INVESTIGATING PHENOTYPIC EFFECTS OF INTRON DELETION AND SISRNA IDENTIFICATION IN CYANIDIOSCHYZON MEROLAE by Maryam Ghaffarzadeh B.A., Shahid Beheshti University of Medical Science, 2011 M.Sc., Shahid Beheshti University of Medical Science, 2018 THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE IN BIOCHEMISTRY UNIVERSITY OF NORTHERN BRITISH COLUMBIA October 2024 © Maryam Ghaffarzadeh, 2024 Abstract In eukaryotic cells, the production of messenger RNA (mRNA) from precursor mRNA involves the removal of non-coding sequences known as introns. Originally considered “junk DNA” with no function, introns have recently been found to play a number of roles, including helping cells adapt to stress. It has been shown that the Last Eukaryotic Common Ancestor (LECA) had a genome characterized by relatively high intron density compared to modern eukaryotes. Post-LECA evolutionary trends in eukaryotes have largely involved intron loss, however, it appears to be rare for eukaryotes to lose all of their introns. The remaining introns in an intronpoor organism either are selected for their function or will eventually be lost. This thesis focuses on the red alga Cyanidioschyzon merolae (C. merolae), which has a minimalistic splicing system of only 39 introns across 38 genes. Therefore, due to the small number of introns, C. merolae presents an excellent opportunity to study intron evolution and function. My research investigates the role of specific introns in C. merolae by deleting them and observing the effects on growth under normal and stress conditions. The findings reveal that while most introns can be removed without impacting growth, certain strains with deleted introns show altered growth rates, indicating that these introns are integral to optimal cellular function. Particularly under nutrient scarcity, some intron deletions are notably deleterious. One known function of introns is to harbour stable intronderived small RNAs (sisRNAs). I present the first demonstration of sisRNAs in an alga, and investigate their function under normal conditions and heat stress. Strikingly, I find that sisRNAs are polyadenylated under heat stress, suggesting a specific adaptive response. Consistent with this possibility, I observe growth defects in strains deleted for sisRNA-containing introns. This work establishes C. merolae as a valuable system for investigating intron evolution and function, and broadens our understanding of the functional roles of introns and sisRNAs. II Table of Contents Abstract ............................................................................................................................... ii List of Tables ..................................................................................................................... VI List of Figures: ................................................................................................................ VII List of Abbreviations ......................................................................................................... IX Note on gene references in this work .............................................................................. XII Acknowledgments .......................................................................................................... XIV CHAPTER I-Introduction .................................................................................................. 1 1.1 Pre-mRNA splicing ........................................................................................................ 2 1.1.1 Splicing in humans .................................................................................................................. 5 1.2 Intron evolution ............................................................................................................. 8 1.3 Intron function ............................................................................................................ 11 1.3.1 Intronic Cis-elements ............................................................................................................ 12 1.3.2 Intronic non-coding RNAs .................................................................................................... 16 1.4 Thesis scope and significance ...................................................................................... 17 1.4.1 C. merolae .............................................................................................................................. 17 CHAPTER II: Evaluating the Impact of Intron Deletion on C. merolae Growth Under Normal and Nutritional Stress Conditions ............................................. 19 2.1 Introduction................................................................................................................. 20 2.2 Materials and Methods ............................................................................................... 22 2.2.1 Designing integration constructs for homologous recombination ....................................... 22 2.2.2 Intron deletion ....................................................................................................................... 25 2.2.1 Construction of vectors containing two homology arms .................................................................... 27 2.2.4 C. merolae URA transformation ........................................................................................... 30 2.2.4.1 Day of transformation...................................................................................................................... 32 2.2.4.2 Day after transformation ................................................................................................................. 33 2.2.4.3 Testing for homologous recombination (Colony PCR) .................................................................... 33 2.2.4.3.1 Quick genomic DNA isolation ....................................................................................................... 34 2.2.5 Sequencing............................................................................................................................. 36 2.2.6 Southern blotting .................................................................................................................. 38 2.2.6.1 Restriction Enzyme Digest ............................................................................................................... 38 2.2.6.2 Probe Production and Labeling ....................................................................................................... 38 2.2.6.3 Southern Gel and Transfer .............................................................................................................. 40 2.2.6.4 Hybridization ................................................................................................................................... 40 2.2.6.5 Washes.............................................................................................................................................. 41 2.2.6.6 Immunological Detection ................................................................................................................. 41 2.2.7 Growth test at 42 C ............................................................................................................... 43 2.2.8 Fitness Test ............................................................................................................................ 45 III 2.2.8.1 T1 mTFP transformation ................................................................................................................. 45 2.2.8.2 Fitness test set up ............................................................................................................................. 46 2.3 Results and Discussion ................................................................................................ 49 2.3.1 Intron deletion ....................................................................................................................... 49 2.3.1.1 Intron deletion by homologous recombination (HR) ....................................................................... 49 2.3.1.2 Confirmation of DNA construct with two homology arms for intron deletion ............................... 50 2.3.1.3 Confirmation of the full-length transformation amplicon. ............................................................. 53 2.3.1.4 Confirmation of successful transformation for making ∆i of S262, E034, D067, and R289 ........... 55 2.3.2 Southern blotting .................................................................................................................. 60 2.3.3 Growth Rate Analysis at 42 C ............................................................................................... 63 2.3.4 Fitness test ............................................................................................................................. 75 2.3.4.1 T1 and T1::mTFP growth test at 42 C ............................................................................................. 75 2.3.4.2 Fitness test for ∆10i and R289∆i ...................................................................................................... 76 2.4 Conclusion ................................................................................................................... 84 CHAPTER III: Discovery of polyadenylated sisRNAs in Cyanidioschyzon merolae .............................................................................................................................. 85 3.1 Introduction................................................................................................................. 86 3.1.1 sisRNA function..................................................................................................................... 89 3.2 Materials and Methods ............................................................................................... 97 3.2.1 C. merolae culturing .............................................................................................................. 97 3.2.2 RNA extraction...................................................................................................................... 97 3.2.3 Fluorescent Northern Blot .................................................................................................... 98 3.2.3.1 Denaturing Polyacrylamide Northern blotting .............................................................................. 102 3.2.3.1.1 Preparing 6% Urea Denaturing Polyacrylamide Gel .................................................................. 102 3.2.3.1.2 Pre-hybridization and hybridization ........................................................................................... 102 3.2.3.1.4 Washing ........................................................................................................................................ 103 3.2.3.1.5 Detection, Analysis, and Stripping ............................................................................................... 103 3.2.3.2 Fluorescent Polyacrylamide Northern on Poly A+ RNA ................................................................ 103 3.2.3.3 Denaturing agarose Northern blot.................................................................................................. 104 3.2.3.3.1 Preparation of Denaturing Agarose Formaldehyde Gel and running the gel ............................. 104 3.2.3.3.2 Transferring and Detection .......................................................................................................... 104 3.2.4 RT-qPCR ............................................................................................................................. 105 3.2.4.1 Primer Design for RT-qPCR Experiment ....................................................................................... 106 3.2.4.2 cDNA synthesis................................................................................................................................ 109 3.2.4.3 RT-qPCR ......................................................................................................................................... 111 3.2.4.3.1 Determining the annealing temperature ...................................................................................... 111 3.2.4.3.2 Primer Efficiency Determination ................................................................................................. 111 3.2.4.3.3 Target and reference gene expression .......................................................................................... 111 3.2.4.3.4 Reference gene stability ............................................................................................................... 112 3.2.5 Growth test at 57 C ..................................................................................................113 IV 3.3 Results and Discussion ...............................................................................................114 3.3.1 Confirmation of sisRNAs by Northern blotting ................................................................. 114 3.3.2 Denaturing Polyacrylamide Northern blotting using polyA+ RNA .................................. 127 3.3.3 Agarose Northern blotting .................................................................................................. 129 3.3.5 Growth test at 57 C ............................................................................................................. 138 Conclusion ....................................................................................................................... 147 CHAPTER IV: Discussion .............................................................................................. 149 4.1 General Discussion ................................................................................................................. 150 4.2 Future Directions ................................................................................................................... 156 References ....................................................................................................................... 158 Appendix ......................................................................................................................... 165 V List of Tables Table 1. Primer sequences for making two homology arms of plasmid constructs. ..................... 24 Table 2. RE digestion regarding the confirmation plasmids with two homology arms. .............. 29 Table 3. Preparation of media for culturing C. merolae. .............................................................. 31 Table 4. Primers and PCR product sizes of E034, D067, R289, and S262 for confirmation of intron deletion. ............................................................................................................................... 35 Table 5. Primers and PCR product sizes of E034, D067, R289, and S262 for confirmation of URA marker deletion. .................................................................................................................... 36 Table 6. DNA oligonucleotides were used to amplify and sequence the desired gene. ................. 37 Table 7. Features of Probes and Restriction Enzymes for Southern blotting............................... 39 Table 8. Co-cultures setup. ............................................................................................................ 47 Table 9. Growth test results for intron deletion strains 42 C. ....................................................... 64 Table 10. Distribution of WT and ∆is in the co-cultures in the logarithmic and saturation phases. ............................................................................................................................................ 80 Table 11. DNA oligonucleotides were used regarding the fluorescent and agarose Northern for the desired genes. ......................................................................................................................... 101 Table 12. DNA oligonucleotides were used for amplifying the desired genes for RT-qPCR. ..... 108 Table 13. Growth test at 57 C. ..................................................................................................... 139 VI List of Figures: Figure 1. Pre-mRNA Splicing process and spliceosome assembly. .......................................... 3 Figure 2. Consensus sequences used in the classification of major versus minor introns. ..... 7 Figure 3. Intron abundance in large genomes. .......................................................................10 Figure 4. Schematic of alternative splicing. ............................................................................13 Figure 5. Splicing occurs cotranscriptionally and affects different steps of transcription. ...15 Figure 6. Homologous recombination for intron deletion. .....................................................23 Figure 7. A summary of the intron deletion process and confirmation. .................................26 Figure 8. Plate layout for doing the growth test in the 48-well plate. .....................................44 Figure 9. The map of the plasmid containing the mTFP. .......................................................46 Figure 10. Fitness test set up for co-culture of WT-∆10i and WT- R289∆i.............................48 Figure 11. Confirmation of homology arm insertion in plasmids. ........................................52 Figure 12. Confirmation of the full-length transformation amplicon. ...................................54 Figure 13. Confirmation of intron deletion for S262. .............................................................56 Figure 14. Confirmation of intron deletion for E034. .............................................................57 Figure 15. Confirmation of intron deletion for D067..............................................................58 Figure 16. Confirmation of intron deletion for R289..............................................................59 Figure 17. Confirmation of intron deletions by Southern blotting. ........................................62 Figure 18. Comparison of doubling times between T1 (WT) and ∆i strains. .........................65 Figure 19.Comparison of R289∆i growth to T1 at 42 C. ........................................................67 Figure 20. Comparison of ∆10i growth to T1 at 42 C. ............................................................68 Figure 21. Comparison of D067∆i growth to T1 at 42 C. .......................................................69 Figure 22. Lag time of T1 at 42 C ...........................................................................................72 Figure 23. Lag time of S270∆i at 42 C.....................................................................................73 Figure 24. Lag time of Q117∆i at 42 C ....................................................................................74 Figure 25. 42 C growth assay for T1::mTFP compared with T1. ...........................................76 Figure 26. Microscopy images for the fitness test. ..................................................................78 VII Figure 27. Fitness test comparing the fraction of WT in the co-cultures for ∆10i and R289∆i. ..............................................................................................................................................................81 Figure 28. Biogenesis of Stable Intronic Sequence RNAs (sisRNAs). ....................................88 Figure 29. long-read transcriptomic data of C. merolae sisRNAs. .........................................93 Figure 30. Short-read transcriptomic data of C. morolae sisRNAs. .......................................96 Figure 31. Fluorescent Northern blotting. ..............................................................................99 Figure 32. RT-qPCR. ............................................................................................................. 106 Figure 33. Primer pair representative location for doing RT-qPCR. ................................... 107 Figure 34. Different cDNA synthesis methods for RT-qPCR................................................ 110 Figure 35. Confirmation of sisRNAs at 42 C. ....................................................................... 115 Figure 36. Effect of temperature on J129 sisRNA expression. ............................................. 117 Figure 37. Effect of temperature on K142 ncRNA expression ............................................. 119 Figure 38. Effect of temperature on Q270 sisRNA expression. ............................................ 121 Figure 39. Effect of temperature on K260 sisRNA expression. ............................................ 123 Figure 40. Effect of temperature on E034 intron expression. ............................................... 125 Figure 41. PolyA+ RNA Northern blotting. .......................................................................... 128 Figure 42. Agarose northern blotting for Q270. ................................................................... 130 Figure 43. Polyadenylation vs accumulation. ....................................................................... 132 Figure 44. RNA levels of regions within sisRNA containing genes as determined by RTqPCR. ................................................................................................................................................. 135 Figure 45. RNA levels of intronic sequences as determined by RT-qPCR. .......................... 137 Figure 46. Growth test for Q270∆i at 57 C. .......................................................................... 140 Figure 47. Growth test for J129∆i at 57 C. ........................................................................... 142 Figure 48. Growth test for K260∆i at 57 C. .......................................................................... 144 Figure 49. Growth test for E034∆i at 57 C. ........................................................................... 145 VIII List of Abbreviations ∆I: The strain with intron ∆i: The intron deletion strain ∆is: Intron deletion strains 3´ss: 3´ splice site 3´UTR: 3´Untranslated region 5´ss: 5´splice site 5´UTR: 5´Untranslated region 5-FOA: 5 Fluorotic acid Bp: Branch point C. merolae (Cm): Cyanidioschyzon merolae ciRNAs: circular intronic RNAs DT: Doubling time EJC: exon-exon junction complex FASTA: Fast-All g.DNA: Genomic DNA GOI: Gene of interest GSP: Gene-specific primers hnRNPs: heterogeneous nuclear ribonucleoproteins ICG: Intron-containing gene ILS: Intron-Lariat Spliceosome IX IR: Intron retention ISE: Intron splicing enhancer ISS: Intron splicing silencers LECA: Last eukaryotic common ancestor LIC: Ligation-independent cloning lncRNA: Long non-coding RNA miRNAs: and microRNAs mRNA: mature messenger RNA MRP: RNAse for mitochondrial RNA processing mTFP: monomeric Teal fluorescent protein NCBI: National Center for Biotechnology Information NF: normalization factors NMD: Non-sense mediate decay NRT: no reverse transcriptase NTC: no-template control OD: Optical density ODU: Optical density unit OMPdecase: Orotidine-5’-phosphate decarboxylase OPRTase: Orotate phosphor ribosyl transferase oSDR: oligonucleotides by Stephen Rader PEG: Polyethylene glycol X piRNAs: piwi-interacting RNAs PolyA+: polyadenylation-selected RNA Pre-mRNA: Pre-messenger RNA pSR: The plasmid by Stephen Rader (The plasmid backbone in Rader lab) PTCs: Premature termination codons RE: Restriction enzyme RFU: relative fluorescence units RT-qPCR: reverse transcriptase quantitative PCR scaRNAs: small Cajal body-associated RNAs sisRNAs: stable intronic sequence RNAs sisRNAs: stable intronic sequence RNAs SnoRNA: small nucleolar RNA snoRNAs: small nucleolar RNAs snRNA: small nuclear RNA snRNPs: small nuclear ribonucleoproteins Tm: melting temprature WT: Wild type YPD: Yeast Extract Peptone Dextrose YsnRNAs: Yeast snRNAs XI Note on gene references in this work C. merolae genes are named serially, starting with the species CM, for Cyanidioschyzon merolae, then the chromosome letter (A, B, C… T), then the serial number of the gene along the chromosome, ending with a letter designating whether the gene is protein-coding (C), transcribed (T), or a hypothetical transcript (Z). To simplify the gene references and make it easier to follow this work, I refer to the genes below using only the chromosome letter and serial number, as shown in column 2 of Table 1. If an intron is deleted from a gene, the gene name is followed by ∆i. Table I. Intron-containing gene function. Intron containing gene Intron containing gene Intron deletion strain Inferred Function CMS262C S262 S262C∆i 60S ribosomal protein L23 CMR289C R289 R289C∆i NADH dehydrogenase I (ComplexI) iron-sulfur protein 75kDa subunit N-terminal fragment CMD067C D067 D067C∆i Probable prohibitin protein CME034C E034 E034C∆i Similar to calmodulin CMJ129C J129 J129C∆i Histone deacetylase CMK260C K260 K260C∆i Probable mitochondrial processing peptidase alpha subunit CMQ270C Q270 Q270C∆i Mitochondrial chaperonin hsp60 precursor CMK142T K142T - Unannotated transcript CMS270C S270 S270∆i chaperonin containing TCP1, subunit4 (delta) CMS342C S342 S342∆i V-type ATPase V1 subunit A CMQ382C Q382 Q382∆i similar to U3 snoRNP component Utp15p CMC008C C008 C008∆i NADH dehydrogenase I(Complex I) alpha subcomplex 7 (B14.5a) CMQ117C Q117 Q117∆i Eukaryotic translation initiation factor eIF-1A XII CMR350C R350 R350∆i Similar to CCAAT-binding transcription facto CMS311C S311 S311∆i Similar to bacterioferritin comigratory protein CMC053C C053 C053∆i CMO094C O094 O094∆i 60S ribosomal protein L35 3-isopropylmalate dehydratase small subunit Additionally, ∆6i and ∆10i are strains with six (R350(2), S311, C053, S262, and Q270) and 10 (R350(2), S311, C052, S262, Q270, D067, C008, Q117, and O094) intron deletions, respectively. XIII Acknowledgments To my supervisor, Dr. Stephen Rader, and my advisor, Dr. Martha Stark, I give my deepest respect and heartfelt gratitude. Together, Stephen and Martha have provided unparalleled training, both at the bench and in fostering my critical thinking abilities. Their supportive and encouraging environment has ignited my curiosity at every stage of my research. These years of study have been truly rewarding. I also wish to extend my sincere gratitude to my supervisory committee members: Dr. Sarah Gray, Dr. Kendra Furber, and Dr. Andrea Gorrell. Their insightful questions and valuable feedback have significantly enriched this thesis. I am also so thankful to Patrick Geertz for his assistance in performing the RT-qPCR experiment and analysis. I would also like to acknowledge Viktor Slat, who did the initial bioinformatic work. I am thankful to the University of Northern British Columbia for the financial support that allowed me to pursue my studies. A special thank you goes to my husband, Ehsan Miyanehsaz, for his unwavering support and wise counsel throughout the years. My deepest appreciation goes to my parents, Seddigheh Ghannadzadeh and Hossein Ghaffarzadeh, for their love and guidance, and to my siblings for bringing joy during challenging times. Lastly, to my expecting boy, thank you for bringing a new sense of purpose and joy into my life. Your presence has been a source of inspiration and motivation throughout this journey. I eagerly await the day we meet. XIV CHAPTER I-Introduction 1 1.1 Pre-mRNA splicing During eukaryotic gene expression, genes are first transcribed into pre-messenger RNAs (pre-mRNAs), in which the coding information (exons) is interrupted by introns. To produce mature messenger RNAs (mRNAs) with an uninterrupted protein coding sequence, introns are excised from pre-mRNAs by the splicing process, which is carried out by a large macromolecular machine called the spliceosome1. The spliceosome is intricately assembled for each intron within pre-mRNA through a detailed, step-by-step process utilizing small nuclear ribonucleoproteins (snRNPs), each comprising small nuclear RNA (snRNA) and various proteins. This assembly involves five principal snRNPs: U1, U2, U4, U5, and U6, each identified by its snRNA component, coming together with the pre-mRNA2,3. This process is marked by critical RNA-recognition and structural remodeling events, alongside significant alterations in protein composition, to form the spliceosome's active site. Initially, the U1 and U2 snRNPs align with the 5′ splice site (5′-SS) and the branch point (BP) on the pre-mRNA, respectively, forming the foundation for the A complex. The assembly progresses with the incorporation of the U4/U6-U5 tri-snRNP, within which U6 snRNA forms extensive base pairs with U4 snRNA. This leads to the formation of the pre-B complex, where U1 snRNP is still attached to the 5′-SS. However, the activity of ATPase Prp28 triggers the release of U1 snRNP, establishing a stable B complex. For catalytic activation to occur, ATPase Brr2 detaches the U4 snRNA, enabling U6 snRNA to assume a catalytically active configuration in tandem with U2 snRNA and to pair with the intron's 5′ end4 (Figure 15,6). 2 Figure 1. Pre-mRNA Splicing process and spliceosome assembly. The spliceosome assembles in a highly ordered manner, becomes activated to form the active site, and undergoes extensive remodeling to carry out the branching and exon ligation reactions. It then releases mRNA (ligated exons) and disassembles. ATPases from the DEAD-box, DEAH-box, and Ski-2 families (red) play crucial roles in these remodeling processes. Abbreviations: BP, branch point; ILS, intron-lariat spliceosome; NTC, Prp19associated complex; NTR, Prp19-related complex; snRNP, small nuclear ribonucleoprotein; SS, splice site. This Figure was obtained from “RNA Splicing by the Spliceosome”, Max E Wilkinson and Clément Charenton with permission. 3 Analysis of the base pairing between U6 and U2 snRNAs with the pre-mRNA has unveiled RNA sequences reminiscent of those in group II self-splicing introns, hinting at structural similarities in their active sites. Following the B complex's remodeling by Brr2 and stabilization by a protein complex linked with Prp19 (named the NTC) and NTC-related factors, the Bact complex emerges, encompassing U2, U5, and U6 snRNAs. ATPase Prp2 then facilitates the alignment of branching factors with the 5′-SS and BP, leading to the branching catalysis by the B* complex and the subsequent formation of the C complex. Following the initial catalytic reaction, ATPase Prp16 restructures the spliceosome to permit the separation of branching factors and the alignment of the 3′ splice site (3′-SS) for the next step, supported by exon ligation factors. The U5 snRNA aligns the 5′ and 3′ exons, allowing the spliceosomal C* complex to execute the second catalytic step. Finally, ATPase Prp22 releases the mRNA, leading to the formation of an Intronlariat spliceosome (ILS) from Post splicing complex (P), which is followed by the spliceosome disassembly by ATPases Prp43 and Brr2, preparing its components for another round of splicing1,7,8. For splicing to occur, both cis and trans components are required. Cis-elements are located within the transcript, while trans-acting elements, including spliceosome proteins, splicing repressors, and activators, recognize cis-elements9. Four important cis-elements define an intron: 5' SS, BP, 3' SS, and the polypyrimidine tract. These elements are all short, conserved sequences. For splicing to occur, the spliceosome must first recognize these short sequences within the premRNA from among many similar sequences, then precisely remove the intron and ligate the exons together. In addition, spliceosome association can be prevented or enhanced via cis-acting regulatory RNA elements, such as intronic or exonic splicing enhancers and silencers. These regulatory elements are bound by trans-acting factors, such as serine/arginine-rich (SR) proteins 4 and heterogeneous nuclear ribonucleoproteins (hnRNPs). Together, these transcript elements, snRNAs, and proteins compose a precise and complex machine that is necessary for accurate gene expression10. As splicing is crucial to the expression of nearly all human genes, alterations in RNA splicing through disruption of either cis- or trans-acting factors can result in abnormal cellular metabolism and/or function9. 1.1.1 Splicing in humans Most of our knowledge of pre-mRNA splicing comes from studying yeast Saccharomyces Cerevisia (S. cerevisiae) and human cells. While only 5% of yeast genes contain introns, over 95% of human genes are intron-containing. Despite having a relatively modest number of proteincoding genes, around 25,000, which is similar to that of many other animals, humans exhibit remarkably complex gene architecture and utilization. This complexity is largely attributed to the process of pre-mRNA splicing. Typically, human genes consist of small exons (about 145 base pairs on average) interspersed with large introns (averaging around 3,400 base pairs) that need to be excised from nascent RNAs through splicing. With an average of eight such large introns per human gene, the bulk of a pre-mRNA transcript is intronic, necessitating the removal of most of the transcript to produce a mature mRNA11,12. The timing and manner of this intron removal provide numerous opportunities for the regulation of gene expression4. Humans utilize two distinct spliceosomes for this purpose: the major (U2 type) and the minor (U12 type) spliceosomes (Figure 213). The major spliceosome, which involves U1, U2, U4, U5, and U6 snRNPs, processes more than 99% of human introns featuring the conventional ‘GU’ at the 5' end and ‘AG’ at the 3' end splice site sequences (major introns). Conversely, the minor spliceosome, comprising U11, U12, U4atac, U5, and U6atac snRNPs, targets less than 1% of introns with alternative splice site consensus sequences. Each intron removal is facilitated by a uniquely assembled spliceosome, 5 functioning as single turnover enzymes that organize sequentially on nascent pre-mRNAs, indicating that every intron is excised by its specifically assembled spliceosome 4,14. 6 A B Figure 2. Consensus sequences used in the classification of major versus minor introns. A) The splice site selection by the respective components of the major and minor spliceosomes has been illustrated. The snRNAs of the major (U1 and U2) and minor (U11 and U12) spliceosomes are shown base-pairing with their cognate consensus sequences. In the center, near the major and minor intron labels, consensus sequences are depicted as nucleotide frequency plots, where the size of each nucleotide indicates its frequency at that genomic position. B) The remaining core snRNAs unique to major (U4 and U6) and minor (U4atac and U6atac) intron splicing, as well as the shared U5 snRNA are shown. This Figure was obtained from “Introns: the “dark matter” of the eukaryotic genome”, by Kaitlin N Girardini, which is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY).13. 7 1.2 Intron evolution Introns come in different types. In addition to eukaryotic pre-mRNA introns, some bacterial and organellar genomes contain self-splicing Group I and II introns15–17. Internal open reading frames in these two groups of introns encode proteins that help to remove introns from their host transcripts and intron propagation to new sites via reverse transcription (self-splicing introns)15. Long-standing research has shown that spliceosomal introns and Group II self-splicing introns share many characteristics. Similarities between the two intron types can be seen in the splice site recognition, branching mechanism, stereochemical course of the splicing reaction, and the existence of related RNA domain structures. A homolog of the intron-encoded protein, IEP, which is required for the proper folding of Group II introns, promotes RNA recognition and splicing in the spliceosome18. It is also clear that some of the spliceosomal snRNAs have a similar function and secondary structure with Group II introns18. These findings support the assertion that Group II introns and spliceosomal introns share a common evolutionary origin. It appears that at an early stage in eukaryotic evolution, autocatalytic splicing was replaced by splicing mediated by the spliceosome. Indeed, Group II introns split into catalytically inert spliceosomal introns and catalytically active RNA fractions during this transition, followed by the degradation of the reverse transcriptase open reading frame within introns19. The evolutionary origins of the exon-intron structure in eukaryotes have been a topic of extensive and ongoing debate. It has long been debated when in evolution, the exon-intron structure of eukaryotes arose. The introns-early hypothesis proposes that eukaryotic genes received almost all of their introns from their prokaryotic ancestors, and the structural differences among homologous genes were the result of differences in intron loss20. Based on this scenario, the extant prokaryotes have lost all of their primordial introns, as well as their spliceosome, as a result of 8 genome streamlining. There is a strong connection between the intron-early and exon theory of gene origin and evolution, which posits that the ancestor of modern protein-coding genes were RNA molecules encoding exon-sized peptides, and also containing non-coding regions. By recombination between such RNA molecules, the first protein-coding genes were formed in which exons are interrupted by introns. Consequently, introns were regarded as a necessary condition for the emergence of protein-coding genes. In contrast, the introns-late hypothesis argues that introns are a eukaryotic novelty and that new introns have been continuously appearing throughout evolution. Bacteria and archaea did not have introns or spliceosomes in such a scenario 19. According to later adaptations of the introns-early hypothesis, introns are still considered to be primordial features, despite the fact that new introns can emerge alongside those that are ancient 21. Eventually, these hypotheses were merged into a concept known as numerous introns early in the evolution of eukaryotes19. Reconstructions of the ancient genomes reveal that the last eukaryotic common ancestor (LECA) had intron densities that were higher than most modern eukaryotes 22,23. According to a study of 99 eukaryotic genomes, intron density can be as low as 0.1 introns/kb in S. cerevisiae 24 or as high as approximately six introns per kb of the coding sequence in mammals or almost nine introns per gene on average19 (Figure 3). Variation in intron densities can be attributed in large part to remarkable differences in intron gain or loss dynamics24. Despite the fact that some intron positions (10–40%) are conserved between highly divergent eukaryotes, these studies have shown that the number and placement of most introns are dynamic during evolution24. 9 Figure 3. Intron abundance in large genomes. This Figure is modeled based on the information from the ‘Introns: Good Day Junk Is Bad Day Treasure’ by Julie Parenteau and Sherif Abou Elela, published in the journal of Cell Press11. 10 These "presence/absence" variations are most likely caused by intron loss, which can occur either when reverse transcription of a spliced RNA, followed by homologous recombination, erases the intron, or through genomic deletion. Additionally, a variety of mechanisms have been proposed for gaining new introns. Single nucleotide changes, for example, can generate new splice sites (and therefore introns) and cause "intron sliding" or alternative splicing events. Introducing a new exon and splitting an intron into two smaller ones are possible outcomes of "exonization" within a large intron. These introns gain mechanisms are based on the splicing of pre-existing introns and represent the diversification of introns in the intron genome, as opposed to creating de novo introns at sites without introns previously. There are two main pathways by which de novo introns are created. The first is intron transposition, in which an intron at one location is copied and inserted at another. The transposition of introns, known as introner elements, may have led to an expansion of intron repeats. DNA damage repair or non-autonomous DNA transposons may also result in intron transposition. The second mechanism, intronization, refers to the process of creating introns by mutations arising either from drift or other changes to sequences so that the splicing machinery recognizes them as introns. Mutations may accumulate in the transcripts that allow them to be recognized by the splicing machinery and spliced. It is believed that this process occurs gradually over evolutionary time, creating sequences that possess both exon and intron properties that are frequently alternatively spliced through weak splicing signals 25. 1.3 Intron function It is obvious that intron removal from pre-mRNAs is important, but it cannot explain how and why introns have persisted in genomic DNA throughout evolution. Given the energy required to transcribe, remove, and decay introns in every mRNA, introns impose an enormous metabolic burden on eukaryotic cells. Gene transcription costs double when only a 5-kb intron is present in 11 a typical eukaryotic gene26. The process of removing introns is also very complicated and costly, as it requires the coordination of hundreds of proteins. In fact, the spliceosome is one of the largest ribonucleoprotein complexes in the cell. An intron burden could be tolerated by multicellular organisms with large genomes and stable cellular environments. However, it is harder to understand how single cells with compact genomes could tolerate this energetic burden if introns' only function is removal27. Several studies have investigated the selective advantages that introns provide to eukaryotes, which contribute to overcoming their energetic cost 28, including harboring various cis-elements as well as non-coding RNAs (ncRNAs). Cis-acting regulatory sequences inside introns, which are selectively recognized by complementary trans-acting factors, can be attributed to known molecular functions including shaping alternative splicing, gene expression enhancement, and mRNA stability as further detailed in the following paragraphs 29,30. 1.3.1 Intronic Cis-elements Besides constitutive splicing, most eukaryotic cells have the ability to splice pre-mRNAs alternatively, which incorporates different coding regions into mature mRNAs, generating multiple mRNA variants from a single gene and promoting proteomic diversity10,31,32 (Figure 433). Introns have an active role in regulating alternative splicing by recruiting intronic splicing enhancers (ISEs) and silencers (ISSs). These factors bind to intronic elements and activate or suppress the use of particular splice sites34. 12 Figure 4. Schematic of alternative splicing. In this representation, lines signify introns, which are excised from the pre-mRNA during splicing, while boxes represent exons, which are joined together to form the mature mRNA. Alternative splicing permits the inclusion of various exon combinations in the final mRNA, enabling the generation of multiple proteins from a single gene. These mature mRNAs are subsequently translated into proteins, with distinct exon arrangements resulting in different proteins, as exemplified by Protein 1 and Protein 2 in the schematic. This Figure was obtained from “Chemical Modulation of Pre-mRNA Splicing in Mammalian Systems”, by Robert E. Boer with permission. 13 Introns play a vital role in transcriptional regulation, which can be categorized into splicing-independent and splicing-dependent types. Splicing-independent regulation occurs due to the presence of enhancer or promoter elements within the intron, allowing these introns to affect transcription even if splicing does not occur. Splicing-dependent regulation, on the other hand, requires functional introns within the gene's transcribed region to enhance transcription when properly spliced35. The splicing process is facilitated by splicing factors recruited by the carboxyterminal domain (CTD) of RNA Polymerase II (RNAPII). This assembly stabilizes transcription factors at the promoter, primes nucleosomes with activation marks, and influences transcription elongation through histone modifications. Additionally, splicing factors enhance transcription termination by recruiting termination factors and altering elongation marks 35 (Figure 5). 14 Figure 5. Splicing occurs cotranscriptionally and affects different steps of transcription. A) An intron within the transcribed region is flanked by exon I and exon II, with the 5′ and 3′ splice sites serving as locations for spliceosome assembly during transcription. (B) Splicing factors are recruited to the intron cotranscriptionally by the carboxy-terminal domain (CTD) of RNA Polymerase II (RNAPII). This spliceosome assembly helps stabilize general transcription factors (GTFs) at the promoter and primes nucleosomes with activation marks (H3-K9 acetylation and H3-K4 trimethylation) for initiation. Additionally, splicing factors interact with transcription elongation factors to influence nucleosome modifications (H3-K36 trimethylation), promoting elongation. They also enhance transcription termination by recruiting termination factors and removing elongation marks that impede effective termination. This Figure was obtained from “Gene Architecture Facilitates IntronMediated Enhancement of Transcription”, Katherine Dwyer, which is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY)35. 15 Introns also can control mRNA stability by modulating the nonsense-mediated decay mechanism (NMD)36,37. NMD is thought to serve as an mRNA surveillance mechanism to prevent the synthesis of truncated proteins. The main NMD substrates are transcripts that have acquired Premature Termination Codons (PTCs). The retention of introns (IRs) may result in PTCs, causing degradation of mRNA by NMD38. On a normal, non-PTC-containing mRNA, the exon-exon junction complexes (EJC) are removed in the first round of translation but when an intron is located at least 50–55 nucleotides downstream of a termination codon, the EJC positioned near the exonexon junction is not displaced by translating ribosomes, resulting in the EJC remaining associated with mRNA, which is the signal for degradation by NMD 39,40. This is consistent with the observation that introns in the 3’ -UTR may significantly enhance NMD40. 1.3.2 Intronic non-coding RNAs Additionally, introns can harbor several kinds of noncoding functional RNA genes, such as small nucleolar RNA (snoRNA), piwi-interacting RNAs (piRNAs), and long noncoding RNAs (lncRNAs)41–43. While introns are usually degraded after splicing, these ncRNA genes are processed from spliced introns. ncRNAs located in introns are considered to play an important role in the auto-regulation of host gene expression30,44. ncRNAs modulate gene expression through two mechanisms: transcriptional gene silencing, which takes place in the nucleus where ncRNAs are involved in chromatin changes, and post-transcriptional gene silencing, which occurs in the cytoplasm where ncRNAs direct the RNA-induced silencing complex to target mRNAs to be cleaved or translationally inhibited45. Recent studies have unveiled the role of intronic sequences as stable intronic sequence RNAs (sisRNAs). SisRNAs are traditionally known as intronic sequences retained during splicing and not immediately degraded. Additional variations have been 16 identified, such as intronic sequences with polyadenylation or the inclusion of exonic sequences in truncated transcripts46. sisRNAs are surprisingly stable and not subject to standard degradation pathways. While the mechanisms behind their production remain unclear, some have demonstrated functionality. They exert control over genes at the levels of DNA, RNA, and proteins, frequently engaging in autoregulatory feedback loops to ensure cellular homeostasis in both normal and stressful conditions46. 1.4 Thesis scope and significance The aim of this thesis is to examine the functions of introns and to elucidate the underlying mechanisms that contribute to the retention of certain introns while others are lost. Given the complexity of the spliceosome and the high density of introns in other model organisms, I will use a new model organism with only a handful of remaining introns and a reduced spliceosome. 1.4.1 C. merolae Cyanidioschyzon merolae (C. merolae) is an extremophilic unicellular red alga that possesses a cellular architecture that includes a single mitochondrion, a single chloroplast, and a few other membranous organelles47. It grows in acidic (pH 0.2–4), sulfate-rich volcanic hot springs with temperatures up to 57 C48,49. It has a simple genome with very little genetic redundancy - in total, 5,331 genes, of which 4,775 are protein-coding49,50 . The lack of a rigid cell wall also makes it relatively easy to isolate or extract the contents of C. merolae cells51. When compared with most other eukaryotic algae, C. merolae cells are quite small (1.5-3 µm in diameter). A genome of 16.5 Mbp is divided into 20 chromosomes. The small cell size and the minimal number of organelles make it suitable for biochemical and omics analyses. 17 In addition to being the first fully sequenced eukaryotic algal genome, C. merolae's nuclear genome is also the first fully assembled eukaryotic genome without gaps48,49. C. merolae has only 3952 introns in 38 genes49,53,54, and compared to other organisms, it has a simplified spliceosome – four snRNAs (U2, U4, U5, U6) and 49 core splicing proteins55,56. By contrast, S. cerevisiae and humans contain five snRNAs (U1, U2, U4, U5, and U6) and over 100 and 200 spliceosomal proteins, respectively57,58. Therefore, due to the small number of introns and a reduced spliceosome, C. merolae presents an excellent opportunity to study intron evolution and function. 18 CHAPTER II: Evaluating the Impact of Intron Deletion on C. merolae Growth Under Normal and Nutritional Stress Conditions 19 2.1 Introduction The emergence of introns marked a significant evolutionary advancement, distinguishing eukaryotic genomes from prokaryotic ones. Historically, introns were considered mere sequences excised to generate spliced transcripts that encode functional products 13. However, increasing evidence suggests that introns have crucial roles including regulating gene expression, mRNA stability, regulation of splicing, regulation of non-sense mediated decay (NMD) mechanism, harboring several kinds of noncoding functional RNA genes13,27,30,34,36,59. Reconstructions of ancestral genomes have shown that the LECA had a genome characterized by relatively high intron density compared to modern eukaryotes. Post-LECA evolutionary trends in eukaryotes have largely involved intron loss, with notable exceptions in certain lineages like plants and animals, which have experienced a net increase in introns22. Despite the overwhelming phylogenetic trend towards intron loss, it appears to be rare for eukaryotes to lose all of their introns, which is one of the evolutionary mysteries of organisms with few introns. The remaining introns in an intron-poor organism probably perform a function or will eventually be lost52. Despite the prevalence of introns in eukaryotic genomes and their recognized importance in gene regulation, much about introns remains to be explored. This research aims to determine whether the maintenance of introns in C. merolae, an acidophilic red alga, whose genome sequence revealed only 39 introns is due to their function or whether they will gradually be lost 60. The objectives of the chapter are the following: 20 AIM I: Deleting four introns in C. merolae (S262, D067, E034, and R289) and performing the cell growth rate assessment to determine the impact of four intron deletions on the growth of C. merolae. AIM II: Performing a growth competition analysis (fitness test) for two ∆i (R289∆i and ∆10i) to determine the effect of introns on C. merolae's growth under starvation conditions. HYPOTHESIS I: The doubling time of cells would change if deleted introns had an important function in the cell. HYPOTHESIS II: R289∆i and ∆10i would not survive under conditions of nutrient deprivation if the deleted introns play an important role in cell survival under nutrient starvation. 21 2.2 Materials and Methods 2.2.1 Designing integration constructs for homologous recombination The coordinate for each gene of interest (GOI) from the NCBI database was obtained, expanded the selection +/- 5000 bp in the FASTA file. Then a new SnapGene map (a software tool for visualizing and analyzing DNA constructs) of the locus was made, annotated with exons and introns using the Cm Intron Database file from the Rader lab (the database that provides information on C. merolae genes) and the 5’ and 3’ UTRs for the GOI were annotated using longread and short-read sequencing data. Primers A/D, E/F, and H/I were designed using NCBI Primer BLAST (as shown in Figure 6 and Table 1). The primers were added to the map, with SwaI restriction enzyme LIC arms added to primers A and D, and PacI restriction enzyme LIC arms added to primers E and F. Overlapping primers B and C were designed and added to the map, starting with the exon 1 part of primer B and the exon 2 part of primer C. The melting temperature (Tm) was adjusted to 64-66 C using the Q5 NEB calculator, ensuring that the Tm of the overlap matched the melting temperature (Tm) of primers A and D with the high-fidelity Q5 polymerase. The pSR887 (the plasmid backbone designed by our lab, identified as the plasmid by Stephen Rader) was used as the backbone to create the plasmid for deleting the intron. The region encompassed by the AD primers was inserted into the SwaI restriction enzyme site and the region by the EF primers into the PacI restriction enzyme site of pSR887. The 2 bp overhang for PacI was deleted before inserting the region, and a G or C was added to the plasmid sequence if needed. Then a SnapGene map of the ∆i locus containing the Ura 5.3 gene was created. It should be mentioned that the DNA construct for S262 was created by a former student in the lab, and in this study, the transformation for this strain was performed. Additionally, the DNA constructs for the 22 transformation and deletion of Q270 and Q117, designated as pSR1013 and pSR1014, respectively, were created for future students in the lab. Figure 6. Homologous recombination for intron deletion. The DNA constructs for each intron deletion include two homology arms and the Ura5.3 auxotrophic marker. Different primer pairs for making the DNA constructs were designed: AB, CD, and EF. After transforming the plasmid into C. merolae, the intron and URA marker will be excised from the genome through the first and second homologous recombination events, respectively. 23 Table 1. Primer sequences for making two homology arms of plasmid constructs. TACTTCCAATCCCACGAGGAGAAATTAACT is the ligation independent cloning (LIC) site for the forward primer and TTATCCACTTCCCACG is the LIC site for the reverse primer. oSDR refers to the oligonucleotides in our lab which are identified as oligonucleotides by Stephen Rader. Column 4 presents the primers region as shown in Figure 6. oSDR # Orientation Gene Region Sequence (5′ to 3′) 2143 Reverse Q270 B GATTGTTGCGACTTGTTTGATTTCATTCTTGCCG Annealing Tm C (Q5) 65 2144 Forward Q270 C AAATCAAACAAGTCGCAACAATCAGTGCA 64 2145 Forward Q270 A CTTATCTCAATATTTGACGAGGGTACAGAGCTTCAA 65 2146 Reverse Q270 D AACAATCACCAATTTGGACCTTGTTCTGGCGATAC 64 2147 Forward Q270 E AGTTGAAGTATGTTACAATATGGATATGTATTGAAGATGCG 63 2148 Reverse Q270 F ATGTTAAGTGGATTACTCATCGCTGGTTTCTGCATT 63 2149 Reverse Q270 H GCGTCGCCAAGGATATGTTA 66 2150 Forward Q270 I GGAAGCATTCTAGGTACCGC 66 2151 Forward Q117 A CTTATCTCAATATTTGACTCCATCTGTCGCAACTGT 67 2152 Reverse Q117 D AACAATCACCAATTTGGCCCTGGACTCGGAAAATA 67 2153 Reverse Q117 F ATGTTAAGTGGATTACGTCAATGTAGGTCGTGAGCG 67 2154 Forward Q117 E AGTTGAAGTATGTTACGGTTTGCGGGATTTCCAAGA 67 2155 Reverse Q117 B TGTCCATATTCTTGCCCGTCCTCCTTG 65 2156 Forward Q117 C GACGGGCAAGAATATGGACAAGTTTTGCGCATG 67 2157 Forward Q117 I GCTCCGCATGAAGTACGTTG 66 2158 Reverse Q117 H GGTAGCCAGCAATTTGTCCA 66 2159 Forward R289 A CTTATCTCAATATTTGTGCCTGTTTGCGTCAAAGTA 66 2160 Reverse R289 D AACAATCACCAATTTGCAATCTCCACGTTGACCAGC 66 2161 Reverse R289 F ATGTTAAGTGGATTACGGTCTGATGGATGGTCCGAA 67 2162 Forward R289 E AGTTGAAGTATGTTACATACGTGCCGCGATTCTCA 68 2163 Forward R289 C CCAATATGCGACCAAGGTGGCGAATGTC 67 2164 Reverse R289 B CCACCTTGGTCGCATATTGGACAGTCCAGC 67 2165 Forward R289 I ACACGTCATCCGTTCGTTG 66 2166 Reverse R289 H CTTGGGCGACTAGTGTCAATC 66 2419 Reverse R289 Intron TCGAAACGGTTAGTATTTTGCTCT 64 2175 Forward E034 A CTTATCTCAATATTTGTCGGCAAAATCCCGAGACAT 67 2176 Reverse E034 D AACAATCACCAATTTGTCCAGCGAACGCAAGAAAC 67 2177 Reverse E034 F ATGTTAAGTGGATTACGTGAGCAAAGCGGAATGCTT 66 2178 Forward E034 E AGTTGAAGTATGTTACAACGCATGCACTGAGAACT 67 2179 Reverse E034 B CAAAGTTTACAGAACCTTTACCATCCGAGTCCGC 67 2180 Forward E034 C CGGATGGTAAAGGTTCTGTAAACTTTGAAGAGTTCCTCG 68 2181 Forward E034 I CATTCCGTGAAATGGGCCAG 68 24 2182 Reverse E034 H TGACCGCGTCAAAACCAGTA 68 2245 Forward E034 A AGACATTCGCAAGCTC 60 2246 Reverse E034 F GAGCACGTGAGTCTTC 60 2183 Reverse D067 A ATGTTAAGTGGATTACAGAAAAAGCGCCCTAGCGTA 67 2184 Forward D067 D AGTTGAAGTATGTTACGAAACCAGGAGCGAGCAAAG 67 2185 Forward D067 F CTTATCTCAATATTTGAGTGAGCACATCCTCAACGA 67 2186 Reverse D067 E AACAATCACCAATTTGTCCTGGATGCGGGTATCTTG 67 2187 Forward D067 B CCGGTGTCCTCCTTCGACATTATAGAGCGC 68 2188 Reverse D067 C CCTTCTACAAGTACGCGCTCTATAATGTCGAAGGAGG 66 2189 Reverse D067 I GGTAGCAGTCACACCACAAG 66 2190 Forward D067 H GTCTGATGCTCGGTTGTCTT 66 2418 Forward D067 Intron TCCCCAGTTCGCTTATGCTT 66 2.2.2 Intron deletion A summary of the intron deletion process and confirmation is shown in Figure 7. 25 Figure 7. A summary of the intron deletion process and confirmation. This figure illustrates the process of constructing DNA with two homology arms, linearizing it by PCR, and then transforming it into C. merolae. After confirming intron deletion using PCR on the colonies grown on plates, the samples are sent for sequencing, and a Southern blot is performed for final confirmation. 26 2.2.1 Construction of vectors containing two homology arms In order to generate the AB, CD, and EF products, the polymerase chain reaction (PCR) using Q5 High-Fidelity DNA Polymerase (New England Biolabs), and a Thermal Cycler (BioRad) was carried out. Each reaction mixture contained 1X Q5 Reaction Buffer, 200 µM dNTPs, 0.5 µM of each primer, 5 ng of template DNA, and 1 U of Q5 High-Fidelity DNA Polymerase in a total volume of 50 µl. Thermocycling conditions consisted of an initial denaturation at 98 C for 30 sec, 35 cycles of denaturation at 98 C for 5 sec, annealing and extension at 72 C followed by a final extension at 72 C for 2 min. The annealing temperature and extension time were different based on each primer pair and product size (20 sec/kb of product size), respectively. Then, the gel purification of the AB and CD products was carried out using the Omega Gel Extraction Kit, Cat# D250033407 according to the manufacturer’s protocol. Then, the AD arm was produced by performing high-fidelity Q5 polymerase Splicing by overhang extension (SOEing) PCR using the AB + CD product. Subsequently, the AD and EF arms were cloned into the backbone plasmid pSR887 using the Ligation Independent Cloning (LIC) technique61. This method creates long cohesive sticky ends on both the vector and the insert that results in the generation of desired molecules after annealing the insert and the vector. For this purpose, pSR887 with genes of interest was digested with the SwaI (New England Biolabs, #R0604S) and PacI (New England Biolabs, #R0L) restriction enzymes. pSR887 consists of two restriction sites, one for PacI and one for SwaI. The PacI restriction site is located at the 3ʹ and SwaI restriction site is located at the 5ʹ end of the plasmid. In summary, 500 ng of DNA plasmid was digested with 5 µL of SwaI or 5 µL of PacI restriction enzyme (New England Biolabs). BSA was used in each of SwaI and PacI digestion reactions. Incubation of PacI and SwaI digestion reaction was performed at 37 C and 25 C 27 respectively for 3 hours. Afterward, heat inactivation of the enzyme was carried out at 75 C for 20 minutes. Next, the digested products were treated with 0.5 µL of T4 DNA polymerase in a 20 µL mixture reaction containing 50 mM Tris-HCl pH 8.0, 10 mM MgCl2, 100 ng/µL BSA, 0.1 M DTT and 0.5 µL of each 50 mM dCTP and 50 mM dGTP for PacI and SwaI digests respectively. SwaI and PacI plasmids were mixed and were heated at 65 C for five minutes then the reaction was allowed to cool at room temperature. Two µL of 25 mM EDTA was added and 5 µL of the reaction was transformed into 50 µL of Escherichia coli (E. coli) DH5alpha. It underwent heat shock at 42 C for 45 seconds, followed by 2 minutes on ice. Afterward, 150 μL of Luria Bertani (LB) media (1% bacto-tryptone, 0.5% yeast extract, 1% NaCl, and 0.1% glucose, with the pH adjusted to 7.0) was added. Finally, the mixture on LB plates containing either carbenicillin or ampicillin at a 25 mg/mL concentration was plated. After 16-18 hours, plasmids purification was performed using the Plasmid DNA Mini Kit II, Cat# 694527771-28) according to the manufacturer’s protocol and subjected them to restriction reactions with the appropriate restriction enzymes for each gene, which can ascertain the efficiency of the transformation. The LIC was repeated to create the plasmid with both 5 ʹ and 3 ʹ ends. Colonies were screened and RE digested again to confirm the final plasmid. The newly created plasmids alongside the selected restriction enzymes for confirming the efficiency of transformation for the AD and EF inserts are shown in Table 2. 28 Table 2. RE digestion regarding the confirmation plasmids with two homology arms. REs with their expected product sizes for each plasmid are shown. Genes of interest Plasmid RE Predicted size of plasmid after RE digestion Q270 pSR1013 Sac I+Spe I 1638+3049+3969 bp Q117 pSR1014 NheI +Xho I 1959+2324+3227 bp R289 pSR1015 XhoI+KpnI 2330+2519+3042 bp E034 pSR1017 SacI+SpeI 1438+2933+3542 bp D067 pSR1018 Bst Xi 3443+4754 bp 29 2.2.4 C. merolae URA transformation For the transformation into C. merolae (Cm), the high-fidelity Q5 polymerase PCR was performed for the AF product ensuring a full-length product and determining yield, followed by additional reactions to obtain 1 pmole of AF. The AF PCR product was then transformed into the T1 strain of Cm. Then, the transformation was plated on Modified Allen's medium with glycerol (MA2G), and selected colonies for screening PCRs to identify successful transformed colonies after 10-15 days, constituting the first screening. The positive colonies were plated from this first screening on MA2GU (MA2G+uracil) + 5-(Fluoroorotic acid (5FOA) for a second screening to eliminate the marker. Cells were diluted 2-3 days before transformation to have an actively dividing culture that will have an OD750 nm< 3.0 one day before transformation and 0.8mg/mL 5-Fluoroorotic acid (5FOA) to MA2GU (MA2G+ uracil) added to ensure a pure culture of T1 cells (no WT cells). One day before transformation, cells were diluted to OD750 nm ~0.2 in 50 mL aiming to transform cells at OD750 0.8-1.0. The preparation of media for culturing C. merolae is shown in Table 362. 30 Table 3. Preparation of media for culturing C. merolae. This information was obtained from “The basics of cultivation and molecular genetic analysis of the unicellular red alga Cyanidioschyzon merolae”, Yuki Kobayashi 62. Solution Component Final concentration in MA2 Solution I (10× for MA2) (NH4)2SO4 MgSO4·7H2O 500× A6 minor salts H2SO4 40mM 4mM 2X - KH2PO4 8 mM CaCl2 1 mM FeCl3 Na2EDTA H3 BO3 MnCl2-4H2O ZnCl2 Na2MoO4-2H2O CoCl2-6H20 CuCl2 0.1 mM 0.08 mM 92 µM 18 µM 1.54 µM 3..2 µM 0.34 µM 0.6 µM Solution II (100× for MA2) Solution III (1000×) Solution IV (250×) 500× A6 minor salts 31 2.2.4.1 Day of transformation The OD 750 nm of the culture was checked, and the heat block was set to 100 C. Fresh Polyethylene glycol (PEG) 4000 was prepared by dissolving 0.9 g of PEG and 750 µL of MA-I in a 2 mL tube at 42 C with occasional vortexing; this preparation should be initiated first. Subsequently, 1.5 mL of 1x MA-I was placed into a 42 C water bath then 50 mL of MA2G aliquoted into either a 50 or 100 mL graduated cylinder for each transformation. Following this, 6 µL of sonicated salmon sperm DNA was boiled per transformation for 5 minutes and then placed on ice. Once the cells achieved an OD 750 nm of 0.8-1.0, 40 mL of cells were spun down at 2000xg for 10 minutes, washed with 1 mL of warm MA-I buffer, transferred to an Eppendorf tube, spun down again at 2000xg for 1 minute, and removed all liquid. Approximately 100 µL of cells remained and were resuspended in about 100 µL of warm MA-I. I subjected the total final volume to 200 µL (200x concentrated), which was sufficient for 8 transformations (with 40 mL at OD 750 nm 1 equating to 20 x 10^7 cells/200 µL; 25 µL of cells per transformation results in 2.5 x 10^7 cells/transformation). While the cells were being spun, a mixture consisting of 1 pmol of linear DNA (PCR product with 0.5-1.5 Kb homology arms) + H2O to 84 µL + 10 µL 10x MA-I + 6 µL 10 mg/mL boiled sonicated salmon sperm DNA was prepared at room temperature in an Eppendorf tube. Then, 25 µL of cells and 125 µL of PEG were added to the DNA mixture, and mixed quickly by flicking the wrist to ensure that the PEG and the cells/DNA were thoroughly mixed. The cells form clumps upon contact with the PEG. Immediately, 1 mL of warm MA2G was added, and quickly diluted to 50 mL of warm MA2G in a graduated cylinder, and the culture was grown for 24 hours with continious light and 2% bubbling CO2 at 42 C. 32 2.2.4.2 Day after transformation The required amount of cornstarch (1.2 ml of 50% v/v) was washed three times with MA2G and then resuspended as a 20% v/v slurry in MA2G. Approximately 2.5 mL of this 20% slurry was needed per square plate (equating to 1 mL of 50% slurry/plate). The cornstarch slurry was placed in a sterile trough and spotted 16 µL x 144 spots on the square plate. One day (24 hours) after transformation, cells were spun down at 2000xg for 10 minutes, resuspended in 300 µL MA2G, and spotted 10 µL of cells onto cornstarch spots following a dilution guideline. Then, 10 µL of WT cells were spotted on a few spots around the plate as the nurse cell and single-wrapped the plated in grafting tape and incubated right side up with light and CO2 for 10-14 days. 2.2.4.3 Testing for homologous recombination (Colony PCR) Ten to fourteen days after plating the transformant, 48-96 colonies were picked to perform a Taq DNA Polymerase PCR to determine colonies with successful transformation and intron deletion (Table 4). The Taq DNA Polymerase with Standard TaqBuffer (New England Biolabs) and a Thermal Cycler (Bio-Rad) was used. Each reaction mixture contained 1X Standard Taq Reaction Buffer, 200 µM dNTPs, 0.5 µM of each primer, 1 µl of template DNA (either WT genomic DNA or colony suspension), and 1 U of Taq DNA Polymerase in a total volume of 20 µl. Thermocycling conditions consisted of an initial denaturation at 95 C for 5 min, 35 cycles of denaturation at 95 C for 20 sec, annealing for 20 sec, and extension at 68 C followed by a final extension at 68 C for 5 min. The annealing temperature and extension time were chosen based on the primer pairs and PCR product length (1min/kb). The PCR products were run alongside 100 bp/1kb DNA Ladders (New England Biolabs) on the agarose gel containing ethidium bromide at 120 V for 60 minutes and visualized on a FluorChem Q(ProteinSimple). 33 Then those positive colonies went under 5FOA selection and plated on MA2GU+5FOA for the second homologous recombination to lose the selection marker (URA 5.3). After 10-15 days, when colonies had grown, they were picked, extracted the genomic DNA, and performed PCR to identify the final positives (Table 5). 2.2.4.3.1 Quick genomic DNA isolation One to two OD Units (1 ODU: 1 ml cell with an OD 750 nm of 1) were spun down and frozen the pellet for 10 minutes at -80 C. Then, the pellet was resuspended in 200 µl of Edward buffer (200 mM Tris-HCl with pH 7.5, 250 mM NaCl, 25 mM EDTA, and 0.5% SDS), vortexed for 5 seconds, and added 200 µl of isopropanol and mixed by inversion. After centrifuging for 5 minutes at 13,000 rpm, the supernatant was decanted, and the tubes were air-dried. Then, 100 µl of H2O was added to the tubes and resuspended the pellet. Finally, the tubes were centrifuged for 1 minute at 13,000 rpm to sediment insoluble material63. After transferring the supernatant to the clean tubes, the DNA was diluted 1:10 or 1:100 with ddH2O for PCR. 34 Table 4. Primers and PCR product sizes of E034, D067, R289, and S262 for confirmation of intron deletion. Confirmation of intron deletion Genes of interests Primer Name Orientation and Sequence OSDR2181 Forward CATTCCGTGAAATGGGCCAG oSDR819 Reverse CGAGCAAAATCTTCCCATTGGACG 60.7 C Forward CATTCCGTGAAATGGGCCAG Reverse GCTTTGAAGCCGCTCTTGTACCCTGAC Forward GGTAGCAGTCACACCACAAG Reverse CCCCCACGAGGCGATTAAAA 68 C oSDR2165 Forward ACACGTCATCCGTTCGTTG 66 C FUB50 Reverse TCGATCGGAACCAAACACCA 67 C oSDR668 Forward GGCGATTGCTGAAGCCGCTGAGG 60.4 C oSDR669 Reverse 56.2 C E034 OSDR2181 oSDR1795 oSDR2189 D067 FUB75 R289 S262 Size of products (bp) Annealing temperature (Q5) 68 C GGCGATATGGTCCTGGTTACG 35 67 C WT ∆i 968 818 602 No band 738 653 803 733 66 C 68 C 489 249 Table 5. Primers and PCR product sizes of E034, D067, R289, and S262 for confirmation of URA marker deletion. Confirmation of URA marker deletion Genes of interests Primer Name Size of products (bp) Orientation and Sequence Annealing temperature (Q5) 68 C oSDR2181 Forward CATTCCGTGAAATGGGCCAG oSDR2182 Reverse TGACCGCGTCAAAACCAGTA 68 C oSDR2189 Forward GGTAGCAGTCACACCACAAG 66 C oSDR2190 Reverse GTCTGATGCTCGGTTGTCTT 66 C oSDR2165 Forward ACACGTCATCCGTTCGTTG 66 C oSDR2166 Reverse CTTGGGCGACTAGTGTCAATC 66 C oSDR1869 Forward TACCAAGGTTTGCCACCCAG 68 C oSDR1870 Reverse TGCTTCATACTTGTGAACGCTG 65 C E034 D067 R289 S262 WT ∆i 1785 1639 2022 1937 1886 1816 1940 1703 2.2.5 Sequencing Sequencing confirmed the successful deletion of those four introns. Sequencing of each gene was done by the UNBC genetic facility. Gene specific primers (Table 6) were used to sequence the exon-exon junctions, allowing for the assessment of accurate intron removal. The results of sequencing were analyzed by 4Peaks software (http://nucleobytes.com/index.php/4peaks). 36 Table 6. DNA oligonucleotides were used to amplify and sequence the desired gene. oSDR refers to the oligonucleotides in our lab which are identified as oligonucleotides by Stephen Rader. Gene oSDR# Sequence (5′ to 3′) S262 oSDR669 GGCGATATGGTCCTGGTTACG D067 oSDR817 GCGGTTCCCGTTATCGTCTGGTGGG E034 oSDR818 GAAGCATTCAACCTGTTCGATCGCG R289 oSDR858 GGCAACTGCCGCATGTGCTTGGTTG 37 2.2.6 Southern blotting To confirm the successful deletion of introns in C. merolae, Southern blotting was employed, in addition to sequencing, for its high specificity and sensitivity, allowing for the detection of specific DNA sequences within a complex mixture 64. This technique provided an additional layer of validation to the sequencing results by confirming the structural changes in the genomic DNA. By visualizing the DNA fragments, I was able to compare the sizes before and after the deletion, ensuring that the intron deletions occurred as expected. 2.2.6.1 Restriction Enzyme Digest Ten µg of genomic DNA was digested with the chosen restriction enzyme (RE) (New England Biolabs). Proper restriction enzymes were selected to produce bands that allow clear detection of both WT and modified DNA on a Southern blot, ensuring a detectable size difference between the two. The digestion was carried out for over 16 hours using 10 U per µg of each RE in a total volume from 100-400 µL depending on the DNA concentration. Then, to evaluate the digestion and determine the optimal running time for good separation of the ladder within the expected band size range (2-3 hours at 120V), 250 ng of each sample was run on a 0.7% agarose gel alongside 250 ng of uncut DNA. Then, the remaining digested DNA (9759 ng) was ethanolprecipitated with glycogen and resuspended in 9 µL of ddH2O and 1 µL of 10x loading buffer (0.1 M Tris-HCl, 0.2% SDS, 0.1% Bromophenol blue, 0.1% Xylene cyanole, and 30% Glycerol). 2.2.6.2 Probe Production and Labeling One of the ~500 bp homology arms used in the transformation plasmids was selected as the probe for each gene and PCR amplified using the high-fidelity Q5 polymerase. A total of 1 µg of each probe was required. After verifying the product on an agarose gel, a PCR cleanup kit and ethanol precipitation were used to obtain 1 µg of DNA in the maximum volume of 16 µL. Probes 38 were labeled using PCR DIG Probe Synthesis Kit, Cat # 11585614910, Roche) according to the manufacturer’s protocol. The features of probes and REs used for doing this southern blotting are shown in Table 7. Table 7. Features of Probes and Restriction Enzymes for Southern blotting. One of the primer pairs of homology arms of plasmids made for homologous recombination was chosen as a probe for each gene. The expected size, length, GC%, annealing temperature, and optimum temperature of probes along with Restriction enzymes and the expected size of their products are shown. Tm (C) Genes Probes (oSDR) Length (bp) GC% S262 1859+1860 521 54 73.1 D067 2183+2187 537 59 75.1 E034 2175+2179 524 56 74 Q270 2145+2143 603 55 73.3 C008 2249+2250 531 56 74 Q117 2154+2153 575 54 73 39 T optimum (C) RE WT/∆i (bp) 48.1-53.1 PstI 1694/1457 50.1-55.1 KpnI+XbaI 852/767 49-54 PstI 1765/4328 48.3-53.3 KpnI 2221/1956 48-53 PstI 1361/1146 48-53 PstI 1415/1159 2.2.6.3 Southern Gel and Transfer Since any residual ethanol in the DNA samples mentioned in 2.2.6.1 can cause the samples to float out of the wells, the tubes were placed in a 70 C heat block for 10 minutes to evaporate any remaining ethanol before running the southern gel. Then, samples (WT and two biological replicates of each gene), along with 5 µL of the DIG-labeled DNA MW marker VII mixed with 1 µL of 6X loading dye (0.1 M Tris-HCl, 0.2% SDS, 0.1% Bromophenol blue, 0.1% Xylene cyanole, and 30% Glycerol). were loaded onto a 0.7%, 0.75 cm thick agarose gel without ethidium bromide, in fresh buffer, and run at 120V for 3 hours. Then, the gel was rinsed with ddH2O and covered with 500 mL of denaturation buffer (43.85 g NaCl (1.5M), 10 g NaOH (0.5M), ddH2O up to 500 mL) and gently agitated for 30 minutes. Following this, the gel was rinsed with ddH2O again, covered with 500 mL of neutralization buffer (43.85 g NaCl (1.5M), 30.25 g Trizma base (0.5M), ddH2O up to 400 mL, pH adjusted to 7.5 with concentrated HCl), gently agitated for 30 minutes, and rinsed with ddH2O. Then samples were capillary transferred to the Hybond N+ Membrane (Amersham Hybond TM -N+, GE Healthcare) overnight (16-20 hours). 2.2.6.4 Hybridization The next day, the membrane was removed, UV crosslinked using the auto cross-link setting on the Stratalinker and rinsed in ddH2O. Then, 23.2 g of DIG Easy Hyb granules (CAT # 11585614910, Roche) were weighed out into a 50 mL conical tube, added 32 mL of ddH2O in two 16 mL aliquots, inverted, and mixed at 42 C until dissolved. The hybridization temperature was calculated using the formula: Tm= 49.82 + 0.41 (% GC) - (600/length in bp of probe); Optimum temperature (Topt)=Tm - 20 to 25 C. Each membrane was pre-hybridized for 30 minutes in the 5 mL hybridization buffer (for enhancing probe binding, reducing non-specific binding, and 40 stabilizing the probe and target DNA) at Topt in a hybridization oven. The appropriate amount of DIG-labeled DNA probe was denatured at 95 C for 5 minutes and then rapidly cooled on ice. About 25 ng/mL DIG Easy Hyb buffer was used, requiring 125 ng of probe per 5 mL. Then, the prehybridization solution was poured off, the pre-warmed hybridization buffer and denatured DIGlabeled DNA were added. Next, the mixture was incubated overnight with rotation. 2.2.6.5 Washes The membrane was briefly rinsed in approximately 5 mL of 2X SSC (Saline-Sodium Citrate), 0.1% SDS (Sodium Dodecyl Sulfate), and washed twice for 5 minutes each in approximately 20 mL of 2X SSC, 0.1% SDS at room temperature under constant rotation. This was followed by two washes of 15 minutes each in 0.5X SSC, 0.1% SDS at 65 C with rotation. 2.2.6.6 Immunological Detection All these steps were performed in a 37 C hybridization oven. The membrane briefly was rinsed for 1-5 minutes in 10 mL of washing buffer (0.1 M maleic acid, 0.15 M NaCl; pH 7.5 (20 C); 0.3% (v/v) Tween 20), incubated twice for 15 minutes each in 10-20 mL of blocking solution (prepared by diluting a 10× blocking solution (vial 6, Roche) 1:10 with maleic acid buffer). The membrane was then incubated for 30 minutes in 5-10 mL of antibody solution (Anti-DigoxigeninAP (vial 4, Roche) was centrifuged for 5 minutes at 10,000 rpm and then diluted 1:10,000 in fresh blocking solution). Next, the membrane was washed three times for 10 minutes each in 20 mL of washing buffer, followed by a single 5-minute wash in 20 mL of maleic acid buffer (0.1 M maleic acid, 0.15 M NaCl; adjusted to pH 7.5 with NaOH (20 C)). It was then equilibrated for 2-5 minutes in 10 mL of detection buffer (0.1 M Tris-HCl, 0.1 M NaCl, pH 9.5). Next, 1 mL of Chemiluminescent Substrate for Alkaline Phosphatase (CSPD) per 100 cm² of the membrane was applied and incubated for 5 minutes at room temperature, and 10-30 minutes 41 at 37 C to enhance the luminescent reaction. Finally, the membrane was exposed to the imager (ChemiDOC TM MP imaging system, Bio-Rad) for 5-30 minutes (chemiluminescent detection). In this study, the Southern blot for C008∆i and Q117∆i, which were made by other lab members was also performed. 42 2.2.7 Growth test at 42 C The growth rates were measured to determine whether the cells after removing introns grew faster or slower than the WT (T1 in my thesis). The growth rates of different C. merolae ∆i were analyzed by measuring their doubling times and comparing them with the WT (T1). For this experiment, each strain was grown initially in a cylinder in the incubator under standard conditions (42 C, 2% CO2, and continuous white light). For this initial growth period, a selective growth with 0.8 mg/ml 5-FOA was conducted. Once the cultures had grown sufficiently and appeared quite green, they were diluted to achieve approximately the same OD 750 nm (target OD 750 nm = 1-3) on the following day, when every single culture was diluted to an appropriate OD 750 nm (0.03-0.05) in 15-20 ml MA2GU. The OD 750 nm was measured and distributed 700800 µl aliquots into 9 wells of the 48-well plate. Each strain occupies 9 wells, allowing two strains with their duplicate biological replicates per 48-well plate with one well reserved for the blank (MA2GU). To minimize evaporation, I distributed ddH2O between the wells and into two external columns of the plate. Cells were grown in MA2GU liquid medium in the 48-well plates (NUNCLON, CAT 150687) shaken at 250 rpm under continuous white light illumination (135 μmol photons m−2 s−1) at 42 C65. The growth curves were recorded by measuring OD 750 nm values at a wavelength of 750 nm using the BioTek Synergy Neo2 plate reader for 72-hour time points. The Doubling time (DT) was calculated based on the slope of the growth curve during the exponential growth phase of the cells. Generally, our lab recorded that the doubling time for C. merolae T1 strain is approximately 12 hours under optimal conditions. so each strain had two biological replicates, with nine technical replicates to ensure accuracy and reproducibility. Additionally, for all intron deletion strains, there were two biological replicates. This means that for each growth test, there 43 were nine technical replicates for T1, and 18 replicates (two biological replicates, each with nine technical replicates) for each ∆i (Figure 8). Figure 8. Plate layout for doing the growth test in the 48-well plate. Each ∆i strain had two biological replicates (∆i.1 and ∆i.2), with nine technical replicates per biological replicate. This setup resulted in each ∆i strain occupying 18 wells on a 48-well plate, as indicated in green. One well was designated as a blank (MA2GU media in my experiment), and the remaining wells, along with the spaces between them, were filled out with ddH₂O to minimize evaporation. 44 2.2.8 Fitness Test To more thoroughly investigate the effect of introns on cell growth under starvation conditions, a competitive growth assay was conducted using a mixture of WT and ∆i cells. This assay was carried out in a rich medium (MA2GU), where the survival of both WT and ∆i cells were monitored during both the logarithmic (OD 750 nm <3) and saturation phases (OD 750 nm >30) of growth via cell counting under fluorescent microscopy. 2.2.8.1 T1 mTFP transformation To differentiate between WT and ∆i under microscopy, a plasmid containing the monomeric Teal Fluorescent Protein (mTFP) (Figure 9) was introduced, which was obtained from our collaborator Prof Kyle Lauersen at King Abdullah University of Science and Technology (KAUST), in Jeddah, into our WT (T1) of C. merolae. In that plasmid, mTFP has been linked to the H166C (DNA gyrase subunit B), which is located in the plastid66. Normally, C. merolae exhibits red autofluorescence from chlorophyll within the chloroplast 49,51,62. Through this transformation, the WT cells displayed both cyan and red fluorescence, whereas ∆i exhibited only the red fluorescence of the chloroplast. This modification allows for a clear distinction between the two strains during microscopy. The mTFP transformation was conducted following the Cm transformation protocol previously described, with the only difference being the use of MA2GU instead of MA2G, due to T1 lacking uridine synthase. 45 Figure 9. The map of the plasmid containing the mTFP. The linear map of the plasmid containing the mTFP that has been inserted into C. merolae. In this plasmid, mTFP, which is linked to H166C (DNA gyrase subunit B) and expressed using the pCPCC promoter, has been introduced between the D184C and D185C loci. Additionally, this plasmid confers chloramphenicol resistance due to the presence of chloramphenicol acetyltransferase (CAT) under the control of the pAPCC promoter. 2.2.8.2 Fitness test set up WT (i.e. T1 transformed with a plasmid containing the mTFP fluorescent protein), and two ∆i, R289∆i and ∆10i, were co-cultured in a series of batch cultures (Table 8 and Figure 10), which were performed in two biological replicates. I started the experiment with 50 ml co-cultures (with each type of cell being added in an equal volume of 25 ml) at the OD750 nm of 0.025. The cocultures were grown for 14 days, diluted back to OD 750 nm 0.025, grown for 14 days again, and so on, for a total of three cycles of growth. Two 100 ul samples were taken at each cycle, one after 2 days (during the logarithmic phase, when OD 750 nm < 3), and the other just prior to dilution (during the saturation phase, when the OD 750 nm >30) for counting. For microscopy, 10 µl of each culture was placed on a microscopic slide with a cover slip. After allowing the cells to settle 46 for 15 minutes, counting was commenced on five fields on each slide including the four corners and the center. Two channels for this microscopy were utilized: one for Texas Red to capture the red autofluorescence of chloroplasts of both WT and ∆i, with specific excitation and emission wavelengths of 586/96 and 603/15, and another for mTFP to detect the WT, employing an excitation wavelength of 420/50 nm and an emission wavelength of 470/92 nm. In this study, the ZEISS Fluorescent microscope, ZEN blue edition 5 image processing program was utilized. Table 8. Co-cultures setup. Culture Contents 1 WT. mTFP.3 + ∆10i 32.3 2 WT. mTFP.3 + ∆10i 37.22 3 WT. mTFP.5 + ∆10i 32.3 4 WT. mTFP.5 + ∆10i 37.22 5 WT. mTFP.3 + R289∆i. 26.3 6 WT. mTFP.3 + R289∆i. 14.14 7 WT. mTFP.5 + R289∆i. 26.3 8 WT. mTFP.5+ R289∆i. 14.14 47 Figure 10. Fitness test set up for co-culture of WT-∆10i and WT- R289∆i. The co-culture of WT-∆10i and WT- R289∆i with their biological replicates were grown in 50 ml cylinders with an equal volume of each strain (25 ml of each) at the OD750 nm of 0.025 in the incubator under standard conditions (2% CO2 with constant light at 42 C). The co-cultures were grown for 14 days, diluted back to OD 750 nm 0.025, grown for 14 days again, and so on, for a total of three cycles of growth . 48 2.3 Results and Discussion 2.3.1 Intron deletion In this study, four introns in C. merolae were deleted using homologous recombination. This allowed me to assess whether their deletion impaired possible functions associated with these introns. 2.3.1.1 Intron deletion by homologous recombination (HR) The first step to determine whether the maintenance of the small number of introns in C. merolae is due to their functional importance is to examine whether deleting C. merolae’s introns affects its growth. In this study, introns were removed from the C. merolae genome individually by means of homologous recombination (HR). HR refers to the exchange of genetic information between DNA molecules that share sequence homology and can happen at any site within a region of homology. A gene targeting technique makes use of this cellular process to exchange DNA between the genome and a defined exogenous repair DNA molecule (see Figure 6). The strategy allows precise modifications to be made to target genes 67. Because HR in Cm is not particularly efficient, the inclusion of a selectable marker on the modifying DNA molecule is essential in order to obtain correctly altered strains. Therefore, the DNA constructs for each intron deletion also contain the URA5.3 auxotrophic marker. The URA5.3 gene encodes a fused protein containing an orotate phosphoribosyl transferase (OPRTase) domain and an orotidine-5'-phosphate decarboxylase (OMPdecase) domain, which synthesizes uridine-5'-monophosphate from orotate. URA5.3 is introduced as the selection marker into the uracil-auxotrophic strain to select the transformant.68. Cm T1, a Ura5.3 complete deletion mutant strain was used as the host strain for this nuclear DNA transformation experiment. T1 cells cannot grow in the absence of exogenous uracil due to the absence of uridine synthase. After doing the transformation and first HR, a 49 secondary internal homologous recombination procedure was performed with 5-fluoroorotic acid to eliminate the URA5.3 gene69. When the URA5.3 gene is present in the C. merolae genome, the compound 5-Fluoroorotic acid (5-FOA) is converted to a toxic product resulting in cell death, but in the absence of this gene, 5-FOA is harmless to the cells. In this way, 5-FOA counterselection yields an intron-deleted strain without extra genomic material at the locus 70. Five introns were previously deleted by former students in the lab (K245, S311, R350 (2 introns), and C053). In this study, four additional introns were chosen to delete (S262, D067, E034, and R289) based on their host gene's elevated expression level and assessed the effect of their elimination on the cell growth rate. 2.3.1.2 Confirmation of DNA construct with two homology arms for intron deletion High-fidelity Q5 PCR generated a linear construct for transforming into C. merolae. The full-length transformation amplicon (PCR product) of pSR946 (S262C), 1013 (Q270C), 1014 (Q117C), 1015 (R289C), 1017 (E034C), and 1018 (D067C) is shown in Figure 11. 50 A B 51 C Figure 11. Confirmation of homology arm insertion in plasmids. Confirmation of plasmids with homology arms for A) pSR1013 and pSR104. B) pSR1015 and pSR1017. C) pSR1018 visualized with agarose gel electrophoresis. Plasmid names are listed above lanes with the expected size of the genes, which are listed below the lanes. 1 kb ladder NEB was used as the size marker. Table 5 shows the REs and the predicted sizes for each plasmid. 52 2.3.1.3 Confirmation of the full-length transformation amplicon. High-fidelity Q5 PCR generated a linear construct for transforming into C. merolae. The full-length transformation amplicon (AF PCR product) of pSR946, 1013, 1014, 1015, 1017, and 1018 is shown in Figure 12. A B 53 D C F E Figure 12. Confirmation of the full-length transformation amplicon. Determining the presence of the full-length transformation amplicon (AF PCR products) for A) pSR946 (4719 bp), B) pSR1013 (5778 bp), C) pSR1014 (4632 bp), D) pSR1015 (5013), E) pSR1017 (4876 bp), and F) pSR1018 (5402 bp) visualized with agarose gel electrophoresis. Plasmid names are listed above lanes with the expected size of the genes, which are listed below the lanes. 1 kb ladder NEB was used as the size marker. 54 2.3.1.4 Confirmation of successful transformation for making ∆i of S262, E034, D067, and R289 High-fidelity Q5 PCR on genomic DNA (g.DNA) confirmed successful transformations and intron deletion. (Figures 13-16). 55 A B Figure 13. Confirmation of intron deletion for S262. PCR confirmation of successful intron deletion for S262∆i. A) intron deletion and B) URA marker deletion shown by PCR from g.DNA for two positive colonies of S262∆I visualized with agarose gel electrophoresis. Gene names are listed above each lane with the expected size of the genes, which are listed below the lanes. 1 kb and 100 bp ladders NEB were used as the size markers. 56 A B C Figure 14. Confirmation of intron deletion for E034. Determining of successful intron deletion for E034∆i strain. A and B) intron deletion and C) URA marker deletion after g.DNA extraction of two positive colonies for E034∆i visualized with agarose gel electrophoresis. Gene names are listed above each lane with the expected size of the genes, which are listed below the lanes. 1 kb and 100 bp ladder NEB were used as the size markers. 57 A B Figure 15. Confirmation of intron deletion for D067. Determining of successful intron deletion for D067∆i. A) intron deletion and B) URA marker deletion after g.DNA extraction of two positive colonies for D067∆i visualized with agarose gel electrophoresis. Gene names are listed above each lane with the expected size of the genes, which are listed below the lanes. 1 kb and 100 bp ladders NEB were used as the size markers. 58 A B Figure 16. Confirmation of intron deletion for R289. Determining of successful intron deletion for R289∆i. A) intron deletion and B) URA marker deletion after g.DNA extraction of two positive colonies for R289∆i visualized with agarose gel electrophoresis. Gene names are listed above each lane with the expected size of the genes, which are listed below the lanes. 1 kb and 100 bp ladders NEB were used as the size markers. 59 2.3.2 Southern blotting Southern analysis confirmed that targeted introns were successfully deleted (Figure 17). It should be noted that the Southern blotting for R289∆i was not successful. B A 60 C D 61 E Figure 17. Confirmation of intron deletions by Southern blotting. Confirmation of intron deletions for A) D067∆i, B) S262∆i and C008∆i, C) Q270∆i, D) Q117∆i, and E034∆i utilizing Southern blotting. Gene names are listed above each lane with the expected size of the genes, which are listed below the lanes. The DNA MW ladder is indicated as well. 62 2.3.3 Growth Rate Analysis at 42 C A growth assay was conducted to evaluate the effects of intron deletion on cellular growth. Our lab previously relied on graduated cylinders and tube cultures for growth test assays. I developed an efficient plate assay utilizing 48-well plates, which significantly enhanced the simplicity and reproducibility of growth assays, surpassing the traditional methods of growing cells using cylinders or tube cultures previously employed in our lab. This advancement represented an important improvement in my experimental approach. This required optimizing several variables, including the shaking speed, minimizing evaporation, starting with the appropriate OD 750 nm to allow sufficient time for data collection before cell saturation, and adjusting the light intensity. The development of the plate assay significantly improved my ability to measure doubling times (DTs) for my strains as well as those created by other lab members. The growth test experiment involved comparing the growth rates of ∆i cells with that of the T1 reference, which is the parental strain from which all ∆I strains were generated (T1 lacks the URA5.3 gene and is therefore not exactly wild type). Additionally, to assess the contribution of introns to cell function, our lab initiated large-scale intron deletions in C. merolae with the ultimate goal of creating an intron-free model eukaryote. I conducted growth tests on two of these strains as well, ∆6i, a strain with six (R350(2), S311, C052, S262, and Q270), and ∆10i, a strain with 10 (R350(2), S311, C052, S262, Q270, D067, C008, Q117, and O094) introns deleted respectively (Table 9) to assess the impact of these changes on cellular functions and to evaluate the growth rates of cells lacking multiple introns. The comparison of the DT the ∆i strains with T1, along with their corresponding p-values are shown in Table 9 and Figure 18. 63 Table 9. Growth test results for intron deletion strains 42 C. ∆i1 and ∆i2 represent two biological replicates of each ∆i strain. DTs are doubling times in hours with standard deviation. Column 2 is the average DT of the two ∆I strains. Column 5 presents the results of a one-way ANOVA with multiple comparisons using Tukey’s range test comparing the DT of T1 with the average DT of ∆i1 and ∆i2. Column 6 shows one-way ANOVA with multiple comparisons using Tukey’s range test analysis comparing the DTs of ∆i1 and ∆i2 to assess if there is a change in growth rates between the two replicates. Column 7 summarizes the overall comparison of growth rates between T1 and ∆is. Columns 8 and 9 represent the lag time of ∆i1 and ∆i2. Strains DT (h) DT ∆i1(h) DT ∆i2(h) P value T1v∆i P value ∆i1v∆i2 ∆i growth relative to T1 R289∆i 10.6 10.6±0.01 10.6±0.00 1.8Î10-4 0.61 Faster K260∆i 10.8 10.3±0.02 11.2±0.01 0.32 0.04 Equivalent S342∆i 11.0 10.9±0.02 11.2±0.01 0.65 0.65 Equivalent Q382∆i 11.1 11.2±0.01 11.1±0.01 0.16 0.9 Equivalent S270∆i 11.3 11.3±0.01 11.2±0.01 0.24 0.83 Equivalent C008∆i 11.4 11.9±0.01 11.0±0.01 0.05 0.01 Equivalent T1 11.6±0.02 Q117∆i 11.7 11.7±0.03 11.8±0.01 0.2 0.38 Equivalent D067∆i 11.8 12.2±0.02 11.3±0.00 0.13 0.14 Equivalent E034∆i 12.0 12.5±0.01 11.6±0.02 0.01 0.02 Slower J129∆i 12.0 11.2±0.01 12.6±0.03 1.7Î10-4 1.5Î10-4 Slower S262∆i 12.6 12.1±0.02 13.0±0.02 2.3Î10-4 0.02 Slower Q270∆i 13.0 13.0±0.00 13.0±0.00 0.03 0.95 Slower ∆6i 13.1 12.8±0.01 13.3±0.02 7Î10-6 0.11 ∆10i 14.2 14.2±0.01 14.2±0.01 1.3Î10-5 0.38 Lag ∆i1 (h) 8 Lag ∆i2 (h) 6 12 7 12 12 2 6 0 0 12 14 10 64 Slower Slower 17 17 0 2 12 4 6 7 8 8 14 14 8 8 16 14 Figure 18. Comparison of doubling times between T1 (WT) and ∆i strains. Doubling times of T1 (labeled WT) and the average DT of two biological replicates. Each strain has nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test was performed. The numbers on the right side of the figure represent the p-values. Whiskers show the standard deviation. 65 The DT for T1 was 11.6 hours under normal growth conditions. R289∆i exhibited a faster growth rate than T1, with a doubling time of 10.6 hours and p-value of 1.8Î10-4. Several ∆is demonstrated slower growth rates compared to T1, including J129∆i, E034∆i, S262∆i, Q270∆i, ∆6i, and ∆10i, with DTs of 12, 12, 12.6, 13, 13.1, and 14 hours, respectively. Strains such as K260∆i, S342∆i, Q382∆i, S270∆i, C008∆i, Q117∆i, and D067∆i, showed doubling times that were not significantly different from T1 (p > .05). The growth curves for three ∆i strains with representative growth are shown in Figures 1921. R289∆i was the only ∆i to demonstrate faster growth. ∆10i exhibited the slowest growth. D067∆i showed no change from T1. Measurements across biological and technical replicates were highly reproducible. 66 Figure 19.Comparison of R289∆i growth to T1 at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and R289∆i (gray and orange for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. Whiskers show the standard deviation. A oneway ANOVA with multiple comparisons using Tukey’s range test with a p-value of 1.83Î10-4 showed that R289∆i grew faster than T1. 67 Figure 20. Comparison of ∆10i growth to T1 at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and ∆10i (gray and orange for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. Whisker shows the standard deviation. A oneway ANOVA with multiple comparisons using Tukey’s range test with a p-value of 1.3Î10-5 showed that Δ10i grew slower than T1. 68 Figure 21. Comparison of D067∆i growth to T1 at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and D0670∆i (gray and orange for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. Whiskers show the standard deviation. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.13 showed that D067∆i grew insignificantly different from T1. 69 Half of the introns tested, including those in genes K260, S342, Q382, S270, C008, Q117, and D067, can be removed with little impact on growth under normal conditions. The growth comparison between T1 and D067∆i, showcasing an example of a ∆i that exhibited no change in growth compared to T1, is shown in Figure 21. Conversely, several ∆is displayed slower growth rates compared to the T1 including J129∆i, E034∆i, S262∆i, Q270∆i, ∆6i, and ∆10i (Figure 20) showing increased DTs of 12, 12, 12.6, 13, 13.1, and 14 hours respectively. While all of these are statistically significant, it is unclear whether the difference between 11.6 and 12 h is biologically meaningful, but certainly a strain with a 14 h DT grows noticeably more slowly than the WT. The decrease in growth rates (p-values < 0.05) indicates that these introns contribute to optimal cellular function and growth under standard conditions. It is important to note that the growth test is a phenotypic assessment and does not elucidate the precise functions and effects of the deleted introns. This test primarily indicates that the deleted introns play measurable roles in cellular function, as evidenced by the observed impact on growth when these introns are removed. The fact that only a few introns in C. merolae are important for normal growth raises the question of why the remaining introns have been retained. It should be noted that the growth test results presented in Table 9 indicate that, in certain ∆i strains (including K260∆i, C008∆i, J129∆i, E034∆i, and S262∆i), a statistically significant difference in DT was observed between two biological replicates (p < 0.05), although the magnitude of the differences was not large. This may indicate that the replicates are growing differently, despite Southern blotting confirming that these biological replicates are genetically identical. The observed differences may indicate the emergence of mutations during the homologous recombination process, meaning these strains may no longer qualify as true biological replicates. Consequently, 70 it is recommended that the transformation for these ∆i strains be repeated to ensure true biological replicates and to determine which growth rate is the correct one for each strain. Additionally, this study observed slower growth rates of the Δ6i and Δ10i with a p-value of 0.03, with six and ten intron deletions respectively. While it is tempting to attribute this solely to the loss of specific intron functions, cumulative effects likely play a significant role. This pattern, where Δ10i grows slower than Δ6i, which in turn grows slower than the WT, underscores how the number of introns deleted can impact cell physiology and growth dynamics. Introns frequently harbor regulatory elements, such as enhancers and silencers, that modulate gene expression at both the local level (affecting the host gene) and the global level (influencing other genes within the genome)71. Deletion of multiple introns can disrupt regulatory networks, resulting in gene misregulation. The extent of disruption correlates with the number of introns deleted, which may account for the observed slower growth in the Δ10i compared to the Δ6i. Furthermore, in other organisms introns facilitate the export of mRNA from the nucleus to the cytoplasm and contribute to mRNA stability72. The loss of multiple introns may compromise these processes, resulting in decreased protein synthesis or increased mRNA degradation, further impairing cellular function and vitality. Additionally, each intron deletion may cause subtle perturbations in cellular metabolism. While a single deletion might have a minimal effect, the cumulative impact of multiple deletions can impose a significant metabolic burden. This burden can lead to reduced energy production and efficiency, which is critical for an organism like C. merolae which relies on efficient metabolic processes to thrive in high-stress environments. In addition to determining the DT of strains, which reflects their exponential growth, the 42 C growth assay also allowed me to measure the lag phase of the strains. As shown in Table 9, 71 intron removal altered the duration of the lag phase in varying ways. While strain T1 exhibited a lag time of 10 hours, this period varies among ∆i strains, ranging from 0 to 17 hours. Lag time was calculated by first plotting the exponential growth of the strains. A horizontal line was then drawn from the Y-axis (lnOD) to intersect the exponential phase trendline, and the corresponding value on the X-axis was used to determine the lag time. Figures 22-24 illustrate the lag phase for different strains. T1 had a lag time of 10 hours, Q117∆i a lag time of 17 hours, and D067∆i exhibiting no detectable lag phase. Figure 22. Lag time of T1 at 42 C The scatter plot with a linear trendline shows the exponential growth of T1 (orange). A horizontal line (blue) was drawn from the Y-axis (lnOD) to intersect the exponential phase trendline, and the corresponding value on the X-axis (10 h) represents the lag time. 72 A B Figure 23. Lag time of S270∆i at 42 C The scatter plots with a linear trendlines show the exponential growth of S270∆i.1 (A) and S270∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (0 h) represent the lag time. 73 A B Figure 24. Lag time of Q117∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of Q117∆i.1 (A) and Q117∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (17 h) represent the lag time. 74 2.3.4 Fitness test In S. cerevisiae, it has been observed that cell growth with an intron deletion is generally impaired under nutrient-depleted conditions. This effect of introns on growth is not related to the expression of the host gene and persists even when the translation of the host mRNA is blocked 73. To test the effect of nutrition-depleted conditions on the intron-deleted C. merolae strains and detect the consequences of growing cells in a competitive environment, I conducted a competitive growth test using ∆10i and R289∆i. The reason for choosing these two was that in a previous growth test performed at 42 C, R289∆i was the only ∆i that grew faster than the parent strain with all introns present (T1), and ∆10i had the slowest growth in comparison. These two ∆is exhibited faster and slower growth, respectively (see Table 9 and Figure 18). In this experiment, I selected them to assess potential differences in their growth between the logarithmic phase, characterized by rich nutritional conditions, and the saturation phase, where competition for limited resources occurs, and to determine which strain, WT or ∆i, would outcompete the other under conditions of nutritional stress. 2.3.4.1 T1 and T1::mTFP growth test at 42 C In order to differentiate WT (T1) cells from ∆i cells, Teal Fluorescent Protein (mTFP) was transformed into the T1 (referred to below as T1::mTFP). Prior to using it, a growth test was conducted to compare T1::mTFP with T1 to ensure that the presence of the fluorescent protein had not affected the growth rate of T1. This test, performed under the same conditions as other growth tests (42 C in a 48-well plate), showed no change between the growth rates of T1 and T1::mTFP with DTs of 11.6 and 11.7 respectively (P=0.07, Figure 25). 75 Figure 25. 42 C growth assay for T1::mTFP compared with T1. The scatter plot with exponential trendlines presented on a logarithmic scale shows the growth of T1 (blue) and T1::mTFP (gray and orange). Whiskers show the standard deviation. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.07 shows that there is no growth rate difference between T1 and T1::mTFP. 2.3.4.2 Fitness test for ∆10i and R289∆i WT (T1::mTFP) and ∆is were co-cultured in a series of batch cultures, along with their biological replicates, as described in the Materials and Methods section. Samples were collected during the logarithmic phase (OD 750 nm < 3) and saturation phase (OD 750 nm > 30) across a total of three cycles. The fraction of WT to total cells was calculated by averaging the cell counts from five fields of view on the microscopic slide. WT cells were identified by both mTFP and 76 Texas Red fluorescence, while the total cell count was measured by Texas Red fluorescence alone because of the red auto-fluorescent of the chloroplast. Then, the WT fraction in each phase of growth was determined by dividing the average number of WT cells by the average number of total cells. The microscopy images regarding the fitness test are shown in Figure 26. 77 A B C D Figure 26. Microscopy images for the fitness test. A) and B) Fluorescence micrographs of the co-cultures of WT-R289∆i and WT-∆10i, respectively. In these images, the yellow cells represent WT (T1::mTFP), characterized by both red autofluorescence and cyan fluorescence, whereas the red cells are ∆i, marked by only red autofluorescence, as indicated by white arrows. C) and D) Cells in the logarithmic and saturation phases, respectively. These images highlight that chloroplasts maintain a well-defined round shape during the logarithmic phase but exhibit various shapes in the saturation phase. 78 In this study, WT cells exhibited a lower fraction in the WT-R289∆i co-culture, with average fractions of 0.35 and 0.33 in the logarithmic and saturation phases, respectively (Table 10). As indicated in Figure 24A, there is no significant change in this fraction between the logarithmic and saturation phases, with a P-value of 0.59. The distribution of WT and R289∆i in this co-culture aligns with the growth test results at 42 C, where R289∆i demonstrated faster growth than WT (see Table 9 and Figure 18). In WT-∆10i co-cultures, WT cells show average fractions of 0.76 and 0.92 for the logarithmic and saturation phases, respectively (Table 10). Based on Figure 27A, this is a significant change in the WT fraction in this co-culture between the logarithmic and saturation phases of growth, with a P-value of 0.01. According to Figure 27B, following inoculation, WT cells initially displayed a higher growth rate. However, after the first dilution, the fraction of WT cells decreases, indicating that ∆10i cells are outcompeting them in growth. After the final dilution, the WT fraction increased again. While ∆10i cells occasionally exhibited a higher growth rate than WT cells, the overall trend favors faster growth of WT cells. The most pronounced increase in WT cell fraction occurred immediately after inoculation, and subsequent differences in growth rates between WT and ∆10i are less evident, complicating a straightforward interpretation of these results. 79 Table 10. Distribution of WT and ∆is in the co-cultures in the logarithmic and saturation phases. In the WT-∆10i co-culture, WT cells show average fractions of 0.76 and 0.92 for the logarithmic and saturation phases, respectively. In the WT-R289∆i co-culture, WT cells showed a lower fraction with average fractions of 0.35 and 0.33 in the logarithmic and saturation phases, respectively. Cell type Growth Phase Average fraction of WT Average fraction of ∆i WT-∆10i Logarithmic 0.76 0.23 WT-∆10i Saturation 0.92 0.07 WT-R289∆i Logarithmic 0.35 0.64 WT-R289∆i Saturation 0.32 0.67 80 A B Figure 27. Fitness test comparing the fraction of WT in the co-cultures for ∆10i and R289∆i. A) Average fraction of WT in co-cultures. No significant change in the WT fraction between the logarithmic and saturation phases in the WT- R289∆i co-culture, with a P-value of 0.59. There is a significant difference in the WT fraction in the WT-∆10i co-culture between the logarithmic and saturation phases of growth, with a P-value of 0.01. B) The fraction of wild type in each co-culture, measured before each dilution and two days after each dilution, is plotted. Lines are colored to indicate periods of saturated growth (orange), returning to and exiting rapid growth (blue), and initial post-inoculation (black). Dashed vertical lines indicate the dilution of cultures. 81 Through a comprehensive deletion of all 295 introns from S. cerevisiae, it was determined that only a minimal subset (five introns) is essential for growth in nutrient-rich media74. Conversely, under nutrient-depleted conditions, the majority of intron-deleted cells exhibited compromised growth. This phenomenon appeared to be independent of the function of the protein encoded by the host gene. Instead of modulating host gene expression, introns enhance resistance to starvation by repressing ribosomal protein genes, which are downstream targets of the nutrientsensing TORC1 and PKA pathways74. In this study, a competitive growth assay was developed to elucidate the significance of C. merolae introns in cellular maintenance under nutritional stress. Previous works have demonstrated that a competitive growth assay is a powerful tool for detecting slight differences in fitness74–76, but as this was the first use in C. merolae, the ∆10i and R289∆i were selected based on prior growth assessments at 42 C, which revealed that R289∆i was the only ∆i to exhibit faster growth than the WT, whereas ∆10i demonstrated the slowest growth among all strains. The objective was to investigate the differential growth patterns between the logarithmic phase, characterized by abundant nutrients, and the saturation phase, marked by competition for limited resources, to ascertain whether introns play a role in adaptation to nutrient depletion. No change between the two growth phases was observed for R289∆i in this experiment. This could be because a single intron does not make a measurable contribution to starvation adaptation by itself, or because different introns have different functions and not all of them work in the same way. Additionally, the results indicated that ∆10i, which previously exhibited a larger DT compared to WT in nutrient-rich media (see Table 9 and Figure 18), occasionally exhibited a higher growth rate than WT cells in the co-culture. However, the overall trend favors faster growth of WT cells, which may imply a role for the ten deleted introns in the ∆10i’s ability to withstand 82 starvation. They can act as enhancers of gene repression during starvation. Similarly, in yeast, it has been shown that introns support growth under starvation conditions by promoting the repression of ribosomal protein genes75. However, considering these results, it is not possible to draw a clear conclusion about the results of this experiment. In this experiment, both R289∆i and ∆10i strains survived during the saturation phase, contradicting my hypothesis that the deleted introns are important for cell survival under nutrient starvation. If the introns were important for survival under nutrient deprivation, strains lacking these introns would not have survived under these conditions. 83 2.4 Conclusion In this chapter, four C. merolae‘s introns were deleted utilizing HR and explored their functional significance by assessing the growth impact of deleting specific introns under both normal and nutritional stress conditions. Half of the introns in C. merolae can be deleted without impacting growth under normal conditions, however, R289∆i, S262∆i, Q270∆i, J129∆i, E034∆i, Δ6i, and Δ10i showed different DTs than WT. The changes in growth rates (p-values < 0.05) indicate that these introns likely contribute to optimal cellular function and growth under standard conditions, albeit in different directions: removal of the R289 intron makes the cells grow faster, suggesting the intron is deleterious. This is consistent with the view that introns pose a metabolic burden to the cells, requiring energy for their maintenance and splicing. Conversely, the other ∆is grew more slowly, arguing that their introns provide a positive benefit to the cells that more than counterbalances the metabolic cost. Subsequently, a fitness test was conducted under starvation conditions for R289∆i and ∆10i, which highlighted the importance of introns in stress resilience. No change was observed between logarithmic and saturation phases for R289∆i in this experiment. Additionally, ∆10i (the strain with ten deleted introns), which exhibited slower growth compared to WT in rich media at 42 C in the plate growth assay, did not show a consistent growth pattern in mixed culture with WT. 84 CHAPTER III: Discovery of polyadenylated sisRNAs in Cyanidioschyzon merolae 85 3.1 Introduction In higher eukaryotes, most protein-coding genes contain one or more non-coding introns within their coding sequences. These introns are spliced out of the primary transcript, usually forming a lariat structure where the 5′ end is connected to the 3′ acceptor splice site. After being debranched into a linear form, these introns are typically degraded quickly77. Although often considered 'junk' sequences, some introns have significant roles, such as encoding small Cajal body-associated RNAs (scaRNAs), small nucleolar RNAs (snoRNAs), and microRNAs (miRNAs), which are involved in various cellular functions46. Recently, a new class of ncRNA known as stable intronic sequence RNAs (sisRNAs) has been characterized within introns. It is worth mentioning that the term "sisRNA" merely identifies the origin of the RNA and does not constitute a functional category. Thus, once the function of a sisRNA is understood, it no longer fits the classification of a sisRNA. sisRNAs play some functions in organisms such as mouse, viruses, Drosophila, and Arabidopsis. Unlike most RNAs, sisRNAs are surprisingly stable, avoiding the rapid degradation common to many RNAs46. They are often polyadenylated and may include exonic sequences, 5′ caps, and polyA tails. Although our understanding of sisRNAs is still limited, some have been found to be involved in the regulation of gene expression via various mechanisms46. sisRNAs can be generated either by a splicing-dependent or splicing-independent mechanism (Figure 2846). In the splicing-dependent mechanism, sisRNAs are transcribed as part of pre-mRNA transcripts and are later processed into linear or circular RNAs through splicing and trimming events. Circular RNAs can also form through back splicing, where the 5′ end of the upstream exon is covalently bonded to the 3′ end of the downstream exon, trapping the sisRNA in between. In the splicing- 86 independent mechanism, sisRNAs are transcribed from intronic or exonic promoters and terminate at polyadenylation sites46. 87 A B Figure 28. Biogenesis of Stable Intronic Sequence RNAs (sisRNAs). (A) Splicing-dependent mechanism: intronic lariats from pre-mRNA splicing form linear or circular sisRNAs. Some linear sisRNAs can re-circularize, and exon back-splicing can generate circular sisRNAs with both exonic and intronic sequences. (B) Splicing-independent mechanism: sisRNAs are transcribed from promoters in host genes or introns, producing polyadenylated sisRNA transcripts with exonic sequences. UV irradiation promotes widespread polyadenylation and the use of alternative last exons, generating additional sisRNAs 46. This Figure was obtained from “Stable Intronic Sequence RNAs (sisRNAs): An Expanding Universe”, by Seow Neng Chan and Jun Wei Pek with permission. 88 3.1.1 sisRNA function It has been shown that in Drosophila melanogaster, several linear sisRNAs are known to downregulate gene expression78,79. These sisRNAs share similar predicted secondary structures, each containing an exposed 3’ tail. It is hypothesized that the exposed 3’ tail plays a crucial role as a guide sequence to locate its target through complementary base-pairing, leading to target degradation78,79 . Conversely, some sisRNAs, such as circular intronic RNAs (ciRNAs), have been reported to enhance the transcription of their host genes46,80 . These ciRNAs accumulate at the transcription start site of the gene locus and colocalize with the RNA polymerase II (Pol II) complex, presumably to enhance the transcriptional rate of the gene. sisRNAs also may activate an enhancer in the intronic region, forming a positive feedback loop that boosts gene transcription46,80 . Besides binding to RNA and DNA, sisRNAs can also interact with proteins46. Among other things, they have been shown to regulate the activity of RNA-binding proteins by sequestering them from their usual functions. This regulation can profoundly affect gene expression, as RNA-binding proteins are often involved in a wide range of RNA processing activities46. Although sisRNAs have been identified in various organisms, very little is known about their functional significance. It has been demonstrated that the accumulation of intronic lariats is detrimental to cells. This observation suggests that introns that remain relatively stable postsplicing may not merely be artifacts, as such stability would have been naturally selected against throughout the course of evolution81. In C. merolae with only 39 introns, long-read transcriptomic data showed that 19 introns contain sisRNAs at 42 C in total RNA samples (i.e. polyadenylated + unpolyadenylated RNAs), but notably not in polyadenylated libraries, indicating that the sisRNAs are not polyadenylated at 42 C54. Additionally, short-read transcriptomic analysis from 89 polyadenylation-selected RNA (polyA+) libraries of samples subjected to heat stress at 57 C revealed that those introns had accumulated peaks closely matching the location of sisRNA sequences identified in the long-read sequencing data54. These particular introns may be important for rescuing the cells under heat stress conditions. K142T, for instance, contains a distinct region within the intron that stands out in the RNA-seq data. This region has been identified based on sequence alignments as the non-coding RNase MRP. The MRP RNA is the scaffold of a conserved eukaryotic ribonucleoprotein complex that is essential in precursor ribosomal RNA (pre-rRNA) processing and cell cycle regulation. This complex is involved in the cleavage of 5.8S rRNA82. Although MRP is a known ncRNA and not classified as a sisRNA, it is included in this analysis due to its significant changes in response to heat stress and its classification as an intronically encoded ncRNA. Long-read and short-read transcriptomic data of C. merolae's sisRNAs, respectively, are illustrated in Figures 29 and 30. AIM I: Validate transcriptomic observation of sisRNA and their upregulation under heat stress for J129, K142, Q270C, and K260. AIM II: Test the hypothesis that certain sisRNAs (J129, Q270C, and K260) help cells adapt to heat stress. HYPOTHESIS I: Northern blotting and RT-q PCR will confirm the presence of J129, K142, Q270C, and K260 sisRNAs in C. merolae. HYPOTHESIS II: sisRNAs in J129, Q270C, and K260 are involved in adaptation to heat stress in C. merolae, and their elimination from the genome would lead to impaired cell growth. 90 Long-read transcriptomic data of C. merolae's sisRNAs, is illustrated in Figure 29. A) J129 B) K142 91 C) Q270 D) K260 92 E) E034 Figure 29. long-read transcriptomic data of C. merolae sisRNAs. Unpublished long-read transcriptomic data of C. merolae's sisRNAs for four sisRNA-containing genes. A) J129, B) K142, C) Q270, D) K260), and one control gene without a sisRNA (E) E034). The dashed lines in the gene model at the top of each panel represent the intron in the transcripts. The comparison of the intronic regions of each gene (highlighted within the red box) between the top trace, representing polyA+ RNA, and the bottom trace, representing total RNA, reveals the presence of sisRNAs at 42 C in the total RNA. Viktor Slat, unpublished data; reproduced with permission54. 93 Short-read transcriptomic data of C. merolae's sisRNAs, is illustrated in Figure 30. A) J129 B) K142 94 C) Q270 D) K260 95 E) E034 Figure 30. Short-read transcriptomic data of C. morolae sisRNAs. Unpublished short-read transcriptomic data for four sisRNA-containing genes. A) J129, B) K142, C) Q270, D) K260), illustrates upregulation of intron reads at heat stress, and one control gene without a sisRNA E) E034). The orange graphs display the read depth at 42 C (optimal growth temperature for C. merolae), dark purples are 55 C for one hour, and light purples are 55 C for 24 hours. Viktor Slat, unpublished data; reproduced with permission 54. 96 3.2 Materials and Methods Note: the work described in this chapter was carried out in collaboration with my labmates Viktor Slat, who did the initial bioinformatic work revealing the presence of sisRNAs, and Patrick Geertz, who developed the qPCR-based assay for polyadenylation. 3.2.1 C. merolae culturing The 10D C. merolae strain (NIES-1332), obtained from the Microbial Culture Collection at the National Institute for Environmental Studies in Tsukuba, Japan (mcc.nies.go.jp/) was cultured, in MA2G media in glass graduated cylinders. Six biological replicates were incubated at 42 C with 2% CO2 bubbled directly into the tubes using an air pump and illuminated with 90 μmol photons m−2 s−1 of light from a Verilux Happy Light for 24 hours. Cells were collected during logphase growth at an OD750 nm of 1.5-2, then aliquoted into 1.5 mL microcentrifuge tubes with 2 OD units in each. Then, two sets of three technical replicates were placed in a water bath at 42 C and 57 C for one hour. After incubation, cells were centrifuged at 16,100 x g for 1 minute, snapfroze with liquid nitrogen to inactivate RNase activity, and the pellets were then stored at -80 C. 3.2.2 RNA extraction Pellets resuspended in cold phenol lysis buffer (0.5 M NaCl, 0.2 M Tris HCl (pH 7.5), 0.01M EDTA, 1 % SDS) to lyse the cells. The cell lysate was sonicated two times, in ten-second bursts, at 2-3 Watts to shear genomic DNA and break down membranes. Three acid phenol/chloroform (pH 4.5) extractions were performed (centrifuged at 15000 x g for one minute), followed by two chloroform extractions (centrifuged at 16100 x g for one minute) to extract total RNA. Then, samples were ethanol-precipitated and washed in 70 % ethanol. RNA was resuspended in 15 µl ddH2O and stored at -80 C for further use. To check the quality of the RNA, 97 500 ng of the RNA was loaded on a 1.5% agarose/1% bleach gel to ensure it was not degraded. Two bands corresponding to the 28S rRNA and two bands corresponding to the 18S rRNA should be observed if it is not degraded. The upper band of the 28S rRNA should display approximately twice the intensity of the upper band of the 18S rRNA. 3.2.3 Fluorescent Northern Blot Northern blotting was performed to validate transcriptomic observation regarding the presence of sisRNAs in K142T, Q270C, J129, and K260C genes and their accumulation under heat stress conditions. Two control genes were for this experiment, E034C, with no sisRNA feature or accumulation as the negative sisRNA control, and J101C, which encodes a member of the Hsp20/alpha-crystallin family of proteins, as a Heat-Shock inducible gene control. The principle of the process of fluorescent Northern blotting is shown in Figure 31. 98 Figure 31. Fluorescent Northern blotting. Following the pre-culturing of the C. merolae WT in MA2G medium and subsequent incubation in a water bath at 42 C and 57 C for one hour, RNA was extracted. The RNA molecules were then separated and detected using the Northern blotting technique. 99 For this method, RNA was detected indirectly using 5' biotinylated oligos (probes), which were subsequently bound by streptavidin conjugated to a near-infrared dye 88 (IRDye 800CW). These probes were designed to be 26–45 nucleotides in length, with a melting temperature (Tm) of 78–90 C and GC content of 47–62%. These probe sequences were subjected to a Nucleotide BLAST against the C. merolae genome to ensure specificity for the target gene. As these probes were designed to bind sisRNAs within introns, they can potentially recognize not only the specific sisRNAs but also their corresponding coding introns and the pre-mRNA containing these introns. Consequently, mRNA that has lost the intron cannot be detected using these sisRNA-specific probes. For the E034 gene, which lacks a corresponding sisRNA, the probe was designed to target a region within the intron. Yeast snRNAs (YsnRNAs) were utilized as a molecular size ladder to determine the size of C. merolae sisRNAs in this study. S. cerevisiae (YSR 309 stored at -80 C) was cultured in yeast extract peptone dextrose (YPD) media, followed by RNA extraction. Specific probes were designed targeting its snRNAs for this purpose (Table 11). In this Table, SDR refers to the oligonucleotides in our lab which are identified as oligonucleotides by Stephen Rader. 100 Table 11. DNA oligonucleotides were used regarding the fluorescent and agarose Northern for the desired genes. The names, sequences, and target regions of probes are provided. It should be noted that all of these probes have reverse orientation. oSDR # Gene Region Sequence (5′ to 3′) 2520 J129C sisRNA AGCTAGCGGTTGCGGATTCCGGAGTAC 2479 K142C MRP ACTCCCGCTCTTATTGGGGTTCTGTGGTT 2478 Q270C sisRNA CTTCAGGATTTCATCGGAGGGAGGCG 2599 K260C sisRNA GTGGCAGCCTCATCAAGGTGGACTGTGGTAC 2600 E034C Intron CTCCACCAGGCCCCTTGGTTAGTTTC 2536 J101C - TTCGCGACTGTTCGCATTCGCTTGCATGAGGCTCGTTCGT 2532 J129C Exon GTAGGGTATCATGTCATCGAGAACGACACCGGCTGCGAGC 2534 Q270C Exon CGCGACCAACCTTGTCCATAGCCTCTGCAATGAGTTCGCC 2535 K142C Exon GTGGATAACATGCCTCATCGCTATGCGCGGCGTCAAGCTG 2546 Sc U1snRNA U1 GGCCCCAGCTCCCCTAACACCAATTTGAATTTGGTGTCAAAC 2547 Sc U4snRNA U4 AGCGAACACCGAATTGACCATGAGGAGACGGTCTGG 2548 Sc U5 snRNA U5 GCAAGCCCACAGTAACGGACAGCTTTACCTGTTTCTATGGAG 2549 Sc U6 snRNA U6 CGGTTCATCCTTATGCAGGGGAACTGCTGATCATCTCTGTATTG 2550 Sc U2snRNA U2 TCCCGCGTTGGACATAAACGGCTCGGAAAGACAGGGAAG 101 3.2.3.1 Denaturing Polyacrylamide Northern blotting 3.2.3.1.1 Preparing 6% Urea Denaturing Polyacrylamide Gel A 15 mL 6%/7M Urea denaturing Polyacrylamide Gel (2.25 mL of 40% acrylamide (19:1), 6.3 g of urea, 750 µL of 20x TBE, and 7.3 mL of H2O) was prepared. Then, freshly prepared 10% Ammonium Persulfate (150 µL) and 15 µL of tetramethyl ethylene diamine (TEMED) were added and the gel was poured using a 10 mL disposable pipette. A 6% denaturing polyacrylamide gel was poured for RNAs with less than 1000 nucleotides and pre-ran for 15 minutes at 400 V in 1x TBE to remove urea from the wells, equilibrate the gel, and ensure uniform ion distribution). Then, equal volumes of RNA (1-10 µg) with 2x formamide loading buffer were mixed, keeping the total volume under 20 µL. Next, samples were denatured at 65 C for 3 minutes immediately before loading on the gel, quickly spun down, loaded using an elongated gel-loading tip, and ran the gel at 400 V for 45-90 minutes depending on the size of the RNA, and transferred the samples to the Hybond+ nylon membrane (Amersham Hybond TM -N+, GE Healthcare) using a semi-dry electroblotter (Panther Semidry Electroblotter HEP-3 Owl). The transfer was done for 30-45 minutes at 2.5 mA/cm², and the membrane was cross-linked immediately using the auto cross-link setting on the Stratalinker. 3.2.3.1.2 Pre-hybridization and hybridization For pre-hybridization, 5-10 mL of ULTRAhyb™–Oligo Buffer (cat# AM8670, Fisher) was preheated to 42 C until completely resolubilized, then applied on the blots and incubated at 42 C for 30 minutes. For hybridization, 5 pmol/mL of biotinylated oligonucleotide was added directly into the pre-hybridization buffer and incubated overnight. 102 3.2.3.1.4 Washing The blot was quickly rinsed with 5-10 mL of wash buffer (2x SSC, 0.5% saline-sodium citrate (SDS), stored at 37 C) to remove most unhybridized probes, washed with about 20-25 mL buffer for 30 minutes at 42 C, discarded the washing buffer., and added 5 mL of blocking buffer to the blot followed by incubation for 1 hour at room temperature. In the dark, 0.5 µL Streptavidin-IRDye 800CW conjugate was added into the blocking buffer. Incubation was done for 30 minutes at room temperature in the dark, and the buffer was discarded. Then, the blot was quickly rinsed with 5-10 mL of wash buffer to remove the unbound dye and again washed with 20-25 mL wash buffer three times for 5 minutes each at room temperature. The final wash was performed with 1x PBS for 5 minutes at room temperature. 3.2.3.1.5 Detection, Analysis, and Stripping The blot was detected using the Imager (ChemiDOC TM MP imaging system, Bio-Rad) with the IR-Dye 800 CW setting and captured the image with auto exposure. Bio-Rad Image Labs software was used for measuring the levels of relative fluorescence units (RFU). If reprobing was required, the membrane was stripped before drying. For this, about 50 mL of the 0.2% SDS buffer was heated in the microwave to almost boiling, and the blot was incubated with that 0.02 %SDS for 10 minutes at room temperature, with rotation. This step was repeated. Then, the blot was rinsed with about 50 mL 2x SSC and then with water. Next, the blot was re-exposed on the imager (ChemiDOC TM MP imaging system, Bio-Rad) to ensure all probes were removed. 3.2.3.2 Fluorescent Polyacrylamide Northern on Poly A+ RNA To detect the polyA+ RNA directly, the Poly A Tract mRNA Isolation System IV kit (Cat# 0000604679, Promega, USA) was used according to the manufacturer’s protocol for isolating the polyA+ RNA, precipitation and loading on the polyacrylamide Northern blotting. 103 3.2.3.3 Denaturing agarose Northern blot Denaturing agarose Northern blotting was performed to validate the sequencing data regarding the changes in the J129, K142T, Q270C, K260, and E034C transcripts under the heat stress condition. 3.2.3.3.1 Preparation of Denaturing Agarose Formaldehyde Gel and running the gel One g of agarose was weighed and dissolved in 87mL of milliQ water by boiling it until melted and added 10mL of 10X MOPS buffer (0.2M MOPS, 50mM Sodium Acetate, 10mM EDTA, pH adjusted to 7.0 with glacial acetic acid) and 3mL of 37% formaldehyde. Once the gel was solidified, the running buffer (1X MOPS + 7% formaldehyde (1L) was poured into the electrophoresis chamber, and the gel was allowed to equilibrate for 15-30 minutes. Sample preparation followed the same protocol as for denaturing polyacrylamide gel Northern blots. Then, samples (30µg of each RNA sample) were loaded onto the gel and run at 50V for 6-8 hours in the fume hood. 3.2.3.3.2 Transferring and Detection The transfer process for the agarose Northern blot was performed using capillary transfer, as described previously for Southern blotting in Chapter II. The detection process followed the same protocol as was used for the urea denaturing polyacrylamide gel mentioned above. 104 3.2.4 RT-qPCR RT-qPCR was conducted to validate transcriptomic data regarding the presence of sisRNAs in J129, K142, and K260 genes and to investigate their accumulation or polyadenylation under heat stress conditions (Figure 32 illustrates the process of RT-qPCR). E034, with no sisRNA feature or accumulation, also was selected to act as a control. Q401, H226, and M167 were chosen as reference genes for this experiment. The stability of each reference gene was analyzed by Patrick Greetz using the GeNorm software by analyzing normalization factors (NF). 105 Figure 32. RT-qPCR. Following the pre-culturing of the C. merolae WT in MA2G medium and subsequent incubation in a water bath at 42 C and 57 C for one hour, RNA was extracted, treated with DNase, and used to synthesize cDNAs. The resulting cDNAs were then utilized for RT-qPCR analysis. 3.2.4.1 Primer Design for RT-qPCR Experiment Three primer pairs were designed for each gene (Figure 33), one primer pair amplified the exon-exon region, one amplified the exon-intron junction, and one amplified a region of the sisRNA. Primer sequences were designed using Primer BLAST (National Institute of Health, United States) to be 70-200 bp long and anneal between 50 C – 60 C (Table 14). Sequences also were designed to contain 40 – 60 % GC content to ensure optimal binding and similar melt 106 temperatures between primer pairs. Primer dimer and hairpin formation were assessed using Oligoanalyzer software (Integrated DNA Technologies, Coralville, Iowa) for each sequence. I selected sequences with Gibb’s free energy (ΔG) values of less than -6.5 kJ/mol for homodimer and hetero dimer formation and annealing temperatures greater than hairpin melt temperatures. The sequences of primers used are Tabulated in Table 12. Figure 33. Primer pair representative location for doing RT-qPCR. A schematic of the three primer sets for RT-qPCR amplifying the: 1) exon-exon region (for mRNA), 2) exon-intron junction (for pre-mRNA), and 3) region of the sisRNA. 107 Table 12. DNA oligonucleotides were used for amplifying the desired genes for RT-qPCR. The names, sequences, target regions, and efficiencies of the forward and reverse primers are provided. Cm gene name Primer name FUB93 Orientation Region Primer sequences (5′ to 3′) Forward Intron TGTCCGTGGACGTATTCAC FUB110 Reverse Exon 2 AGAGTGTCGGTGGTAAGCAA FUB94 Reverse Exon 2 GCGATCCTGAATCTGGTCAA FUB113 Forward Exon 2 ACCTGCTTCAGTTCCTTGGAC FUB146 Forward Intron AATAAGAGCGGGAGTGCTG FUB147 Reverse Intron CCAGAGTAAGCCCCATTGTG FUB97 Forward Intron AACCGTTTCATCAGTGCGAA FUB98 Reverse Exon 1 GGAAACCGCACAGAAGCAG FUB124 Forward Exon 2 TTGAGGATTCCCCCTGTTTTG FUB125 Reverse Exon 2 TGATAGCCACGTCGCAGAAA FUB132 Forward Intron CAGGCTTCCTCTACCGAGAC FUB133 Reverse Intron GCAGCACAACCGTTACTCAC FUB180 Forward Intron AGATGTTGCTTCTCGCTCG FUB181 Reverse Intron GAAAAGGGTTCCACTCTGCC FUB182 Forward Exon 1 GCAAGGACACGCAATTACAA FUB183 Reverse Intron GTTGCCACCAGCGAAAATAA FUB184 Forward Exon 2 GATTCGGGTCTGTTTGGGAT FUB185 Reverse Exon 2 GTTTGATTCTTGGCTCGCAC FUB210 Forward Intron AGTGTCAGGGTACAAGAGCG FUB211 Reverse Intron GCCCCTTGGTTAGTTTCTGC FUB214 Forward Exon 2 ACAAGCGTTTCGTGTGTTTG FUB215 Reverse Exon 2 TAAGCAGCCGAGCAAAATCT FUB216 Forward Intron CCCATGTGTTCGTCCCTATG FUB217 Reverse Exon 2 GCGTGCTGTGTCATTAGCG Q401 18s rRNA FUB154 Forward Exon 2 AAACGGCTACCACATCCAAG FUB155 Reverse Exon 2 TGTCACTACCTCCCTGAGTC H226 eF1a FUB158 Forward Exon 2 GTTTGAGGCTGGTATCTCGT FUB159 Reverse Exon 2 ATCGTCCATCTTGTTCACCG M167 Glyceraldehyde-3phosphate dehydrogenase (GADPH) FUB 162 Forward Exon 2 TCAACCACGACACCTACTCC FUB 163 Reverse Exon 2 TACCAAACGCCTCATCCAGA K142 RNase MRP in intron J129 Histone deacetylase K260 Mitochondrial processing peptidase alpha subunit E034 Similar to calmodulin 108 Efficiency (%) 90.50 92.70 90.00 101.60 99.70 94.20 97.20 92.60 96.10 99.40 94.40 99 102.10 96.80 96.10 3.2.4.2 cDNA synthesis DNase treatment was performed for the extracted RNA to remove genomic DNA contamination using the TURBO DNA-free kit (cat#AM1907, ThermoFisher Scientific, Waltham, MA) according to the manufacturer’s protocol. The concentration and purity of samples were assessed using a Nanodrop (cat#ND-ONE-W, Thermofisher Scientific, Waltham). RNA samples with an A260/A280 ratio between 1.9-2.2 and an A260/A230 ratio between 1.8-2.2 were considered clean of cell contaminants and organic solvents. A 1 % bleach, 1.5 % agarose gel with 1 μg of total RNA from each biological replicate was run at 150 V for 50 minutes to assess RNA degradation. Samples with clear doublet 28 S and 18 S bands and little smearing and no g.DNA contamination at the top of the wells were used for the experiment. cDNA was synthesized using the iScript Reverse Transcription Supermix (cat#1708890, BioRad Laboratories, Hercules, CA), and OdTs as well as GSPs using the BioRad Reliance Select cDNA synthesis kit (cat#12012802, BioRad Laboratories, Hercules, CA) according to the manufacturer’s protocols (Figure 34). A no reverse transcriptase (NRT) sample for each biological replicate was included to determine the amount of gDNA contamination. Then, biological replicates were pooled at each temperature together to generate a single sample for each temperature condition. 109 Figure 34. Different cDNA synthesis methods for RT-qPCR . Schematic various cDNA synthesis methods: 1) Oligo dT Primers (OdT): specifically amplify the polyadenylation tail. 2) gene-Specific Primers (GSP): exclusively amplify the gene body. 3) iScript Reverse Transcription: employs both oligo dT and random hexamer primers to synthesize cDNA from the gene body and the polyadenylation tail. 110 3.2.4.3 RT-qPCR 3.2.4.3.1 Determining the annealing temperature The annealing temperature of primer pairs was determined using a BioRad T100 Thermal Cycler (BioRad Laboratories, Hercules, CA), by running the thermal gradients from 48 – 61.5 C for all primer pairs using 5 ng of cDNA in each reaction in a total of 20 µL volume. The annealing time was chosen based on the length of the PCR products. A no-template control (NTC) was also included in each reaction at a temperature closest to the predicted annealing temperature. Then, PCR product samples were run on a 2 % agarose gel at 150 V for 50 minutes. The temperature corresponding to the thickest band, with the least amount of primer dimer formation, and nonspecific bands was considered as the annealing temperature. 3.2.4.3.2 Primer Efficiency Determination Primer efficiency determination was performed by Patrick Greetz using CFX Maestro software (BioRad Laboratories, Hercules, CA). Primers with efficiencies between the accepted 90110 % and with r2 greater than 0.980 were chosen for further analysis. The concentration at the middle of the standard curve was chosen as the experimental dilution for each primer. The melt peaks generated by CFX Maestro software were analyzed to ensure that a single melt peak was produced from each primer pair, indicating no non-specific binding. 3.2.4.3.3 Target and reference gene expression Target sisRNAs and reference genes were tested using three biological replicates at 42 C and 57 C of each cDNA type, each running in triplicate on clear 96 well plates (cat#HSP9601, BioRad Laboratories, Hercules, CA). Fluorescent dye and enzyme were supplied in LUNA 2X Master mix (cat#M3003E, New England Biolabs, Ipswich, MA). NTC and NRT were run in triplicate for each primer pair. As exon-exon boundaries could not be used to reduce gDNA 111 contamination and sisRNAs contain potential secondary structures, samples with a Cq in NRT or NTC wells lower than 33 were excluded to ensure robust data analysis. 3.2.4.3.4 Reference gene stability The stability of reference genes was analyzed by Patrick Greetz by comparing the change in Normalization Factor (NF) between biological groups using GeNorm provided by CFX Maestro software and kept if the mean standard deviations (M-values) were lower than 0.5. The reference genes stability was analyzed by comparing the change in NF between biological groups. 112 3.2.5 Growth test at 57 C The growth rates for strains with introns deleted in Q270, J129, K260 (sisRNA strains), and the E034C strain (as a control) were measured to determine whether these cells grew faster or slower than the WT under heat stress condition. The growth rates were analyzed and compared to the T1 (the parental strain from which the ∆is were made; it is wild type except for a deletion of the URA5.3 gene). It should be noted that our lab has not yet succeeded in deleting the intron in the K142T gene. Consequently, there was no K142∆i available for conducting growth tests. For this experiment, each strain was initially grown in a cylinder in an incubator under standard conditions (42 C, 2% CO2, and continuous white light). For this initial growth period, a selective growth with 0.8 mg/ml 5-FOA was conducted. Once the cultures had grown and appeared quite green, they were diluted to achieve approximately the same OD 750 nm (target OD 750 nm = 1-3) in 24 hours, when every culture needed to be diluted to an appropriate OD (0.03-0.05) in 15-20 ml MA2GU. The OD 750 nm was measured and distributed 700-800 µl aliquots into 9 wells of the 48-well plate. Each strain occupied 9 wells, allowing four strains per 48-well plate (two strains with their duplicate biological replicates), with one well reserved for the blank (MA2GU). To minimize evaporation, ddH2O was distributed between the wells and into the external columns of the plate. Cells were grown in MA2GU liquid medium in the 48-well plates (NUNCLON, Cat 150687) shaken at 250 rpm under continuous white light illumination (135 μmol photons m−2 s−1) at 57 C. Growth curves were recorded by measuring OD 750 nm values at a wavelength of 750 nm using the BioTek Synergy Neo2 plate reader. Then, the growth rates were calculated from the slope of the growth curves during the cells' linear growth phase. Each strain were tested in two biological and nine technical replicates to ensure accuracy and reproducibility. This means that for 113 each growth test, nine technical replicates were used for T1, and 18 replicates were used for each ∆i. 3.3 Results and Discussion 3.3.1 Confirmation of sisRNAs by Northern blotting Northern blotting was performed to confirm the presence of sisRNAs at 42 C in J129∆I, Q270∆I, K142∆I, and K260∆I, as identified in long-read sequencing data54 (see Figure 29), and to validate the short-read sequencing data, which indicated intron accumulation in these genes under heat stress conditions54 (see Figure 30). In this experiment, the gene J101 was chosen as a heat shock control as it has been demonstrated that this gene, which encodes a small heat shock protein, is transcribed only when cells are exposed to increased temperatures83. E034 was also selected as a negative control since it lacks a sisRNA, and sequencing data did not show any increase in intron expression. C. merolae cells were initially exposed to 42 C and 57 C for one hour, as well as 42 C and 60 C for 30 minutes, to assess the effect of higher temperatures on sisRNA. Subsequently, the experiments were continued just at 42 C and 57 C to ensure consistency with transcriptomic data as suggested by transcriptomic data. sisRNA bands with higher intensities should be observed at 57 C or 60 C compared to 42 C if accumulation occurs under heat stress. The presence of sisRNAs at 42 C is shown in Figure 35. 114 A) J129 sisRNA B) K142 ncRNA (MRP) C) Q270 sisRNA 441nt 127nt 123nt D) K260 sisRNA E) E034 277nt Figure 35. Confirmation of sisRNAs at 42 C. Northern blotting shows the presence of sisRNAs at 42 C with sizes closely matching the predictions from the long-read sequencing data for A) J129 (127nt), B) K142 (441nt), C) Q270 (123nt), D) K260 (277nt), and E) no sisRNA for E034, the control gene without a sisRNA. U5 is C. merolae U5 (450nt). Lanes 1-3 are technical triplicates. YsnRNA is S. cerevisiae total RNA probed for the snRNAs U6 (112 nt), U4 (160 nt), U5 (179 and 214 nt), U1 (568 nt), and U2 (1175 nt). 115 The results of Northern blotting regarding the presence of sisRNA in J129∆I, K142∆I, Q270∆I, K260∆I, and the E034∆I (as a control) under normal and heat stress conditions are shown in Figures 36-40. The J129 sisRNA, measuring 127 nucleotides, was detected under all temperature conditions (Figure 36). This result is the first confirmation of long-read transcriptomic data demonstrating the presence of a sisRNA at 42 C, and together these data constitute the first report of sisRNAs in any alga. However, in contrast to short-read transcriptomic data showing intron accumulation, there was no change in band intensity between 42 C and 57 C, as well as 42 C and 60 C, with P values of 0.5 and 0.8, respectively, when band intensities were calculated using C. merolae U5 snRNA as the loading control to normalize the data (i.e. to compensate for unequal loading or transfer of RNA samples). From this, I concluded that the sisRNA is stable under all temperature conditions but in contrast to short-read transcriptomic data, there is no change (accumulation) for this sisRNA under heat stress. It should be noted that the band corresponding to J101, which was expected to be detected under heat stress as one of the heat-inducible genes confirming our heat stress condition, was only observed at 57 C. The failure of this positive control for heat shock may indicate that the 60 C sample did not experience heat shock for some reason, but the extended incubation time of 30 minutes at 60 C in my experiment exceeds the duration used in previous studies, 20 minutes at 60 C83,84. This prolonged exposure may have led to mRNA degradation. Nevertheless, the heat shock response was clearly induced at 57 C, and there was no change in the sisRNA, so I conclude that sisRNA expression is not increased under heat stress. 116 A) J129 sisRNA B) Figure 36. Effect of temperature on J129 sisRNA expression. A) Denaturing Northern analysis of J129 sisRNA (127nt) expression at 42 C, 57 C, and 60 C as indicated. Samples are in three technical replicates. YsnRNA: S. cerevisiae snRNAs used as a size ladder. U5 is C. merolae U5snRNA (450nt) as a loading control. J101 (720nt) is a heat shock control. The expected transcript size is shown on the left. B) Quantification of the data in (A). Paired t-tests with p-values of 0.55 and 0.88 show no difference between sisRNA intensity at 42 C and 57 C as well as 60 C. Whiskers show standard deviation. 117 The K142 ncRNA (MRP), 441 nucleotides, was detected under all temperature conditions (Figure 37). The size of the sisRNA remained consistent across all conditions, and there was no change in band intensity between 42 C and 57 C when band intensities were calculated using C. merolae U5 snRNA as the loading control to normalize the data and verify equal loading of RNA samples. From this, again I concluded that the sisRNA is stable under all temperature conditions. Still, in contrast to short-read transcriptomic data, there is no change in this ncRNA under heat stress. It should be noted that the band corresponding to J101 was detected at 57 C as expected. 118 B) J101 and U5 A) K142 ncRNA C) Figure 37. Effect of temperature on K142 ncRNA expression A) Denaturing Northern analysis of the K142 ncRNA (441nt) expression at 42 C and 57 C as indicated. Samples are in three technical replicates. YsnRNA: S. cerevisiae snRNAs used as a size ladder. U5 is C. merolae U5snRNA (450nt) as a loading control. J101 (720nt) is a heat shock control. The expected transcript size is shown on the left. (B) Quantification of the data in (A). A paired t-test with a p-value of 0.55 shows no difference between sisRNA intensity at 42 C and 57 C. Whiskers show standard deviation. 119 The Q270 sisRNA, 123 nucleotides, was detected under all temperature conditions (Figure 38). The size of the sisRNA remained consistent across all conditions, and there was no change in band intensity between 42 C and 57, with a P value of 0.5, when band intensities were calculated using C. merolae U5 snRNA as the loading control to normalize the data and verify equal loading of RNA samples. It should be noted that the band corresponding to J101 was detected at 57 C as expected. This results again indicate that the sisRNA is stable under all temperature conditions but in contrast to short-read transcriptomic data, there is no accumulation for that under heat stress. 120 B) Q270 sisRNA A) J101 and U5 C) P=0.55 Figure 38. Effect of temperature on Q270 sisRNA expression. A) Denaturing Northern analysis of Q270 sisRNA (123nt) expression at 42 C and 57 C as indicated. Samples are in three technical replicates. YsnRNA: S. cerevisiae snRNAs used as a size ladder. U5 is C. merolae U5snRNA (450nt) as a loading control. J101 is a heat shock control. The expected transcript size is shown on the left. (B) Quantification of the data in (A). A paired t-test with a p-value of 0.55 shows no difference between sisRNA intensity at 42 C and 57 C. Whiskers show standard deviation. 121 The K260 sisRNA, 277 nucleotides, was detected under all temperature conditions (Figure 39). The size of the sisRNA remained consistent across all conditions, and there was no change in band intensity between 42 C and 57 with a P value of 0.23 when band intensities were calculated using C. merolae U2 snRNA as the loading control. The Northern blot analysis of K260, similar to J129, Q270, and K142, revealed that the sisRNA remains stable across various temperature conditions as seen for the other sisRNAs I tested and in contrast to what we saw in short-read transcriptomic data. 122 A) K260 sisRNA B) A) E034 Figure 39. Effect of temperature on K260 sisRNA expression. (A) Denaturing Northern analysis of K260 sisRNA (277nt) expression at 42 C and 57 C as indicated. Samples are in three technical replicates. YsnRNA: S. cerevisiae snRNAs used as a size ladder. U2 is C. merolae U2snRNA (131nt) as a loading control. The expected transcript size is shown on the right. (B) Quantification of the data in (A). A paired t-test with a p-value of 0.24 shows no difference between sisRNA intensity at 42 C and 57 C. Whiskers show standard deviation. 123 No band was detected for the E034 intron (150nt) under normal (42 C) and heat stress conditions (57 C and 60 C). However, the band corresponding to the C. merolae U5 snRNA was detected in all conditions confirming the presence of RNA on the blot and ruling out technical issues with this Northern blotting (Figure 40). 124 E034 intron Figure 40. Effect of temperature on E034 intron expression. Denaturing Northern analysis of E034 expression at 42 C and 57 C as indicated. YsnRNA: S. cerevisiae snRNAs used as a size ladder. U5 is U5 C.merolae snRNA(450nt) as a loading control. No band corresponding to E034 intron was observed. 125 Polyacrylamide Northern blotting using total RNA was performed to validate the long-read sequencing data observations regarding the presence of sisRNAs at 42 C and the short-read sequencing data showing intron accumulation for J129, K142, Q270, and S270 under heat stress. The long-read data provides preliminary evidence for the existence of sisRNAs, indicating their presence at 42 C exclusively in total RNA samples, not in polyA+ samples, which suggests that sisRNAs at this temperature are unadenylated. Northern blotting definitively confirms the presence of these sisRNAs, corroborating the long-read sequencing data. However, the short-read sequencing observations have not been confirmed by Northern blotting. If sisRNA exhibits accumulation at higher temperatures, alterations in intensity, indicative of the elevated expression level should be discernible at 57 C relative to 42 C. As shown in Figures 36-40, there was no change in the intensity of sisRNAs across different temperatures, demonstrating consistent expression of intronic sisRNAs regardless of the temperature. Additionally, no band corresponding to the E034C intron was detected, confirming that the E034C intron does not encode a sisRNA and is rapidly degraded after splicing. In contrast, the sisRNAs from J129, K142, Q270, and K260 are stable and not immediately degraded after splicing, possibly because they perform some function in the cells. This observation aligns with findings in Xenopus tropicalis that sisRNAs are stable for at least 1–2 days46. The absence of sisRNA accumulation observed in this Northern blot analysis suggests that the short-read sequencing data may be indicating the process other than the accumulation of sisRNAs. Given that the RNA sample for short-read transcriptomic analysis was polyA+ RNA, it is possible that sisRNAs are being polyadenylated under heat stress rather than accumulated. Previous studies have shown that polyadenylation can stabilize sisRNAs within the cell46. If sisRNAs are indeed being polyadenylated, size variations would be expected between the samples 126 at 42 C and those under heat stress. However, no such size differences were detected in this Northern blot analysis. 3.3.2 Denaturing Polyacrylamide Northern blotting using polyA+ RNA Since total RNA Northern blot analysis revealed no differences in the size or expression of sisRNAs between normal and elevated temperatures, a Northern blotting using polyA+ RNA was conducted. This experiment was consistent with transcriptomic data, which was derived from polyadenylated RNA samples. Polyadenylation might occur in only a small subset of sisRNAs, making it undetectable in total RNA samples. PolyA+ RNA was selected using a kit (Methods), and Northern blotting was performed for J129, Q270, K142, K260, and E034. A single band at roughly 160 nucleotides was observed in all genes tested, including E034C which does not contain a sisRNA, suggesting non-specific binding. The polyA+ purified RNA Northern blotting for E034, serving as an example representative of other genes of interest is illustrated in Figure 41. This figure indicates the presence of non-specific bands across all temperature conditions. 127 Figure 41. PolyA+ RNA Northern blotting. Denaturing Northern analysis of sisRNA expression of polyA+ RAN at 42 C and 57 C as indicated show the presence of non-specific bands across all temperature conditions. YsnRNA: S. cerevisiae snRNAs used as a size ladder. A non-specific band around 160nt was observed in all conditions. 128 3.3.3 Agarose Northern blotting Transcriptomic analyses (see Figure 30) revealed not only an increase in intron reads but also alterations in exon reads and splicing levels under heat stress for those genes. For instance, Q270C exhibited an increase in exon reads, although this was not as pronounced as the increase in intron reads, and there was a reduction in splicing levels after one hour of exposure to heat stress. This suggests enhanced expression of its transcript and a higher production of pre-mRNA at elevated temperatures. Conversely, K142 and K260 displayed decreases in exon reads and splicing levels, indicating reduced transcript levels and an increase in pre-mRNA production under heat stress conditions. J129 showed no change in exon levels but exhibited decreased splicing. Agarose Northern blot analysis using total RNA samples was performed to investigate the observed alterations in exon reads and splicing levels in the sequencing data. A distinct band was detected for Q270, but smeary bands for J129, and I could not detect any band for K142 and K260. I repeated these experiments twice, yielding consistent results. Based on Figure 42A, there is a shift in the size of the transcript at elevated temperatures for Q270, from approximately 2.5 kb at 42 C to around 3 kb at 57 C. The expected sizes of the mRNA and pre-mRNA are 2.5 kb and 3 kb, so I interpret the shift in mobility to be due to a decrease in splicing. This shift reveals an increase in pre-mRNA levels under heat stress compared to 42 C, aligning with transcriptomic data that show a decrease in splicing during heat stress. However, in contrast to transcriptomic observation indicating increasing exon expression under heat stress, there was no change in exon expression under heat stress as measured by the intensity of the bands (Figure 42B). 129 A) Q270 transcript B) Figure 42. Agarose northern blotting for Q270. (A) Denaturing Northern analysis of transcript expression at 42 C and 57 C as indicated. Samples are in two technical replicates. ssRNA: single-strand RNA used as a size ladder. Expected transcript sizes are shown on the left. A shift in the size of transcript was observed from 2.5kb at 42 C to 3kb at 57 C.(B) Quantification of the data in (A). A paired t-test with a p-value of 0.24 shows no difference between the transcript intensity at 42 C and 57 C. Whiskers show standard deviation. 130 3.3.4 RT-qPCR The absence of sisRNA accumulation observed in the Northern blot analysis, despite the findings from the short-read sequencing, suggests that short-read sequencing data may reflect sisRNA polyadenylation under heat stress (Figure 43). This inference arises from the fact that the RNA sample for short-read transcriptomic analysis was polyA+ RNA so an 131 increase in polyadenylation would look like an increase in expression. However, no size differences indicative of sisRNA polyadenylation were detected in Northern blot analysis. A B) B) Figure 43. Polyadenylation vs accumulation. A) Polyadenylation of sisRNA (same number of sisRNA with poly-A tail). B) more transcription of sisRNA (increasing in the transcription of sisRNA). 132 Due to the ambiguity of Northern blotting results regarding whether sisRNAs were polyadenylated or accumulated under heat stress, reverse transcriptase quantitative PCR (RTqPCR) was conducted on three regions within three sisRNA-containing genes (sCGs), J129, K142, and K260 (Figure 44 A, B, and C.) and one intron containing gene (ICG) without a sisRNA feature, E034 as a control, (Figure 44 D). In this study, Q401, H226, and M167 were chosen as reference genes85,86. Based on Figure 41, all sisRNA levels increased (> five-folds) at 57 C, with P values < 0.05 compared to 42 C, mirroring the pattern observed in transcriptomic data but in contrast to the Northern blotting results mentioned earlier. No change in expression was observed in E034C, which was chosen as the negative control in this experiment as it does not contain any sisRNA. However, transcriptomic data showed a decrease in the expression of this gene under heat stress. Additionally, to better understand whether the observed increase in sisRNA expression is due to an increase in host gene expression under heat stress or if it is independent of changes in gene expression or premRNA levels, the expression of exons (which are present in pre-mRNA and mRNA) and exon-intron junctions (pre-mRNA) was assessed independently for exon-exon and exonintrons. Based on Figure 41, exon expression (corresponding to total RNA) decreased in all genes, and pre-mRNA levels decreased in all genes except J129. Although expression 133 increased slightly in J129, the change in pre-mRNA levels is much smaller than the increase observed in sisRNA levels. 134 A) J129 B) K142T C) K260 D) E034 Figure 44. RNA levels of regions within sisRNA containing genes as determined by RTqPCR. Cells grown at 42 C (white) and 57 C (grey) for one hour were used for RNA extraction. The genes were tested: A) J129, B) K142, C) K260, and D) E034. Data shows the geometric mean (geomean)± SD from three biological replicates. P values were calculated using a two-tailed Student’s t-test. 135 To determine if sisRNAs were polyadenylated or if they accumulated under heat stress, another RT-qPCR was performed using oligo DT (OdT), gene-specific primers (GSP), and iScript. OdT primers. GSPs, designed specifically for introns (sisRNAs), reverse transcribe from the 3’ end of each sisRNA. iScript utilizes both random hexamer and OdT primers, enabling the reverse synthesis of the entire gene from the polyA tail, as well as of other transcripts such as sisRNAs, truncated RNAs, and other non-polyadenylated molecules. Therefore, if an increase in expression is observed using both iScript and OdT, but not with GSPs, it would indicate polyadenylation of the sisRNA. Conversely, if an increase in expression is observed using both iScript and GSP (intron-specific), this suggests the accumulation of sisRNA. According to Figure 45, a greater increase of sisRNAs was observed in OdT cDNA samples, which represents the polyA+ samples (Figure 45 A, B, and C), compared to iScript and GSP samples. This indicates sisRNA polyadenylation. Conversely, polyA+ intronic sequences (detected with OdTs) decreased in E034, however, total RNA levels remained the same (Figure 45 D). In sCGs, sisRNAs levels changed more dramatically using iScript cDNA than with OdTs. iScript uses a combination of both random hexamers and oligodTs, thus targeting both polyadenylated and total RNA. Therefore, it is counterintuitive that iScript cDNA shows a greater change in sisRNA levels. However, if sisRNA polyA tails were shorter than the designed primer length for the polyA tail, cDNA synthesis might have been inefficient, affecting the detection of the change in expression. Together, these may suggest that polyadenylated sisRNAs represent a small fraction of the total and that the polyA tail is short. 136 A) J129 B) K142 C) K260 D) E034 Figure 45. RNA levels of intronic sequences as determined by RT-qPCR. Cells subjected to heat stress (57 C) and control growth (42 C) conditions for one hour were used for RNA extraction. cDNA was synthesized using ISR, ODT, and GSP. A) J129, B) K142, C) K260, and D) E034 expression was used as an internal control. Data shows the geometric mean (geomean) ± SD from three biological replicates. P values were calculated using a two-tailed Student’s t-test. 137 This study validated the polyadenylation of sisRNAs, initially observed in short-read transcriptomic data, through RT-qPCR using various cDNA synthesis primers. Additionally, sisRNAs were found to undergo polyadenylation independently of their host genes, indicating their stability under both normal and heat stress conditions, even as the expression of their host genes was downregulated. These results are consistent with the patterns observed in the transcriptomic data. Northern blot analysis, on the other hand, showed the presence of sisRNAs at both normal (aligning with long-read transcriptomic data) and heat stress. Still, it did not support the findings from short-read transcriptomics and did not reveal any variation in the size of sisRNA as would be expected if they were polyadenylated. The failure to observe sisRNA polyadenylation in the Northern blot may be due to only a small fraction of sisRNAs becoming polyadenylated at the higher temperature, making them undetectable by this method. 3.3.5 Growth test at 57 C The presence of sisRNAs at normal conditions (42 C) and their polyadenylation under heat stress, raise the question of whether their presence is important for the cell under heat stress. One way to test this would be to ask whether their loss from cells affects cell growth. The hypothesis is that if they are important for maintaining the cell under heat stress, a strain lacking the sisRNA will exhibit reduced tolerance to heat stress, resulting in either cell death or slower growth compared to the WT. Therefore, a growth test at 57 was performed as described in the materials and methods section for Q270∆i, J129∆i, and K260∆i, with E034∆i strains serving as a negative control. The growth rates of these ∆is were compared to the T1. The growth rates for T1 and ∆is at 57 C are shown in Table 13. 138 Table 13. Growth test at 57 C. The growth rates for T1 alongside the average growth rate for ∆i1 and ∆i2. ∆i1 and ∆i2 represent two biological replicates of ∆is. Columns 3 and 4 detail the growth rate for each biological replicate of ∆is. Column 5 presents the results of a paired t-test comparing the growth rate of T1 with the average growth rate of ∆i1 and ∆i2, aimed at determining if there is a change in growth between T1 and ∆is. A one-way ANOVA with multiple comparisons using Tukey’s range test was performed and a p-value of less than 0.05 was used to indicate statistical significance. Column 6 summarizes the overall comparison of growth rates between T1 and ∆is. Strain Average growth rate Growth rate ∆i 1 Growth rate ∆i 2 P-value T1v∆i Compare to T1 T1 8Î10-4 Q270 -9.3Î10-5 -3Î10-4 1.2Î10-4 2.2Î10-4 Slower (dead) K260 0.00 0.00 0.00 0.00 Faster J129 8.6Î10-4 0.00 -4.2Î10-4 0.00 Inconsistent E034 9Î10-4 0.00 5.6Î10-4 0.06 Equivalent 139 The Q270∆i did not grow at 57 C (Figure 46). This suggests that the sisRNA in the Q270 intron might play an important role in enabling cells to withstand heat stress. A P= 2.24Î10-4 B Figure 46. Growth test for Q270∆i at 57 C. A) The Scattered plot with linear trendlines shows the growth of T1 (blue) and Q270∆i (green and orange for ∆i.1 and ∆i.2) at 57 C as indicated. Each strain had nine technical replicates. B) Quantification of the data in (A) A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 2.24Î10-4 showed Q270∆i death at 57 C. Whiskers show standard deviation. 140 The J129∆i showed different growth between the two biological replicates: ∆i.1 grew faster than T1 while ∆i.2 did not grow at all at this temperature (Figure 47). Since it is unclear which one was the correct phenotype for this ∆i, I cannot make any inferences about whether or how the J129 sisRNA contributes to heat adaptation. This discrepancy is consistent with the growth test at 42 C, where the DTs were 10.6 and 11.9 for ∆i.1 and ∆i.2, respectively, compared to 11.6 for T1. Although the exact reason for the discrepancy between these ∆is is unclear, a possible genetic mutation during or following intron removal might have affected the growth of one strain. Fresh J129∆i thawed from -80 C showed the same growth patterns, indicating the problem originated with the generation of these ∆is. 141 A P= 0.001 B) B Figure 47. Growth test for J129∆i at 57 C. A) The Scattered plot with linear trendlines shows the growth of T1 (blue) and J129∆i (orange and green for ∆i.1 and ∆i.2) at 57 C as indicated. Each strain had nine technical replicates. B) Quantification of the data in (A). ∆i.1 and ∆i.2 showed a discrepancy in growth rate at 57 C with an average p-value of 0.001 from T1. Whiskers show standard deviation. 142 The K260C∆i initially grew faster than T1 but stopped growing sooner, around 40 hours (Figure 48). This is an interesting observation on both counts: the fast initial growth suggests neither the intron nor the sisRNA are important for the immediate response to heat stress and the faster cessation of growth could indicate that either the intron or the sisRNA contributes to the medium-term adaptation to heat, but the T1 eventually stops growing as well. 143 A B) B Figure 48. Growth test for K260∆i at 57 C. (A) The Scattered plot with linear trendlines shows the growth of T1 (blue) and K260∆i (green and orange for ∆i.1 and ∆i.2) at 57 C as indicated. Each strain had nine technical replicates. K260∆i initially grew faster than T1 but stopped growing sooner than T1 with a p-value of P= 0.004. B) Quantification of the data in (A). Whiskers show standard deviation. 144 The E034∆i (Figure 49) exhibited no change in growth compared to T1 (Table 13), supporting my view that the preceding observations are due to the removal of the sisRNAs rather than the intron itself. A B Figure 49. Growth test for E034∆i at 57 C. A) The Scattered plot with linear trendlines shows the growth of T1 (blue) and E034∆i (green and orange for ∆i.1 and ∆i.2) at 57 C as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.06 showed no difference between the growth rate of E034∆i and T1. B) Quantification of the data in (A). Whiskers show standard deviation. 145 Although the growth defects at 57 C for the Q270∆i is evident, this phenotypic growth test does not reveal the underlying cause. These results suggest that this intron or the sisRNA might play important roles in enabling cells to withstand heat stress. Studies have demonstrated that sisRNAs are biologically active ncRNAs that serve as an additional layer of gene regulation within the cell. They regulate host gene expression, function as molecular sinks for RNA-binding proteins, and influence protein translation81. Additionally, they regulate genes at the DNA, RNA, and protein levels, often participating in autoregulatory feedback loops to maintain cellular homeostasis under both normal and stress conditions46. For instance, K142 contains the ncRNase MRP, an essential ribonucleoprotein complex involved in the processing of precursor ribosomal RNA (pre-rRNA) and the regulation of the cell cycle. This complex is crucial for the cleavage of 5.8S rRNA82. Research in yeast has shown that the removal of stable introns under stress conditions can be detrimental, leading to abnormally low growth rates 74. Furthermore, research has shown that ncRNAs within intronic sequences, rather than the introns themselves, contribute to the phenotype, thereby enabling the cell to respond to salt stress87. 146 Conclusion This chapter focused on resolving whether heat stress induces polyadenylation or accumulation of sisRNAs in C. merolae. sisRNAs, a newly characterized class of ncRNA, have been found to play important roles in gene regulation across various organisms. In this study, I explored the presence and behavior of sisRNAs in C. merolae under heat stress, with a focus on the specific genes: J129, K142, Q270, and K260; E034 served as a negative control. Northern blotting using total RNA revealed that sisRNAs from J129, K142, Q270, and K260 are stable, i.e. they persist at 42 C and 57 C, unlike the introns that do not contain sisRNAs. As predicted, the E034C intron did not contain a sisRNA, as confirmed by the Northern blotting as well. Although Northern blotting confirmed long-read transcriptomic data showing the presence of sisRNAs at 42 C, it could not validate short-read transcriptomic data regarding the accumulation of sisRNAs under heat stress: there was not any difference in their sisRNA intensities between 42 C and 57 C. Importantly, the RNA sample for short-read transcriptomic analysis was polyA+, suggesting that polyadenylation of sisRNAs under heat stress could explain their apparent accumulation. Therefore, RT-qPCR was performed to differentiate between accumulation or polyadenylation for sisRNA seen in short-read transcriptomic data under heat stress. The results confirmed that sisRNAs are indeed polyadenylated. This raises the question of whether such polyadenylation has a functional consequence, particularly in the adaptation to heat. This will provide an important direction for future research. The polyadenylation of sisRNAs under heat stress and the growth defects observed in ∆is highlight the functional importance of these introns and their sisRNAs. These results support the hypothesis that introns and sisRNAs are important for cellular adaptation to environmental stress 147 and therefore could contribute to gene regulation, RNA processing, and overall cellular homeostasis. 148 CHAPTER IV: Discussion 149 4.1 General Discussion C. merolae exhibits a remarkably compact genome, comprising only 39 introns and a limited set of core splicing proteins. This is in contrast to S. cerevisiae, which has introns in approximately 5% of its transcripts, and Homo sapiens, where over 95% of transcripts contain introns88. Additionally, 49 splicing proteins have been found in C. merolae compared to almost 90 in budding yeast and around 140 in Homo sapiens56. Furthermore, C. merolae only contains four snRNAs (U2, U4, U5 and U6), since no candidates for the U1-associated proteins or U1 snRNA were found56. The reduced number of splicing proteins and introns defines C. merolae as a simpler organism from a splicing perspective, making it a useful system in which to study intron function. Several studies have investigated introns’ selective advantages to eukaryotes, seeking to explain what compensates for the energetic cost of maintaining intronic DNA, transcribing it, and splicing it out of pre-mRNA. Cis-acting regulatory sequences inside introns, which are selectively recognized by complementary trans-acting factors, have been associated with known molecular functions including shaping alternative splicing, gene expression enhancement, and mRNA stability. Additionally, introns can harbor several kinds of noncoding functional RNA genes, such as snoRNAs, piRNAs, and lncRNAs. ncRNAs located in introns are sometimes involved in the auto-regulation of host gene expression, in addition to their trans-acting functions29,30. This research aimed to determine whether the retention of a small number of introns in C. merolae, in the context of massive intron loss over evolutionary time, is due to some important function for which they have been selected or whether they will eventually be lost like all the others. A first reasonable step toward answering this question was to examine whether deleting C. merolae’s introns affects its growth rate. For this, HR was employed with plasmid constructs to delete specific introns (D067, S262, E034, and R289) chosen based on their host gene expression 150 level. This method allowed precise excision of introns, ensuring minimal disruption to surrounding genomic regions. To assess the impact of intron deletion on cellular growth, 42 C growth tests were conducted on the ∆is. I developed a new plate assay protocol using 48-well plates to enhance the efficiency and reproducibility of this experiment. This required optimizing several variables, including shaking speed, evaporation control, starting OD 750 nm to allow sufficient time for data collection before cell saturation, and light intensity adjustment. The development of this plate assay significantly improved the convenience and accuracy of measuring DTs for C. merolae ∆i strains compared to the previously used methods involving graduated cylinders and tube cultures. Using this assay, the growth test was performed for the four ∆is generated in this study– D067∆i, S262∆i, E034∆i, R289∆i – as well as some other previously generated ∆is, namely K260∆i, S342∆i, Q382∆i, S270∆i, J129∆i, C008∆i, Q117∆i, Q270∆i, ∆6i, and ∆10i, for which no data was available. The question of whether introns are a burden or an advantage for eukaryotic cells including C. merolae has been debated. Findings from this study offer evidence supporting both perspectives of this hypothesis. In half of the cases, introns in C. merolae could be removed individually without affecting growth under normal conditions. This finding shows that many introns are not important for basic cellular processes. However, the deletion of certain introns had significant effects on growth rates. ∆is such as S262∆i, Q270∆i, J129∆i, E034∆i, ∆6i, and ∆10i exhibited slower growth rates, suggesting that the introns in these strains are important for optimal cellular function. Notably, the slower growth observed in ∆10i (with ten introns deleted) compared to single-intron deletions and ∆6i suggests that introns in the genome act cumulatively. Conversely, some introns 151 seem to impose a burden on rapidly growing cells. The R289∆i, for instance, exhibited a faster growth rate compared to the WT. Introns can also play a significant role in cellular responses to environmental stress. A systematic individual deletion of all introns from S. cerevisiae revealed that only a minority of introns are required for growth in the rich medium, but, in most cases, cells with an intron deletion are impaired when nutrients are depleted76. This is independent of the effect of their host mRNA function. Rather than regulating the expression of the host gene, introns promote resistance to starvation via the repression of ribosomal protein genes downstream of the nutrient-sensing TORC1 and PKA pathways73. To test whether C. merolae introns play a similar role in adaptation to nutrient depletion, a novel fitness test was developed for C. merolae in this study, based on one developed in yeast, that was performed on two ∆i: ∆10i (with ten intron deletions) and R289∆i. The experiments involved comparing their growth during the logarithmic phase (when sufficient nutrients were available) and the saturation phase (when nutrients were limited and cells competed for survival). The results revealed that R289∆i exhibited a higher growth compared to the WT in both the logarithmic and saturation phases. However, the difference between the two phases was not statistically significant. The ∆10i strain occasionally exhibited a higher growth rate than WT cells in the co-culture. However, the overall trend favors faster growth of WT cells in both the logarithmic and saturation phases. These findings underscore the role of those deleted introns in stress resilience, suggesting that they may play some functions in enhancing cellular survival under adverse conditions. It is widely accepted that splicing in higher eukaryotes enhances protein diversity by generating multiple mRNAs from a single gene locus89. Since alternative splicing in C. merolae seems to be rare as it has just one intron in each gene (except one gene with two introns), we 152 hypothesize that functional C. merolae introns may have been retained because they contain sisRNAs. A high-throughput RNA sequencing study conducted in 2012 on the Xenopus tropicalis oocyte nucleus revealed intronic sequences that are abundant and exhibit unusual stability, persisting for at least 1–2 days90. Further experiments revealed that these intronic sequences constitute a novel class of non-coding RNAs, referred to as stable intronic sequence RNAs (sisRNAs). Since their discovery, sisRNAs have been identified across a range of organisms, including viruses, yeast, Arabidopsis, Drosophila, and mice. These sisRNAs have been implicated in the regulation of gene expression through diverse mechanisms46. Additionally, recent studies have documented the presence of polyadenylated intronic sequences that exhibit remarkable stability and resist degradation through canonical pathways, including nonsense mediated decay (NMD)46,78. In C. merolae also, the Rader lab’s transcriptomic analysis from polyA+ libraries of samples subjected to heat stress demonstrated an apparent upregulation of 19 introns. These introns contain sisRNAs as seen in long-read sequence data. For instance, K142 contains a distinct region within its intron that is prominently highlighted in RNA-seq data. This region has been identified as the RNase MRP, which plays a role in the cleavage of 5.8S rRNA82. In order to discover potential functional RNA structures within introns in C. merolae, first Northern blotting was performed to validate our sequencing data showing stable sisRNA expression under normal conditions (Chapter 3). This was the first demonstration of sisRNAs in algae and is an exciting expansion of our understanding of the intron landscape in C. merolae. Next intron upregulation under heat stress conditions was tested for J129, K142, Q270, and K260. Northern blotting did not reveal any difference in size or intensity of sisRNAs between normal and a heat stress condition, but it is possible that only a small fraction of the sisRNA is polyadenylated, which could be below detection limits. To further test the hypothesis that heat stress leads to 153 polyadenylation, another Northern blotting was performed using polyA+ RNA to align with transcriptomic data, which was also derived from polyA+ RNA. However, a single band at approximately 160 nucleotides was observed in all genes tested, including E034, across all temperature conditions, and was considered a non-specific band. Due to the ambiguity of the Northern blotting results concerning whether sisRNAs were polyadenylated or accumulated under heat stress, RT-qPCR was conducted for those specific genes, which confirmed their polyadenylation. The results were consistent with findings from other studies indicating that the polyadenylation of sisRNAs helps protect them from degradation and stabilizes them within the cell46. Although the mechanisms of sisRNA production are not yet fully understood, their stability and resistance to rapid degradation suggest they play important functional roles in the cell. Additionally, the assessment of expression for exons and exon-intron junctions showed that the observed increase in sisRNA expression is independent of changes in their gene expression or premRNA levels, suggesting stability under both control and heat stress conditions. Considering their presence under normal conditions (42 C) and their polyadenylation under heat stress, it prompts the question of whether sisRNAs are important for cellular function under heat stress and if their absence impacts cell growth. Therefore, a growth test at 57 C was conducted comparing the growth rate of Q270∆i, J129∆i, and K260∆i, with E034∆i serving as a negative control as it has no sisRNA. The E034∆i exhibited no change in growth compared to the WT at either temperature. This suggests that the intron in E034 does not have functional importance under heat stress conditions. At 57 C, the J129∆i displayed different growth rates between biological replicates, consistent with observations from growth tests at 42 C. While the exact cause of this variability is unclear, it is possible that a genetic mutation following intron removal affected the growth of one of the J129∆is. The Q270∆i, which demonstrated slower growth at 42 C, exhibited 154 cell death at 57 C. This finding underscores the functional significance of the presence of this intron or sisRNA under both normal and heat stress conditions, particularly under heat stress, in which losing the intron and sisRNA led to cell death. The K260∆i did not show a change in growth compared to WT at 42 C indicating that under normal conditions the intron or sisRNA neither help nor hurt the cell. Upon shifting the temperature to 57C, the ∆i initially grew faster, suggesting neither the intron nor the sisRNA is important for the immediate response to heat stress. However, this ∆i stopped growing sooner than WT (T1) after the shift, suggesting that the intron or the sisRNA may play a role in the medium-term adaptation to heat stress, but the T1 eventually stopped growing as well. This temperature was clearly too high for survival, but it appears T1 was less sensitive to heat. One possible explanation is that there is a beneficial effect from losing the intron probably due to reduced energetic costs to the cell, which allows faster initial growth after the heat shift, but a detrimental effect from losing the sisRNA at longer time points. However, I cannot draw a strong conclusion about this ∆i as it did not exhibit a consistent pattern of growth at 57 C. Although the Q270∆i and K260∆i showed different growth than T1 at 57 C, this phenotypic growth test does not elucidate the underlying cause. In other organisms, sisRNAs can play a significant role in fine-tuning gene expression, which is important for maintaining cellular homeostasis and developmental stability in both normal and stress conditions 91. Studies in yeast have indicated that removing stable introns under stress conditions can be detrimental, causing abnormally slow growth rates74. Moreover, it has been shown that ncRNAs found within intronic sequences, rather than the introns themselves, are important for determining phenotype and promoting intron retention, thereby enabling the cell to adapt to salt stress effectively87. However, for R289 and S262, which do not contain sisRNAs, the growth test at 42 C for the ∆is revealed faster and slower growth compared to the WT for these two ∆is, respectively. This suggests that 155 not all intron functions in C. merolae are related to sisRNA activity. Some introns may exert their functional roles through different mechanisms, such as harboring enhancers or silencers that modulate gene expression71. 4.2 Future Directions Growth tests indicated that most introns in C. merolae can be lost individually without impacting growth under normal conditions; this finding suggests that many introns are not essential for basic cellular processes. This observation raises the question of whether they are important under adverse conditions. Therefore, to determine the range of conditions under which introns could provide a benefit to the cell, growth tests under various conditions, including osmotic stress, CO2 stress, and different nutritional stress conditions, should be conducted. Although this study detected sisRNAs under normal conditions and their polyadenylation under heat stress, the fraction of sisRNA relative to total RNA remains unknown. To determine the fraction of sisRNA in the transcriptome, Northern blotting could be done to quantitatively measure the amount of each sisRNA, using a serial dilution of T7 in vitro transcribed RNA as a standard. Additionally, it is yet to be elucidated if polyadenylation occurs under other stress conditions besides heat stress. Thus, Northern blotting and RT-qPCR could be performed on samples subjected to various adverse conditions, including salt stress and CO2 stress. The experiments presented in this thesis point to the importance of introns and their sisRNAs in C. merolae, but do not provide insights into the mechanism(s) by which they benefit the cell. Broadly, introns or sisRNAs could act in cis by regulating the expression of their host genes. For example, S262 encodes a ribosomal protein and R289 a subunit of Complex I in the 156 electron transport chain. It would not be surprising if the cell were sensitive to changes in the expression level of these fundamentally important proteins. Alternatively, introns or sisRNAs could act in trans in a variety of ways: base pairing to targets to regulate their expression, sequestering proteins to keep them away from their other substrates, changing the localization of targets in the cell, etc. In addition, my observation of polyadenylation of some sisRNAs under heat stress again suggests a functional consequence, possibly relating to stability or cellular localization. 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The scatter plot with exponential trendlines shows the growth of T1 (blue) and S342∆i (green and orange for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.65 showed no difference in doubling time (DT) of S342∆i and T1. Whiskers show the standard deviation. P=0.16 Figure A3. Q3820∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and Q382∆i (orange and gray for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.16 showed no difference in doubling time (DT) of Q382∆i and T1. 166 P=0.05 Figure A4. C008∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and C008∆i (orange and green for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.05 showed no difference in doubling time (DT) of C008∆i and T1. Whiskers show the standard deviation. P=0.24 Figure A5. S270∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and S270∆i (orange and gray for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.24 showed no difference in doubling time (DT) of S270∆i and T1. 167 Figure A6. S270∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and J129∆i (orange and green for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 1.73Î10-4 showed a significant difference in the doubling time (DT) of that J129∆i and T1. Whiskers show the standard deviation. P=0.2 Figure A7. Q117∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and Q117∆i (green and orange for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.2 showed no difference in the doubling time (DT) of Q117∆i and T1. Whiskers show the standard deviation. 168 P= 0.01 Figure A8. E034∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and E034∆i (gray and orange for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.007 showed that E034∆i grew slower than T1. Whiskers show the standard deviation. P= 2.3Î10-4 Figure A9. S262∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and S262∆i (orange and green for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 2.3Î10-4 showed that S262∆i grew slower than T1. Whiskers show the standard deviation. 169 P= 0.03 Figure A10. Q270∆i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and Q270∆i (orange and gray for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 0.03 showed that Q270∆i grew slower than T1. Whiskers show the standard deviation. P= 7Î10-6 Figure A11. ∆6i growth test at 42 C. The scatter plot with exponential trendlines shows the growth of T1 (blue) and ∆6i (orange and green for ∆i.1 and ∆i.2) as indicated. Each strain had nine technical replicates. A one-way ANOVA with multiple comparisons using Tukey’s range test with a p-value of 7Î10-6 showed that ∆6i grew slower than T1. Whiskers show the standard deviation. 170 Figure A12. Lag time of Q382∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of Q382∆i.1 (A) and Q382∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (2 and 6 h) represent the lag time. 171 Figure A13. Lag time of D067∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of D067∆i.1 (A) and D067∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (0 and 2 h) represent the lag time. 172 Figure A14. Lag time of E034∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of E034∆i.1 (A) and E034∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (12 and 4 h) represent the lag time. 173 Figure A15. Lag time of Q270∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of Q270∆i.1 (A) and Q270∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (14 h) represent the lag time. 174 Figure A16. Lag time of S262∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of S260∆i.1 (A) and S260∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (8 h) represent the lag time. 175 Figure A17. Lag time of R289∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of R289∆i.1 (A) and R289∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (8 and 7 h) represent the lag time. 176 Figure A18. Lag time of ∆6i at 42 C The scatter plots with exponential trendlines show the exponential growth of ∆6i.1 (A) and ∆6i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (8h) represent the lag time. 177 Figure A19. Lag time of C008∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of C008∆i.1 (A) and C008∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (12 and 14 h) represent the lag time. 178 Figure A20. Lag time of S342∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of S342∆i.1 (A) and S342∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (12h) represent the lag time. 179 Figure A21. Lag time of K260∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of K260∆i.1 (A) and K260∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (12 and 7 h) represent the lag time. 180 Figure A22. Lag time of ∆10i at 42 C The scatter plots with exponential trendlines show the exponential growth of ∆10i.1 (A) and ∆10i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (16 and 14 h) represent the lag time. 181 Figure A23. Lag time of J129∆i at 42 C The scatter plots with exponential trendlines show the exponential growth of J129∆i.1 (A) and J129∆i.2 (B). Horizontal lines (blue) were drawn from the Y-axis (lnOD) to intersect the exponential phase trendlines, and the corresponding values on the X-axis (6 and 7 h) represent the lag time. 182