyanidioschyzon merolae. Plant Mol. Biol. 59:375-385.

Moreira, D., Le Guyader, H. & Philippe, H. 2000. The origin of red algae and the

evolution of . Nature 405:69-72.

Muñoz-Gómez, S.A., Mejía-Franco, F.G., Durnin, K., Colp, M., Grisdale, C.J.,

Archibald, J.M. & Slamovits, C.H. 2017. The new red algal subphylum

Proteorhodophytina comprises the largest and most divergent plastid genomes

known. Curr. Biol. 27:1-8.

Salomaki, E.D., Kwandrans, J., Eloranta, P. & Vis, M.L. 2014. Molecular and

morphological evidence for Sheathia gen. nov. (Batrachospermales, Rhodophyta)

and three new species. J. Phycol. 50:526-542.

Sheath, R.G. 1984. The biology of freshwater red algae. In Round, F.E. & Chapman, D.J.

[Eds] Progress in Phycological Research, vol. 3. Biopress Ltd, Bristol UK, pp.

89-157. 16 Vis, M.L., Saunders, G.W., Sheath, R.G., Dunse, K. & Entwisle, T.J. 1998. Phylogeny of

the Batrachospermales (Rhodophyta) inferred from rbcL and 18S ribosomal DNA

gene sequences. J. Phycol. 32:341-350.

Vis, M.L., Necchi, O. Jr., Chiasson, W.B. & Entwisle, T.J. 2012. Molecular phylogeny of

the genus Kumanoa (Batrachospermales, Rhodophyta). J. Phycol. 48:750-758.

17 CHAPTER 2: ADDING TO THE FRESHWATER RED ALGAL DIVERSITY IN

NORTH AMERICA: LYMPHA MUCOSA GEN. ET SP. PROV.

(BATRACHOSPERMALES, RHODOPHYTA).

Introduction

The red algal order Batrachospermales contains the greatest diversity of freshwater taxa and is distributed worldwide in streams and rivers (Sheath 1984, Kumano

2002). Current taxonomic research of this order has been focused on rectifying the paraphyly of the genus Batrachospermum Roth (Vis et al. 1998, Entwisle et al. 2009,

2016, Salomaki et al. 2014). For example, Salomaki et al. (2014) proposed a new genus

Sheathia Salomaki & M.L. Vis based on molecular data for species formerly classified in

Batrachospermum section Helminthoidea. Likewise, Entwisle et al. (2016) proposed an expanded Nothocladus Skuja to accommodate many Australian endemics that had previously been included in Batrachospermum. In addition, a new genus Nocturama

Entwisle & M.L. Vis was established for Australian specimens not closely related to

Nothocladus, but previously attributed to Batrachospermum (Entwisle et al. 2016).

These studies, as well as others, have made significant progress in our understanding of the systematics of the Batrachospermales. However, further taxonomic revision is required for the resolution of the paraphyly of Batrachospermum sensu lato, and new taxa are likely to be discovered with additional research.

North America has been shown to be the continent with the greatest batrachospermalean richness of genera and sections of Batrachospermum (Vis 2016).

However, only the genus Tuomeya is endemic to the continent with all others having members on other continents (Vis 2016). Thirty-eight infrageneric taxa were reported by 18 Sheath & Vis (2015) with five additional taxa described by Chapuis et al. (2017),

bringing the total to forty-three. North America appears to have a mix of both endemic

taxa and more cosmopolitan taxa with seventeen (40%) of those taxa being unique to this

continent (Sheath & Vis 2015, Chapuis et al. 2017). The cosmopolitan taxa, such as

Sirodotia suecica Kylin, have been recorded from Africa, Australia, Europe, and North

America based on specimens with molecular data (Lam et al. 2012). Documentation with

molecular data has been important in determining the biogeography of some members of

the Batrachospermales. For example, Sheathia boryana (Sirodot) Salomaki & M.L. Vis

(= Batrachospermum boryanum) was thought to be widespread in both North America

and Europe, but the sequence data showed that samples from the two continents were

genetically distinct, necessitating new species epithets to distinguish the North American specimens (Salomaki et al. 2014). The biogeographic patterns of these genera are diverse and further study of North American streams and rivers are expected to reveal more species and generic diversity.

In the southeastern United States, recent surveys have uncovered freshwater red algal diversity with new taxa proposed. Kumanoa holtonii M.L. Vis, Necchi, W.B.

Chiasson et Entwisle was described from a stream in Alabama (Vis et al. 2012), as well as Volatus personatus I.S. Chapuis & M.L. Vis and V. ulterior I.S. Chapuis & M.L. Vis from streams in Tennessee and Georgia, respectively, and Batrachospermum naiadis I.S.

Chapuis & M.L. Vis from Virginia (Chapuis et al. 2017). The current study continues this documentation of freshwater red algal diversity of the region with a study of specimens collected from streams in Kentucky and Alabama, USA. When the sequence data from the rbcL and COI -5P genes were phylogenetically analysed with previously published 19 data, they were determined to have a unique placement within Batrachospermales. The combination of these data and morphological assessment resulted in the proposal for

Lympha mucosa gen. et sp. prov.

Methods

Sample Collection

Samples were collected from the Kinniconick Creek, Kentucky (KY), and Hill

Creek, Alabama (AL), USA. Specific conductance of the stream water was measured with a Waterproof ECTestr low (Oakton, Vernon Hills, Illinois USA), pH with a

Waterproof pHTestr 30 Double Junction (Oakton), and temperature was taken with a thermometer. The algal collection was subdivided for DNA extraction, morphological examination and herbarium vouchers. Material for DNA extraction was cleaned of visible epiphytes, blotted dry, and immediately desiccated in silica gel. For morphological examination, material was placed in calcium carbonate-buffered 2.5% glutaraldehyde solution for preservation. The remaining material from each sample was used to create herbarium vouchers deposited in the New York Botanical Garden Herbarium (NY) and the Floyd Bartley Herbarium (BHO) at Ohio University.

DNA Extraction and Amplification

Silica-dried samples were ground in liquid nitrogen using a mortar and pestle.

DNA was extracted with a NucleoSpin® Plant II (Macherey-Nagel, Düren, Germany) kit following the manufacturer’s protocol. A region of the plastid-encoded ribulose 1,5- biphosphate (rbcL, 1,282 bp) gene and mitochondrial-encoded, 5’ region of the 20 cytochrome c oxidase subunit I (COI-5P, 664 bp) gene were PCR amplified. For the rbcL

marker, two different PCR systems were used as follows: 19 µL dH2O, 25 µL

AmpliTaqGold master mix (Applied Biosystems, Carlsbad, CA, USA), 2.5 µL each of

the F160 and rbcLR primers (Vis et al. 1998) and 1 µL of DNA for a total reaction

volume of 50 µL, or 32.75 µL dH2O, 5 µL 10X buffer, 4 µL dNTP, 4 µL MgCl2, 1.5 µL each of the above primers, 0.75 µL ExTaq (Takara Bio USA, Inc., Mountain View, CA,

USA) and 1 µL of DNA for a total reaction volume of 50 µL. The PCR parameters were as follows: initial denaturing at 95°C for 1 min; 35 cycles of 93°C for 30 s, 50°C annealing for 30 s and 68°C elongation for 1 min; and a final elongation at 72°C for 10 min. For the COI-5P, the PCR mixture consisted of reagents and volumes used for the rbcL marker, but with 2.5 µL/1.5 µL each of the GazF1 and GazR1 primers (Saunders

2005). The PCR parameters were as follows: initial denaturing at 94°C for 1 min; 40 cycles of 94°C for 1 min, 50°C annealing for 1.5 min, and 72°C extension for 1 min; and a final extension at 72°C for 5 min. Successful PCR products were purified with the

UltraClean™ PCR Clean-up DNA purification kit (Mo Bio, Carlsbad, CA, USA) following the manufacturers protocol.

Phylogenetic Analyses

The purified PCR products were sequenced at the Ohio University Genomics

Facility using the PCR amplification primers. For rbcL, additional internal primers were used for sequencing. Sequences generated from AL specimens were sequenced with the internal primers F650 (Salomaki et al. 2014) and R897.test (5’- 21 CGTGAGTATGTTGAATTACCTGC-3’) and specimens from KY with F650 and

R897.3 (Johnston et al. 2014). The raw data were edited and assembled using either

Geneious V7.1.9 software (Kearse et al. 2012) or Sequencher 5.2.4 (Gene Codes Corp.,

Ann Arbor, Michigan, USA). In addition, GenBank was searched for other batrachospermalean taxa (n=31) with representative COI-5P and rbcL sequences for phylogenetic analyses, and two species in the Thoreales were incorporated as outgroup taxa. Each individual gene and a concatenated alignment (1,946 bp) were subjected to

Bayesian Inference analysis (BI) using MrBayes v3.2 (Ronquist et al. 2012) and

Maximum Likelihood (ML) using RAxML (Stamatakis 2006) in Geneious plug-ins. For both analyses, genes and codon positions were partitioned (6 partitions) using the GTR + gamma model. The BI consisted of two independent simultaneous runs of one cold and three incrementally heated chains and 3 x 106 generations with sampling every 100 generations and the first 250 generated trees removed as burn-in to determine the posterior probabilities (pp). The ML analyses were conducted with the same model parameters and 1000 bootstrap support (bs) replicates.

Morphological Measurements

The morphological characters used in previous studies were measured to compare these specimens with members of Batrachospermum and other related genera (Table 2-1)

(Vis et al. 1995, Necchi and Vis 2012, Salomaki et al. 2014). At least fifteen measurements were taken for whorl diameter, fascicle cell number, carposporophyte and carposporangium dimensions, and spermatium diameter, and at least seven measurements were taken for carpogonium dimensions. The characters were observed and documented 22 using a BX40 Olympus microscope with an attached SC20 camera system (Olympus

American Inc., Center Valley, PA, USA).

Results

Molecular Results

Specimens collected from both locations were used to generate rbcL (1,282 bp) and COI-5P (664 bp) sequences. All unique sequences were deposited in GenBank.

Sequences for both markers were generated from additional specimens collected from both locations to confirm sequence similarity among specimens. Between the two locations, the rbcL sequence differed at 8 sites (0.6% divergent) and the COI-5P at 23 sites (3.5% divergent). The sequences for the two locations differed by 5.5% in the rbcL from the nearest sequence (Volatus carrionii), and 8.9-9.0% in COI-5P.

Phylogenetic trees inferred from BI and ML analyses of both the rbcL and COI-

5P showed similar topologies such that the two regions were concatenated for further analyses. Although both Bayesian and ML analyses were conducted, only the tree generated from a concatenated alignment of the ML analyses is presented with the statistical support from both analyses (Fig. 2-1). Both the BI and ML analyses placed this novel genetic group (herein Lympha mucosa gen. et sp. prov.) as sister to the recently described genus Volatus (Chapuis et al. 2017) with high support (96 bs/1.0 pp). Although the relationship of these genera with Tuomeya is weakly supported (<70 bs/0.8 pp), the larger clade composed of these, Batrachospermum sensu stricto, Paralemanea, , and was highly supported (100 bs/1.0 pp).

23 Morphological Results

Specimens from the KY population ranged in color from light green in the full sunlight to dark blue-green in shaded areas. The AL location was heavily shaded and the thalli collected were dark blue-green to almost black in color. Morphometric and qualitative characters were similar among specimens from the two locations (Fig. 2-2,

Table 2-1). The thalli had small whorls, which became indistinct in older parts of the thallus and the carposporophytes protruded from the whorls (Fig. 2-2 A-D, F-H). This difference in whorls and carposporophytes could be seen in the measurements of each with the whorls being primarily <200 µm and the carposporophyte diameters >200 µm

(Table 2-1). The thalli were densely compact as seen in cross-section (Fig. 2-2 E).

Carpogonia were clavate to elongate-lanceolate (Fig. 2-2 J, K). The specimens from the two locations did differ in mean carpogonium length with longer measurements from the

KY location (Table 2-1). The spermatangia and carposporangia appearance and measurements were characteristic for batrachospermalean taxa (Fig. 2-2 I, L, Table 2-1).

The suite of characters shown by these specimens is not present in any one genus or section of Batrachospermum. Therefore, a new genus and species are proposed.

Taxonomy

Lympha gen. prov. J.R. Evans, I.S. Chapuis et M.L. Vis

Description. Thalli ranging in color from light green in bright sunlight to dark blue-green to almost black in heavy shade. Thalli monoecious with dense, elongate pear- shaped or obconical or compressed whorls, becoming indistinct in older parts. Secondary fascicles very abundant. Spermatangia terminal or subterminal on primary fascicles or 24 adjacent to the involucral filaments. Cortical cells of the main axis, appressed to the axis and cylindrical. Carpogonial branches straight, few-celled, arising from the pericentral or proximal cell. Involucral filaments short and scarce. Carpogonia with clavate to elongate- lanceolate trichogynes. Carposporophyte axial, subspherical to spherical, and protruding from the whorl, carposporangia ovoid.

Etymology. The generic name is to honor the ancient Roman deity of fresh water,

Lympha. She was the deity to whom one prayed to maintain the water supply.

Lympha mucosa sp. prov. J.R. Evans, I.S. Chapuis et M.L. Vis

Description. Thalli ranging in color from light green in bright sunlight to dark blue-green to almost black in heavy shade. Thalli monoecious with dense, elongate pear- shaped or obconical or compressed whorls, becoming indistinct in older parts. Mature whorls 154-223 µm in diameter. Primary fascicles composed of 4-7 cells. Secondary fascicles very abundant. Cortical cells of the main axis, appressed to the axis and cylindrical. Spermatangia (5.5-8 µm in diameter) terminal or subterminal on primary fascicles or adjacent to the involucral filaments (see below). Carpogonial branches composed of 1-3 cells, straight, arising from the pericentral or proximal cell. Involucral filaments short and scarce. Carpogonia with clavate to elongate-lanceolate trichogynes, 9-

13 µm in diameter and 32-55 µm long. Carposporophyte axial, 1 per whorl, dense, sub- spherical to spherical, protruding from the whorl, 173-305 µm in diameter and 151-247

µm high; gonimoblast filaments composed of 2-3 cells; carposporangia ovoid, 14-20 µm in diameter and 23-31 µm in length.

Holotype. USA, Kinniconick Creek, Lewis County, Kentucky, N 38º29'48", W 25 83º15'26" Coll. M.L. Vis, W.B. Chiasson & O. Necchi Jr., 06.viii.2008, Holotype NY

Voucher Barcode 02137800, Isotype BHO A-176.

Topotypes. Coll. A. Vandenbroek & P. Contraras. 17.ix.2009. N 38º29'48", W

83º15'26", BHO A-407. Coll. M.L. Vis & D. Gonzalez. 17.vi.2014. N 38º29'48", W

83º15'26", BHO A-1248.

Representative DNA barcode. GenBank KM593865 (rbcL), KM593873 (COI-

5P).

Etymology. The species epithet is in recognition of the large quantities of cell wall

polysaccharides that give the thallus a slimy feel and makes this species particularly

difficult for extraction of DNA and RNA.

Specimens examined. In addition to the type material, a second population was

examined. Hill Creek, Alabama, USA. 33.0536° N, 87.18726° W. Coll. M.L. Vis, A.

Rambani & A. Vandenbroek. 25.vi.2009. BHO A-0175 (GenBank accession rbcL

KM593850, COI-5P KM593868).

Distribution. This species has been collected in two locations of the southeastern

United States, in Kentucky and Alabama, USA.

Remarks. Stream conditions recorded at the KY site while collecting the holotype

were as follows: water temperature 25°C, pH 7.3, and specific conductance of 140 µS

cm-1. Water conditions when topotypes were collected were as follows: water

temperature 19, 29°C, pH 6.6, 8.3 and specific conductivity 110, 120 µS cm-1. The AL site had a water temperature of 30°C, pH 8.1, and specific conductance of 230 µS cm-1

26 Discussion

In the phylogenetic analyses, Lympha was most closely related to Volatus.

However, these two genera are easily distinguished based on morphology. The genus

Volatus is similar to Kumanoa in that members of both genera produce curved carpogonial branches (Chapuis et al. 2017). In contrast, L. mucosa produces straight carpogonial branches. The three species of Volatus have been reported from North

America. Volatus carrionii has a known distribution in both Europe and eastern Canada, while V. personatus and V. ulterior have been collected from streams in Georgia and

Tennessee, USA. It is interesting that L. mucosa, V. personatus and V. ulterior occur in the southeastern US, a region known for biodiversity of aquatic organisms (Boschung and Mayden 2004, Williams et al. 2008). All three taxa are currently only known from one to two stream sites, but it is probable that more locations will be discovered with further thorough surveying of this region.

Although not closely related in the phylogenetic analyses, Lympha is morphologically most similar to members of Batrachospermum sections Virescentia and

Turfosa. Shared attributes include short, straight carpogonial branches, axial carposporophytes that are typically 1-2 per whorl and large in comparison to the whorl.

However, most species of section Virescentia have well-developed whorls and carpogonia with stalked cylindrical trichogynes (Kumano 2002). Section Turfosa is more similar to Lympha in thallus habit with dense, compressed whorls becoming indistinct in older parts and carpogonium/trichogyne dimensions and shape. However, Section

Turfosa is characterized by the occurrence of two gonimoblast filament types

(determinate and indeterminate) in the carposporophytes, which is not present in Lympha. 27 The southeastern US is a rich region for freshwater batrachospermalean diversity.

As noted above, Lympha mucosa and two species of Volatus have only been collected from this area. Likewise, Kumanoa holtonii has only been reported from a single stream in Alabama. Although Tuomeya americana has a restricted distribution to eastern North

America, it is quite commonly collected in the southeast (Kaczmarczyk et al. 1992).

Beyond these taxa with restricted distributions, there are many more with wider geographic distributions that inhabit this region with at least four Batrachospermum species, three Sheathia species, four Kumanoa species as well as at least one

Paralemanea species (Vis and Sheath 1992, 1997, Sheath et al. 1994a, b, c, Necchi and

Vis 2012, Salomaki et al. 2014). One likely reason for the diversity of the area for freshwater reds is that parts of the southeastern US served as a putative southern

Pleistocene glacial refugium for plants and animals (Hewitt 1996, Church et al. 2003,

Zamudio & Savage 2003, Griffin & Barrett 2004). House et al. (2010) hypothesized that

Alabama was a southern refugium for Batrachospermum gelatinosum due to the greater genetic diversity in COI-5P within those populations than in more northern locations.

These findings suggest that the southeastern US may be a ‘hotspot’ of batrachospermalean diversity in North America and further studies are warranted.

As a natural classification for batrachospermalean taxa is achieved, our understanding of the known distribution for these species is also subject to change. The once global distribution of Batrachospermum is now been largely restricted to the northern hemisphere, with new generic descriptions to accommodate most taxa previously assigned to the genus that occur in the southern hemisphere (e.g., Entwisle et al. 2016). Likewise, new genera are being proposed for many northern hemisphere taxa 28 (e.g., Salomaki et al. 2014, Chapuis et al. 2017). The proposal of Lympha mucosa gen. et

sp. prov. supports the vast amount of genetic variation observed in species traditionally

allied to Batrachospermum sensu lato, and contributes to a monophyletic

Batrachospermales and diversity of this order on North America.

References

Boschung, H.T., Jr. & Mayden, R.L. 2004. of Alabama. Smithsonian Books,

Washington D.C., USA. 736 pp.

Chapuis, I.S., Necchi Jr., O., Zuccarello, G.C., Xie, S.L., Aboal, M., Sánchez Castillo,

P.M. & Vis, M.L. 2017. A new genus, Volatus and four new species of

Batrachospermum sensu stricto (Batrachospermales, Rhodophyta). Phycologia

56:454-468.

Church, S.A., Kraus, J.M., Mitchell, J.C., Church D.R. & Taylor, D.R. 2003. Evidence

for multiple Pleistocene refugia in the postglacial expansion of the eastern tiger

salamander, Ambystoma tigrinum tigrinum. Evolution 57:372-383.

Entwisle, T.J., Vis, M.L., Chiasson, W.B., Necchi, O. & Sherwood, A.R. 2009.

Systematics of the Batrachospermales (Rhodophyta) – a synthesis. J. Phycol.

45:704-715.

Entwisle, T.J., Johnston, E.T., Lam, D.W., Stewart, S.A. & Vis, M.L. 2016. Nocturama

gen. nov., Nothocladus s. lat. and other taxonomic novelties resulting from the

further resolution of paraphyly in Australasian members of Batrachospermum

(Batrachospermales, Rhodophyta). J. Phycol. 52:384-396. 29 Griffin, S.R. & Barrett, S.C.H. 2004. Post-glacial history of Trillium grandiflorum

(Melantiaceae) in eastern North America: inferences from phylogeography. Am.

J. Bot. 91:465-473.

Hewitt, G.M. 1996. Some genetic consequences of ice ages, and their role in divergence

and speciation. Biol. J. Linnean Soc. 58:247-276.

House, D.L., Vandenbroek, A.M. & Vis, M.L. 2010. Intraspecific genetic variation of

Batrachospermum gelatinosum (Batrachospermales, Rhodophyta) in eastern

North America. Phycologia 49:501-507.

Johnston, E.T., Lim P.E., Buhari, N., Keil, E.J., Djawad, M.I. & Vis, M.L. 2014.

Diversity of freshwater red algae (Rhodophyta) in Malaysia and Indonesia from

morphological and molecular data. Phycologia 53:329-341.

Kaczmarczyk, D., Sheath, R.G. & Cole, K.M. 1992. Distribution and systematics of the

freshwater genus Tuomeya (Rhodophyta, ). J. Phycol.

28:850-855.

Kearse, M., Moir, R., Wilson, A., Stones-Havas, S., Cheung, M., Sturrock, S., Buxton, S.,

Cooper, A., Markowitz, S., Duran, C., Thierer, T., Ashton, B., Mentjies, P. &

Drummond, A. 2012. Geneious Basic: an integrated and extendable desktop

software platform for the organization and analysis of sequence data.

Bioinformatics 28:1647-1649.

Kumano, S. 2002. Freshwater red algae of the world. Biopress Ltd, Bristol, 375 pp.

Lam, D.W., Entwisle, T.J., Eloranta, P., Kwandrans, J. & Vis, M.L. 2012.

Circumscription of species in the genus Sirodotia (Batrachospermales, 30 Rhodophyta) based on molecular and morphological data. Eur. J. Phycol. 47:42-

50.

Ronquist, F., Teslenko, M., Van Der Mark, P., Ayres, D.L., Darling, A., Hohna, S.,

Larget, B., Liu, L., Suchard, M.A. & Huelsenbeck, J.P. 2012. MrBayes 3.2:

Efficient Bayesian phylogenetic inference and model choice across a large

model space. Syst. Biol. 61:539-542.

Salomaki, E.D., Kwandrans, J., Eloranta, P. & Vis, M.L. 2014. Molecular and

morphological evidence for Sheathia gen. nov. (Batrachospermales,

Rhodophyta) and three new species. J. Phycol. 50:526-542.

Saunders, G.W. 2005. Applying DNA barcoding to red macroalgae: a preliminary

appraisal holds promise for future applications. Phil. Trans. R. Soc. B. 360:1879-

1888.

Sheath, R.G. 1984. The biology of freshwater red algae. In Round, F.E. & Chapman, D.J.

[Eds] Progress in Phycological Research, vol. 3. Biopress Ltd, Bristol UK, pp.

89-157.

Sheath, R.G., Vis, M.L. & Cole, K.M. 1994a. Distribution and systematics of

Batrachospermum (Batrachospermales, Rhodophyta) in North America. 5.

Section Aristata. Phycologia 33:404-414.

Sheath, R.G., Vis, M.L. & Cole, K.M. 1994b. Distribution and systematics of

Batrachospermum (Batrachospermales, Rhodophyta) in North America. 4.

Section Virescentia. J. Phycol. 30:108-117. 31 Sheath, R.G., Vis, M.L. & Cole, K.M. 1994c. Distribution and systematics of

Batrachospermum (Batrachospermales, Rhodophyta) in North America. 6.

Section Turfosa. J. Phycol. 30:872-884.

Sheath, R.G. & Vis, M.L. 2015. Red algae. In Wehr, J.D., Sheath, R.G. & Kociolek, J.P.

[Eds] Freshwater Algae of North America second edition. Elsevier Academic

Press, San Diego. pp. 237-264.

Vis, M.L. 2016. Biogeography of river algae. In Necchi, O. Jr. [Ed] River Algae. Springer

Nature, pp. 219-244.

Vis, M.L. & Sheath, R.G. 1992. Systematics of the freshwater red algal family

Lemaneaceae in North America. Phycologia 31:164-179.

Vis, M.L. & Sheath, R.G. 1997. Biogeography of Batrachospermum gelatinosum

(Batrachospermales, Rhodophyta) in North America based on molecular and

morphological data. J. Phycol. 33:520-526.

Vis, M.L., Sheath, R.G. & Entwisle, T.J. 1995. Morphometric analysis of

Batrachospermum section Batrachospermum (Batrachospermales, Rhodophyta)

type specimens. Eur. J. Phycol. 30:35-55.

Vis, M.L., Necchi, O. Jr., Chiasson, W.B. & Entwisle, T.J. 2012. Molecular phylogeny of

the genus Kumanoa (Batrachospermales, Rhodophyta). J. Phycol. 48:750-758.

Vis, M.L., Saunders, G.W., Sheath, R.G., Dunse, K. & Entwisle, T.J. 1998. Phylogeny of

the Batrachospermales (Rhodophyta) inferred from rbcL and 18S ribosomal

DNA gene sequences. J. Phycol. 32:341-350. 32 Williams, J.D., Bogan, A.E. & Garner, J.T. 2008. Freshwater mussels of Alabama and

the Mobile basin in Georgia, Mississippi, and Tennessee. The University of

Alabama Press, Tuscaloosa, USA. 960 pp.

Zambudio, K.R. & Savage, W.K. 2003. Historical isolation, range expansion, nd

secondary contact of two highly divergent mitochondrial lineages in spotted

salamanders (Ambystoma maculatum). Evolution 57:1631-1652. 33 Table 2-1. Morphometrics (mean ± SD) of vegetative and reproductive characters for Lympha mucosa gen. et sp. prov. from

Kinniconick Creek, KY, and Hill Creek, AL, USA. For each population, at least fifteen measurements were taken for all characters except for carpogonial dimensions, which had seven measurements.

Taxon Whorl Spermatangium Carpogonium Carpogonium Carposporophyte Carposporophyte Carposporangium Carposporangium location diameter diameter (µm) diameter (µm) length (µm) diameter (µm) length (µm) diameter (µm) length (µm)

(µm)

KY 192 ± 31 6.4 ± 0.9 11.1 ± 2.2 48.9 ± 6.5 252 ± 53 212 ± 35 16.6 ± 2.2 25.1 ± 2.5

AL 174 ± 20 6.7 ± 1.2 11.7 ± 0.6 36.8 ± 4.5 206 ± 33 179 ± 28 17.6 ± 1.9 28.1 ± 3.1

34

Figure 2-1 Maximum likelihood (ML) phylogeny of Lympha mucosa gen. et sp. prov. with other genera of Batrachospermales. The phylogeny is based on rbcL and COI-5P concatenated sequence data with support values given as bootstrap/posterior probability

(pp) from Bayesian inference. Asterisks (*) indicates full support in both analyses, and dashes (-) indicate support <70% bootstrap/0.8 pp. Outgroup taxa (Thoreales) not shown.

GenBank accession numbers provided in Appendix A. 35

Figure 2-2 Morphological characteristics of Lympha mucosa gen. et sp. prov. (A) Habit of thallus. (B) Habit of thallus with conspicuous carposporophytes (arrows). (C) Dense, elongate pear-shaped whorls with abundant secondary fascicles. (D) Confluent obconical whorls with abundant secondary fascicles. (E). X-section of thallus showing tightly packed cortical cells (arrows) with radiating short-celled fascicle that compose the whorl. (F) Immature, spherical carposporophyte (arrow) that is axial and protruding from the whorl. (G) Mature, dense, hemispherical carposporophyte (arrow) protruding from whorl and carpogonium (arrowhead) on side branch. (H) Mature, dense, spherical carposporophytes with one protruding from the whorl (arrow) and one seemingly at the 36 termination of a branch (arrowhead). (I) Spermatangia (arrows) at a fascicle tip. (J) Immature carpogonium with a clavate trichogyne

(arrow) on a short-celled straight carpogonial branch (arrowhead). (K) Mature carpogonium with an elongate-lanceolate trichogyne

(arrow) on a short-celled straight carpogonial branch (arrowhead). (L) Compact carposporophyte with ovoid carposporangia (arrows) at branch tips. Scale bars represent: A, 5 mm; B, 200 µm; C, D &; F, 100µm; E, 20 µm; G, H, 150 µm; I-K, 10 µm; L, 30 µm. 37 CHAPTER 3: PLASTID GENOMICS AND RELATIVE EXPRESSION ANALYSIS

OF LIGHT-HARVESTING GENES IN LYMPHA MUCOSA EXPOSED TO LOW AND

HIGH IRRADIANCES

Introduction

Like other oxygenic, photosynthetic organisms, red algae have two photosystem

(PS) reaction centers that serve as the sites of light-harvesting and photophosphorylation,

a critical process for the generation of usable cellular energy (Lepetit and Dietzel 2015).

The pigment composition of PS I is fairly uniform in all lineages, being primarily

composed of chl. a, and a specialized chl. a (P700) molecule that serves as the active reaction center (Gantt 1990). Although most photosynthetic eukaryotes contain other chlorophylls that comprise the light-harvesting antenna of PS II, red algae differ by

utilizing light-harvesting phycobiliprotein complexes known as phycobilisomes (Gantt

1990). The differences in the absorption spectra of these unique phycobilins relative to

chlorophylls allow red algae to take advantage of energy from wavelengths poorly

absorbed by most green algae and plants (Gantt 1990).

In the red algae, other apoproteins that play an important role in light harvesting

for photosynthesis include high-light-inducing proteins (HLIP), one-helix proteins

(OHP1), two-helix stress-enhanced proteins (SEP), and chl a/b binding-like proteins

(RedCAP)(Engelken et al., 2010). All of these proteins are part of the Light-Harvesting-

Complex-like (LHC-like) superfamily, which serve critical functions in light-harvesting

and for photoprotective mechanisms in all photosynthetic eukaryotes (Engelken et al.

2010). However, some of these proteins are found only in eukaryotic lineages that

obtained their chloroplast organelle from an ancestral red alga, such as the RedCAP 38 proteins (Sturm et al. 2013). In diatoms, RedCAP proteins are localized near the light- harvesting antenna of PS I and their regulation is affected by irradiance (Sturm et al.

2013).

The ~5% of red algae that occur in freshwater environments have a worldwide distribution, particularly in streams with low organic pollution and little canopy modification (Sheath 1984). The strictly freshwater order Batrachospermales is widespread, but most taxa occur in first- to third-order streams (Entwisle et al. 2009,

Sheath and Vis 2015). The distribution and biogeographic patterns in the

Batrachospermales are diverse, but there are limited physiological and genetic data for factors that may affect distributions, phenology, and dispersal (Vis 2016).

Light regime is an important environmental factor that controls the growth, distribution, and seasonality of freshwater red algae (Sheath 1984, Necchi et al. 1999). In temperate regions, many streams have seasonal changes in light regimes related to deciduous tree canopy cover that affect red algal distribution and seasonality. Drerup and

Vis (2014) noted differences in the phenology of Batrachospermum gelatinosum from geographically close temperate streams with the gametophyte thalli being spring ephemerals in the smaller stream and perennial in the larger stream. The spring ephemerals had growth inhibited with the onset of summer in the smaller streams, likely due to greater canopy cover, and consequently a lack of sufficient light for basic photosynthetic requirements.

Previously, ecological investigations of the effect of light regime on growth and photosynthetic rates in freshwater red algae have concluded that the Batrachospermales are primarily shade-adapted. These investigations included transplant experiments of two 39 Batrachospermum spp. from shaded sites that disintegrated in the first few days of

exposure to open sites (Parker et al. 1973); and ecophysiological studies of

photosynthetic rates in several batrachospermalean taxa that become light saturated at

irradiances < 250 mol photon m-2 s-1 (e.g., Leukart and Hanelt 1995, Necchi and Zucchi

2001, Necchi 2005, Drerup et al. 2015). However, there are several species that occur

naturally in consistently higher light intensities, some as high as 2,400 mol photon m-2 s-

1 (Bautista and Necchi 2007), and different taxa occurring in the same stream can vary in

their response to changes in light intensities (Drerup et al. 2015). As well, species that

were previously considered shade-adapted (e.g., Parker et al. 1973) have since been

shown to acclimate to a much wider range of light (Aigner et al. 2017). Hence, it is probable that red algal photosystems have evolved adaptations for a wide range of light,

and species can exhibit substantial differences in their capacity for photoacclimation.

The term “acclimation” refers to the ability of an organism to respond to

environmental changes within the limits of its genome, whereas “adaptation” refers to

responses that have formed through alterations to the genome over generations (Gantt

1990). Studies have shown various types of photoacclimation utilized by freshwater red

algae including differential adjustments to pigment content and distinct photosynthetic characteristics, which have been documented in culture and field studies of eight freshwater red algal taxa (Kaczmarcyk and Sheath 1991, Bautista and Necchi 2007,

Drerup et al. 2015). Potential adjustments for photoacclimation in red algae include changes in the size or number of photosystems and/or phycobilisomes on the thylakoid, and changes in photopigment content, specifically the ratio of phycocyanin and phycoerythrin (Gantt 1990, Kaczmarcyk and Sheath 1991, Bautista and Necchi 2007). 40 Adjustments of light-harvesting machinery at varying irradiances include

measurable differences in chl. a and total pigment content, which generally decrease in concentration at higher light intensities (Kaczmarcyk and Sheath 1991, Bautista and

Necchi 2007). In addition, the dissimilarities in these adjustments among different taxa are representative of two photoacclimation strategies, shade-adapted or sun-adapted algae

(Bautista and Necchi 2007). During long-term exposure (4 days), Bautista and Necchi

(2007) reported spp. had characteristics of shade-adapted algae (e.g., changes in the size of PS units) and Compsopogon caeruleus had some characteristics of sun-adapted algae (e.g., changes in the number of PS units). Although not statistically supported, these differences were correlated with gross morphology [tuft morphology

(i.e., Audouinella) versus filamentous morphology (i.e., Compsopogon)], which has been observed in field-collected specimens (Drerup et al. 2015). However, for freshwater red algae, no genetic data are available for discerning the cellular mechanisms that regulate

these observed differences in light-harvesting machinery.

The majority of photosynthetic studies for freshwater red algae have been at the

whole-plant level, but understanding photoacclimation strategies and other

photosynthetic processes likely involves determining the changes that occur at both the

individual and molecular levels (Talarico and Maranzana 2000). The kinetics of

photoacclimation have been studied in the unicellular red alga, Rhodella violacea, which

included measurements of mRNA transcript abundance for four plastid-encoded genes

involved in P700 chl. a and phycobiliprotein biosynthesis (Ritz et al. 2000).

Phycobiliprotein gene expression showed an inverse relationship with increasing light

quantity, but the psaA gene encoding the P700 chl. a. was upregulated with increasing 41 irradiance (Ritz et al., 2000). These results are indicative of transcriptional regulation as a potential mechanism for making cellular adjustments related to photoacclimation in red algae, which occurs more frequently in their plastids than in green algae and plants

(Minoda et al. 2005).

Red algal plastid genomes are among the largest of sequenced taxa (i.e., over

1Mbp, Muñoz-Gómez et al. 2017), but most are in the 170-200 Kbp range (e.g.,

Verbruggen and Costa 2015, Lee et al. 2016a, b, Costa et al. 2016, Hughey et al. 2017).

These genomic features, combined with ultrastructure, make red algal plastids the most reminiscent of the cyanobacterial progenitor to the organelle (McFadden 2001,

Janouškovec et al. 2013, Muñoz-Gómez et al. 2017). Although these genomes are reduced compared to extant cyanobacterial species, their gene content is two-three times greater than most green algae and plants (i.e., <100 genes, Barkan 2011), often encoding

>200 genes, many of which are related to photosynthetic and other basic cellular processes.

Sequencing red algal plastid genomes has provided sufficient data for studies of plastid evolution (e.g., Lee et al. 2016a, b) and phylogenomics (e.g., Costa et al. 2016). In addition, these data are useful for genetic regulatory studies of photosynthetic processes, such as photoacclimation. The goal of the current study was to generate genetic data related to photoacclimation in a freshwater red alga that occurs in a wide range of irradiances. Lympha mucosa is a mucilaginous, filamentous taxon in the

Batrachospermales that occurs abundantly throughout the summer months in both shaded and open sites in a single stream. Previously, Drerup et al. (2015) reported high photosynthetic efficiencies and saturation points for this taxon (as Batrachospermum sp.), 42 but little is known of the adjustments at the molecular level. Therefore, the plastid

genome was sequenced and utilized as a tool to determine the DNA sequence of light- harvesting genes to be utilized in a laboratory experiment of light acclimation. The gene expression for five plastid-encoded, and one nuclear-encoded, light-harvesting genes from sun-acclimated and shade-acclimated thalli in sun and shade conditions were compared to determine transcriptional regulation of these genes. It was hypothesized that the underlying mechanism controlling photoacclimation in freshwater red algae is transcriptionally regulated. Therefore, genes encoding components of the red algal light harvesting machinery were expected to show differential expression after exposure to varying irradiances.

Methods

Sample Collection and Experimental Design

Samples of Lympha mucosa were collected from the Kinniconick Creek, KY in

July 2016. Site location and conditions, including water temperature, pH, specific conductance, and light quantity at the river midsection (open site) and edge (shaded site)

were recorded (Table 3-1). Stream water specific conductance was measured with a

Waterproof ECTestr low (Oakton, Vernon Hills, Illinois USA), pH with a

Waterproof pHTestr 30 Double Junction (Oakton) and temperature with a thermometer.

Light quantity was measured using a Li-250A light meter equipped with a Li-192

underwater quantum sensor (Li-Cor, Lincoln, Nebraska USA). Twelve samples were

collected from the river midsection (i.e., sun-acclimated) and the river edge (shade- 43 acclimated) for a total of 24 samples, and these were stored in bags on ice for transport.

Additional samples were collected for DNA extraction.

In the laboratory, samples for DNA extraction were desiccated in silica gel. All other samples (~ 1g wet weight each) were immediately put in Corning culture flasks with 35 mL filtered stream water and placed on a Lab-line orbital shaker (Lab-line

Instruments Inc., Dubuque, Iowa USA) at a medium setting to simulate water flow

(Appendix B). The shaker was located in a Conviron CMP6050 walk-in growth chamber

(Controlled Experiments Ltd., Winnipeg, Manitoba, Canada) equipped with 60-W cool- white fluorescent and/or 40-W incandescent bulbs. Sun- and shade-acclimated samples were split equally into two treatments of low (<20 mol photon m-2 s-1) or high (220

mol photon m-2 s-1) light, resulting in four treatment groups: high light (HL)/Sun- acclimated (SA), HL/shade-acclimated (SH), low light (LL)/SA, LL/SH. The quantity of light in the growth chamber was set to high light conditions, and low light conditions were achieved by covering flasks with two layers of 50% shade cloth. Light conditions for each treatment were verified using the Li-250A light meter. All other growth chamber conditions were kept constant: temperature 22C, relative humidity 50-70% and day length 16:8 light:dark. Photoacclimation can take a few days to a few weeks in red algal cells (e.g., Gantt 1990) and it is difficult to culture batrachospermalean longer than one week. Therefore, samples were allowed to grow in the growth chamber for 72 h to allow enough time to acclimate to new light conditions, but without deterioration to the thalli. Following the 72 h growth period, all samples were flash frozen in liquid nitrogen and stored in a -80C freezer until RNA extraction.

44 DNA Extraction and Sequencing

Silica-desiccated samples had DNA extracted with a NucleoSpin® Plant II DNA kit (Macherey-Nagel) following the manufacturer’s protocol. Due to large quantities of polysaccharides that became evident during extraction, a pre-precipitation modification used previously for other polysaccharide-rich algae (e.g., Dos Reis Falcão et al. 2008) was needed to successfully isolate pure DNA from the sample. During the lysis step,

0.2M KOAc was added to the buffer containing sample and mixed. This mixture was

incubated for 10 minutes, followed by centrifugation at 12,000 x g for 10 minutes. A

large quantity of polysaccharides and KOAc were pelleted, and the supernatant

containing DNA was transferred for purification. Quality of the isolated DNA was

examined by visualizing 20 L DNA in a 0.6% agarose gel stained with ethidium

bromide and measuring the 260/280 ratio using a NanoDrop Spectrophotometer (Thermo

Scientific, Waltham, MA, USA), both of which indicated high-quality DNA.

Isolated DNA was processed and sequenced using an Illumina MiSeq platform

(Illumina Inc., San Diego, CA, USA), with the following steps performed by the Ohio

University Genomics Facility (OUGF) personnel. DNA was initially quantified using 1

L of DNA on a Qubit® 2.0 fluorometer (Thermo Scientific). The paired-end library was

then prepared with a Nextera™ XT DNA Sample Prep kit following the manufacturer’s

protocol for tagmentation and library enrichment. The DNA library was purified with

Agencourt AMPure® XP purification beads (Beckman Coulter Life Sciences,

Indianapolis, IN USA). After a final quality check with an Agilent 2100 Bioanalyzer

(Agilent Technologies, Santa Clara, CA USA), the recovered DNA library was used as

input for bridge amplification and cluster generation following the standard Illumina 45 protocol. Paired-end sequencing was completed using Nextera™ sequencing primers with an Illumina MiSeq Reagent Kit v3 (600 cycle). An additional sequencing run using an Illumina MiSeq Reagent Kit v2 (300 cycle) was required to compensate for bacterial contamination that overloaded the first round of sequencing. Raw data were uploaded to the Illumina BaseSpace server with removal of adaptor sequences.

Plastid Assembly and Annotation

Raw reads were assessed for per base sequence quality, content, length distribution, and Kmer content with FastQC (Babraham Bioinformatics; https://www.bioinformatics.babraham.ac.uk/projects/fastqc/). Low-quality reads (

(https://clcbio.com), and with the SPAdes 3.10.0 assembler plug-in for Geneious 10.1.3

(Kearse et al. 2012). For the CLC assembly, reads were assembled with default parameters (word size = 20, bubble size = 50) with a minimum contig length of 1000 bp and paired distances auto-detected. For the SPAdes assembly, multi-cell data option was chosen with careful mode set to reduce mismatches and short indels. Contiguous sequences (contigs) generated from the De Novo assemblies were made into a local

BLAST database. The complete plastid genome sequence for the batrachospermalean taxon, Kumanoa americana (GenBank accession NC031178; Lee et al. 2016a), was used as a query to find possible plastid matches in the contig database. Contigs with matches to K. americana were remapped with the filtered L. mucosa reads using either the “map 46 reads to contigs” option in CLC with a similarity fraction of 0.85-0.9 and length fraction of 0.8-0.85, or the “map to reference” option in Geneious v.10.1.3 with medium

sensitivity/fast, until the circular nature of the molecule was verified. A final remapping

was completed with Bowtie2 (Langmead and Salzberg 2012) to verify the entire plastid

sequence.

Putative ORFs and protein-coding predictions were completed using ORF finder

and live annotation and prediction options in Geneious v.10.1.3, and using the tBLASTx

tool (e-value 1.0e-06) with codon translation table 11 (Bacterial, Archaeal, and Plant

Plastid Code). Gene annotations were performed with Pfam 30.0 (Finn et al. 2016,

http://pfam.xfam.org/) and MFannot (http://megasun.bch.umontreal.ca/cgi-

bin/mfannot/mfannotInterface.pl). Annotations for rRNA and tRNA genes were detected

using RNAmmer (Lagesen et al. 2007, http://www.cbs.dtu.dk/services/RNAmmer/) and

tRNAdb (Chan and Lowe 2009, http://gtrnadb2009.ucsc.edu/). Intronic regions were

identified using RNAweasel (http:www.theplantlist.org). Final annotations were

embedded and visualized on the L. mucosa plastid genomic sequence using Geneious v.10.1.3 and will be submitted to GenBank. The L. mucosa plastid genome was compared

to other sequenced freshwater taxa in the Nemaliophycidae (n=3) for conservation, gene gain/loss, and rearrangements using a progressive Mauve alignment (Darling et al. 2010) in Geneious v10.1.3.

RNA Extraction

Frozen Lympha mucosa samples for total RNA isolation were processed within four months of collection to reduce the potential for sample degradation. Total RNA was 47 isolated using a NucleoSpin® RNA Plant Kit (Macherey-Nagel) following the manufacturer’s protocol, with similar modifications used in the DNA extraction. Isolation of RNA was attempted with other silica column-based kits (Qiagen RNeasy, Zymo

Quick-RNA™ MiniPrep, MoBio RNA Isolation with polysaccharide inhibitor), but each required similar steps to purify RNA from samples. Therefore, NucleoSpin® was used for extraction of all experimental samples. Initially, a 0.2M KOAc pre-precipitation step was completed as outlined above; however, for RNA samples the incubation period was performed at -20C, followed by 4C centrifugation at 12,000 x g for 10 minutes to reduce RNA degradation. RNA in the supernatant layer was then removed and

precipitated on column with the addition of 0.8 volume 100% EtOH. DNA digestion was

performed on-column with a 15-minute incubation period. The absorbance ratio of each

RNA sample was measured using a NanoDrop spectrophotometer, and quantity/quality

was measured using an Agilent 2100 Bioanalyzer and given an RNA Integrity Number

(Schroeder et al. 2006) (Appendix C). RNA samples were stored in a -80C freezer.

Gene Selection and RT-qPCR Primer Design

The L. mucosa plastid genome generated in this study was used to design specific

primers for gene expression analyses. Target genes chosen for study of photoacclimation

were based on their function in light harvesting, and in their use in previous studies (Ritz

et al. 2000, Engelken et al. 2010, Lepetit and Dietzel 2015). Subunits encoding both

photosystem reaction centers (psaA, photosystem I P700 chlorophyll a apoprotein A1;

psbA, photosystem II protein D1), and a gene encoding a photophosphorylated electron

acceptor (petF, PetF ferredoxin I) were targeted. For the light-harvesting phycobilisome 48 protein complex, subunits encoding each of the main pigment-binding proteins were

chosen (apcA, allophycocyanin; cpcA, phycocyanin; cpeA, phycoerythrin). In addition to plastid-encoded genes, sequence data for the nuclear-encoded HV60 were mined from the

L. mucosa high-throughput sequencing data using tBLASTx searches and the one

available red algal HV60 CDS sequence in GenBank (Griffithsia japonica, GenBank

accession AY123144). The endogenous controls chosen as reference genes in the RT-

qPCR analysis were based on available literature of stable gene expression for nuclear-

encoded red algal genes (eRF3, ethylene-responsive transcription factor 3; IF4A-1,

eukaryotic translation initiation factor 4A1; Kowalczyk et al. 2014), and for plastid-

encoded genes in vascular plant groups (accD, acetyl-coenzyme A carboxylase carboxyl

transferase subunit beta; rps3, ribosomal protein subunit 3; Cortleven et al. 2009). These

genes are stably expressed in at least fifteen different environmental conditions, including

changes in light regime. Reference genes were explored in each genomic compartment to

account for regulatory differences that occur in separate genomes of the same cell

(Cortleven et al. 2009).

Primer sets for each gene were designed near the three-prime region, when

possible, using NCBI Primer-BLAST (Ye et al. 2012) (Fig. 3-1). Specifications were set

with an optimal melting temperature (Tm) of 60C, maximum Tm difference of 1C

between primer pairs, and GC content of 40-60%. The specificity of these synthetic

oligonucleotides was tested in silico on the L. mucosa plastid genome and through a

BLASTn search on the contig database. Primer sets were tested for amplification using

gDNA (given there were no intronic regions in these genes to use for primer design).

These products were purified using an UltraClean™ PCR Clean-up DNA purification kit 49 (Mo Bio, Carlsbad, CA, USA) and Sanger sequenced at the OUGF to determine

successful binding to the correct genomic region. Successful primer sets were used for

RT-qPCR (Table 3-2). All nuclear gene sequences will be submitted to GenBank.

cDNA Synthesis and RT-qPCR Conditions

A total of three biological replicates were used for each treatment and consisted of

RNA samples that were the highest quality (Appendix C). RNA samples were

synthesized into cDNA using the iScript™ cDNA Synthesis Kit (Bio-Rad, Hercules,

California USA). Each reaction consisted of 4 L 5X iScript reaction mix, 1 L iScript

reverse transcriptase, and 15 L RNA template for a total volume of 20 L. The PCR parameters used for cDNA synthesis followed the manufacturer’s protocol. The cDNA samples were stored in a -20C freezer.

For RT-qPCR, PowerUp™ SYBR™ Green Master Mix (Thermo Fisher

Scientific, Waltham, MA USA) was used with ROX as a passive reference dye.

Amplification was performed on either an Agilent Stratagene Mx3000P qPCR machine

(Agilent Technologies, Santa Clara, CA USA) or a Bio-Rad CFX96 Touch™ Real-Time

PCR Detection System (Bio-Rad Laboratories, Hercules, CA, USA). Initially, each gene

product had cDNA concentration and primer efficiencies optimized by performing a 5-

fold dilution series to produce a cycle threshold (Ct) range for each target gene. Each

reaction consisted of 10 L SYBR master mix (with ROX), 1 L of each 0.5-0.75 M

forward and reverse primer, 6 L nuclease-free dH2O, and 2 L diluted cDNA.

Standardization of genes was performed with technical duplicates. Amplification specificity (generation of a single product) was obtained by heating the samples from 50 55C to 95C following each amplification run. All successfully optimized genes were

included for relative expression profiling using 10 L SYBR master mix (with ROX), 1

L of each 0.75 M forward and reverse primer, 6 L nuclease-free dH2O, and 2 L

cDNA diluted 1:5 or 1:25 (depending on target gene). PCR parameters were as follows: 2

minutes at 50C for Uracil-DNA Glycosylase (UDG) activation, 2 minutes initial

denaturation at 95C for initial denaturation, and 40 cycles of 15 seconds denaturation at

95C, 15 seconds anneal at 58C, and 1 minute extension at 72C. All samples were amplified with technical triplicates and each gene set had NT (no template) controls performed on the same plate. NRT (no reverse transcriptase) controls were performed on

RNA samples using PCR amplification with parameters and primer concentrations listed above.

Gene Expression Analysis

Raw Ct data for each target gene were compiled and normalized to two of the

chosen endogenous controls (eRF3 and rps3) separately to calculate Ct values for each

sample replicate. Although analyses of each target gene were performed with each endogenous control, the analyses discussed are based on plastid-encoded target genes

normalized to the plastid-encoded reference gene (i.e., rps3), and the single nuclear-

encoded target gene was normalized to the nuclear-encoded reference gene (i.e., eRF3).

The data for each target gene were tested for normality using a Shapiro-Wilks test in R.

Relative expression of each target gene was calculated as the fold change between each

treatment using the Ct method on a log2 scale (Schmittgen and Livak 2008). 51 Differences in expression between each pair of treatments were assessed for statistical significance using a two-way Analysis of Variance (ANOVA) with a post hoc Tukey

HSD.

Results

Plastid Genome

A total of 186,825 paired-end reads were assembled with an average sequencing

depth of 146 reads to generate the  189,825 bp Lympha mucosa plastid genome (Fig. 3-

2). Approximately 24 short intergenic regions will require Sanger sequencing to complete

the genome, but no predicted genes were missing based on comparison to other

Nemaliophycidae plastid genomes (Fig. 3-3). The genome had 243 genes and was AT-

rich (GC content 28.4%). Overall, 201 protein-coding genes were annotated and included

eight open-reading frames (ORFs) and 23 members in the ycf gene family, which largely

encode hypothetical proteins or transcription factors (Fig. 3-2, Minoda et al. 2005). A

whole-genome alignment of the L. mucosa plastid genome and the only other sequenced

freshwater members of the Nemaliophycidae (Appendix D) resulted in seven local co-

linearized blocks (LCBs), with the exception of Balbiania investiens (only included in six

LCBs)(Fig. 3-3b). These freshwater red algal plastid genomes were highly conserved and

only differed slightly in the rearrangement in genomic architecture with L. mucosa and B.

investiens having LCB 6 before 7 (yellow and green), and the other two taxa (Kumanoa

americana and Thorea hispida) showing an inverse arrangement of the two LCBs (Fig. 3-

3b). Likewise, L. mucosa and B. investiens had two inverted copies of the ribosomal

operon, while the other two taxa only had one copy. In comparison to the other 52 freshwater taxa, L. mucosa shared 181 protein-coding genes, including one ORF (Fig. 3-

3a). Three protein-coding genes were unique to L. mucosa, ycf37 and two ORFs (ORF20,

ORF102), and no genes were shared exclusively by the two members of the

Batrachospermales (L. mucosa and K. americana) (Fig. 3-3a). The pbsA (heme oxygenase) and dfr (drug sensory protein) genes were annotated in L. mucosa and B. investiens, but were not detected in K. americana and T. hispida (Fig. 3-3). Like the marine members of the Nemaliophycidae, L. mucosa, K. americana and T. hispida have a complete compliment of the chl gene family, its associated group II intron, and petP, but these features were not present in B. investiens (Fig. 3-2, 3-3a). Three genes in the ycf family (ycf34, ycf35, and ycf46) that are found in marine representatives of the

Nemaliophycidae (Costa et al. 2016, Lee et al. 2016a) were not annotated for the L. mucosa plastid genome, or for the other freshwater taxa.

A double-copy ribosomal operon in the L. mucosa plastid genome was present as an inverted repeat (IR) region consisting of the 5S, 16S, and 23S rRNAs, and two tRNAs.

The first IR is transcribed on the plus strand and occurs between psaM and rps6; the second IR is transcribed on the minus strand and occurs between chlL and ycf27 (Fig. 3-

2). Independent remapping of the intergenic space next to each IR and their respective gene neighbors resolved their placement in the full L. mucosa assembly. The IR arrangement and overall genomic architecture is most like the B. investiens (Balbianiales) plastid genome and most unlike K. americana even though L. mucosa is in the same order Batrachospermales (Fig. 3-3, Appendix D). In addition to the tRNAs in the

ribosomal operons, 29 other tRNAs encoding all 20 amino acids were annotated

throughout the genome (not shown). Of these, there were multiple copies for alanine, 53 arginine, glycine, isoleucine, leucine, methionine, serine, threonine, and valine. Finally,

two highly conserved group II introns in the Nemaliophycidae were detected in a tRNA

for Methionine and in the chlB gene with each being 1,898 bp and 1,949 bp in length,

respectively (Fig. 3-2).

Gene Expression Analyses

Transcription for six of the seven target genes was successfully quantified using

primers designed from the L. mucosa plastid sequence data. The gene apcA (encoding

allophycocyanin pigment binding protein) was discarded from the analyses due to a

failure to obtain ~100% PCR efficiency during optimization of the PCR conditions.

Optimization of all other target genes had PCR efficiencies of 90-120% and R2 of 0.989-

0.999 (Appendix E). Melt curves showed amplification of a single product for each target. The two analyses for each target gene using both nuclear- and plastid-encoded reference genes showed statistically similar relative expression in each treatment comparison, except in two instances, and all treatments were normally distributed with the exception of the plastid-encoded cpeA normalized to the nuclear-encoded eRF3 (p <

0.02). For plastid target genes, only log2-fold changes calculated from data normalized to

rps3 are shown (Figs. 3-4 to 3-6, 3-8), and the data shown for the only nuclear gene

HV60 were normalized to the nuclear-encoded eRF3 (Fig. 3-7).

The target genes encoding subunits of PS I (psaA and petF) were differentially

expressed at low and high light. For psaA, relative expression in high-light treatments

was significantly lower (p < 0.001) in sun- and shade-acclimated thalli (Fig. 3-4). No

significant differences were detected when examining acclimation type alone; however, 54 the two acclimation types had a significant differential response to the two light

treatments (p < 0.05). There was significantly greater differential regulation of psaA in

shade-acclimated (50-fold downregulated, p < 0.0001) than sun-acclimated thalli (8-fold

downregulated; p < 0.01)(Fig. 3-4). The gene encoding PetF ferredoxin (petF) was

differentially expressed in high and low light treatments, but not between sun- and shade- acclimated thalli. In both types of thalli, petF was significantly downregulated (2-fold, p

< 0.05) at high light (Fig. 3-5).

No significant differences were observed in the regulation of psbA (subunit of PS

II), but there were biologically relevant differences. At high light, there was a 3-fold downregulation in sun-acclimated versus shade-acclimated thalli (Fig. 3-6). Contrary to the greatly down-regulated psaA at high light, psbA expression may be upregulated in shade-acclimated thalli exposed to high light (Fig. 3-6). However, other treatment comparisons show little to no change (< 2-fold change) in psbA transcription. For the nuclear-encoded HV60 normalized to eRF3, there was a 3-fold significant downregulation in sun-acclimated thalli at high light, but this was not observed in shade- acclimated thalli (Fig. 3-7). HV60 was downregulated in shade-acclimated thalli, but was only represented as a single fold-change with substantial variation around the mean. The amount of variation observed was caused by several outliers in the HV60 data collected

from shade-acclimated individuals in both light treatments (x̅ = 0.78, σx̅ = 0.30) (Fig. 3-

7).

The genes encoding phycobilisome proteins, cpcA and cpeA, showed an inverse in

expression between light treatments. At high light, cpcA (phycocyanin) was upregulated

(2-fold) and cpeA (phycoerythrin) was downregulated (2-4-fold), but only fold changes in 55 cpeA were significantly differentiated (p < 0.05; Fig. 3-8). Although there were no

significant differences between acclimation types alone, there was an interesting

difference based on light treatment. At low light, sun-acclimated thalli downregulated the

expression of cpcA and cpeA relative to shade-acclimated thalli, but these genes were

differentially regulated in the same acclimation-type at high light. At high light in sun-

acclimated thalli, cpcA was downregulated and cpeA was upregulated (Fig. 3-8).

Discussion

Freshwater Nemaliophycidae Plastid Genomes

Red algal and glaucophyte plastids are unique in that light harvesting is accomplished with only one chlorophyll type (chl. a) and phycobilisomes serve as light- harvesting antenna for Photosystem II (PS II)(Gantt 1990, Busch et al. 2010). The ultrastructure and development of these plastids is more like cyanobacteria than to other photosynthetic eukaryotes. Moreover, the genomes of red algal plastids are among the largest sequenced and are evolutionarily stable, indicating they are most likely the closest extant representatives of the ancestral cyanobacterium that gave rise to the organelle

(Janouškovec et al. 2013, Muñoz-Gómez et al. 2017). In this study, sequencing and assembly of the Lympha mucosa plastid genome was a useful tool to design highly specific primers for targeted light-harvesting genes. These primers were used for quantification of relative expression in L. mucosa photosynthetic genes using RT-qPCR.

However, these data also contribute to the increasing dataset of red algal plastid genomes, which is critical for comparative studies of plastid evolution and its implications for red algal physiology and ecology. 56 In terms of genome size for sequenced plastids in the Nemaliophycidae, L.

mucosa is only second to the marine alga palmata, and it is the largest of

sequenced freshwater taxa (Lee et al. 2016, Evans et al. unpublished, this study). Overall,

freshwater red algal plastid genomes are highly conserved, with a > 94% similar gene

content. Although most taxa (excluding T. hispida) contain unique protein-coding genes, their functions are unknown and further study is required to understand their role in the

plastid. None of these genes were absent from marine species in the subclass.

The one unique gene in L. mucosa relative to other freshwater red algal plastids,

ycf37 has an unknown function and is lost in different taxa across the Florideophyceae

(e.g., Salomaki et al. 2015, Costa et al. 2016, Lee et al. 2016a, b, Verbruggen and Costa,

2016). The gene ycf37 is retained in two marine orders (, Costa et al. 2016;

Palmariales, Lee et al. 2016a) of the Nemaliophycidae, but the amino acid translation is

highly divergent (27-18% similarity) among these taxa, including L. mucosa. Therefore,

ycf37 may represent a pseudogene with no putative function. A transcriptional

investigation would be required to test this hypothesis.

Highly conserved red algal plastid genomes suggest that differences in

photoadaptive strategies between genera, and even species, may be regulated at the

transcriptional level, or above. Two of the four known transcription factors encoded in

other red algal plastids (ycf27, 29, Minoda et al. 2005) are encoded in L. mucosa and all

other sequenced orders of Nemaliophycidae (Costa et al. 2016, Lee et al. 2016a).

Complete protein synthesis within the plastid makes these transcription factors ideal for

activating rapid transcriptional responses of plastid-encoded genes, and this may

contribute to the greater degree of transcriptional regulation in red algal plastids relative 57 to green algal and land plant chloroplasts (Minoda et al. 2005). The results from the

relative expression study presented here, which were made possible by utilizing the

plastid genome as a tool, show an active role in transcriptional regulation for several light-harvesting genes after exposure to low and high light.

Differentially Expressed Genes in Light Treatments

High light prompted substantial differentiation in the relative expression of psaA for both sun- and shade-acclimated thalli, but the 50-fold downregulation in shade- acclimated thalli was the largest response observed in this study. Shade-acclimated thalli were collected from a site in the stream receiving 76 mol photons m-2 s-1, and those

thalli exposed to lower light (< 20 mol photons m-2 s-1) for three days had psaA

upregulated. Differences in the regulation of psaA transcription indicates there may be

adjustments to Photosystem I (PS I), and an upregulation of psaA in shade-acclimated

thalli at low light was likely to further increase the number of PS I units. Alternatively,

the shade-acclimated individuals exposed at a higher light intensity may have had many

PSI units embedded on the thylakoid surface. Upon immediate introduction to notably

higher light regimes, an excess number of PS I units capturing light can result in reactive

oxygen species accumulation and proteins involved in photoprotection may be

overwhelmed (Vass, 2012). Therefore, having too many PS I units at high light can be

damaging to the cell. Significant downregulation in psaA at high light may be supported by decreases in chl. a content of cultured freshwater red algae in similar conditions

(Bautista and Necchi, 2007), as chl. a in red algae is predominantly located in PS I

(Gantt, 1990). Likewise, psaA transcripts produced from sun-acclimated thalli were 58 upregulated 7-fold at low light, indicating a need for the sun-acclimated thalli to increase

the number of PS I units per cell through changes occurring at the transcriptional level.

Interestingly, PS I in red algae differs from other photosynthetic groups and has four

protein subunits involved with oxygen evolution, but the overall role of these subunits is

unknown (Grouneva et al. 2013). Modifications based on differential transcription in

genes making up these subunits may provide insights for the evolution of red algal PS I

adaptations to a dynamic range of light.

The PS II apoprotein gene psbA was not significantly different among any of the

treatments, but the regulation of transcripts at different light intensities was biologically relevant. psbA appears to be upregulated in sun-acclimated algae at low light, but it is

also upregulated in shade-acclimated algae at high light. This result differs from the

differential expression of psaA in shade-acclimated thalli at high light, in which psaA was

greatly downregulated. PS II can become inactivated by a relatively short burst of excess

light, and similar pressures exist between the two photosystems for acclimating to drastic

seasonal fluxes in light (Adir 2005, Necchi 2005, Grouneva et al. 2013). In red algae, the

structure of PS II differs from green algae and plants because phycobilisomes form the

main light-harvesting antenna (Gantt 1990). Photosystem II units in green plastids can

only absorb shorter wavelengths of light, whereas the structure of red algal

phycobilisomes allows for shorter (phycocyanin) and longer (phycoerythrin) wavelength

absorption (Gantt 1990). These differences may influence differential strategies between

organisms bearing green and red plastids, such that changes in the ratio of phycocyanin

and phycoerythrin may provide red algal PS II with a unique adaptation to changes in

light quantity and quality. Furthermore, the minor differences in transcription observed in 59 psbA relative to psaA suggest that PS I must undergo greater modifications based on

changes in light intensity than PS II and that any adaptations related to photoacclimation

in PS II are not controlled at the transcriptional level.

Previously, ecological studies have suggested changes in the ratio of

phycoerythrin to phycocyanin as a mechanism for photoacclimation to highly variable

light intensities, which would likely impact PS II (Kaczmarcyk and Sheath 1991).

Although the data from the current study do not serve as undisputable evidence for

changes in the ratio of pigment-binding proteins, there were interesting patterns in the

differential expression of cpcA and cpeA at high light. In high light, cpcA was upregulated 2-fold and cpeA was downregulated 4-fold. This difference in regulation indicates a change in the ratio of the main pigment-binding proteins that form light-

harvesting rods of the phycobilisome. Unfortunately, only cpeA was significantly

differentiated between light treatments, but the presented correlation between the two

genes provides preliminary evidence for adaptive strategies produced by changes in the

phycobilisome, which could also provide a mechanism for adaptation of PS II to variable

light conditions. These data corroborate a documented decrease in total pigment content

in other studies (Ritz et al. 2000, Aigner et al. 2017), and in the observed disappearance

of phycoerythrin hexamers in the light-harvesting rods of the phycobilisome in Rhodella

violacea during exposure to higher irradiances (Ritz et al. 2000). Increased replication for

expression studies and incorporation of techniques for photopigment isolation may

increase our understanding of phycobilisome modifications that allow freshwater red

algal species to occupy highly variable light environments. 60 The petF data supported significant downregulation in sun- and shade-acclimated thalli exposed to high light. The PetF ferredoxin is involved with photophosphorylation in PS I and serves a role in the photoreduction of NADP+ (Richard et al. 2000, Jacobs et al. 2008), suggesting it has a close functional relationship with PSI and consequently their transcriptional regulation may be coupled. Ferredoxins in the green alga

Chlamydomonas reinhardtii also play an important role as electron acceptors during anaerobic metabolism, with transcription likely anaerobically induced (Happe and Naber

1993, Jacobs et al. 2008). A dual function of petF in red algal plastids may also be

present with potential evidence in the highly reduced plastid of a red algal parasite. The

plastid genome of the parasitic Choreocolax polysiphoniae has lost all genes associated

with photosynthesis except petF (Salomaki et al. 2015). The retention of petF in all red

plastid genomes may indicate other roles of ferredoxin in the organelle. Importantly,

regardless of its potential multiplicative role in the plastid, the current study shows petF

transcription is significantly affected by light. Investigation of petF transcriptional

regulation in other environmental conditions, such as during metabolic stress, may

provide insights for PetF ferredoxin function and regulation in the plastid.

It is important to have an appropriate endogenous control for studies of relative gene expression, and potential differences in transcriptional activity among different genomic compartments warrants the use of a reference gene encoded in the same compartment for data normalization (e.g., Cortleven et al. 2009). In this study, the one nuclear-encoded gene HV60, which encodes the RedCAP apoprotein in the LHC-like superfamily (Engelken et al. 2010), was normalized to the nuclear-encoded eRF3.

RedCAP is localized near the surface of PS I in centric and pennate diatoms (Grouneva et 61 al. 2011, Ikeda et al. 2013). However, other studies show RedCAP is only peripherally

localized to PSI, and it may transfer excitation energy to PSII as well (Gundermann et al.

2013, Sturm et al. 2013). There were only significant differences in HV60 transcript abundance when comparing sun-acclimated L. mucosa thalli exposed to low and high light (2.7-fold) and not in shade-acclimated thalli. HV60 was downregulated at high light, which was the same response observed for expression of genes encoding subunits of the photosystems (i.e., psaA and psbA) in the same treatments. These results corroborate a previous study demonstrating downregulation in RedCAP protein abundance at higher light in diatoms (Sturm et al. 2013). Some LHC proteins are specifically involved with photoprotection of the photosystem units and are believed to be among the first proteins of this function in photosynthetic eukaryotes (Richard et al. 2000), having derived from cyanobacterial one-helix proteins (Sturm et al. 2013). Moreover, the original function of the LHC family is theorized to be photoprotective, with the evolution of light-harvesting roles occurring later (Montane and Kloppstech, 2010). The biological significance of concerted regulation of HV60 and both photosystem genes at high light suggests a potential role of RedCAP in modifying light-harvesting units to adapt to changing light conditions. However, there appears to be a difference in RedCAP activity based on the site the thalli originated from, such as open or shaded sites, but the variation in the data from shade-acclimated thalli limits inference from these results. Moreover, measuring the relative gene expression of HV60 in samples culled at different time points after exposure to light treatments should be used to test for any short-term transcriptional changes that may induce photoadaptive or protective responses to varying light, and in understanding differential transcription in thalli growing in open and shaded sites. 62 Ecophysiological studies of several freshwater red algal species have identified distinct photoacclimation strategies utilized by various species. Photosynthetic responses based on light response curves ranging from 20 to 427 mol photons m-2 s-1 have identified at least two strategy types for freshwater red algal species that are correlated with gross morphology; species or life histories with a tuft morphology became light saturated more quickly and had lower photosynthetic efficiency than the mucilaginous, filamentous morphology (Necchi and Zucchi, 2001; Drerup et al. 2015). Lympha mucosa is filamentous and has a high Pmax and photosynthetic efficiency (as Batrachospermum sp., Drerup et al. 2015), and its distribution and abundance in different light environments within the same stream indicates it is highly successful in a wide range of light intensities. Therefore, L. mucosa is not only adapted for shaded environments, it may be adapted to light environments. These measurable differences in photoacclimation may play an important role in shaping the biogeographic and dispersal patterns of various freshwater red algal species, with some species able to more successfully exploit a larger range of habitats with more variable light regimes.

This study provides evidence for transcriptional regulation of light-harvesting genes in a freshwater red alga, but there remains the possibility for post-transcriptional modifications as a mechanism for photoacclimation, which was not tested in this study.

This type of regulation has been shown in the green alga Chlamydomonas reinhardtii, in which petF transcription was uncoupled from protein synthesis/degradation, and differences in transcription were not represented by changes at the protein level (Richard et al. 2000). Another limitation is in the differences of light quality the organism may be exposed to in the field. For example, thalli were exposed to <20 mol photons m-2 s-1, 63 which was close to field measurements for the shaded collection site (76 mol photons m-2 s-1), but the spectral quality of light reaching these individuals may differ.

Kaczmarczyk and Sheath (1991) showed differences of phycobiliprotein content in

Sheathia boryana (as Batrachospermum boryanum) after exposure to increased levels of green light. Combining other ecological studies involving genetic and proteomic data will further elucidate regulatory mechanisms for acclimating to light in this early-diverging group of photosynthetic eukaryotes.

Implications of this Research

Genetic and biochemical studies of photosynthetic processes in many algal lineages have been limited with the exception of green algae and diatoms (e.g., Koziol et al. 2007, Neilson and Durnford 2010, Sturm et al. 2013, Duanmu et al. 2017). For the unicellular red alga Rhodella violacea, Ritz et al. (2000) observed differences in the total number of thylakoids and total pigment content, as well changes in the phycobilisome through alterations in rod composition (ratio of phycocyanin to phycoerythrin) after exposure to higher irradiances. To date, these changes have only been observed in this taxon so it is unknown if other red algae, especially multicellular taxa like Lympha mucosa, also utilize this strategy. It is important to note the recent discovery of highly differentiated plastid genomes in the earlier-diverging subphylum of Rhodophyta

(Proteorhodophytina, includes R. violacea) compared to more recently radiated taxa

(Eurhodophytina, including L. mucosa) (Muñoz-Gómez et al. 2017). In addition, R. violacea is related to the taxon with the largest plastid genome (1.13 Mbp), Corynoplastis japonica (Muñoz-Gómez et al. 2017). These genomes are intron-rich and contain other 64 genetic elements (i.e., transposable elements), making them highly divergent from other mesophilic red plastid genomes, like L. mucosa.

The ecophysiological evidence for photoacclimation in freshwater red algae includes adjustments in light-harvesting structures and total pigment content, but there have been no genetic, biochemical, or physiological data available to determine underlying cellular mechanisms for this strategy. In this study, the changes in the relative transcript abundance in psaA, petF, and cpeA suggest some level of transcriptional regulation of photoacclimation in freshwater red algae. However, there are hundreds of genes involved with light harvesting that are encoded in the plastid and nuclear genomes of photosynthetic eukaryotes, including the Rhodophyta (Engelken et al. 2010; Grouneva et al. 2013). In the nucleus, many of these genes are part of the LHC-like superfamily and have functions in light-capture and photoprotection (Sturm et al. 2013). Originally, a total

RNA-seq study encompassing all mRNA transcripts produced in the cells of a freshwater red alga was proposed for this study, which would have permitted the discovery of all transcripts encoding light-harvesting apoproteins differentially expressed in different light treatments and thallus acclimation-types. Lympha mucosa proved difficult for RNA isolation, as it was later determined to contain a substantial concentration of polysaccharides within its cell wall. A series of modifications to a commercial RNA extraction kit (NucleoSpin®) adapted from protocols used in Dos Reis Falcāo et al.

(2008), for agar-producing red algae, allowed for removal of excess polysaccharide content and isolation of purified RNA. Although absorbance values (260/280) were well within the acceptable range for high-quality RNA (1.9-2.0), further analysis of the samples showed there was enough degradation in the extracted RNA (RIN < 8) that a 65 RNA-seq study could not be reliably performed. Therefore, RT-qPCR was chosen given

the RNA quality, but a plastid genome became necessary to target gene sequences. In the

future, an RNA-seq study of L. mucosa could provide a large dataset to be mined for all

potential genes involved with photoacclimation. In addition, the size and depth of such a

dataset would contain data that are useful for other questions regarding photosynthesis in

freshwater red algae, such as the photoprotective functions of the large LHC family.

References

Adir, N. 2005. Elucidation of the molecular structures of components of the

phycobilisome: reconstructing a giant. Photosynth. Res. 85:15-32.

Aigner, S., Holzinger, A., Karsten, U. & Kranner, I. 2017. The freshwater red alga

Batrachospermum turfosum (Florideophyceae) can acclimate to a wide range of

light and temperature conditions. Eur. J. Phycol. 52:1-12.

Barbrook, A.C., Howe, C.J., Kurniawan, D.P. & Tarr, S.J. 2010. Organization and

expression of organellar genomes. Phil. Trans. R. Soc. B 365:785-797.

Barkan, A. 2011. Expression of plastid genes: organelle-specific elaborations on a

prokaryotic scaffold. Plant Phys. 155:1520-1532.

Busch, A., Nield, J. & Hippler. 2010. The composition and structure of photosystem I-

associated antenna from Cyanidioschyzon merolae. Plant J. 62:886-897.

Bustin, S.A. 2000. Absolute quantification of mRNA using real-time reverse transcription

polymerase chain reaction assays. J. Mol. Endocrinol. 25:169-193.

Chan, P.P. & Lowe, T.M. 2009. GtRNAdb: A database of transfer RNA genes detected in

genomic sequence. Nucl. Acids Res. 37(Database issue):D93-D97. 66 Cortleven, A., Remans, T., Brenner, W.G. & Valcke, R. 2009. Selection of plastid- and

nuclear-encoded reference genes to study the effect of altered endogenous

cytokinin content on photosynthesis genes in Nicotiana tabacum. Photosynth.

Res. 102:21-29.

Costa, J.F., Lin, S.M., Macaya, E.C., Fernández-Garcia, C. & Verbruggen, H. 2016.

Chloroplast genomes as a tool to resolve red algal phylogenies: a case study in the

Nemaliales. BMC Evol. Biol. 16:205.

Darling, A.E., Mau, B. & Perna, N.T. 2010. progressiveMauve: multiple genome

alignment with gene gain, loss and rearrangement. PLoS one 5:e11147.

Dos Reis Falcão, V., Pedroso Tonon, A., Cabral Oliveira, M. & Colepicolo, P. 2008.

RNA isolation method for polysaccharide rich algae: agar producing Gracilaria

tenuistipitata (Rhodophyta). J. Appl. Phycol. 20:9-12.

Drerup, S.A. & Vis, M.L. 2014. Varied phenologies of Batrachospermum gelatinosum

gametophytes (Batrachospermales, Rhodophyta) in two low-order streams. Fottea

14:121-127.

Drerup, S.A., Gonzalez, D.A. & Vis, M.L. 2015. Photosynthetic characteristics of some

common temperate freshwater red algal taxa (Rhodophyta). Phycologia 54:609-

616.

Duanmu, D., Rockwell, N.C. & Lagarias, J.C. 2017. Algal light sensing and

photoacclimation in aquatic environments. Plant Cell Environ. Accepted, DOI

10.1111/pce.12943 67 Engelken, J., Brinkmann, H. & Adamska, I. 2010. Taxonomic distribution of the

extended LHC (light-harvesting complex) antenna protein superfamily. BMC

Evol. Biol. 10:233.

Entwisle, T.J., Vis, M.L., Chiasson, W.B., Necchi, O. & Sherwood, A.R. 2009.

Systematics of the Batrachospermales (Rhodophyta) – a synthesis. J. Phycol.

45:704-715.

Finn, R.D., Coggill, P., Eberhardt, R.Y., Eddy, S.R., Mistry, J., Mitchell, A.L., Potter,

S.C., Punta, M., Qureshi, M., Sangrador-Vegas, A., Salazar, G.A., Tate, J. &

Bateman, A. 2016. The Pfam protein families database: towards a more

sustainable future. Nucl. Acid. Res. 44(Database issue):D279-D285.

Gantt, E. 1990. Pigmentation and photoacclimation. In Biology of Red Algae [Eds] Cole,

K.M. & Sheath, R.G. Cambridge University Press, Cambridge, UK. pp203-219.

Grouneva, I., Rokka, A. & Aro, E.M. 2011. The thylakoid membrane proteome of two

marine diatoms outlines both diatom-specific and species-specific features of the

photosynthetic machinery. J. Proteome Res. 10:5338-5353.

Grouneva, I., Gollan, P.J., Kangasjärvi, S., Suorsa, M., Tikkamen, M. & Aro, E.M. 2013.

Phylogenetic viewpoints on regulation of light harvesting and electron transport in

eukaryotic photosynthetic organisms. Planta 237:399-412.

Gundermann, K., Schmidt, M., Weisheit, W., Mittag, M. & Büchel, C. 2013.

Identification of several sub-populations in the pool of light harvesting proteins in

the pennate diatom Phaeodactylum tricornutum. BBA Bioenergetics 1827:303-

310. 68 Hughey, J.R., Hommersand, M.H., Gabrielson, P.W., Miller, K.A. & Fuller, T. 2017.

Analysis of the complete plastomes of three species of Membranoptera

(Ceramiales, Rhodophyta) from Pacific North America. J. Phycol. 53:32-43.

Hunsperger, H.M., Randhawa, T. & Cattolico, R.A. 2015. Extensive horizontal gene

transfer, duplication, and loss of chlorophyll synthesis genes in the algae. BMC

Evol. Biol. 15:16.

Jacobs, J., Pudollek, S., Hemschemeier, A. & Happe, T. 2008. A novel, anaerobically

induced ferredoxin in Chlamydomonas reinhardtii. FEBS Letters 583:325-329.

Janouškovec, J., Liu, S.L., Martone, P.T., Carrè, W., Leblanc C., Collèn, J. & Keeling,

P.J. 2013. Evolution of red algal plastid genomes: ancient architectures, introns,

horizontal gene transfer, and taxonomic utility of plastid markers. PLoS one

8:e59001.

Kaczmarczyk, D. & Sheath, R.G. 1991. The effect of light regime on the photosynthetic

apparatus of the freshwater red alga Batrachospermum boryanum. Cryptogam.

Algol. 12:249-263.

Kearse, M., Moir, R., Wilson, A., Stones-Havas, S., Cheung, M., Sturrock, S., Buxton, S.,

Cooper, A., Markowitz, S., Duran, C., Thierer, T., Ashton, B., Mentjies, P. &

Drummond, A. 2012. Geneious Basic: an integrated and extendable desktop

software platform for the organization and analysis of sequence data.

Bioinformatics 28:1647-1649.

Koziol, A.G., Borza, T., Ishida, K., Keeling, P., Lee, R.W. & Durnford, D.G. 2007.

Tracing the evolution of the light-harvesting antennae in chlorophyll a/b-

containing organisms. Plant Phys. 143:1802-1816. 69 Kowalczyk, N., Rousvoal, S., Hervé, C., Boyen, C. & Collén, J. 2014. RT-qPCR

normalization genes in the red alga Chondrus crispus. PLoS one 9:e86574.

Ikeda, Y., Yamagishi, A., Komura, M., Suzuki, T., Dohmae, N., Shibata, Y., Itoh, S.,

Koike, H. & Satoh, K. 2013. Two types of fucoxanthin-chlorophyll-binding

proteins 1 tightly bound to the photosystem I core complex in marine centric

diatoms. BBA Bioenergetics 1827:529-539.

Lagesen, K., Hallin, P., Rødland, E.A., Stærfeldt, H.H., Rognes, T. & Ussery, D.W.

2007. RNAmmer: consistent and rapid annotation of ribosomal RNA genes. Nucl.

Acid. Res. 35:3100-3108.

Lam, D.W., Entwisle, T.J., Eloranta, P., Kwandrans, J. & Vis, M.L. 2012.

Circumscription of species in the genus Sirodotia (Batrachospermales,

Rhodophyta) based on molecular and morphological data. Eur. J. Phycol. 47:42-

50.

Langmead, B. & Salzberg, S.L. 2012. Fast gapped-read alignment with Bowtie 2. Nature

Methods 9:357-359.

Lee, J., Cho, C.H., Park, S.I., Choi, J.W., Song, H.S., West, J.A., Bhattacharya, D. &

Yoon, H.S. 2016a. Parallel evolution of highly conserved plastid genome

architecture in red seaweeds and seed plants. BMC Biol. 24:75.

Lee, J., Kim, K.M., Yang, E.C., Miller, K.A., Boo, S.M., Bhattacharya D. & Yoon, H.S.

2016b. Reconstructing the complex evolutionary history of mobile plasmids in red

algal genomes. Sci. Rep. 6:23744.

Lepetit, B. & Dietzel, L. 2015. Light signalling in photosynthetic eukaryotes with ‘green’

and ‘red’ chloroplasts. Environ. Exper. Bot. 114:30-47. 70 Leukart, P. & Hanelt, D. 1995. Light requirements for photosynthesis and growth in

several macroalgae from a small soft-water stream in the Spessart Mountains,

Germany. Phycologia 34:528-532.

Liu, L.N., Aartsma, T.J., Thomas, J.C., Lamers, G.E., Zhou, B.C. & Zhang, Y.Z. 2008.

Watching the native supramolecular architecture of photosynthetic membrane in

red algae: topography of phycobilisomes and their crowding, diverse distribution

patterns. J. Biol. Chem. 283:34946-34953.

McFadden, G.I. 2001. Primary and secondary endosymbiosis and the origin of plastids. J.

Phycol. 37:951-959.

Minoda, A., Nagasawa, K., Hanaoka, M., Horiuchi, M., Takahashi, H. & Tanka, K. 2005.

Microarray profiling of plastid gene expression in a unicellular red alga,

Cyanidioschyzon merolae. Plant Mol. Biol. 59:375-385.

Montane, M. & Kloppstech, K. 2000. The family of light-harvesting-related proteins

(LHCs, ELIPs, HLIPs): was the harvesting of light their primary function? Gene

258:1-8.

Muñoz-Gómez, S.A., Mejía-Franco, F.G., Durnin, K., Colp, M., Grisdale, C.J.,

Archibald, J.M. & Slamovits, C.H. 2017. The new red algal subphylum

Proteorhodophytina comprises the largest and most divergent plastid genomes

known. Curr. Biol. 27:1-8.

Necchi, O. Jr. 2005. Light-related photosynthetic characteristics of freshwater

rhodophytes. Aquat. Bot. 82:193-209. 71 Necchi, O. Jr. & Zucchi, M.R. 2001. Photosynthetic performance of freshwater

Rhodophyta in response to temperature, irradiance, pH and diurnal rhythm.

Phycol. Res. 49:305-318.

Necchi, O. Jr., Branco, C.C.Z. & Branco, L.H.Z. 1999. Distribution of Rhodophyta in

streams from São Paulo State, southeastern Brazil. Arch. Hydrobiol. 147:73-89.

Neilson, J.A. & Durnford, D.G. (2010). Structural and functional diversification of the

light-harvesting complexes in photosynthetic eukaryotes. Photosyn. Res. 106:57-

71.

Odom, O., Shenkenberg, D.L., Garcia, J.A. & Herrin, D.L. 2004. A horizontally acquired

group II intron in the chloroplast psbA gene of a psychrophilic Chlamydomonas:

In vitro self-splicing and genetic evidence for maturase activity. RNA 10:1097-

1107.

Parker, B.C., Samsel, S.G.L. & Prescott, G.W. 1973. Comparison of microhabitats of

macroscopic subalpine stream algae. Am. Midl. Nat. 90:143-153.

Richard, C., Quellet, H. & Guertin, M. 2000. Characterization of the LI818 polypeptide

from the green unicellular alga Chlamydomonas reinhardtii. Plant. Mol. Biol.

42:303-316.

Ritz, M., Thomas, J., Spilar, A. & Etienne A. 2000. Kinetics of photoacclimation in

response to a shift to high light of the red alga Rhodella violacea adapted to low

irradiance. Plant Phys. 123:1415-1426.

Rorbach, J., Bobrowicz, A., Pearce, S., & Minczuk, M. 2014.Polyadenylation in bacteria

and organelles. Methods Mol. Biol. 1125:211-227. 72 Schmitter, J.M., Jacquot, J.P., de Lamotte-Guery, F., Beauvallet, C., Dutka, S., Gadal, P.,

Decottignies, P. 1988. Purification, properties and complete amino acid sequence

of the ferredoxin from a green alga, Chlamydomonas reinhardtii. Eur. J. Biochem.

172: 405-412.

Schmittgen, T.D. & Livak, K.J. 2008. Analyzing real-time PCR data by the comparative

CT method. Nature Protocols 3:1101-08.

Schoreder, A., Mueller, O., Stocker, S., Salowsky, R., Leiber, M., Gassmann, M.,

Lightfoot, S., Menzel, W., Granzow, M & Ragg, T. 2006. The RIN: an RNA

integrity number for assigning integrity values to RNA measurements. BMC Mol.

Biol. 7:3.

Shi, C., Shuo, W., Xia E.H., Jiang, J.J., Zeng, F.C., & Gao, L.Z. 2016. Full transcription

of the chloroplast genome in photosynthetic eukaryotes. Sci. Rep. 6:30135.

Sturm, S., Engelken, J., Gruber, A., Vugrinec, S., Kroth, P.G., Adamska, I. & Lavaud, J.

2013. A novel type of light-harvesting antenna protein of red algal origin in algae

with secondary plastids. BMC Evol. Biol. 13:158.

Talarico, L. & Maranzana, G. 2000. Light and adaptive responses in red macroalgae: an

overview. J. Photochem. Photobiol. B 56:1-11.

Vandesompele, J., De Preter, K., Pattyn, F., Poppe, B., Van Roy, N., De Paepe, A.&

Speleman, F. 2002. Accurate normalisation of real-time quantitative RT-PCR data

by geometric averaging of multiple internal controls. Genome Biol. 3:R0034.1-

R0034.11.

Vass, I. 2012. Molecular mechanisms of photodamage in the photosystem II complex.

Biochimica et Biophysica Acta 1817:209-217. 73 Ye, J., Coulouris, G., Zaretskaya, I., Cutcutache, I., Rozen, S. & Madden, T. 2012.

Primer-BLAST: a tool to design target-specific primers for polymerase chain

reaction. BMC Bioinformatics 13:134.

74 Table 3-1 Site conditions for the Kinniconick Creek, KY. All samples of Lympha mucosa were collected on July 20, 2016 at midday.

Coordinates, Collectors, Water pH Specific Midsection total Edge total

Voucher Temperature Conductance light (mol light (mol

(C) (S cm-1) photon m-2s-1) photon m- N 38.496667, 29 6.6 110 1132 76.21

W 83.257222,

J.R. Evans & M.L. Vis,

BHO A-1418

Table 3-2 Primer sets that were used in RT-qPCR for each target and reference gene in

Lympha mucosa. All primers listed were successfully amplified with L. mucosa gDNA.

Primer sequence, expected Tm (C), and GC content (%) are shown.

Gene Primer Primer Sequence (5’-3’) Expected Tm (C) GC Content (%)

TARGET GENES apcA

apcAF1 ATGACTGCAACGTGTTTGCG 60.32 50.00

apcAR1 GCACGTAAGCCTTCTGCAAC 60.11 55.00 cpcA

cpcAF1 TAACAAGCAGCTCGCAACGA 60.60 50.00

cpcAR1 CTCATCCATGGGTCCAGTTGAA 60.03 50.00

cpeA

cpeAF1 TGGCACAGGACCTTTAGACG 59.68 55.00 75 Table 3-2 continued

Gene Primer Primer Sequence (5’-3’) Expected Tm (C) GC Content (%)

cpeAR1 CTACACCTGCTTGAGCAGACA 60.00 52.38

HV60

LCAPF1 ATGTCATGTTCGGCTGGCTT 60.32 50.00

LCAPR1 AAGTGGATGTGAGCGACGAG 60.11 55.00 petF

petFF1 TTGGATGCAGCGGAAGATCAA 60.34 47.62

petFR1 CAGACTGGTCAACCGAACCTT 60.20 52.38 psaA

psaAF1 GGGCACATTTTGTATGGGCTTT 60.03 45.45

psaAR1 AATGTGCAACTCCGACTGCT 60.25 50.00 psbA

psbAF1 AGTCAAGGGCGCGTAATCAA 60.04 50.00

psbAR1 GGTGCAACTAAAGCAACTGGG 60.00 52.38

REFERENCE GENES eRF3

eRF3F1 TGAAAGTCAGGGCAACGGAA 59.82 50.00

eRF3R1 GGGCACAATGGTAAAGGGGA 59.96 55.00 rps3

rps3F1 GCCAATAGACGTGCTTCTGTG 59.61 52.38

rps3R1 AACAGCTAGACCAGGAATTGTCC 60.31 47.83

76

Figure 3-1 Map of potential target and reference genes for expression analysis of

Lympha mucosa photoacclimation. Primer-binding regions are represented by arrow pairs of the same color, and the predicted sequence product in grey.

77

Figure 3-2 The  189,825 bp Lympha mucosa plastid genome. The genome contains 243 genes with an average GC content of 28.45%. The genome encodes 201 protein-coding genes (function indicated by color), 6 rRNA genes that form two inverted ribosomal operons (red), and 33 tRNA genes encoding anti-codons for all 20 amino acids (not shown). Two highly conserved group II introns (chlB, tRNA-Met) were detected.

78

Figure 3-3 Comparison of the L. mucosa plastid genome to all other sequenced plastid genomes of freshwater Nemaliophycidae (Rhodophyta). (A) The Venn diagram compares protein-coding genes among the four plastid genomes. Lympha mucosa contains the greatest number of protein-coding genes, of which 181 are shared with the other four genomes. (B) Mauve alignment of seven co-linear blocks (distinguished by color) shared among the four plastid genomes. One major inversion is present between green and yellow blocks (LCBs 6 and 7). 79

Figure 3-4 Log2 fold change (mean ± SE) of psaA relative expression normalized to rps3 (using the ΔΔCt method). Bars on the left of the vertical line are downregulated and bars on the right are upregulated. The treatment comparisons are listed on the left side: from top to bottom, sun-acclimated (SA) relative to shade-acclimated (SH) individuals at high light (HL) (white bar), SA relative to SH individuals at low light (LL) (black bar), SA individuals at HL relative to LL (light grey bar), and SH individuals at HL relative to LL

(dark grey bar). Significance within treatment comparisons is denoted by (*) and significantly differences between bars denoted by a letter. psaA is differentially downregulated at HL in SH thalli (p < 0.001) and SA thalli (p < 0.05). 80

Figure 3-5 Log2 fold change (mean ± SE) of petF (PetF ferredoxin) relative expression normalized to rps3 (using the ΔΔCt method).

Bars on the left of the vertical line are downregulated and bars on the right are upregulated. The treatment comparisons are listed on the left side: from top to bottom, sun-acclimated (SA) relative to shade-acclimated (SH) individuals at high light (HL) (white bar), SA relative to SH individuals at low light (LL) (black bar), SA individuals at HL relative to LL (light grey bar), and SH individuals at HL relative to LL (dark grey bar). An (*) denotes significance and petF is equally downregulated in SA and SH thalli at HL (p < 0.05). 81

Figure 3-6 Log2 fold change (mean ± SE) of psbA relative expression normalized to rps3 (using the ΔΔCt method). Bars on the left of the vertical line are downregulated and bars on the right are upregulated. The treatment comparisons are listed on the left side: from top to bottom, sun-acclimated (SA) relative to shade-acclimated (SH) individuals at high light (HL) (white bar), SA relative to SH individuals at low light (LL) (black bar), SA individuals at HL relative to LL (light grey bar), and SH individuals at HL relative to LL

(dark grey bar). No significant differences were found. 82

Figure 3-7 Log2 fold change (mean ± SE) of nuclear-encoded HV60 (RedCAP) relative expression normalized to eRF3 (using the

ΔΔCt method). Bars on the left of the vertical line are downregulated and bars on the right are upregulated. The treatment comparisons

are listed on the left side: from top to bottom, sun-acclimated (SA) relative to shade-acclimated (SH) individuals at high light (HL)

(white bar), SA relative to SH individuals at low light (LL) (black bar), SA individuals at HL relative to LL (light grey bar), and SH

individuals at HL relative to LL (dark grey bar). An (*) denotes significance and HV60 was downregulated at HL in SA thalli (p <

0.05). 83

Figure 3-8 Log2 fold change (mean ± SE) in relative expression of genes encoding the main pigment-binding proteins in the phycobilisome. (A) is cpcA (phycocyanin) and (B) is cpeA (phycoerythrin). Data were normalized to rps3 (using the ΔΔCt method).

For each graph, bars on the left of the vertical line are downregulated and bars on the right are upregulated. The treatment comparisons are listed on the left side: from top to bottom, sun-acclimated (SA) relative to shade-acclimated (SH) individuals at high light (HL)

(white bar), SA relative to SH individuals at low light (LL) (black bar), SA individuals at HL relative to LL (light grey bar), and SH individuals at HL relative to LL (dark grey bar). An (*) denotes significance and cpeA was downregulated at HL in all thalli (p <

0.05). Differential expression in cpcA was not significant, but all thalli showed an opposite response at HL compared to cpcA. 84 CHAPTER 4: CONCLUSIONS

The Rhodophyta contains a diverse assemblage of unicellular and multicellular species that occur in environments worldwide (Woekerling 1990). Of these species, the

~5% that occur in freshwater environments are primarily members of the

Batrachospermales, which contains extensive genetic, ecological, and biogeographic diversity (Sheath, 1984, Vis et al. 1998, Sheath and Vis, 2015, Vis 2016). While systematics of the Batrachospermales is ongoing, with approximately one new taxon described each year (Entwisle et al. 2009), recent advances in molecular studies have helped to resolve the paraphyly of the most species rich batrachospermalean genus,

Batrachospermum (e.g., Salomaki et al. 2014, Entwisle et al. 2016, Chapuis et al. 2017).

Here, the proposal of Lympha mucosa gen. et sp. prov. adds to the already extensive documented diversity and endemism of North American freshwater red algal flora. Future studies of the flora in southeastern United States, and worldwide, will continue to resolve these systematic issues and provide a better picture of the diversity within the

Batrachospermales.

We are now starting to understand more about the complexity and diversity of adaptations in photosynthesis and light harvesting of plastid-bearing eukaryotes through genome-level studies (Busch et al. 2010; Lee et al. 2016a; Lee et al. 2016b). Studying red algal plastids provides data for understanding photosynthetic development in an early- diverging, photosynthetic lineage that has retained a plastid highly similar to cyanobacterial architecture with an accompanying genome of similar representation

(Janouškovec et al. 2013; Muñoz-Gómez et al. 2017). Although the overall genome architecture is highly conserved in red algae, differences in transcriptional regulation in 85 these plastids may contribute to differences in photosynthetic strategies and ecological

characteristics, particularly since this type of modification occurs more extensively in red

than green plastids (Minoda et al. 2005). Specific to freshwater taxa in the Rhodophyta,

adjustments in the photosynthetic apparatus may provide highly successful mechanisms

for adapting/acclimating to the dynamic, seasonal range of light that is experienced,

particularly in temperate regions (Kaczmarczyk and Sheath 1991, Necchi, 2005; Aigner

et al. 2017). In this thesis, a dataset for relative expression of several photosynthetic

genes in Lympha mucosa has offered insights into the underlying mechanisms involved

with these adaptations, and provided support for highly plastic responses in the light-

harvesting structures of freshwater red algae. There is now genetic evidence for the role

of transcriptional modifications for adjustments to light-harvesting structures when

exposed to low or high light. These data, combined with the genomic data that were

produced as a tool, will be fruitful in future comparative genetic and ecological studies.

References

Aigner, S., Holzinger, A., Karsten, U. & Kranner, I. 2017. The freshwater red alga

Batrachospermum turfosum (Florideophyceae) can acclimate to a wide range of

light and temperature conditions. Eur. J. Phycol. 52:1-12.

Busch, A., Nield, J. & Hippler. 2010. The composition and structure of photosystem I-

associated antenna from Cyanidioschyzon merolae. Plant J. 62:886-897.

Chapuis, I.S., Necchi Jr., O., Zuccarello, G.C., Xie, S.L., Aboal, M., Sánchez Castillo,

P.M. & Vis, M.L. 2017. A new genus, Volatus and four new species of 86 Batrachospermum sensu stricto (Batrachospermales, Rhodophyta). Phycologia

56:454-468.

Entwisle, T.J., Vis, M.L., Chiasson, W.B., Necchi, O. & Sherwood, A.R. 2009.

Systematics of the Batrachospermales (Rhodophyta) – a synthesis. J. Phycol.

45:704-715.

Entwisle, T.J., Johnston, E.T., Lam, D.W., Stewart, S.A. & Vis, M.L. 2016. Nocturama

gen. nov., Nothocladus s. lat. and other taxonomic novelties resulting from the

further resolution of paraphyly in Australasian members of Batrachospermum

(Batrachospermales, Rhodophyta). J. Phycol. 52:384-396.

Janouškovec, J., Liu, S.L., Martone, P.T., Carrè, W., Leblanc C., Collèn, J. & Keeling,

P.J. 2013. Evolution of red algal plastid genomes: ancient architectures, introns,

horizontal gene transfer, and taxonomic utility of plastid markers. PLoS one

8:e59001.

Kaczmarczyk, D. & Sheath, R.G. 1991. The effect of light regime on the photosynthetic

apparatus of the freshwater red alga Batrachospermum boryanum. Cryptogam.

Algol. 12:249-263.

Lee, J., Cho, C.H., Park, S.I., Choi, J.W., Song, H.S., West, J.A., Bhattacharya, D. &

Yoon, H.S. 2016a. Parallel evolution of highly conserved plastid genome

architecture in red seaweeds and seed plants. BMC Biol. 24:75.

Lee, J., Kim, K.M., Yang, E.C., Miller, K.A., Boo, S.M., Bhattacharya D. & Yoon, H.S.

2016b. Reconstructing the complex evolutionary history of mobile plasmids in red

algal genomes. Sci. Rep. 6:23744. 87 Minoda, A., Nagasawa, K., Hanaoka, M., Horiuchi, M., Takahashi, H. & Tanka, K. 2005.

Microarray profiling of plastid gene expression in a unicellular red alga,

Cyanidioschyzon merolae. Plant Mol. Biol. 59:375-385.

Muñoz-Gómez, S.A., Mejía-Franco, F.G., Durnin, K., Colp, M., Grisdale, C.J.,

Archibald, J.M. & Slamovits, C.H. 2017. The new red algal subphylum

Proteorhodophytina comprises the largest and most divergent plastid genomes

known. Curr. Biol. 27:1-8.

Necchi, O. Jr. 2005. Light-related photosynthetic characteristics of freshwater

rhodophytes. Aquat. Bot. 82:193-209.

Salomaki, E.D., Kwandrans, J., Eloranta, P. & Vis, M.L. 2014. Molecular and

morphological evidence for Sheathia gen. nov. (Batrachospermales, Rhodophyta)

and three new species. J. Phycol. 50:526-542.

Sheath, R.G. 1984. The biology of freshwater red algae. In Round, F.E. & Chapman, D.J.

[Eds] Progress in Phycological Research, vol. 3. BioPress Ltd, Bristol UK, pp.

89-157.

Sheath, R.G. & Vis, M.L. 2015. Red algae. In Wehr, J.D., Sheath, R.G. & Kociolek, J.P.

[Eds] Freshwater Algae of North America second edition. Elsevier Academic

Press, San Diego. pp. 237-264.

Vis, M.L. 2016. Biogeography of river algae. In Necchi, O. Jr. [Ed] River Algae. Springer

Nature, pp. 219-244.

Vis, M.L., Saunders, G.W., Sheath, R.G., Dunse, K. & Entwisle, T.J. 1998. Phylogeny of

the Batrachospermales (Rhodophyta) inferred from rbcL and 18S ribosomal DNA

gene sequences. J. Phycol. 32:341-350. 88 APPENDIX A: GENBANK ACCESSION NUMBERS FOR COI-5P AND RBCL

SEQUENCE DATA OF BATRACHOSPERMALES AND THOREALES USED FOR

PHYLOGENETIC ANALYSES: CHAPTER 2

Taxon COI-5P rbcL

Batrachospermales

Batrachospermum KU672388 FJ386458

brasiliense

B. cayennense KM055325 KM055245

B. dapsile KM593871 KM593855

B. gelatinosum KM055327 GU810833

B. helminthosum KM055328 AF244112

B. macrosporum KT802766 EU106056

B. naiadis KM593872 KM593857

B. pozoazulense KM593867 KM593863

B. shanxiense KM593869 KM593851

B. turfosum KT802767 KJ825960

Kumanoa ambigua EU636724 AY423390

K. americana JN604910 JN589995

K. montagnei KM055331 AY423396

Lemanea fluviatilis KM055333 KM055243

L. fucina KU672391 KJ825958

Nocturama antipodites KT802754 FJ386456 89 Appendix A continued.

Taxon COI-5P rbcL

Batrachospermales

Nothocladus ater KT802757 KT802842

N. kraftii KT802760 FJ386461

N. wattsii KT802765 KT802866

Paralemanea annulata KM055335 GQ285124

Petrohua bernabei KM055336 AY960690

Psilosiphon scoparum KU672392 AF029155

Sheathia americana JX669695 JX669757

S. exigua JX669616 GU457345

S. heterocortica EU636740 DQ393136

Sirodotia delicatula KU672394 DQ646474

S. huillensis EU636739 AF029157

S. suecica KM055337 JF344718

Tuomeya americana KM055330 KM055244

Volatus carrionii KM593866 KM593861

V. ulterior KM593870 KM593852

Thoreales

Nemalionopsis shawii KM055241 KM005141

Thorea hispida KM055237 KM005138 90 APPENDIX B: EXPERIMENTAL DESIGN FOR LYMPHA MUCOSA SAMPLES

EXPOSED TO LOW OR HIGH LIGHT: CHAPTER 3

Sun- and shade-acclimated thalli were equally divided into two light treatments (low light

= < 20 µmol photons m-2 s-1, high light = 220 µmol photons m-2 s-1) and placed on a

Labline orbital shaker in a Conviron CMP6050 walk-in growth chamber for 72h. 91 APPENDIX C: TOTAL QUANTITY, RNA INTEGRITY NUMBER (RIN), AND

ABSORBANCE RATIO (260/280) FOR EACH LYMPHA MUCOSA RNA SAMPLE:

CHAPTER 3

Treatment High Light (SA) High Light (SH)

Sample 361 39 47 44 51 32 40 43 48 33 49 34

ID

(JREN)

Total 215 300 4.37 70.5 600 300 1550 1700 53.15 14.5 350 1150

RNA (ng)

RIN 2.3 3.2 2.3 2.6 7.2 N/A 3.4 4.5 4.3 2.6 N/A 1

Abs. 2.25 2.01 1.79 2.1 1.95 1.81 1.78 1.94 1.56 1.94 1.79 1.87

(260/280)

1Samples in bold were used in subsequent RT-qPCR.

Treatment Low Light (SA) Low Light (SH)

Sample 351 42 41 28 31 50 45 37 38 27 46 30

ID

(JREN)

Total 550 13.95 700 600 150 65.6 301.5 300 350 1100 197.5 750

RNA (ng)

RIN 1 2.3 6.3 1 2.1 2.4 2.4 N/A N/A 4.9 2.6 1.8

Abs. 1.84 2.23 1.92 2.13 2.01 1.87 1.95 1.97 2.15 2.13 1.94 2.11

(260/280)

1Samples in bold were used in subsequent RT-qPCR. 92 APPENDIX D: TWO-WAY ANALYSIS OF VARIANCE TABLES USING DATA

NORMALIZED TO RPS3: CHAPTER 3

HV60

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 2.1000 2.100003 3.1149 0.1156

Acclimation-type 1 0.8008 0.80083 1.1879 0.3075

Light:Acclimation 1 0.1281 0.12813 0.1901 0.6744

Residuals 8 5.3935 0.67418

cpcA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 1.6875 1.68750 1.3267 0.2827

Acclimation-type 1 1.6280 1.62803 1.2799 0.2907

Light:Acclimation 1 0.0901 0.9013 0.0709 0.7968

Residuals 8 10.1758 1.27198

cpeA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 9.9554 9.9554 7.3422 0.02667

Acclimation-type 1 0.0030 0.0030 0.0022 0.96359

Light:Acclimation 1 0.7750 0.7550 0.5568 0.47689

Residuals 8 10.8473 1.3559

petF

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 7.1456 7.1456 7.0014 0.02944

Acclimation-type 1 0.0096 0.0096 0.0094 0.92499 93 Light:Acclimation 1 0.2352 0.2352 0.2305 0.64404

Residuals 8 8.1648 1.0206 psaA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 52.710 52.710 62.4942 4.757e-05

Acclimation-type 1 1.944 1.944 2.3049 0.16744

Light:Acclimation 1 5.810 5.810 6.8887 0.03043

Residuals 8 6.748 0.843 psbA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 0.1680 0.16803 0.1665 0.6940

Acclimation-type 1 2.7840 2.78403 2.7582 0.1353

Light:Acclimation 1 1.5552 1.55520 1.5408 0.2497

Residuals 8 9.0748 1.00935

94 APPENDIX E: TWO-WAY ANALYSIS OF VARIANCE TABLES USING DATA

NORMALIZED TO ERF3: CHAPTER 3

HV60

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 2.39413 2.39413 7.2436 0.02744

Acclimation-type 1 0.11603 0.11603 0.3511 0.56987

Light:Acclimation 1 0.90750 0.90750 2.7457 0.13611

Residuals 8 2.64413 0.33052

cpcA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 1.4421 1.44213 1.4491 0.2631

Acclimation-type 1 0.5208 0.52083 0.5234 0.4900

Light:Acclimation 1 0.8008 0.80083 0.8047 0.3959

Residuals 8 7.9615 0.99518

cpeA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 10.5844 10.5844 5.5338 0.04651

Acclimation-type 1 0.2494 0.2494 0.1304 0.72737

Light:Acclimation 1 0.0752 0.0752 0.0393 0.84776

Residuals 8 15.3016 1.9127

petF

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 7.6800 7.680 17.1825 0.00323

Acclimation-type 1 0.2080 0.208 0.4654 0.51436

Light:Acclimation 1 0.0120 0.012 0.0269 0.87374

Residuals 8 3.5757 0.447 95 psaA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 54.145 54.145 92.9608 1.114e-05

Acclimation-type 1 0.706 0.706 1.2116 0.3030

Light:Acclimation 1 3.297 3.297 5.6606 0.0446

Residuals 8 4.660 0.582 psbA

Df Sum Sq. Mean Sq. F-Value p-value

Light Treatment 1 0.2581 0.2581 0.2692 0.61790

Acclimation-type 1 1.2416 1.2416 1.2950 0.28807

Light:Acclimation 1 3.3920 3.3920 3.5377 0.09677

Residuals 8 7.6706 0.9588 96 APPENDIX F: LIST OF PLASTID GENOMES FOR TAXA IN THE NEMALIOPHYCIDAE: CHAPTER 3

Order Taxon1 Size (bp) GC Content (%) CDS tRNA rRNA Accession

Batrachospermales Lympha mucosa >189.825 28.4 201 33 6 pending

Acrochaetiales Acrochaetium secundatum2 ≥183541 - ≥196 ≥29 ≥0 SAMEA4393238

Balbianiales Balbiania investiens3 186,778 33.9 195 200 6 pending

Batrachospermales Kumanoa americana4 184,025 29.27 198 31 3 KX284725

Nemaliales Nothogenia fastigiata2 182 457 33 193 30 3 SAMEA4358435

Nemaliales Scinaia undulata2 183 795 35.9 197 31 3 SAMEA4478602

Nemaliales Actinotrichia fragilis2 183 324 29.8 197 31 3 SAMEA4357171

Nemaliales Galaxaura rugosa2 181 215 29.6 197 31 3 SAMEA4357173

Nemaliales Tricleocarpa cylindrica2 ≥150 119 - ≥149 ≥25 ≥1 SAMEA4478608

Nemaliales Dichotomaria marginata2 184 395 28.8 197 31 3 SAMEA4357172

Nemaliales Liagora brachyclada2 182 937 33.7 195 31 3 SAMEA4395348

Nemaliales Liagora harveyana2 182 933 33.9 195 31 3 SAMEA4358432

Nemaliales Izziella formosana2 183 248 35 195 31 3 SAMEA4358392

Nemaliales Neoizziella asiatica2 183 313 33.4 195 31 3 SAMEA4358434

Nemaliales Titanophycus setchellii2 ≥183 356 - 195 ≥31 ≥3 SAMEA4478603 97

Order Taxon1 Size (bp) GC Content (%) CDS tRNA rRNA Accession

Nemaliales Helminthora furcellata2 184 585 32.1 195 31 3 SAMEA4393237

Nemaliales Trichogloeopsis pedicellata2 183 497 31.9 194 31 3 SAMEA4358437

Nemaliales Hommersandiophycus borowitzkae2 184 728 32.2 194 31 3 SAMEA4358391

Nemaliales Dermonema virens2 184 997 34.1 195 31 3 SAMEA4357169

Nemaliales Helminthocladia australis2 185 694 32.8 195 31 3 SAMEA4358292

Nemaliales Yamadaella caenomyce2 182 460 35.9 194 31 3 SAMEA4358436

Nemaliales Liagoropsis maxima2 189 564 32.1 195 31 3 SAMEA4358433

Nemaliales Nemalion sp. 2 182 930 35.5 194 31 3 SAMEA4478604

Palmariales Palmaria palmata4 192,960 33.91 204 33 6 KX284726

Palmariales floridula2 ≥182 494 - ≥196 ≥29 ≥1 SAMEA4478606

Thoreales Thorea hispida4 175,193 28.35 193 30 3 KX284714

1 Taxa in bold are freshwater representatives of the Nemaliophycidae. 2 Costa, J.F., Lin, S.M., Macaya, E.C., Fernández-Garcia, C. & Verbruggen, H. 2016. Chloroplast genomes as a tool to resolve red algal phylogenies: a case study in the Nemaliales. BMC Evol. Biol. 16:205. 3 Evans, J.R., St. Amour, N., Salomaki, E., Verbruggen, H. & Vis, M.L. The organellar genomes of the freshwater epiphyte Balbiania investiens (Balbianiales, Nemaliophycidae). In prep. 4 Lee, J., Cho, C.H., Park, S.I., Choi, J.W., Song, H.S., West, J.A., Bhattacharya, D. & Yoon, H.S. 2016. Parallel evolution of highly conserved plastid genome architecture in red seaweeds and seed plants. BMC Biol. 24:75. 98 APPENDIX G: RT-QPCR OPTIMIZATION DATA: CHAPTER 3

2 Gene (Protein) PCR efficiency R Ct range

(%)

apcA (Allophycocyanin) 143.1 0.999 24.07-29.50

HV60 (RedCAP) 82.7 0.999 23.38-34.14

cpcA (Phycocyanin) 104.7 0.995 25.50-30.11

cpeA (Phycoerythrin) 87.1 0.999 17.32-30.04

petF (PetF ferredoxin) 106.6 0.993 20.97-30.04 psaA (Photosystem I alpha subunit) 80.0 0.999 18.36-32.07 psbA (Photosystem II apoprotein) 91.1 0.999 18.22-30.73

eRF3 (Ethylene-responsive 90.4 0.995 29.98-37.41

transcription factor)

rps3 (Ribosomal Protein S3) 100.7 0.993 26.54-33.67

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