MULTIFACETED INVESTIGATION OF RIBONUCLEASE MRP IN
CYANIDIOSCHYZON MEROLAE: HEAT STRESS RESPONSE, CANONICAL rRNA
PROCESSING PATHWAY, STRUCTURAL PREDICTION AND MUTATIONAL
ANALYSIS

by

Sebastian Fumador
BSc., University of Health and Allied Sciences, 2020

THESIS SUBMITTED IN PARTIAL FULFILLMENT OF
THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
IN
BIOCHEMISTRY

UNIVERSITY OF NORTHERN BRITISH COLUMBIA
December 2024

© Sebastian Fumador, 2024

Abstract

RNase MRP is a ribonucleoprotein complex essential for ribosome biogenesis in eukaryotes
including Saccharomyces cerevisiae and humans. Mutations in the single genomic locus
encoding its noncoding RNA component result in cartilage-hair hypoplasia (CHH), a recessively
inherited developmental disorder. In Cyanidioschyzon merolae, the 442-nucleotide RNA
component of RNase MRP is encoded within the intronic region of the CMK142T gene. Under
heat stress conditions, this intronic region accumulates significantly, prompting investigations
into the effects of heat stress on RNase MRP expression and its role in 5.8S rRNA processing.
The impact on 5.8S rRNA processing of deleting the P19 region (∆372–405) and the G162A
mutation via homologous recombination was assessed using Northern blot analysis while
computational analyses were performed to compare structural conservation and protein
composition of RNase MRP in C. merolae with other eukaryotes. Total RNA analysis indicates
that deletion of the P19 region (∆372–405) of MRP significantly alters the stoichiometry of 5.8S
rRNA forms, underscoring its importance in rRNA processing. Additionally, C. merolae adheres
to the canonical rRNA processing pathway and while rDNA transcription is inhibited under heat
stress, the stability of mature 28S and 18S rRNA remains unaffected, indicating the organism's
sophisticated regulatory mechanism in ribosome biogenesis. Computational comparative
genomics analyses revealed conserved structural regions in C. merolae RNase MRP RNA,
highlighting evolutionary conservation. The complex in C. merolae is predicted to comprise five
proteins, fewer than the eleven in S. cerevisiae, reflecting dramatic streamlining of RNA
processing pathways which parallels findings in pre-mRNA splicing. These findings confirm the
conserved function of RNase MRP in C. merolae and raise important questions about why the
levels appear to increase under heat stress.
ii

Table of Contents
Abstract .......................................................................................................................................... ii
Table of Contents ......................................................................................................................... iii
List of Tables ................................................................................................................................ vi
List of Figures .............................................................................................................................. vii
List of Abbreviations .....................................................................................................................x
Acknowledgements .................................................................................................................... xiii
Dedication ................................................................................................................................... xiv
Chapter 1 - Introduction ...............................................................................................................1
1.1 RNase MRP ...............................................................................................................................2
1.2 Diseases Associated with RNase MRP ......................................................................................5
1.3 RNase MRP in Saccharomyces cerevisiae ................................................................................8
1.4 RNase MRP in Humans ...........................................................................................................10
1.5 RNase MRP in Drosophila melanogaster ...............................................................................11
1.6 RNase MRP in Cyanidioschyzon merolae ...............................................................................13
1.7 Heat Stress and Ribosomal RNA Biogenesis ..........................................................................15
1.8 Research Objectives .................................................................................................................16
Chapter 2 - Intronic Accumulation Induced by Heat Stress Does Not Modulate RNase
MRP Expression in C. merolae, Resulting in Unaltered Stoichiometry of 5.8S rRNA .........18
2.1 Introduction ..............................................................................................................................19
2.2 Materials and Methods .............................................................................................................19
2.2.1 Cultivation of C. merolae and Subsequent Heat Stress Treatment at 57 C ..........................19
2.2.2 Total RNA Isolation ..............................................................................................................20
2.2.3 RNA Integrity Assessment ...................................................................................................21
2.2.4 Fluorescence Northern Blot Analysis ...................................................................................21
2.2.4.1. 6%/7M Urea Denaturing Gel Preparation.........................................................................21
2.2.4.2. Denaturing Polyacrylamide Gels for RNA Analysis ........................................................22
2.2.4.3. Membrane Transfer with Semi-Dry Electroblotter/Capillary Transfer ............................23
2.2.4.4. RNA Cross-Linking for Membrane Stabilization Using the Strata Linker System..........24
2.2.4.5. Pre-hybridization...............................................................................................................24
2.2.4.6. Hybridization ....................................................................................................................24
iii

2.2.4.7. Washing Strategies for Enhanced Specificity ...................................................................25
2.2.4.8. Blocking Procedure for Northern Blot Analysis ...............................................................25
2.2.4.9. Fluorescent Labeling of Northern Blot; IRDye 800CW Streptavidin Binding ................25
2.2.4.10. Post-Streptavidin Binding Washing ................................................................................26
2.2.4.11. Detection .........................................................................................................................26
2.2.4.12. Northern Blot Analysis Utilizing Bio-Rad Image Lab 6.1 .............................................26
2.2.4.13. Blot Stripping and Storage for Subsequent Re-probing .................................................27
2.3. Results .....................................................................................................................................28
2.4. Discussion ...............................................................................................................................35
Chapter 3 – Heat Stress Inhibits rDNA Transcription but 18S and 28SrRNA Processing
Remains unaltered as C. merolae Adheres to the Canonical rRNA Processing Pathway .....37
3.1 Introduction ..............................................................................................................................38
3.2 Materials and Methods .............................................................................................................39
3.3. Results .....................................................................................................................................39
3.3.1 18S and 28S rRNA Northerns...............................................................................................39
3.3.2 Detection of Pre-rRNA Intermediates ..................................................................................42
3.3.2.1 Analysis of 5.8S and Processing Intermediates .................................................................43
3.3.2.2 Analysis of ITS1 and Processing Intermediates ................................................................44
3.3.2.3 Analysis of ITS2 and Processing Intermediates ................................................................45
3.3.2.4 28S and Processing Intermediates .....................................................................................47
3.4 Discussion ................................................................................................................................50
Chapter 4 - In Silico Prediction of C. merolae’s MRP RNA: Secondary Structure,
Conserved Regions, and Protein Constituents of the RNase MRP Complex. ........................53
4.1 Introduction ..............................................................................................................................54
4.2 Materials and Methods .............................................................................................................56
4.2.1 Genome and Protein Sequences ............................................................................................56
4.2.2 Alignment of C. merolae MRP RNA with Other Organisms ...............................................56
4.2.3 Identification of Protein Homologs ......................................................................................57
4.2.4 Validation of RNase MRP Protein Homologs via Reciprocal BLAST Searches .................58
4.3. Results .....................................................................................................................................59
4.3.1 Conserved Regions and Predicted Secondary Structure of RNase MRP RNA ....................59
iv

4.3.2 Identification of Protein Constituents of the RNase MRP Complex ....................................66
4.4. Discussion ...............................................................................................................................68
Chapter 5 - Mutational analysis of the RNA component of C. merolae RNase MRP Reveals
a Shift in the Stoichiometry of the Two Forms of 5.8S rRNA. ................................................70
5.1 Introduction. .............................................................................................................................71
5.2 Materials and Methods. ............................................................................................................73
5.2.1 Construction of Plasmid for Mutagenesis .............................................................................73
5.2.2 Construction of Knockout Plasmid .......................................................................................77
5.2.3 Construction of Transient and Integrable Plasmids ..............................................................82
5.2.4 Making of Mutant Plasmids ..................................................................................................86
5.2.5 Inserting Mutated Regions from PSR1127 into Integrable Plasmids ...................................88
5.2.6 Transformation of C. merolae ..............................................................................................93
5.2.7 Analysis of Colonies .............................................................................................................95
5.2.8 Northern Blot Analysis .........................................................................................................96
5.3. Results .....................................................................................................................................97
5.3.1 Plasmid Construction ............................................................................................................97
5.3.2 Northern Blot Analysis .......................................................................................................114
5.3.3 Establishing Plasmid Shuffling System in C. merolae .......................................................117
5.4. Discussion .............................................................................................................................125
Chapter 6 - General Conclusion and Remarks .......................................................................127

v

List of Tables
Table 2.1. Oligonucleotides used in Northern Blot Analysis ........................................................24
Table 3.1. Oligonucleotides used in Northern Blot Analysis ........................................................39
Table 3.2. Observed Bands in Canonical rRNA Processing: Northern Blot Data Summary ........48
Table 3.3. Canonical rRNA Processing Pathway: Expected vs. Observed Bands in C. merolae .50
Table 4.1: Predicted Protein Constituents of RNase MRP in C. merolae Identified Through
Comparative Genomic Analysis ....................................................................................................66
Table 5.1. Summary of NME1 Mutagenesis Data .........................................................................72
Table 5.2. Oligonucleotides used in the construction and sequencing of Plasmid Shuffling or
Direct Replacement vectors ...........................................................................................................92
Table 5.3. Primers Used to Verify Successful Genomic Integration ............................................96

vi

List of Figures
Figure 1.1. RNA components of RNase MRP and RNase P in S. cerevisiae ..................................3
Figure 1.2. Structure of the RNase MRP holoenzyme.....................................................................4
Figure 1.3. Function of the RNase MRP enzyme complex ............................................................6
Figure 1.4. Secondary structure of S. cerevisiae RNase MRP and Shift in the ratio of the 5.8S
rRNAs after either depletion of the RNase MRP RNA (NME1) or Temperature-sensitive
mutations ..........................................................................................................................................9
Figure 1.5. The Secondary Structure of D. melanogaster RNase MRP and 5.8S rRNA Processing
Impaired in dMRP mutants ............................................................................................................13
Figure 1.6. The eukaryotic RNA processing cascade integrates splicing, rRNA processing, and
translation .......................................................................................................................................15
Figure 2.1. Transcriptomic data depicts C. merolae CMK142T intron accumulation ..................19
Figure 2.2. C. merolae rRNA Cassette ..........................................................................................28
Figure 2.3. Isolation and Analysis of Total RNA from C. merolae Wild Type ............................29
Figure 2.4. The RNA component of RNase MRP is stably expressed at 42 C and 57 C ..............30
Figure 2.5. Detection of 5.8S rRNA in C. merolae Using Ethidium Bromide Stained 6% PAGE
........................................................................................................................................................32
Figure 2.6. Northern Analysis of 5.8S rRNA in C. merolae .........................................................33
Figure 2.7. Heat Stress Effect on 5.8S rRNA Intensities using 8% PAGE ...................................34
Figure 3.1. Canonical rRNA Processing Pathways .......................................................................38
Figure 3.2. Heat stress does not impact 18S rRNA processing .....................................................40
Figure 3.3. Heat stress does not impact 28S rRNA processing .....................................................41
Figure 3.4. Detection of 5.8S rRNA on Formaldehyde-Agarose Gel, illustrating intermediates
associated with the Canonical rRNA Processing Pathway ............................................................43
Figure 3.5. Detection of ITS1 on Formaldehyde-Agarose Gel, revealing intermediates
corresponding to the Canonical rRNA Processing Pathway ..........................................................44
Figure 3.6. ITS2 detection on Formaldehyde-Agarose Gel reveals intermediates corresponding to
the Canonical rRNA Processing Pathway......................................................................................45
Figure 3.7. 28S rRNA detection on Formaldehyde-Agarose Gel reveals intermediates
corresponding to the Canonical rRNA Processing Pathway ..........................................................47
vii

Figure 3.8: Pre-rRNA levels reduced at 57 C ................................................................................49
Figure 4.1: Sequence Alignment of C. merolae RNase MRP RNA with Drosophila melanogaster
RNase MRP RNA ..........................................................................................................................61
Figure 4.2: Sequence Alignment of C. merolae RNase MRP RNA with Chlamydomonas
reinhardtii RNase MRP RNA ........................................................................................................62
Figure 4.3: Sequence alignment of C. merolae RNase MRP RNA with Saccharomyces cerevisiae
RNase MRP RNA ..........................................................................................................................63
Figure 4.4: Sequence alignment of C. merolae RNase MRP RNA with Homo sapiens RNase
MRP RNA......................................................................................................................................64
Figure 4.5: Predicted Secondary Structure of C. merolae RNase MRP RNA ...............................65
Figure 5.1. RNA Analysis of NME1 Mutant Strains .....................................................................73
Figure 5.2. Modification of PCR2.1 with Multiple Cloning Sites .................................................74
Figure 5.3. Introduction of CMK142 Region into the Modified pCR 2.1 Vector .........................75
Figure 5.4. Construction of PSR1113 Vector with a Sulfadiazine Marker ...................................78
Figure 5.5. Introducing CMK142 Homology Arms into PSR1113 ...............................................80
Figure 5.6 Introducing CMK142 region into the PacI site of PSR887 ..........................................83
Figure 5.7 Introducing CMK142 region into the PacI site of PSR886 ..........................................84
Figure 5.8 Introducing a Second Homology Arm into the SwaI site of PSR1119 ........................85
Figure 5.9 Introducing a ∆372 - 405 Mutant into PSR1124 ..........................................................89
Figure 5.10 Linear transformation product amplified from PSR1128 ...........................................90
Figure 5.11 Introducing a G162A Mutant into PSR1124 ..............................................................91
Figure 5.12 Linear transformation product amplified from PSR1130 ...........................................91
Figure 5.13. Restriction Digest Analysis of pCR2.1 and pSR1126 ...............................................98
Figure 5.14. Restriction Digest Analysis of Insert DNA and pSR1127 ........................................99
Figure 5.15. Construction and Restriction Digest Analysis of pSR1113 ....................................100
Figure 5.16. Insertion of CMK142-derived DNA fragment into the SwaI site of the pSR1113
vector ...........................................................................................................................................101
Figure 5.17. Construction and Restriction Digest Analysis of pSR1117 ....................................102
Figure 5.18. Construction and Restriction Digest of two shuffle plasmids with distinct selectable
markers .........................................................................................................................................103
Figure 5.19. Construction and Restriction Digest of pSR1124 ...................................................104
viii

Figure 5.20 Predicted secondary structure of RNase MRP RNA in C. merolae with mutation sites
highlighted ...................................................................................................................................105
Figure 5.21 PCR results from site-directed mutagenesis of RNase MRP RNA in C. merolae ...106
Figure 5.22 PCR analysis of mutagenized plasmids compared to a non-mutagenized control,
displayed on a 2% agarose gel stained with ethidium bromide ...................................................107
Figure 5.23 Restriction enzyme digestion and ligation process for constructing pSR1128 and
pSR1130.......................................................................................................................................108
Figure 5.24. Restriction Digest of pSR1128 and pSR1130. ........................................................109
Figure 5.25: Amplification of linear DNA from plasmids used for the transformation of C.
merolae ........................................................................................................................................110
Figure 5.26 Initial colony PCR screening for genomic integration of pSR1128 (∆372–405) .....111
Figure 5.27. Final PCR confirming the successful genomic integration of pSR1128 (∆372–405)
into CMK142 locus ......................................................................................................................112
Figure 5.28 Initial colony PCR screening for genomic integration of pSR1130 (G162A) .........113
Figure 5.29. Final PCR confirming the successful genomic integration of pSR1130 (G162A) into
CMK142 locus .............................................................................................................................114
Figure 5.30 Changes in the stoichiometry of the two forms of 5.8S rRNA following RNase MRP
RNA mutations in C. merolae ....................................................................................................115
Figure 5.31 Amplification of linear DNA from pSR1117 used for the transformation of C.
merolae ........................................................................................................................................118
Figure 5.32. Selection of transformed cells with sulfadiazine .....................................................119
Figure 5.33 Growth of transformed C. merolae cells with Increasing sulfadiazine concentration
......................................................................................................................................................120
Figure 5.34 Initial colony PCR screening for genomic integration of pSR1117 .........................121
Figure 5.35. PCR confirming genomic integration of pSR1117 .................................................122
Figure 5.36. Chloramphenicol resistance in transformed C. merolae cells .................................123
Figure 5.37. Final PCR analysis revealed unsuccessful integration of pSR1117 into the CMK142
locus ............................................................................................................................................124

ix

List of Abbreviations
aa – amino acid(s)
APS – ammonium persulfate
ATP – adenosine triphosphate
bp – base pair(s)
CHH – Cartilage hair hypoplasia
C. merolae – Cyanidioschyzon merolae strain 10D
CR – Conserved Region
CAT – Chloramphenicol Acetyltransferase
dATP – deoxyadenosine triphosphate
dCTP – deoxycytidine triphosphate
dGTP – deoxyguanosine triphosphate
DMSO – dimethyl sulfoxide
DNA – deoxyribonucleic acid
dNTPs – deoxynucleotide triphosphates, consisting of dATP, dCTP, dGTP, and dTTP
dTTP – deoxythymidine triphosphate
EDTA – Ethylenediaminetetraacetic acid
ETS1 – Externally Transcribed Spacer 1
ETS2 – Externally Transcribed Spacer 2
GTP – guanosine triphosphate
Int – Integrands
ITS1 – Internally Transcribed Spacer 1

x

ITS2 – Internally Transcribed Spacer 2
kbp – kilobase pairs
knt – kilonucleotides
LB – Luria broth
MA2 – modified Allen’s 2×
MA2G – MA2 with glycerol
MA2GU – MA2G with uracil
MCS – multiple cloning site
mRNA – mature messenger RNA
MRP – Mitochondrial RNA Processing
NaOAc – sodium acetate
NCBI – National Center for Biotechnology Information
ncRNA – non-coding RNA
Nc – Negative Control
nt – nucleotide(s)
NTPs – nucleotide triphosphates, consisting of ATP, CTP, GTP, and UTP
OD750 – optical density at 750 nm
OMPD – orotidine 5′-monophosphate decarboxylase
OPRT – orotate phosphoribosyltransferase
PCR – polymerase chain reaction
PEG – polyethylene glycol
Pop – Processing of Precursor
pre-mRNA – precursor messenger RNA

xi

RMRP – RNase MRP
pre-tRNA – precursor tRNA
RNA – ribonucleic acid
RNA-seq – RNA sequencing
RNase – Ribonuclease
rpm – revolutions per minute
rRNA – ribosomal RNA
Sd – Sulfadiazine
SDS – Sodium Dodecyl Sulfate
snoRNA – small nucleolar ribonucleic acid
snRNA – small nuclear ribonucleic acid
snRNP – small nuclear ribonucleoprotein
TERT – telomerase reverse transcriptase
tRNA – transfer RNA
UniProt – Universal Protein Resource
UTP – uridine triphosphate

xii

Acknowledgments
I acknowledge that this research was conducted on the traditional, unceded territory of the
Lheidli T’enneh First Nation, part of the Dakelh (Carrier) peoples’ territory. I am grateful for the
opportunity to live, learn, and work on their lands.
I would like to express my deepest gratitude to my supervisor, Dr. Stephen Rader, for his
unwavering support, guidance, and encouragement throughout my research journey. His
expertise, patience, and insightful feedback have been instrumental in shaping both my thesis
and my development as a scientist.
I extend my sincere appreciation to Dr. Martha Stark for her patience in training me and for her
invaluable help with technical challenges and troubleshooting. Her support made an enormous
difference in my lab work.
I would like to thank my Graduate Supervisory Committee, Dr. Andrea Gorrell and Dr. Michael
Preston, for their valuable feedback, advice, and encouragement. Their thoughtful guidance
motivated me throughout this journey, and I appreciate their unwavering support. I would also
like to extend my sincere thanks to my defense chair, Dr. Catharine Schiller and my external
examiner, Dr. Marlene Oeffinger.
I am grateful to my colleagues in the Rader Lab for their friendship, collaboration, and the
stimulating discussions that made my time in the lab both enjoyable and intellectually enriching.
Your camaraderie has been an essential part of my graduate experience.

xiii

Dedication
This thesis is dedicated to my mother, Rosemary Afi Mattey, whose unwavering love, support,
and encouragement have been my constant source of strength.

xiv

Chapter 1 – Introduction

1

1.1 RNase MRP
RNase MRP (ribonuclease for mitochondrial RNA processing) is an essential ribonucleoprotein
endoribonuclease that cleaves RNA substrates in a site-specific manner and comprises a catalytic
RNA moiety and multiple (ten in Saccharomyces cerevisiae) protein components (Karwan et
al.,1991). RNase MRP is an essential eukaryotic enzyme that has been found in practically all
eukaryotes analyzed (Piccinelli et al., 2005). It is localized to the nucleolus and, transiently, to
the cytoplasm (Esakova et al., 2010). RNase MRP appears to have split from the RNase P
lineage early in the evolution of eukaryotes, acquiring distinct substrate specificity and cellular
functions (Piccinelli et al., 2005; Rosenblad et al., 2006; Lopez et al. 2009). The catalytic (C)
domain of RNase MRP RNA (Figure 1.1a) has a secondary structure resembling that of the Cdomain of RNase P (Figure 1.1b) and includes elements forming a highly conserved catalytic
core. The specificity (S) domain of RNase MRP RNA does not have any apparent similarities
with the specificity domain of RNase P (Figures 1.1a and b). Crosslinking studies (Esakova et
al., 2013) indicate the involvement of the RNase MRP S-domain in substrate recognition. Most
of the RNase MRP protein components are also found in eukaryotic RNase P (Chamberlain et
al., 1998); the structures of S. cerevisiae and human RNases P have been determined (Lan, P. et
al., 2018; Wu, J. et al. 2018). S. cerevisiae RNase MRP and RNase P share eight proteins (Pop1,
Pop3, Pop4, Pop5, Pop6, Pop7, Pop8, and Rpp1 (two copies); RNase MRP protein Snm1 has a
homolog in RNase P (Rpr2), while Rmp1 is found only in RNase MRP. Shared proteins bind to
both catalytic (C) and specificity (S)-domains. Yeast RNase MRP proteins Pop1, Pop3, Pop4,
Pop5, Pop6, Pop7, and Pop8 have homologs in human RNase P, while Pop3, Pop4, Pop5, and
Rpp1 have homologs in archaeal RNases P. RNase MRP proteins Pop1, Pop6, Pop7 are also an
essential part of yeast telomerase, where they are involved in the localization of the enzyme and

2

form a structural module that stabilizes the binding of telomerase components Est1 and Est2
(Lemieux et al., 2016; Garcia et al. 2020).

Figure 1.1. RNA components of RNase MRP and RNase P in S. cerevisiae. The catalytic (C-) domains of the two
related enzymes are similar both in their secondary structures and in their folds, whereas the specificity (S-) domains
are distinct. a, b Secondary structure diagrams of the RNase MRP and RNase P RNAs, respectively. c, d Folding of
the RNase MRP and RNase P RNAs, respectively, color-coded as in (a, b). Adapted from Lan et al.,2018.

In a study published in 2020, a cryo-EM structure of the S. cerevisiae RNase MRP holoenzyme
was resolved to a nominal resolution of 3.0 Å, providing new insights into its molecular
architecture (Figure 1.2). It elucidates the overall structural organization of the ribonucleoprotein
(RNP) complex, highlighting the arrangement of its catalytic RNA component, the substrate
binding pocket, and the intricate interactions among RNase MRP components. A significant
aspect of the research is the comparative analysis between RNase MRP and its evolutionary
progenitor, eukaryotic RNase P. This comparison reveals that several proteins common to both
3

RNase MRP and RNase P undergo RNA-driven structural remodeling, enabling these
ribonucleoproteins to function within distinct structural contexts (Figure 1.2) (Perederina et
al.,2020). Notably, while the catalytic center of RNase MRP closely mirrors that of RNase P,
there is a divergence in the topology of the substrate binding pocket, suggesting functional
specialization within these closely related enzymes. These findings contribute to a deeper
understanding of the structural and functional dynamics of RNase MRP, highlighting its role in
cellular processes and its evolutionary relationship with RNase P (Perederina et al.,2020).

Figure 1.2. Structure of the RNase MRP holoenzyme. Protein components (shown as surfaces) are color-coded as
marked; the RNA elements (shown as a cartoon) are color-coded according to Fig.1. Adapted from Perederina et al.,
2020.

4

1.2 Diseases Associated with RNase MRP
There are over 17, 000 human genetic disorders listed in the database of Online
Mendelian Inheritance in Man (OMIM), and majority of these disorders relate to protein-coding
genes, whereas only a few noncoding RNA genes have been linked to genetic diseases. Whilst
noncoding RNA is an RNA molecule that functions without being translated into a protein,
certain diseases are associated with noncoding RNAs. RNase MRP is a noncoding RNA
involved in mitochondrial DNA replication (Figure 1.3a), pre-rRNA processing (Figure 1.3b and
d), and processing of 5′-UTR of CLB2 mRNA (Figure 1.3c).

5

Figure 1.3. Function of the RNase MRP enzyme complex. (a) RNase MRP is involved in the processing of
mitochondrial RNA that functions as a primer for mitochondrial DNA replication in mitochondria. Transcription
starts from the light-strand promoter by mitochondrial RNA polymerase. After transcription of the heavy-strand
origin of replication, the transcript remains bound to the DNA duplex and is cleaved by RNase MRP to form primers
that are used for the initiation of DNA synthesis by DNA polymerase; Adapted from (Shadel et al.,1997). (b, d)
RNase MRP functions in the pre-rRNA processing in S. cerevisiae. The 35S primary transcript is processed into
mature 25S, 18S, and 5.8S rRNAs; Adapted from (Venema and Tollervey 1999). The cleavage sites (A0 through E),
the external transcribed spacers (5′-ETS and 3′-ETS), and the internal transcribed spacers (ITS1 and ITS2) are
indicated. The small white box marks the sequence in the long form of 5.8S rRNA, 5.8SL, that is absent in the short
form, 5.8SS. RNase MRP processes the A3 site in ITS1; Adapted from (Schmitt and Clayton,1992). (c) RNase MRP
processes the 5′-UTR of CLB2 mRNA in cytoplasmic temporal asymmetric MRP (TAM) bodies. CLB2 mRNA
normally disappears rapidly as cells complete mitosis. RNase MRP mutations have an exit-from-mitosis defect and a
late anaphase delay. RNase MRP specifically cleaves the CLB2 mRNA in its 5′-UTR to allow rapid 5′ to 3′
degradation by the Xrn1 nuclease. Degradation of the CLB2 mRNA by RNase MRP provides a novel way to
regulate the cell cycle that complements the protein degradation machinery; Adapted from (Allison and Yong 2006).

Mutations to the single genomic locus for this noncoding transcript cause inviable yeast (Shadel
et al. 2000), embryonic lethality in mice (Rosenbluh et al. 2011), and a spectrum of severely
debilitating human diseases (Ridanpää et al. 2001), harkening to MRP RNA’s essential role in
6

biology. Among these conditions is Cartilage Hair Hypoplasia (CHH), a pleiotropic human
disease (Hirose et al.,2006). Two categories of mutations involving RNase MRP have been
identified in patients with CHH. The first type is when an insertion, duplication,
or triplication occurs at the promoter of the RNase MRP gene between the TATA box and the
transcription initiation site. This causes the initiation of RNase MRP to be slow, or to not occur
at all. The second category consists of mutations that are in the transcribed RNA made by the
RNase MRP. Patients with CHH have been identified to have over 70 different mutations in the
RNA transcript made by RNase MRP, whereas around 30 distinct mutations have been identified
in the promoter region of the RNase MRP gene. Most CHH patients have a combination of either
a promoter mutation in one allele along with an RNase MRP RNA mutation in the other allele, or
a combination of two RNase MRP RNA mutations in both alleles. The fact that there is not often
a mutation in the promoter region in both alleles shows the lethality of not having this RNA
present that is transcribed by RNase MRP (Hermanns et al.,2006).

7

1.3 RNase MRP in Saccharomyces cerevisiae
In Saccharomyces cerevisiae, the RMRP ortholog NME1 (nuclear mitochondrial
endonuclease 1) showed an essential role in cell viability, indicating a nuclear role for RNase
MRP (Schmitt and Clayton 1992). RNase MRP cleaves the pre-rRNA at the B2 cleavage site in
yeast pre-rRNA, which is thought to be the functional equivalent of site 2 in humans (Schmitt
and Clayton 1993). Its secondary structure has been determined (Figure 1.4a). Further,
conditional depletion of the RNA component of the enzyme (Figure 1.4b) showed that this is
responsible for the maturation of 5.8S rRNA (Schmitt and Clayton 1992). It was found that there
was a reversal in the stoichiometry of the two mature forms (long and short) of 5.8S rRNA
(Figure 1.4c), a component of the large ribosomal subunit. In the MRP RNA depleted condition
via the utilization of glucose-repressed GAL1 promoter, the 7-nucleotide-longer version of 5.8S
rRNA was 10 times more abundant than the shorter species lacking this 7-nucleotide sequence at
the 5’ end, and the accumulation of an aberrant rRNA precursor (a defective RNA intermediate
that deviates from the normal processing pathway) (Schmitt and Clayton 1993). These results
contrasted with the normal stoichiometry in which the shorter version of 5.8S rRNA is 10-fold
more abundant than the slightly longer version. Also, the NME1 temperature-sensitive mutants
show the same rRNA processing defect (Figure 1.4d). Literature has revealed that a particular A
to G transition at position 122 in the RNA sequence defines its functional capacity (Shadel et
al.,2000). High-copy suppressor analysis of this point mutation led to the identification of
interacting proteins, and SNM1 was the first identified protein component unique to the RNase
MRP enzyme complex. The protein contains a leucine zipper motif, a zinc-cluster motif, and a
serine/lysine-rich tail (Schmitt and Clayton 1994). Another role has been assigned to the RMRP
by observing a delay in the progression of the cell cycle at the end of mitosis in some nme1
8

mutants (Cai et al., 2002). This is caused by an increase in CLB2 (B-type cyclin) mRNA levels
leading to increased Clb2p (B-cyclin) levels and a resulting late anaphase delay. Normally, the
RNase MRP complex cleaves the 5’ UTR of CLB2 mRNA, which, in turn, causes rapid
degradation of CLB2 mRNA and efficient cell cycle progression (Gill et al., 2004).

A

D

C

B

Figure 1.4. Secondary structure of S. cerevisiae RNase MRP and Shift in the ratio of the 5.8S rRNAs after
either depletion of the RNase MRP RNA (NME1) or Temperature-sensitive mutations: a) Structure of S.
cerevisiae’s RNase MRP (Marcela et al.,2009). (b) Depletion was induced using a glucose-repressed GAL1
promoter. Following the shift to a glucose-containing medium, samples were collected every 4 hours for total RNA
extraction. After gel electrophoresis and transfer to nylon membranes, the RNA was probed for SCR1 (yeast signal
recognition particle RNA, serving as a loading control) and RNase MRP RNA (NME1), or with a probe targeting
the ITS1 region of the rRNA precursor. c) A shift in the ratio of 5.8S rRNAs was observed following the depletion
of RNase MRP RNA (NME1). RNA was isolated from yeast cells grown in glucose and analyzed by ethidium
bromide staining after PAGE. No changes were detected in the tRNA or 5S rRNA profiles; however, a significant
shift in the 5.8S rRNA ratio was evident. d) NME1 conditional mutants show the same rRNA processing defect.
Yeast strains were grown at the permissive temperature (24 C) and then shifted to the nonpermissive temperature
(37 C) for 6 h. Total RNA was prepared, fractionated by PAGE, and then visualized with ethidium bromide (Schmitt
and Clayton 1993).

9

1.4 RNase MRP in Humans
In humans, the role of the RNA component of the RNase MRP complex (RMRP) in prerRNA processing has long been established. However, the precise details of its function were not
fully understood until CRISPR/Cas9-mediated deletions of the RMRP gene provided more
insights. These experiments revealed that RMRP directs the cleavage at site 2 in ITS1 of human
pre-rRNA, highlighting its role in ribosomal RNA maturation (Goldfarb et al., 2017). This
finding solidified RMRP's involvement in processing pre-rRNA, which is essential for ribosome
assembly and cell viability. RMRP is the RNA component of the RNase MRP (ribonuclease
mitochondrial RNA processing) complex, a ribonucleoprotein endonuclease. The enzyme was
first identified in mice for its ability to cleave mitochondrial RNA, which serves as a primer for
mitochondrial DNA replication (Chang et al., 1987). Initially thought to be primarily
mitochondrial, subsequent studies showed that RMRP is nuclear-encoded and predominantly
localized in the nucleolus, suggesting a broader role beyond mitochondria (Reimer et al., 1988).
In humans, RMRP is 267 nucleotides long, sharing 84% sequence homology with the mouse
RMRP gene. Interestingly, the conservation extends beyond the coding region, as approximately
700 nucleotides of the 5’-flanking regions are also conserved, indicating the importance of
regulatory elements for the expression of RMRP (Topper and Clayton, 1990).
The high degree of sequence conservation across species, including humans, mice, rats, cows,
Xenopus, yeast, Arabidopsis, and tobacco, underscores the essential nature of RMRP's function
(Schmitt et al., 1993). The length of the RMRP transcript varies between species, but its
conserved core structure is essential for its function. Structural models of RMRP have revealed a
complex secondary structure, which helps in the assembly and functionality of the
10

ribonucleoprotein complex (Walker and Avis, 2004). These conserved structural elements, such
as stem-loops and internal bulges, are thought to facilitate interactions with the protein subunits
of RNase MRP, which in turn are required for the enzyme's catalytic activity and proper
localization. In addition to its role in rRNA processing, RNase MRP has been implicated in
several other cellular processes, including the regulation of cell cycle progression and
mitochondrial DNA replication. For instance, mutations in the RMRP gene are associated with a
variety of human diseases, including cartilage-hair hypoplasia (CHH), an autosomal recessive
disorder characterized by skeletal dysplasia, immunodeficiency, and increased cancer
susceptibility caused by mutations in the transcribed RNA made by RNase MRP or an insertion
or duplication at the promoter of RNase MRP (Ridanpää et al., 2001). These mutations often
affect the secondary structure of RMRP, leading to disrupted ribonucleoprotein assembly and
altered cleavage activity, which in turn impairs ribosome biogenesis and cellular
proliferation. The nucleolar localization of RMRP, observed through immunolocalization
studies, further underscores its involvement in ribosome biogenesis within the nucleolus (Reimer
et al., 1988). This localization is consistent with its role in pre-rRNA cleavage, where it interacts
with other key factors involved in the maturation of 5.8S, 18S, and 28S rRNAs. Overall, the
RNase MRP complex plays an indispensable role in human cellular function, particularly in
ribosome biogenesis. The continued study of its RNA component, RMRP, is not only important
for understanding fundamental aspects of rRNA processing but also for shedding light on the
molecular underpinnings of human diseases associated with RMRP dysfunction.
1.5 RNase MRP in Drosophila melanogaster
In Drosophila, the expression of the Drosophila ortholog of MRP RNA (CR33682),
which was predicted by a bioinformatics screen for MRP RNA sequences (Piccinelli et al. 2005)
11

has been reported (Figure 1.5). Characterization of a mutant strain shows that Drosophila MRP
(dMRP) is an essential gene. dMRP mutants display a severe impairment in growth, a
characteristic shared with human diseases carrying mutations in this gene (Martin and Li 2007).
These phenotypic defects were attributed to impairments at different stages of rRNA processing
that were observed. These include the classic defect in processing 5.8S rRNA (Figure 1.5a and b)
that has been associated with human and S. cerevisiae RNase MRP mutants (Schmitt and
Clayton 1993; Lygerou et al. 1996; Hermanns et al. 2005; Thiel et al. 2005), as well as a defect
in early rRNA processing similar to a defect reported by Lindahl et al. 2009 in S. cerevisiae.
Expression of dMRP RNA was detected throughout the Drosophila life cycle. This is consistent
with its role in fundamental cellular processes such as ribosome biogenesis, mitochondrial DNA
replication, and cell cycle regulation (Chang and Clayton 1987; Schmitt and Clayton 1993;
Lygerou et al. 1996; Gill et al. 2004; Thiel et al. 2007). Results from Mary et al., (2010) support
the idea that dMRP RNA shares structural and functional homology with conserved MRP RNA
genes previously characterized in other eukaryotes. The first characterization of the ribosomal
RNA processing pathway in Drosophila by Long and Dawid (1980) identified a single form of
5.8S rRNA. However, extensive characterization studies done by Mary et al., (2010) identified
both long and short forms of 5.8S rRNA in normal larvae. The data also reveal similarities in
rRNA processing between dMRP RNA and MRP RNA orthologs in other species. Homozygous
dMRP mutants display a similar change in relative abundances of the two forms of 5.8S rRNAs,
indicating a similar function for this gene in Drosophila. Normally, the pre-rRNA transcript is
cleaved at defined sites in a consistent order to produce a defined set of rRNA intermediates that
are ultimately processed into mature rRNAs.

12

C

Figure 1.5. The secondary structure of D. melanogaster RNase MRP and 5.8S rRNA processing impaired in
dMRP mutants. Total RNA isolated from wild-type (WT) and dMRPEY08633 mutant larvae were separated in a
denaturing polyacrylamide gel and either directly analyzed by staining with ethidium bromide (a) or used for
Northern blotting with probes specific to the dMRP RNA or 5.8 rRNA (b). The two forms of 5.8S rRNA are
indicated (Schneider et al., 2010). (c) Secondary structure of RNase MRP in Drosophila melanogaster (Piccinelli et
al., 2005).

1.6 RNase MRP in Cyanidioschyzon merolae
Cyanidioschyzon merolae (C. merolae) is a thermophilic and acidophilic red alga that thrives in
hot springs, characterized by extreme conditions of 45 C and a pH of 1.5. This microorganism
has a cell length of ~2 µm (Matsuzaki et al., 2004) and a compact genome of approximately 16.5
million base pairs (Nozaki et al., 2007). Interestingly, concerning splicing, C. merolae strain 10D
exhibits a simplified spliceosome, notably lacking the U1 small nuclear ribonucleoprotein
(snRNP) in its spliceosome complex (Stark et al., 2015). Splicing involves the excision of introns
and the ligation of exonic regions to produce mature mRNA, which is essential for protein
13

synthesis (Figure 1.6). However, some of these introns excised during splicing contain snoRNAs
(small nucleolar RNAs), which in turn modify snRNA, tRNAs, and rRNAs (Figure 5). The
reduction in splicing machinery (Stark et al., 2015), raises intriguing questions about the
evolutionary simplification, functional adaptation, and conserved mechanisms of RNase MRP
complex in C. merolae. Also, the organism's ability to thrive in extreme environments raises
questions about the resilience and adaptability of RNase MRP under stress. C. merolae exhibits
an extremely simple cytological genomic architecture, this feature offers advantages in
cytological and biochemical studies (Fujiwara 2017) and positions C. merolae as a valuable
model organism for elucidating the complexities of RNase MRP and its broader implications in
cellular biology. Although RNase MRP has been extensively studied in a wide range of
organisms, relatively little is known about this complex in C. merolae. However, studies have
identified a putative MRP RNA gene in the C. merolae genome, suggesting that this organism
also possesses an RNase MRP complex. It is unclear how the MRP RNA gene in C. merolae is
processed or how the MRP RNA complex functions in this organism, however, given the
conservation of MRP RNA in other eukaryotes and the importance of RNase MRP for ribosome
biogenesis (Piccinelli et al.,2005), it seems likely that the MRP RNA complex in C. merolae
plays a similar role in processing rRNA and maintaining cell growth and proliferation as reported
in other organisms. In C. merolae, the MRP RNA is located in the intronic region of the noncoding CMK142T gene, as in D. melanogaster and C. elegans. Repeated efforts in our lab to
delete the intronic region from the CMK142T gene have been unsuccessful (Rader Lab
unpublished data), suggesting that MRP is essential.

14

Figure 1.6. The eukaryotic RNA processing cascade integrates splicing, rRNA processing, and translation.
The spliceosome, composed of snRNAs and proteins, excises introns from pre-mRNA, releasing mature mRNA and
introns. Some introns harbor snoRNAs, which subsequently modify snRNAs, tRNAs, and rRNAs. RNase P cleaves
pre-tRNA, while RNase MRP targets rRNA. The ribosomal complex, formed by rRNAs, facilitates the interaction
between tRNAs and mature mRNAs during translation. Image adapted from Woodhams et al., 2007.

1.7 Heat Stress and Ribosomal RNA Biogenesis
Ribosomal RNAs (rRNAs) are the essential structural and functional components of ribosomes
involved in protein synthesis. The rRNA gene clusters, referred to as rDNA, differ slightly across
species: 35S in yeast, 45S in plants, and 47S in mammals. These rDNA units encode the 18S,
5.8S, and 25S rRNAs (with 28S rRNA in mammals). Each rDNA unit comprises external
transcribed spacers (5'ETS and 3'ETS) and the 18S, 5.8S, and 25S/28S rRNA sequences, which
are interspersed by internal transcribed spacers (ITS1 and ITS2). These rDNA units are
transcribed by RNA polymerase I (Pol I) within the nucleolus, generating a single precursor
transcript - 35S in yeast, 45S in plants, and 47S in mammals. This precursor rRNA undergoes a
15

series of processing steps, including exonucleolytic and endonucleolytic cleavages, to remove
the ETS and ITS regions, ultimately yielding the mature 18S, 5.8S, and 25S/28S rRNAs.
Additionally, specific RNA modifications occur at designated positions during this processing to
ensure proper ribosome function (Sharma and Lafontaine 2015; Henras et al. 2015; Sloan et al.
2017; Tomecki et al. 2017). Environmental and cell stress conditions induce known changes in
nucleolar morphology and functions (Boulon et al. 2010; Hayashi and Matsunaga 2019; Kalinina
et al.,2018), however, the impact of heat stress on the processing of pre-rRNAs remains poorly
investigated. In mammals, a short heat shock inhibits pre-rRNA transcription and processing into
mature rRNAs (Ghosha and Jacob 1996), while 40 min exposure at 43 C causes accumulation of
30SL pre-RNAs from the ITS1-first pathway (Coccia et al.,2017). Heat stress is known to inhibit
rDNA transcription in animal cells (Ghosha and Jacob 1996; Coccia et al.,2017), whereas in
Drosophila, heat shocks increase RNA pol I transcription of retrotransposons located in rDNA
clusters (Raje et al., 2018).
In Arabidopsis thaliana, studies have shown that heat stress disturbs nucleolar structure, inhibits
pre-rRNA processing, and provokes imbalanced ribosome profiles. Upon heat stress, precursors
of 18S, 5.8S, and 25S RNAs are rapidly undetectable in A. thaliana (Darriere et al., 2022).
1.8 Research Objectives
Considering the complexity of elucidating the essential function of MRP RNA in human cells, a
much simpler organism with fewer components such as C. merolae studied in the Rader Lab is
of considerable interest in investigating the function of RNase MRP in ribosome biogenesis. It
has been reported in S. cerevisiae that at a nonpermissive temperature in temperature-sensitive
mutants of the MRP RNA, there is a reduction in the catalytic activity of RNase MRP leading to
a defect in the synthesis of the two forms of 5.8S rRNAs required in protein synthesis (Schmitt
16

and Clayton 1993). However, in C. merolae, the intronic region of the non-coding CMK142T
gene that houses MRP, turns out to be the most accumulated when exposed to heat stress at 57 C
(Rader Lab unpublished data). This raises the possibility that the accumulation of RNase MRP
during heat stress at 57 C in C. merolae may result in a 5.8S ribosomal RNA processing defect.
Also, building upon the literature review addressing the impact of heat stress on rDNA
transcription and processing of mature rRNA, it's interesting to verify whether heat stress
impacts these processes in C. merolae.
The objectives of this thesis are fourfold: First, to determine if there is a defect in 5.8S rRNA
processing via Northern blot analysis, potentially due to hypothesized modulation of RNase
MRP catalytic activity under heat stress and heat stress impact on mature rRNAs (28S and 18S).
Second, to verify whether C. merolae subscribes to the canonical rRNA processing pathway and
to evaluate the impact of heat stress on the precursors of this pathway. Third, to employ
bioinformatics tools to predict the secondary structure of C. merolae RNase MRP, identify
conserved regions through comparative genomics, and predict the protein constituents of the
RNase MRP complex in C. merolae. Finally, to elucidate the function of C. merolae RNase
MRP by conducting mutational analysis of its RNA component, using plasmid shuffling or direct
replacement via homologous recombination.

17

Chapter 2 - Intronic Accumulation Induced by Heat Stress Does Not
Modulate RNase MRP Expression in C. merolae, Resulting in Unaltered
Stoichiometry of 5.8S rRNA

18

2.1 Introduction
In C. merolae, the intronic region of the CMK142T gene, which harbors RNase MRP, appears to
accumulate under heat-stress conditions (Figure 2.1). This chapter delves into the effects of heat
stress on RNase MRP expression and its catalytic function in cleaving ITS1, leading to the
generation of two 5.8S rRNA isoforms. Furthermore, given that heat stress has been shown in
other species to alter ribosomal profiles, the chapter examines the impact of heat stress on mature
5.8S rRNAs in C. merolae. Northern blot analysis was utilized in this chapter to assess the
expression levels of RNase MRP and 5.8S rRNA.

Figure 2.1. Transcriptomic data depicts C. merolae CMK142T intron accumulation. The intronic region of
CMK142T is the first to accumulate under heat stress at 57 C. This intronic region houses the RNA component of
RNase MRP (Schubert Lab unpublished data).

2.2 Materials and Methods
2.2.1 Cultivation of C. merolae and Subsequent Heat Stress Treatment at 57 C
The cultivation protocol for C. merolae was based on the method outlined by Kobayashi et al.
(2010). The cells were grown in liquid MA2G medium, which consists of 40 mM (NH4)2SO4, 8
mM KH2PO4, 4 mM MgSO4, 1 mM CaCl2, 184 µM H3BO3, 100 µM FeCl3, 80 µM Na2EDTA,
19

36 µM MnCl2, 6.4 µM Na2MoO4, 3.08 µM ZnCl2, 1.2 µM CuCl2, 0.68 µM CoCl2, and 50 mM
glycerol. Cultures were maintained at 42 C with 2% CO2 and continuous illumination at 90 µmol
photons·m⁻²·s⁻¹. Wild-type C. merolae cells were grown to an OD750 =1.0 under standard
conditions. Cells were then transferred to a 57 C water bath for 1 h for heat stress or 42 C for
controls.
2.2.2 Total RNA Isolation
RNA extraction was carried out using the cold phenol method with phase-lock gel (PLG) tubes.
After heat stress treatment, the cells were centrifuged at 15,000g for 2 minutes, the supernatant
was discarded and resuspended the cells in 300 µL of cold phenol lysis buffer (200mM TrisHCL, pH7.5, 500mM NaCl, 10mM EDTA, 1%SDS) in a 1.5 mL Eppendorf tube. The cells were
sonicated for 2 bursts of 5-10 seconds, followed by the addition of 300 µL of acid-phenol,
vortexed for 5 seconds, and then spun at maximum speed (12,000-16,000g) for 20-30 seconds.
The mixture of acid-phenol and cell lysate were transferred to a PLG microtube and centrifuged
at 15,000g for 5 minutes. 300 µL of acid-phenol was added with gentle mixing before spinning
again for 5 minutes at 15,000g. The aqueous phase was then transferred to a new Eppendorf
tube, mixed with 300 µL of chloroform, vortexed for 5 seconds, and spun for 5 minutes at
15,000g. The aqueous phase was transferred to a new tube, 1 mL of 100% ethanol was added,
and centrifuged at 4 C for 30 minutes at maximum speed, followed by discarding the
supernatant. The pellet was washed with 180 µL of cold 70% ethanol, centrifuged for 1 minute at
maximum speed, and all liquid was carefully aspirated off. The pellet was allowed to dry for
about 5 minutes, or longer if needed, before being resuspended in 25 µL of dH2O. 1 µL of the
sample was used to determine the concentration with a Nanodrop spectrophotometer.

20

2.2.3 RNA Integrity Assessment
RNA integrity was assessed using a 2% agarose bleach gel. The gel was run at a constant voltage
(120 V) for an hour for better separation of bands. Post-electrophoresis, the gel was stained with
ethidium bromide, and RNA bands were visualized under UV light.
2.2.4 Fluorescence Northern Blot Analysis
Fluorescence Northern Blot Analysis was performed using either a denaturing polyacrylamide
gel or a denaturing formaldehyde agarose gel.
2.2.4.1. 6%/7M Urea Denaturing Gel Preparation
To prepare a 6% denaturing polyacrylamide gel with 7M urea, 6.3 g of urea was weighed into a
50 mL beaker. 2.25 mL of 40% acrylamide (19:1 ratio) was added using a 10 mL disposable
pipette, followed by 7.3 mL of deionized water (dH2O) from a dedicated bottle. After adding
750 µL of 20x Tris-Borate-EDTA (TBE) buffer and a small stir bar, the solution was mixed on a
hotplate/stirrer set to 100 C until the urea completely dissolved. Once dissolved, the mixture was
cooled on ice to slow down polymerization before adding 150 µL of 10% ammonium persulfate
(APS), stirring with a pipette tip. 15 µL of TEMED was then added and stirred again. The
amount of acrylamide and water was changed accordingly for the percentage of gel needed. The
gel was then poured using the same disposable pipette, inserted the comb into the appropriate
depth based on the sample volume, and allowed the gel to polymerize. The leftover gel mix was
solidified, and the solidified acrylamide was discarded safely.
To prepare a 1.5% formaldehyde-agarose gel in a 150-mL volume, 2.25 g of agarose was mixed
with 109.5 mL of water. The agarose was melted in a microwave and cooled to 65 C before
adding 15 mL of 10x MOPS buffer and 25.5 mL of 37% formaldehyde under a chemical fume
21

hood. The agarose mixture was then poured into a gel tray with an inserted comb and allowed to
solidify. Concurrently, sufficient 1x MOPS buffer was prepared for the gel tank reservoirs by
diluting the 10x stock. After placing the gel in the tank and adding the 1x MOPS buffer to
prevent drying, the gel was loaded and run immediately to minimize formaldehyde diffusion. For
RNA sample preparation, 4.7 µL of each RNA sample, containing 10-30 µg of total cellular
RNA, was added to a 1.5-mL microcentrifuge tube. A fresh stock of sample buffer was prepared,
consisting of 660 µL ultrapure formamide, 200 µL 10x MOPS buffer, and 270 µL formaldehyde
(37%). A total of 11.3 µL of this sample buffer was added to each RNA sample, followed by
heating at 60 C for 5 minutes and cooling on ice. Finally, 4 µL of tracking dye was added to
bring the total volume to 20 µL, and the samples were ready for electrophoresis.
2.2.4.2. Denaturing Polyacrylamide Gels for RNA Analysis
6-15% denaturing polyacrylamide gel, suitable for analyzing RNAs smaller than 1000
nucleotides was poured. Denaturing (formaldehyde) agarose gel was used for larger RNAs. The
gel was pre-run for 15 minutes at 400 V in 1X TBE buffer, ensuring that urea was cleared from
the wells and any trapped air was removed from the bottom of the gel using a syringe with a bent
needle. Next, 1–10 µg of RNA was mixed with an equal volume of 2X formamide loading
buffer, with the total volume kept under 20 µL. RNA samples were denatured at 65 C for 3
minutes, quickly spun down, and immediately placed on ice. The wells were cleared again with a
syringe before loading samples using an elongated gel-loading tip, with the tip rinsed in TBE
buffer between samples. The gel was then run at 400 V for 45–90 minutes, depending on the size
of the RNA being analyzed. Samples were analyzed by staining with ethidium bromide or via
Northern blotting.

22

2.2.4.3. Membrane Transfer with Semi-Dry Electroblotter/Capillary Transfer
To prepare for RNA transfer, six pieces of Whatman paper and one piece of Hybond+ nylon
membrane were cut slightly larger than the gel, with the membrane labeled with the date and
experiment identifier. Two pieces of Whatman paper were pre-wetted in 1X TBE, placed on the
semi-dry blotter, and arranged to avoid any trapped air bubbles. The gel was carefully transferred
from the glass plate to the Whatman paper, and excess gel was trimmed. The membrane was then
pre-wetted, aligned with the gel, and covered with three additional pre-wetted Whatman papers.
After ensuring no bubbles were present, the setup was secured for transfer, conducted at 2.5
mA/cm² for 30 - 45 minutes.
For a capillary RNA transfer from formaldehyde agarose gel, a few millimeters of the gel edges
were trimmed with a scalpel to create a flat surface. A nylon membrane was cut approximately 2
mm larger than the gel on all sides, along with six pieces of Whatman 3MM paper and a 2-inch
stack of paper towels matching the membrane size. A capillary gel-transfer system was set up on
an elevated base gel tank, with two pieces of Whatman 3MM paper pre-wetted in 20X SSC
layered on top. The gel was placed upside down relative to its position in the gel tank on top of
the Whatman paper, and excess liquid was blotted off with Kimwipes. The nylon membrane was
pre-wetted with water, then 20X SSC, and carefully positioned on the gel without allowing
movement after contact. Bubbles between the gel and membrane were smoothed out with a 5-mL
glass pipette or a gloved finger. A pre-wetted piece of Whatman 3MM paper was layered on top
of the membrane, followed by three dry pieces of Whatman paper, a 2-inch stack of paper
towels, a glass plate, and a weight. This blotting sandwich was left to transfer overnight.

23

2.2.4.4. RNA Cross-Linking for Membrane Stabilization Using the Strata Linker System
Using forceps, the membrane was transferred, RNA side up, onto a piece of Whatman paper and
immediately cross-linked the RNA to the membrane using the auto cross-link setting on the
Strata linker.
2.2.4.5. Pre-hybridization
The ULTRAhyb™–Oligo Buffer (Invitrogen, AM8663) was preheated to 42 C in a water bath
until fully resolubilized. The hybridization oven was also set to 42 C. Using forceps, the blot(s)
were carefully positioned in a hybridization bottle with the RNA side facing inward. Between 510 mL of the preheated hybridization buffer was added to the bottle, and the assembly was
incubated at 42 C for 30 minutes.
2.2.4.6. Hybridization
5 pmol/mL of biotinylated oligonucleotide was incorporated into the pre-hybridization buffer,
avoiding direct application onto the blot. The hybridization was conducted at 42 C for a duration
ranging from 1 to 24 hours.
Table 2.1. Oligonucleotides used in Northern Blot Analysis
The oligonucleotides ordered are 5’ biotinylated. oSDR2488 was designed by Dr. Martha Stark.
Oligonucleotide

Target

Sequence (5′ to 3′)

oSDR2487

Cm 5.8SrRNA

CGCTGCGAGAGCCTAGATATCCACCG

oSDR2586

Cm 28SrRNA

CGCTATCGGTCTCTCGCCGGTATTTAGCCTTAGGTGAAG

oSDR2587

Cm 18SrRNA

GTTACCATGAATCACCAGAGACCGCCGAGGCGGTTTGG

oSDR2488

Cm RNase MRP

AGCTTTGCTTACCACCGACACTCTCTG

24

2.2.4.7. Washing Strategies for Enhanced Specificity
For the washing step, 2x SSC, 0.5% SDS, stored at 37 C was used. The blot was initially rinsed
with 5-10 mL of wash buffer to eliminate excess unhybridized probe. It was then subjected to a
series of washes with approximately 20-25 mL of buffer for 30 minutes, performing three 5minute washes at 42 C. Subsequently, the oven temperature was lowered to 20 C, and the door
was left ajar while the blot was prepared for blocking.
For cases where non-specific bands were detected, more stringent washing conditions were
employed to enhance specificity: first, a 5-minute wash with 2x SSC and 0.1% SDS at 42 C,
followed by two 20-minute washes with 0.5x SSC and 0.1% SDS at 42 C, and, if necessary, an
additional 20-minute wash with 0.1x SSC and 0.1% SDS.
2.2.4.8. Blocking Procedure for Northern Blot Analysis
Blocking was performed with 5 mL of blocking buffer (Licor, 927-70001), and the blot was
incubated for 1 hour at room temperature (20 C).
2.2.4.9. Fluorescent Labeling of Northern Blot; IRDye 800CW Streptavidin Binding
To minimize the background signal, 100 µL of 10% Tween-20 (to achieve a final concentration
of 0.2%) and 50 µL of 10% SDS (final concentration of 0.1%) were added to the blocking buffer
before the addition of dye. In the dark, 0.5 µL of Streptavidin-IRDye 800CW conjugate (Licor,
926-32230) was introduced into the blocking buffer at a 1:10,000 dilution. For multiple tubes, a
1:100 dilution was prepared, and 50 µL was added to each tube. The blot was then incubated for
30 minutes at room temperature in the dark.

25

2.2.4.10. Post-Streptavidin Binding Washing
The blot was initially rinsed with 5-10 mL of PBST (1X PBS, 0.1% Tween-20) to remove the
majority of unbound dye. Subsequently, it was washed with 20-25 mL of PBST for three
intervals of 5 minutes each at room temperature. Finally, a single 5-minute wash with 1X PBS at
room temperature was performed to remove the detergent, thereby enhancing fluorescence.
2.2.4.11. Detection
For imaging, the blot was placed on a piece of Whatman paper moistened with water and
positioned on an acetate sheet before being placed into the Bio-Rad Imager. If the moistened
Whatman paper caused any blotchiness, it was removed. To prevent the blot from drying, which
would hinder stripping, Saran wrap was used if a long exposure (more than a few minutes) was
necessary. The IR-Dye 800 CW setting was selected, and the image was captured using auto
exposure, with a preview followed by an optimal capture. Manual exposure time was also
adjusted as needed to achieve the desired image quality.
2.2.4.12. Northern Blot Analysis Utilizing Bio-Rad Image Lab 6.1
Analysis of Northern blots was conducted using Bio-Rad Image Lab software 6.1. The Image
Lab 6.1 software was opened, and the image of the Northern blot to be analyzed was imported.
The analysis tools in Image Lab were used to detect bands, define boundaries, subtract
background noise. To determine the relative front, the software’s lane profile feature was
employed. By analyzing the migration distance of the bands in relation to the dye front and
loading wells, the relative front was detected. This measurement was essential for calculating
relative mobility, ensuring accurate comparison between samples. Finally, the bands were
quantified according to the analysis requirements, with options for relative or absolute
26

quantification. Software tools for normalization, annotation, and manual quantification were
employed as needed. The analyzed data, including band tables, was exported to Excel, and the
image was saved in a format suitable for publication.
2.2.4.13. Blot Stripping and Storage for Subsequent Re-probing
The blot was stripped to allow re-probing with a different probe before it dried out.
Approximately 50 mL of 0.2% SDS was heated in a microwave to near boiling and incubated
with the blot in hybridization bottles for 10 minutes at room temperature with rotation. This step
was repeated with boiling SDS. The blot was then rinsed with around 50 mL of 2x SSC followed
by a rinse with water. To ensure complete removal of the probe, the blot was re-exposed on the
Imager for at least as long as the original exposure time. If re-probing was not planned for the
same day, the blot was wrapped in Saran wrap and stored at room temperature. While stripping
may remove a small amount of RNA from the blot, it was possible to strip up to three times
without issue.

27

2.3. Results
RNase MRP is involved in the generation of the two forms of 5.8S rRNA. Mutations in the MRP
RNA have been shown to alter the 10:1(small: large) stoichiometry (Schmitt and Clayton 1992;
Lindahl et al. 2009). I carried out analyses to investigate how the accumulation of the intronic
region of CMK142T (which houses MRP) at 57 C (heat stress conditions) affects MRP
expression and the stoichiometry of the two forms of 5.8S rRNA. I hypothesized that MRP
accumulates in an inactive form under heat stress, which should result in a change in the 5.8S
isoform ratio, so I used northern blotting to look for such a change. An rRNA cassette was
constructed in SnapGene (a sequence viewing and analysis tool) to facilitate the study of
ribosomal RNA (rRNA) processing in Cyanidioschyzon merolae (Figure 2.2). The constructed
rRNA cassette was solely utilized to design specific probes targeting the rRNAs of interest for
Northern blot analysis and does not contain any promoter sequence. The cassette was not meant
to drive the expression of rRNA but solely for primer designs.

Figure 2.2. C. merolae rRNA Cassette. The rRNA sequences of C. merolae were identified and retrieved from the
NCBI database using the appropriate accession numbers 5.8S rRNA (Accession Number: XR_002461615), 18S
rRNA (Accession Number: XR_002461616), 28S rRNA (Accession Number: XR_002461614), ITS1 (Accession
Number: AB158485), and ITS2 (Accession Number: AB158484).

After cultivating C. merolae and subjecting it to heat stress treatment, total RNA was extracted
from the cells. The integrity of the RNA was then analyzed to ensure its suitability for
downstream applications. As one of the major issues affecting the integrity of RNA is the
ubiquitous presence of ribonucleases (RNases), RNA quality was quickly analyzed by adding
28

small amounts of commercial bleach to TBE buffer-based agarose gels prior to electrophoresis
(described in Section 2.2.3). This RNA integrity check was to confirm that the RNA was of high
quality, free from degradation, and appropriate for subsequent analyses such as Northern
blotting. Figure 2.3 below demonstrates that the RNA extracted was of high quality and suitable
for subsequent downstream analysis.
Cm

42C

B

Ladder

A

42C

Ladder
S1

S2

57C
S1

S2

28SrRNA
18SrRNA

28SrRNA
18SrRNA
500bp
200bp
5SrRNA

200bp

tRNA
50bp

tRNA

Figure 2.3. Isolation and analysis of total RNA from Cyanidioschyzon merolae Wild Type: RNA was isolated
from Cm WT. Samples were examined by running 1xTBE, 2% Agarose Bleach gel a) Sample exposed to 42 C and
b) Comparison of samples at two different temperatures 42 C and 57 C. 50bp DNA ladder was used as a size
marker. The two samples in each temperature set in b, are technical replicates.

The total RNA samples were used for polyacrylamide Northern blotting to investigate: the
expression of MRP under optimum temperature and heat stress, to determine whether C. merolae
contains two distinct forms of 5.8S rRNA, and whether heat stress modulates RNase MRP
function by altering the stoichiometric ratio of these 5.8S rRNA forms, as suggested in the
literature (Figures 1.5a,1.5b,1.6a and 1.6b). The RNase MRP in C. merolae had only been
bioinformatically identified in the intronic region of the CMK142T gene without experimental
validation. To confirm the expression, I performed a northern blot on total RNA (Figure 2.4).

29

A

SSRNA
Ladder

42⁰C 57⁰C

9Kb
7Kb
5Kb

3Kb
2Kb

1Kb
Intron
0.5Kb
RNase MRP

B
7000000

Intensity (A.U)

6000000
5000000
4000000
3000000
2000000
1000000
0
Intensity

Intron_57°C

Intron_42°C

MRP_57°C

MRP_42°C

4737568

4808000

6566976

6639568

Figure 2.4. The RNA component of RNase MRP is stably expressed at 42 C and 57 C. A) RNA was isolated
from C. merolae after growth and treatment at 42 C and 57 C temperatures and probed for MRP after running 1.5%
formaldehyde-agarose gel for 6 hours at 50V and transferred to a nylon membrane. B) Quantification of MRP and
Intron intensities using Bio-Rad Image Lab software 6.1.

30

Quantification of RNase MRP expression levels, based on band intensities at both temperatures
(Figure 2.4a and b), confirmed the presence of RNase MRP in C. merolae but does not indicate
the anticipated change in expression at 57 C. This result contrasts with the transcriptomic data,
which indicated an increase in expression at 57 C (Figure 2.1), creating a discrepancy between
the two findings.
Evidence from lab colleagues suggests that the transcriptomic data reflects adenylation of RNase
MRP transcripts rather than their accumulation, and this may cause it to be targeted for
degradation by the exosome, as polyadenylation in non-coding RNAs often signals turnover.
In light of the polyadenylation of MRP, I hypothesized that it becomes inactive under heat stress
and would therefore lead to a change in the 5.8S isoform ratio. To investigate the presence of
two forms of 5.8S rRNA and whether heat stress affects the stoichiometry of these two forms, I
visualized total RNA on an ethidium bromide-stained gel (Figure 2.5). This analysis did not
reveal any hint of the existence of two forms of 5.8SrRNA (see Figure 2.5 below).

31

42C
S1

S2

57C
S3

S1

S2

5.8SrRNA

S3

154nt

5SrRNA

130nt

70-90nt

tRNA

Figure 2.5. Detection of 5.8S rRNA in C. merolae using ethidium bromide-stained 6% PAGE. RNA was
isolated from C. merolae after growth and treatment at 42 C and 57 C temperatures and examined by staining with
ethidium bromide after running 6% PAGE. The three samples in each temperature set are technical replicates.

Since the initial results did not indicate the presence of two forms of 5.8S rRNA, I
proceeded with a 6% PAGE Northern blot analysis run at different time intervals to evaluate the
optimal migration time for different fragment sizes, balance resolution, and prevent the loss of
smaller fragments or over-migration of larger ones (Figure 2.6).

32

A
42C

5.8S

C

B
57C

42C

42C

U4SnRNA
177nt

154nt
5.8S

154nt
154nt

U2SnRNA
131nt

5.8SL
5.8SS

Figure 2.6. Northern analysis of 5.8S rRNA. Northern blots of total RNA separated on a 6% polyacrylamide gel
for (a) 45 minutes, (b) 75 minutes, and (c) 90 minutes. Cells were treated at the indicated temperatures prior to RNA
isolation. In (b), total RNA was probed for Cm U2 and U4 snRNAs for size markers. The three samples in each
temperature set are technical replicates.

Panels b and c of Figure. 2.6 hints at the possibility of a long form of 5.8S in C. merolae
confirming the exhibition of two forms of 5.8S rRNA, mirroring findings in S. cerevisiae and D.
melanogaster wild-type cells (Figures 1.5c,1.6a, and 1.6b).
The third analysis utilized 8% PAGE with a 95-minute run, followed by probing for 5.8S rRNA
after transferring the samples to a nylon membrane. This analysis confirmed the presence of two
distinct forms of 5.8S rRNA but did not reveal any changes in the stoichiometry of these forms
in C. merolae (Figure 2.7). These results suggest that the accumulation of the intronic region of
CMK142T did not impact RNase MRP's expression or catalytic activity, as evidenced by the
unchanged stoichiometry of the two 5.8S rRNA forms.

33

A

57⁰C

42⁰C

5.8SL
5.8SS

Av. Intensity (A.U)

B

20,000,000
15,000,000
10,000,000
5,000,000
0
Av.Intensity

Av. Relative front

C

P-value
P-value==0.91
0.91
P-value > 0.05

5.8S_57°C

5.8S_42°C

15,175,520

14,994,667

0.55

P-value = 0.13
P-value
P-value> =0.05
0.13

0.54

0.53
0.52
0.51
0.5
0.49
Av.Rf

5.8S_57°C

5.8S_42°C

0.54

0.53

Figure 2.7. Heat Stress Effect on 5.8SrRNA Intensities using 8% PAGE. a) RNA was isolated from C. merolae
after growth and treatment at 42 C and 57 C temperatures, probed for 5.8S rRNA after running 8% PAGE for 95
minutes, and transferred to a nylon membrane. b) band intensities and c) mobility (relative front) were quantified
using Bio-Rad Image Lab software 6.1. The three samples in each temperature set are technical replicates.

Quantification of the intensities and relative front (the distance a specific RNA fragment
migrates on the gel relative to the total distance traveled by the dye front) of the bands at these
34

two temperatures gave a P-value > 0.05 signifying that there is no statistical difference in the
relative front and intensities of 5.8S rRNA either at 42C and 57C (Figure 2.7b and c).
2.4. Discussion
The previous observation indicated that the CMK142T intron is predicted to harbor the RNase
MRP RNA, known from the literature to be involved in 5.8S rRNA processing and that the
expression of this intron appears to increase dramatically under heat stress (Figure 2.1). I
therefore, sought to confirm that the MRP RNA is expressed and to test whether the
accumulation at 57 C reflects inactivation, which should be revealed in a change in the 5.8S
isoform ratio as evidenced in S. cerevisiae and D. melanogaster (Schmitt and Clayton 1993;
Schneider et al., 2010; Shadel et al., 2000). I successfully confirmed MRP expression, but
surprisingly did not see the expected change in expression by Northern blotting. My labmates
have evidence that the transcriptomic data reflects adenylation of MRP, rather than
accumulation, causing it to be targeted for degradation by the exosome, as polyadenylation in
non-coding RNAs often signals turnover. The result from the Northern blot implies that RNase
MRP is stable even under heat stress in C. merolae (Figure 2.4). This stability of RNase MRP in
C. merolae may reflect a broader evolutionary adaptation mechanism, allowing organisms to
thrive in fluctuating thermal environments without detrimental effects on cellular processes
(Lefort et al. 2014). Thus, RNase MRP's resilience under heat stress underscores its potential
role in cellular homeostasis during environmental challenges. The data presented in this chapter
also confirms the presence of two distinct forms of 5.8S rRNA but did not reveal any changes in
the stoichiometry of these forms in C. merolae (Figures 2.6c and 2.7). Unlike in S. cerevisiae and
D. melanogaster, the 5.8S isoforms were difficult to detect in my experiments, and I did not
observe any change in their ratio. Another way to detect the two isoforms will be by primer
35

extension. However, these results suggest that the accumulation of the intronic region of
CMK142T did not impact RNase MRP's expression or catalytic activity, as evidenced by the
unchanged stoichiometry of the two 5.8S rRNA forms. This suggests that, unlike in Arabidopsis
thaliana, where heat stress leads to rapid degradation of rRNA (Darriere et al., 2022), heat stress
does not influence 5.8S rRNA processing in C. merolae (Figure 2.7). This difference highlights
the potential for species-specific responses to thermal stress, indicating that the mechanisms
governing rRNA stability and processing may vary significantly across different organisms. I
conclude that either RNase MRP is not involved in 5.8S rRNA processing in C. merolae, or,
more likely, heat does not inactivate MRP.

36

Chapter 3 - Heat Stress Inhibits rDNA Transcription, but 18S and 28S rRNA
Processing Remains Unaltered as C. merolae Adheres to the Canonical rRNA
Processing Pathway

37

3.1 Introduction
In this chapter, I detailed the experiments conducted to determine heat stress effect on rRNAs
and assessed whether Cyanidioschyzon merolae follows the canonical rRNA processing pathway
as observed in other eukaryotes. Northern blot analysis was employed to probe for processing
intermediates in the rRNA transcript. Below is a schematic diagram generated with information
from Li et al. (2021), depicting the expected bands in the canonical rRNA processing pathway
and indicating the C. merolae probes used in this experiment.

Figure 3.1. Canonical rRNA processing pathways. This figure depicts rRNA processing intermediates with names
of relevant processing enzymes and their sites of action. Refer to Table 3.1 for the oligonucleotide sequences and the
target positions on pre-rRNA to which they hybridize.

38

3.2 Materials and Methods
In this chapter, the materials and methods employed are the same as those outlined in Chapter 2,
from sections 2.2.1 to 2.2.4.13, utilizing formaldehyde agarose Northern blotting and Capillary
transfer techniques.
Table 3.1. Oligonucleotides used in Northern Blot Analysis
The oligonucleotides ordered from Integrated DNA Technologies are 5’ biotinylated.
Oligonucleotide

Target

Sequence (5′ to 3′)

oSDR2487

Cm 5.8S rRNA

CGCTGCGAGAGCCTAGATATCCACCG

oSDR2586

Cm 28S rRNA

CGCTATCGGTCTCTCGCCGGTATTTAGCCTTAGGTGAAG

oSDR2587

Cm 18S rRNA

GTTACCATGAATCACCAGAGACCGCCGAGGCGGTTTGG

oSDR2588

Cm ITS1

ACCGCCGTCTTCCCACTGGGGAATAGCACAG

oSDR2589

Cm ITS2

ACGCATGCAGTCTGAGACAGACAGAACCTGCGCGC

3.3. Results and Discussion.
3.3.1 18S and 28SrRNAs Northern
Given that no change in the stoichiometry of 5.8S rRNA was observed in Chapter 2 and
considering the possibility that RNase MRP may be involved in other rRNA processing steps, I
extended the investigation to 18S rRNA, 28S rRNA, and other intermediates in the rRNA
processing pathway. This decision was further motivated by the complex interplay between heat
stress and rRNA processing observed in other organisms, such as Arabidopsis thaliana, where
heat stress disrupts nucleolar structure, inhibits pre-rRNA processing, and alters ribosome
profiles, leading to a rapid reduction in precursors for 18S, 5.8S, and 25S rRNAs (Darrière et al.,
2022). To explore the effects of heat stress at 57 C on 18S and 28S rRNAs, I performed
Northern blotting on total RNA (Figures 3.2 and 3.3).
39

A

SSRNA
Ladder

42⁰C

57⁰C

18SrRNA

B

40,000,000

P-value = 0.8
P-value > 0.05

Av. Intensity (A.U)

35,000,000
30,000,000
25,000,000
20,000,000
15,000,000
10,000,000
5,000,000
0
Av.Intensity

C

18S_57°C

18S_42°C

31,664,444

30,745,535

0.35

P-value = 0.065
P-value > 0.05

Av. Relative front

0.3
0.25
0.2
0.15
0.1
0.05
0

Av.Rf

18S_57°C

18S_42°C

0.3

0.29

Figure 3.2. Heat stress does not impact 18S rRNA processing. a) RNA was isolated from C. merolae after
growth and treatment at 42 C and 57 C temperatures, probed for 18S rRNA after running 1.5% formaldehydeagarose gel for 6 hours at 50 V. b) band intensities and c) relative front (mobility) were quantified using Bio-Rad
Image Lab software 6.1. The samples are technical replicates.

40

A

SSRNA
Ladder

42⁰C

57⁰C

28SrRNA

Av. Intensity (A.U)

B
70,000,000
50,000,000
40,000,000
30,000,000
20,000,000
10,000,000
0
Av.Intensity

Av. Relative front

C

P-value = 0.097
P-value > 0.05

60,000,000

0.245
0.24
0.235
0.23
0.225
0.22
0.215
0.21
0.205
Av.Rf

28S_57°C

28S_42°C

53,918,497

56,029,129

P-value = 0.067
P-value > 0.05

28S_57°C

28S_42°C

0.24

0.23

Figure 3.3. Heat stress does not impact 28S rRNA processing. a) RNA was isolated from C. merolae after
growth and treatment at 42 C and 57 C temperatures, probed for 18S rRNA after running 1.5% formaldehydeagarose gel for 6 hours at 50 V. b) band intensities and c) relative front (mobility) were quantified using Bio-Rad
Image Lab software 6.1. The samples are technical replicates.

41

Figures 3.2 and 3.3 demonstrate that there are no detectable changes in the levels of 18S or 28S
rRNA under heat stress conditions. However, since I previously observed no effects on 5.8S
rRNA (as discussed in Chapter 2), I am unable to draw any conclusions regarding the potential
role of RNase MRP in the processing of these rRNA forms. However, in contrast to findings in
Arabidopsis thaliana where heat stress leads to the disruption of nucleolar structure, inhibition of
pre-rRNA processing, and a rapid decline in detectable levels of 18S, 5.8S, and 25S rRNA
precursors (Darrière et al., 2022), this data reveals that C. merolae exhibits a unique resilience
under similar conditions. Specifically, exposure to 57 C does not affect the processing levels of
18S and 28S rRNA in C. merolae.
3.3.2 Detection of Pre-rRNA Intermediates
The canonical model for processing the primary Pol I transcript begins with the
endonucleases Utp24 and Rnt1 cleaving the 5' ETS and 3' ETS, respectively, from the main
portion of the pre-rRNA, forming the 32S intermediate (Figure 3.1) (Kufel et al., 1999; An et al.,
2018). Utp24 then cleavages at the A2 site within ITS1, separating the rRNA sequences destined
for the 40S and 60S ribosomal subunits. Further processing of ITS1 produces the 5' end of the
5.8S rRNA and the 3' end of the 18S rRNA. The 3' end of the 5.8S rRNA is generated by Las1
cleavage at the C2 site in ITS2, followed by exonucleolytic trimming by the exosome, with the
downstream portion of ITS2 being removed by the exonucleases Rat1 and Xrn1 (Mitchell et al.,
1997). Two pathways lead to forming the 5' end of the 5.8S rRNA. In the major pathway, the
ribozyme RNase MRP cleaves ITS1 at the A3 site, after which exonucleases Rat1 and Rrp17
trim the resulting 5' end to create the “short” 5.8S rRNA (5.8SS).
To investigate potential heat stress effects on additional intermediates in the pre-rRNA
processing pathway and to determine whether C. merolae follows the canonical rRNA
42

processing pathway, I utilized probing for these precursors (ITS1, ITS2, 5.8S, and 28S) on a
1.5% formaldehyde agarose gel and performed a Northern blot analysis.
3.3.2.1 Analysis of 5.8S and Processing Intermediates

B

A
42C

57C

SSRNA Ladder

42C

57C

5Kb

Pre-rRNA
Transcript
A3 Cleavage product

3Kb

ITS1 to a region in
28S

9Kb
7Kb

2Kb
5.8SrRNA

1Kb
0.5Kb

5.8SrRNA

Figure 3.4. Detection of 5.8S rRNA on formaldehyde-agarose gel via Northern, illustrating intermediates
associated with the Canonical rRNA Processing Pathway. RNA was isolated from C. merolae grown at both
temperatures and probed for 5.8S rRNA under two conditions: a) 1.5% Formaldehyde-Agarose gel run for 3 hours
with lower total RNA, and b) 1.5% formaldehyde-agarose gel run for 6 hours with higher total RNA to enhance the
detection of intermediates. oSDR2487 probe was used. The samples are technical replicates.

The oSDR2487 probe is designed to hybridize specifically to the 5.8S rRNA region within the
pre-rRNA transcript (B1L canonical processing site). Therefore, it is expected to detect the fulllength pre-rRNA transcript, the A2/A3 cleavage intermediate, and the mature 5.8S rRNA
species. Figure 3.4 indicates the presence of pre-rRNA and the A3 cleavage product, alongside
43

expected expression levels of 5.8S rRNA. However, a notable decrease in pre-rRNA prominence
at 57 C was observed (Figure 3.4) suggesting reduced expression of pre-rRNA under heat stress
conditions.
3.3.2.2 Analysis of ITS1 and Processing Intermediates
The next step is to investigate cleavage within the ITS1 region, which results in the separation of
the 18S rRNA sequence from the 5.8S-ITS2-28S rRNA sequences (Figure 3.1). The 5.8S rRNA
blot (Figure 3.4b), was stripped and reprobed with oSDR2588 within ITS1.

SSRNA Ladder

42C

57C

9Kb
7Kb

Pre-rRNA Transcript

5Kb

3Kb
2Kb

1Kb

0.5Kb

5.8SrRNA

Figure 3.5. Detection of ITS1 on formaldehyde-agarose gel via Northern, revealing intermediates
corresponding to the canonical rRNA Processing Pathway. RNA was isolated from C. merolae after growth at
both temperatures and probed for 5.8S rRNA after running 1.5% formaldehyde agarose for 6 hours, this blot was
stripped and re-probed for ITS1. oSDR2588 probe was used. The samples are technical replicates.

44

The oSDR2588 probe is designed to hybridize specifically to the ITS1 region within the prerRNA transcript. As such, it is expected to detect the full-length pre-rRNA transcript. The
analysis reveals the detection of the pre-rRNA transcript. This also indicated a notable reduction
in pre-rRNA levels under heat stress (Figure 3.5), corroborating the findings presented in Figure
3.4. This observation provides further evidence of rDNA transcription inhibition in response to
heat stress conditions.
3.3.2.3 Analysis of ITS2 and Processing Intermediates
The ITS1 cleavage is then followed by a cleavage in ITS2, which results in the separation of the
5.8S and 28S rRNA sequences. Subsequent trimming of these intermediates generates the mature
5.8S and 28S rRNAs.

SSRNA Ladder

42C

57C

9Kb
7Kb

Pre-rRNA Transcript
A3 Cleavage product
27S(A3)

5Kb

3Kb
2Kb

1Kb

0.5Kb

5.8SrRNA

Figure 3.6. ITS2 detection on formaldehyde-agarose gel via Northern reveals intermediates corresponding to
the Canonical rRNA Processing Pathway. RNA was isolated from C. merolae after growth at both temperatures
and probed for 5.8S rRNA after running 1.5% formaldehyde agarose for 6 hours; this blot was stripped and reprobed for ITS2. oSDR2589 probe was used. The samples are technical replicates.

45

The oSDR2589 probe is designed to hybridize specifically to the ITS2 region within the prerRNA transcript. It is expected to detect the full-length pre-rRNA transcript as well as the A2/A3
cleavage intermediates generated during rRNA processing. The expected pre-rRNA transcript
and A2/A3 cleavage intermediates were observed. However, the investigation of the internal
transcribed spacer 2 (ITS2) region, similar to the findings observed in the 5.8S and ITS1 results,
demonstrated a significant decrease in pre-rRNA levels under heat stress conditions (Figure 3.6).
This consistent pattern across multiple rRNA processing components provides robust evidence
supporting the inhibition of rDNA transcription during heat stress. The correlation between
reduced pre-rRNA levels in the ITS2 region and the previously observed reductions in the 5.8S
and ITS1 regions reinforces the notion that heat stress adversely affects rDNA transcription.

46

3.3.2.4 28S and Processing Intermediates
Lastly, a probe was designed to target the 5’ end of 28S rRNA to detect other intermediates in
the canonical pathway.
A
SSRNA Ladder

B
42C

SSRNA Ladder

57C

28SrRNA
Band

Pre-rRNA
Transcript
A3/A2
Cleavage
product

5Kb

2Kb

3Kb
Covered for
high
2Kb
Exposure of
Intermediates

1Kb

1Kb

0.5Kb

0.5Kb

3Kb

57C

9Kb
7Kb

9Kb
7Kb
5Kb

42C

5.8S rRNA

Figure 3.7. 28S rRNA detection on formaldehyde-agarose gel via Northern reveals intermediates
corresponding to the Canonical rRNA Processing Pathway. a) RNA was isolated from C. merolae after growth
at both temperatures and probed for 5.8SrRNA after running 1.5% formaldehyde agarose for 6 hours, this blot was
stripped and re-probed for 28S rRNA. b) The 28S band was covered to ensure optimal exposure of other bands.
oSDR2586 probe was used. The samples in each temperature set are technical replicates.

The oSDR2586 probe is designed to hybridize specifically to the 28S rRNA region within the
pre-rRNA transcript. It is expected to detect the full-length pre-rRNA transcript, the A2/A3
cleavage intermediates, and the mature 28S rRNA. The expected bands were observed. The
tabulated results of C. merolae’s Canonical rRNA processing pathway Northerns and the bands
observed are shown in Table 3.2 below.

47

Table 3.2. Observed Bands in Canonical rRNA Processing: Northern Blot Data Summary

42 C
a

57 C

2587

2588

2487

2589

2586

2587

2588

2487

2589

2586

8kb pre-rRNA

✓ ++

✓ ++

✓ ++

✓ ++

✓ ++

✓+

✓+

✓+

✓+

✓+

5.6kb A3
cleavage
product

-

-

✓ ++

✓+++

✓ ++

-

-

✓+

✓+

✓+

4.5kb prerRNA

-

✓+++

-

-

-

-

✓+

-

-

-

28S rRNA

-

-

-

-

✓
+++

-

-

-

-

✓+++

18S rRNA

✓
+++

-

-

-

-

✓
+++

-

-

-

-

5.8S rRNA

-

-

✓+++

-

-

-

-

✓+++

-

-

OLIGO
SIZES

a
oSDR2586

oSDR2587
oSDR2588
18SrRNA

ITS1
845nt

oSDR2589

5.8SrRNA

ITS2
1798nt

28SrRNA

154nt

1791nt

= Detected

oSDR2487

+ = Less Prominent

++ = Medium

48

3418nt

+++ = Prominent

- = Not Detected

Figure 3.8 below presents the quantification of band intensities and the relative front of prerRNA intermediates seen at 42 C and 57 C (Figure 3.4 to Figure 3.7).

A

Pre-rRNA bands at 42 C

Pre-rRNA bands at 57 C

0.14
0.12

Av. Relative front

0.1
0.08
0.06

0.04
0.02
0
Av.Rf

B

Av. Intensity (A.U)

250,000

5.8S

ITS2

ITS1

28S

5.8S

ITS2

ITS1

28S

0.094

0.094

0.095

0.0965

0.0975

0.098

0.1145

0.0935

Pre-rRNA bands at 42 C

Pre-rRNA bands at 57 C

200,000
150,000
100,000
50,000
0

Av.Intensity

5.8S

ITS2

ITS1

28S

5.8S

ITS2

ITS1

28S

53,034

124,399

197,303

46,185

2,820

6,664

8,717

3,762

Figure 3.8: Pre-rRNA level reduced at 57 C: a) relative front (mobility) and b) band intensity of pre-rRNA levels
reduced at 57 C. Bands were quantified using Bio-Rad Image Lab software 6.1.

49

To determine whether Cyanidioschyzon merolae follows the canonical rRNA processing
pathway, Table 3.3 below, compares the expected bands for each probed intermediate in the
canonical pathway with those observed in C. merolae. The presence of all expected bands in C.
merolae supports the conclusion that this organism adheres to the canonical rRNA processing
pathway.
Table 3.3. Canonical rRNA Processing Pathway: Expected vs. Observed Bands in Cyanidioschyzon merolae

Precursor
Probes

Expected rRNA Canonical Processing
Pathway Product
8 kb pre-rRNA
5.6 kb A3 cleavage product
5.8S rRNA (154 b)

C. merolae Intermediate
Product Observed
✓
✓
✓

2587

8 kb pre-rRNA
18S rRNA (1.7 kb)

✓
✓

2586

8 kb pre-rRNA
5.6 kb A3 cleavage product
28S rRNA (3.4 kb)

✓
✓
✓

2589

8 kb pre-rRNA
5.6 kb A3 cleavage product

✓
✓

2588

8 kb pre-rRNA

✓

2487

= Detected

3.4. Discussion.
The stability of 5.8S rRNA (Figure 2.7, chapter 2), 18S rRNA, and 28S rRNA levels (Figures 3.2
and 3.3 respectively) under heat stress in C. merolae contrasts with the observed inhibition of
rRNA maturation in A. thaliana (Darrière et al., 2022), mammals (Ghosha and Jacob 1996;
50

Coccia et al., 2017) and the altered ribosome profiles seen in other eukaryotes. The unaltered
levels of mature 5.8S, 28S, and 18S rRNAs in C. merolae under heat stress despite a reduction in
pre-rRNA levels (Figures 3.5 and 3.6), suggest the presence of a sophisticated regulatory
mechanism that ensures ribosome function is maintained under extreme conditions. This
mechanism prioritizes the stability of mature rRNAs to maintain ribosome function. This could
involve post-transcriptional stabilization or modification of mature rRNAs, protecting them from
degradation during stress. Alternatively, if no such regulation exists, heat stress could impair prerRNA processing, leading to a reduction in both precursor and mature rRNA levels, potentially
disrupting ribosome biogenesis. The maintenance of mature rRNAs under stress indicates a
sophisticated mechanism ensuring ribosomal function under extreme conditions.
This finding implies that C. merolae may possess distinct adaptive mechanisms that ensure the
maintenance of ribosome biogenesis even under extreme environmental stress, highlighting the
evolutionary diversity in stress response pathways across species. One possible explanation is
that mature rRNAs, once synthesized, exhibit high stability and resistance to degradation,
allowing their levels to remain constant even when rDNA transcription is inhibited.
Alternatively, C. merolae may possess an efficient rRNA processing machinery that optimizes
the conversion of available pre-rRNA into mature rRNAs, compensating for the diminished
precursor synthesis. The organism may also regulate rRNA turnover by slowing the degradation
of mature rRNAs during stress, thereby preserving essential ribosomal components. These
adaptive mechanisms, potentially involving specialized pathways that protect and stabilize
rRNAs, highlight the resilience of C. merolae in maintaining cellular functions despite
environmental challenges. This contrasts with organisms such as Arabidopsis thaliana, where
heat stress leads to a rapid decline in rRNA levels, illustrating the evolutionary diversity in stress
51

response strategies. Furthermore, the reduction in pre-rRNA levels at 57 C, suggesting that
rDNA transcription is inhibited in C. merolae under heat stress may be attributed to an effect on
RNA polymerase I or a regulatory element involved in rDNA transcription. These findings align
with observations in Arabidopsis thaliana, where heat stress impedes pre-rRNA processing
(Darriere et al., 2022), and in mammals, where a brief heat shock inhibits both pre-rRNA
transcription and its subsequent processing into mature rRNAs (Ghosha and Jacob, 1996).
Additionally, it is well-established that heat stress inhibits rDNA transcription in animal cells
(Ghosha and Jacob, 1996; Coccia et al., 2017).
Further research is needed to elucidate the molecular basis of this stability, which could offer
insights into the broader mechanisms of stress tolerance in extremophiles.

52

Chapter 4 - In Silico Prediction of C. merolae's MRP RNA: Secondary
Structure, Conserved Regions, and Protein Constituents of the RNase MRP
Complex

53

4.1 Introduction
The RNase MRP complex is an essential ribonucleoprotein enzyme involved in the processing of
precursor rRNA, particularly in the generation of the mature 5.8S rRNA in eukaryotes. Its RNA
component is essential for its catalytic activity, while its associated proteins are necessary for
structural stability and proper function. Studies in well-characterized organisms such as
Saccharomyces cerevisiae, Homo sapiens, Drosophila melanogaster, and Chlamydomonas
reinhardtii have revealed conserved secondary structures within the RNase MRP RNA
component and a common set of protein constituents. In the case of S. cerevisiae, conserved
regions, and domains were identified through structural probing experiments, while the structure
of D. melanogaster MRP RNA and other organisms were derived through comparative genomics
involving S. cerevisiae and Homo sapiens. These features highlight the evolutionary
conservation and functional importance of the complex. However, Cyanidioschyzon merolae, a
red alga with a highly streamlined genome, offers a unique opportunity to investigate how these
structural and functional elements have been conserved or diverged over evolutionary time.
In this chapter, I address two key questions: Does the RNase MRP RNA in C. merolae possess
conserved secondary structural regions comparable to those found in other eukaryotic
organisms? Second, does the RNase MRP complex in C. merolae retain the same protein
constituents, or has it lost some of these proteins as part of its evolutionary adaptation? By
answering these questions, this analysis will not only enhance our understanding of the structural
and functional conservation of RNase MRP in C. merolae but also provide insights into its
evolutionary divergence.
Predicting the secondary structure of RNase MRP RNA is important for identifying conserved
regions that may be functionally important. Conservation of structure often correlates with
54

conservation of function, and by comparing C. merolae to S. cerevisiae, Drosophila, humans,
and other organisms, we can infer which regions of the RNA are essential for catalytic activity.
Additionally, the identification of these regions opens the possibility of mirroring mutational
analysis experiments performed in S. cerevisiae. If similar regions of the RNase MRP RNA are
conserved in C. merolae, mutations performed in these regions in S. cerevisiae may provide
valuable functional insights that could be translated to C. merolae.
Furthermore, the identification of the protein constituents of RNase MRP in C. merolae is of
particular interest because this organism’s reduced genomic content suggests potential
evolutionary streamlining. In other eukaryotes, RNase MRP is composed of several conserved
proteins that are necessary for its assembly and function. Understanding whether C. merolae has
retained all these proteins or has lost some provides important clues to how the RNase MRP
complex may have adapted to function with fewer components in this minimalistic organism.
This analysis also connects to other chapters of this thesis, particularly the investigation into how
RNase MRP functions under heat stress and its role in the processing of 5.8S rRNA. By first
determining whether C. merolae retains key conserved structural and protein elements of RNase
MRP, we can then assess whether these elements are involved in the observed resilience of the
complex under stress conditions. Thus, in this chapter, I built a secondary structure model based
on sequence alignments with MRP from other organisms using RNA folding software and
manual examination of base pairing potential to lay a foundation for the broader study of RNase
MRP function in C. merolae, both in normal conditions and under heat stress, linking structure to
function and evolutionary adaptation.

55

4.2 Materials and Methods
4.2.1 Genome and Protein Sequences
RNase MRP protein sequences were retrieved from NCBI - The National Centre for
Biotechnology Information at the National Library of Medicine and the Universal Protein
knowledgebase (http://www.ncbi.nlm.nih.gov/gene/), Swiss-Prot (http://www.expasy.ch/sprot/),
and UniProt (http://www.expasy.uniprot.org/). The C. merolae genome sequence was obtained
from the NCBI GenBank database. Comparative genomic analysis was conducted with other
model organisms, including Saccharomyces cerevisiae (baker's yeast), Homo sapiens (humans),
Drosophila melanogaster (fruit fly), Caenorhabditis elegans (nematode), and Chlamydomonas
reinhardtii (green alga). The genome and protein sequences of these organisms were also
retrieved from NCBI, UniProt, and Ensembl databases to facilitate comparative analysis. These
sequences were used as references for identifying homologous regions and conserved domains.
4.2.2 Alignment of Cyanidioschyzon merolae MRP RNA with Other Organisms
The MRP RNA sequence of C. merolae was retrieved from the NCBI GenBank database. To
compare this sequence with those from other organisms, homologous MRP RNA sequences were
obtained from RNAcentral and Rfam databases. Organisms used for this comparative analysis
included Saccharomyces cerevisiae (baker's yeast), Homo sapiens (humans), Drosophila
melanogaster (fruit fly), Caenorhabditis elegans (nematode), and Chlamydomonas reinhardtii
(green alga). In addition, the following organism-specific databases were used; the
Saccharomyces Genome Database (Cherry et al., 1997) and the Drosophila Genome Project
(Christie et al., 2003). Multiple sequence alignments were performed using Clustal Omega
v3.13.8 (Sievers and Higgins, 2013) and MUSCLE v3.8.31 (Sievers et al., 2010), allowing for
56

the identification of conserved nucleotides and secondary structure elements across different
species. Pairwise sequence alignments were also conducted using EMBOSS Needle
(http://www.ebi.ac.uk/Tools/psa/emboss_needle/) for more detailed comparisons, highlighting
conserved base-pairing interactions relevant to the secondary structure of MRP RNA. The results
of these alignments were used to infer evolutionary relationships and to validate the structural
predictions made for C. merolae MRP RNA.

4.2.3 Identification of Protein Homologs
To identify homologs of the known protein constituents of RNase MRP in Cyanidioschyzon
merolae, Position-Specific Iterative BLAST (PSI-BLAST) searches (Altschul and Koonin, 1998)
was used. All known protein subunits of RNase MRP were used as queries in these searches,
with an E-value threshold of 0.001 set for inclusion in subsequent PSI-BLAST iterations. In
some cases, multiple PSI-BLAST searches were performed with different query sequences to
maximize the identification of potential homologs. The primary database searched was the NCBI
GenBank protein set (Benson et al., 2006). For proteins not present in this database, sequences
were retrieved from individual genome projects or identified through TBLASTN searches of
genome sequences. These novel sequences were subsequently included in the database used for
PSI-BLAST searches to ensure comprehensive coverage. Additionally, in some instances, further
homologs were identified using Pfam models, which provided supplementary sequence data and
enhanced the search's robustness.

57

4.2.4 Validation of RNase MRP Protein Homologs via Reciprocal BLAST Searches
To identify homologs of RNase MRP protein constituents in C. merolae, I employed a reciprocal
BLAST search strategy based on the approach described by (Ward and Moreno-Hagelsieb 2014).
Initially, forward BLASTP searches were conducted using known RNase MRP proteins from
organisms such as Saccharomyces cerevisiae (yeast), Homo sapiens (humans), Drosophila
melanogaster (fruit fly), Caenorhabditis elegans (nematode), and Chlamydomonas reinhardtii
(green alga) as queries to identify putative orthologs in C. merolae. These searches were
performed using the NCBI BLAST tool (Altschul and Koonin, 1998) with a BLOSUM62 matrix
and an E-value threshold of 0.001 to ensure significant matches. To validate the identified
homologs and resolve ambiguous or unexpected results, reciprocal BLASTP searches were
carried out. In this approach, the identified C. merolae proteins were used as queries against the
protein databases of the original species from which the RNase MRP subunits were first
identified. The same E-value threshold of 0.001 was applied in these reciprocal searches. A C.
merolae protein was confirmed as a true homolog if the reciprocal BLAST search returned the
original query protein or a closely related homolog as the top hit with an E-value smaller than
10^-10. If the top hit did not correspond to the original query protein, all candidate C. merolae
proteins with an E-value threshold of less than 10^-2 were further analyzed against the original
organism's proteome. Additionally, the domain structure of uncertain candidates was examined
using NCBI’s DELTA-BLAST tool to distinguish true orthologs from false positives. This
comprehensive approach increased the likelihood that the identified proteins were indeed
functional homologs of the known RNase MRP components in C. merolae.

58

4.3. Results
In this section, I address the question of whether the RNase MRP RNA in Cyanidioschyzon
merolae retains conserved structural regions and protein constituents found in other eukaryotic
organisms. To explore this, I used comparative computational analysis to predict the secondary
structure of the RNase MRP RNA, alongside bioinformatic tools to identify the corresponding
protein components compared with known RNase MRP structures in organisms such as
Saccharomyces cerevisiae, Homo sapiens, Drosophila melanogaster, and Chlamydomonas
reinhardtii. The secondary structure predictions revealed conserved regions that align with
functional domains in other species, suggesting that C. merolae has maintained functional
aspects of RNase MRP. Similarly, the analysis of protein constituents showed a streamlined set
of core proteins, consistent with the organism's reduced genome but still sufficient for
maintaining RNase MRP activity. The following sections will detail these structural predictions
and protein identifications, and their implications for understanding the evolutionary
conservation and divergence of RNase MRP in C. merolae.
4.3.1 Conserved Regions and Predicted Secondary Structure of RNase MRP RNA
To identify conserved regions, nucleotides, and structural elements within C. merolae MRP
RNA that correspond to these experimentally validated and evolutionarily conserved structures
(Figures 1.5A and 1.6C), I performed a comparative genomics analysis. This approach allowed
for the alignment of C. merolae RNase MRP RNA with the conserved and structurally probed
RNase MRP RNA sequences from S. cerevisiae, D. melanogaster, and other model organisms,
facilitating the identification of evolutionarily conserved features within the C. merolae RNase
MRP RNA structure.

59

Figure 4.1 below illustrates the alignment of C. merolae RNase MRP RNA with D.
melanogaster RNase MRP RNA, highlighting the conserved regions that underpin the prediction
of structural motifs within the RNase MRP RNA. The sequence alignment reveals a 46.4%
similarity between the RNase MRP RNAs of these two organisms. Consistent with existing
literature (Piccinelli et al., 2005), the alignment shows the conservation of the ubiquitous P1, P2,
and P3 helices, as well as the CR-I, CR-IV, and CR-V regions. The P4 helix is again formed by
pairing of elements from the CR-I and CR-V regions.
Figure 4.2 also illustrates the alignment of C. merolae RNase MRP RNA with Chlamydomonas
reinhardtii RNase MRP RNA, highlighting the conserved regions that serve as a foundation for
predicting structural motifs within the RNase MRP RNA. The sequence alignment reveals a
39.7% similarity between the RNase MRP RNAs of these two organisms.
Furthermore, Figures 4.3 and 4.4 illustrates the alignment of C. merolae RNase MRP RNA with
Saccharomyces cerevisiae and Homo sapiens RNase MRP RNA, highlighting the conserved
regions that serve as a basis for predicting structural motifs within the RNase MRP RNA. The
sequence alignment reveals a 39.5% and 40.1% similarity between C. merolae against Sc and
Hm RNase MRP RNAs respectively. The alignment shows the conservation of the Domain 1
region (the CR-I, CR-IV, and CR-V regions). The P4 helix is formed by pairing elements from
the CR-I and CR-V regions.
Lastly, figure 4.5, the predicted secondary structure, was built using manual model building to
manipulate the secondary structure with the predicted conserved regions and I utilized other
bioinformatic tools (Mfold and sfold) to make it domain 2 region.

60

Aligned sequences: 2
# 1: Drosophila Melanogaster (Dm)
# 2: Cyanidioschyzon Merolae (Cm)
# Matrix: EDNAFULL
# Gap penalty: 10.0
# Extend penalty: 0.5
# Length: 515
# Identity:
239/515 (46.4%)
# Similarity:
239/515 (46.4%)
# Gaps:
206/515 (40.0%)
# Score: 444.5
#=======================================
P1 Helices
EMBOSS_001
1 GCCGGTTTGAGTCTTCCATGCTTGTCTCTCG--------GGGCCACAAA.||...||.|||.|.|.|| ||..||||
|.||.||..|
EMBOSS_001
1 --GGGGAAGACTCTGCTAAGC--GTTCCTCGTTATCAGAGCGCTACGTAC
EMBOSS_001
42 -----ACG---AGTTCCTGG---------TAA------CTC------AA|||
.|.||||||
|||
|||
||
EMBOSS_001
47 GGTTTACGGGAGGGTCCTGGATTAGCCCCTAAACCAGGCTCGGTGCGAAC
P3
P4(CR-I)
EMBOSS_001
62 ---------CTGAT--AATGCC------------CTGGGCGAAAGTCCCC
||||| .|||||
|||||.||||||||||
EMBOSS_001
97 AGGTGCGCCCTGATTCGATGCCAGCCGTTTGGCTCTGGGTGAAAGTCCCC
EMBOSS_001
89 GGGC-CTAGGATAGAAAGTATCAAGGT-GTAA--AAAG-TGTGC---ACA
||.| .||||..|||.|||.|| ||| |||| |||| ||.||
.||
EMBOSS_001
147 GGACAGTAGGTCAGAGAGTGTC--GGTGGTAAGCAAAGCTGGGCGTTTCA
EMBOSS_001
131 AAACACCCACCACCCCTG-TGGTGGGTGGTGCA-------TTCGCCTATA
...|.|.|| |||..||| |.|.||| ..||||
..|.||.|||
EMBOSS_001
195 GGTCGCGCA-CACTTCTGCTCGCGGG-ACTGCAACCACAGAACCCCAATA
EMBOSS_001
173 ----------TTCTGCG-------GAATTTCGCCTGGCGTATGGATGAAG
|.|||||
|||..||||.||.|| |||| ||.||
EMBOSS_001
243 AGAGCGGGAGTGCTGCGCGACAGTGAACGTCGCTTGTCG-ATGG-TGCAG
EMBOSS_001
206 AGGATTTTATCC--GAA------------------TCCTTACGCGCCA-G
.|
| |||
|.|||.|||.||| |
EMBOSS_001
291 CG---------CTGGAACCTGCTGCCGCGTTGGGGTGCTTGCGCCCCATG
CR-IV
P2
EMBOSS_001
235 GTTGTCTGCGGAAATCTGCCAGAGT-AATCTTAGATATGG-ACGAG---.|||.|.||||.|..||||
|.| |||| ||..|||| |||||
EMBOSS_001
332 TTTGGCAGCGGCAGCCTGC---ACTGAATC--AGCAATGGAACGAGCGAG
EMBOSS_001
279 -----------TTGGTAGGACTCGGCGGGTGGTGTTCACACACTTTCTCG
||..| ||| |||||
.||||
EMBOSS_001
377 AGCGACCGTCCTTCAT-GGA--CGGCG------------------ACTCG
CR-V
EMBOSS_001
318 TCTGAGAAACCGCCTACACAGAATGGGGCTTACATTGGGAAACTCGGACG
|||
|||.||||||||||
||||.|.|.
EMBOSS_001
406 --TGA------------ACACAATGGGGCTT----------ACTCTGGCA
EMBOSS_001
368 GCGCACTCCCTTTTT
382 Dm
|.|..||||||
EMBOSS_001
432 GTGTGCTCCCT---442 Cm

41 Dm
46 Cm
61 Dm
96 Cm
88 Dm
146 Cm
130 Dm
194 Cm
172 Dm
242 Cm
205 Dm
290 Cm
234 Dm
331 Cm
278 Dm
376 Cm
317 Dm
405 Cm
367 Dm
431 Cm

Figure 4.1: Sequence alignment of C. merolae RNase MRP RNA with Drosophila melanogaster RNase MRP
RNA. Conserved regions within Domain 1 of the MRP structure are highlighted. Drosophila conserved sequences
are highlighted in yellow and Cm in red. The names of the conserved regions are labeled in green.

61

# Aligned sequences: 2
# 1: Chlamydomonas reinhardtii (Cr)
# 2: Cyanidioschyzon merolae (Cm)
# Matrix: EDNAFULL
# Gap penalty: 10.0
# Extend penalty: 0.5
# Length: 474
# Identity:
159/474 (33.5%)
# Similarity:
188/474 (39.7%)
# Gaps:
239/474 (50.4%)
# Score: 356.5
EMBOSS_001
1 ------------GCGCAGUG-----------CGGAGGGCCACCU-CGGUG
||..||.|
|.|||.||.||.: |||:
EMBOSS_001
1 GGGGAAGACTCTGCTAAGCGTTCCTCGTTATCAGAGCGCTACGTACGGTEMBOSS_001
27 UCACUUACGGCAGGAGUCGAGGGGCUGCUGCUUUGAGCGCGGCC-----C
::||||
||||| :.|:|..:: |||
||
|
EMBOSS_001
50 ----TTACGG--------GAGGG--TCCTGGATT-AGC----CCCTAAAC
P3
EMBOSS_001
72 CCGGCUGGGCGC-------UGCAUCCAUGUAUU-GA-GCACAUC-----|.|||:.||.||
:|| .||.:| |:: || || ||.|
EMBOSS_001
81 CAGGCTCGGTGCGAACAGGTGC-GCCCTG-ATTCGATGC-CAGCCGTTTG
P4(CR-I)
P8
EMBOSS_001
107 -----GGGCGAAAGUCCCCGGGCGACGGGGCAGAGAGUGCC-------AG
|||.|||||:||||||.|....||.|||||||:|.|
||
EMBOSS_001
128 GCTCTGGGTGAAAGTCCCCGGACAGTAGGTCAGAGAGTGTCGGTGGTAAG
EMBOSS_001
145 C------------------------------------------CCGCAAG
|
|.||||.
EMBOSS_001
178 CAAAGCTGGGCGTTTCAGGTCGCGCACACTTCTGCTCGCGGGACTGCAAC
EMBOSS_001
153 GGCAGGACCCC-------------------CGCG--------CGUCGUUU
..|||.|||||
||||
||:||.::
EMBOSS_001
228 CACAGAACCCCAATAAGAGCGGGAGTGCTGCGCGACAGTGAACGTCGCTT
EMBOSS_001
176 UUACGA-----------------------CCGCG--GGGCAGCUCGCCAU
.: |||
||||| |||..||:.||
EMBOSS_001
278 GT-CGATGGTGCAGCGCTGGAACCTGCTGCCGCGTTGGGGTGCTTGC--EMBOSS_001
201 AGCAACA-GUUAUGUGGC------------CAGUG-----GGGAUACAAC
||..|| |::
:|||
||.:|
|..|:..|||
EMBOSS_001
324 -GCCCCATGTT---TGGCAGCGGCAGCCTGCACTGAATCAGCAATGGAAC
CR-V
EMBOSS_001
233 ---------CCACC------------------------AACACAACGGGG
|.|||
|||||||.||||
EMBOSS_001
370 GAGCGAGAGCGACCGTCCTTCATGGACGGCGACTCGTGAACACAATGGGG
EMBOSS_001
250 CUUACUCCCGCACUGACG-----267 Cr
|::||:|..|||.:| .|
EMBOSS_001
420 CTTACTCTGGCAGTG-TGCTCCCT
442 Cm

26 Cr
49 Cm
71 Cr
80 Cm
106 Cr
127 Cm
144 Cr
177 Cm
152 Cr
227 Cm
175 Cr
277 Cm
200 Cr
323 Cm
232 Cr
369 Cm
249 Cr
419 Cm

Figure 4.2: Sequence alignment of C. merolae RNase MRP RNA with Chlamydomonas reinhardtii RNase
MRP RNA. P3, P4(CR-I and CR-V), and P8 regions 1 of the MRP structure are highlighted. Chlamydomonas
conserved sequences are highlighted in yellow and Cm in red. The names of the regions are labeled in green.

62

# Aligned sequences: 2
# 1: Saccharomyces cerevisiae (Sc)
# 2: Cyanidioschyzon Merolae (Cm)
# Matrix: EDNAFULL
# Gap penalty: 10.0
# Extend penalty: 0.5
# Length: 521
# Identity:
206/521 (39.5%)
# Similarity:
206/521 (39.5%)
# Gaps:
260/521 (49.9%)
# Score: 278.5
EMBOSS_001
1 ----AATCCATGAC----CAAAG----AATCGTCACAAATCGAAGC---||
|||
|.|||
..||||
.|||..|||
EMBOSS_001
1 GGGGAA-----GACTCTGCTAAGCGTTCCTCGT----TATCAGAGCGCTA
EMBOSS_001
35 --------TTACAAAA------TGGAGTA------AAATTTTTTTTACTC
||||...|
||||.||
|||
.|
EMBOSS_001
42 CGTACGGTTTACGGGAGGGTCCTGGATTAGCCCCTAAA----------CC
EMBOSS_001
65 AG-----------------------TAAT---ATGC------TTTG---||
|.||
||||
||||
EMBOSS_001
82 AGGCTCGGTGCGAACAGGTGCGCCCTGATTCGATGCCAGCCGTTTGGCTC
P3
P4(CR-I)
P8
EMBOSS_001
79 -GGTTGAAAGTCTCCCACCAATTCGTATGCGGAAA--ACGTAATGAGATT
||.||||||||.|
||||.| |.||.| ||||.|
EMBOSS_001
132 TGGGTGAAAGTCCC---------------CGGACAGTAGGTCA-GAGAGT
EMBOSS_001
126 --------TAA--AAA--------TTTTA----------AATTGT----||| |||
|||.|
|.||.|
EMBOSS_001
166 GTCGGTGGTAAGCAAAGCTGGGCGTTTCAGGTCGCGCACACTTCTGCTCG
EMBOSS_001
143 ------TTAAATC-----AACTCATTAA-----GGAG-GATGC----CCT
|..||.|
|.|.||.|||
|||| |.|||
|..
EMBOSS_001
216 CGGGACTGCAACCACAGAACCCCAATAAGAGCGGGAGTGCTGCGCGACAG
EMBOSS_001
172 TG---GGTATTCTGCTTCTTGA---------CCTGGTACCT-CT-----A
||
|
|| ||||.|.||
.||||.|||| ||
.
EMBOSS_001
266 TGAACG----TC-GCTTGTCGATGGTGCAGCGCTGGAACCTGCTGCCGCG
EMBOSS_001
204 TTGCAGGGTACTGG-------TGTTTTCTTCGGTACTGGATTCCGTTTGT
||| ||||.||.|
|||||
||.|..||
|.|..||.
EMBOSS_001
311 TTG--GGGTGCTTGCGCCCCATGTTT-----GGCAGCGG---CAGCCTGC
CR-IV
EMBOSS_001
247 ATGGAATCTAAACCATAGTTATG--ACGATTGC-----------TCTTTC
|..|||||
||..||| |||| ||
||.|||
EMBOSS_001
351 ACTGAATC--------AGCAATGGAACGA--GCGAGAGCGACCGTCCTTC
CR-V
EMBOSS_001
284 CCGTGCTGGA-------TCGAGTAACCCAATGGAGCTTACTATTCT---.||||
|||.| |||.||||||.||||||
|||
EMBOSS_001
391 -----ATGGACGGCGACTCGTG-AACACAATGGGGCTTAC---TCTGGCA
EMBOSS_001
323 ---TGGTCCATGGATTCACCC
340 Sc
||.|||.|
EMBOSS_001
432 GTGTGCTCCCT---------442 Cm

34 Sc
41 Cm
64 Sc
81 Cm
78 Sc
131 Cm
125 Sc
165 Cm
142 Sc
215 Cm
171 Sc
265 Cm
203 Sc
310 Cm
246 Sc
350 Cm
283 Sc
390 Cm
322 Sc
431 Cm

Figure 4.3: Sequence alignment of C. merolae RNase MRP RNA with Saccharomyces cerevisiae RNase MRP
RNA. The alignment highlights conserved regions that provide a foundation for predicting structural motifs within
the RNase MRP RNA. The sequence similarity between the RNase MRP RNAs of these two organisms is 39.5%.
Conserved regions within Domain 1, including CR-I, CR-IV, and CR-V, are evident. The P4 helix is observed to
form through pairing elements from the CR-I and CR-V regions.

63

# Aligned sequences: 2
# 1: Homo sapiens (Hm)
# 2: Cyanidioschyzon Merolae (Cm)
# Matrix: EDNAFULL
# Gap penalty: 10.0
# Extend penalty: 0.5
# Length: 469
# Identity:
192/469 (40.9%)
# Similarity:
192/469 (40.9%)
# Gaps:
226/469 (48.2%)
# Score: 335.0
EMBOSS_001
1 ---------------------------------------------TGGTT
.||||
EMBOSS_001
1 GGGGAAGACTCTGCTAAGCGTTCCTCGTTATCAGAGCGCTACGTACGGTT
P2 helices
EMBOSS_001
6 CGTGCTGAAGG--CCTGTAT-----CCT-----AGGCT--------ACA|.|.|.||| ||||.||
|||
|||||
|||
EMBOSS_001
51 --TACGGGAGGGTCCTGGATTAGCCCCTAAACCAGGCTCGGTGCGAACAG
P3
P4(CR-I)
EMBOSS_001
35 -----CACTGAGGACTCTGTTCCTCCCCTTTCCGC-CTAGGGGAAAGTCC
|.||
||.||..|.||..||.||| .|| ||.||.||||||||
EMBOSS_001
99 GTGCGCCCT---GATTCGATGCCAGCCGTTT-GGCTCTGGGTGAAAGTCC
EMBOSS_001
79 CCGGAC-CTCGGGCAGAGAGTGCCACGTGCATACGCA--------CGT-|||||| .|.||.|||||||||.| .||| .||.|||
|||
EMBOSS_001
145 CCGGACAGTAGGTCAGAGAGTGTC-GGTG-GTAAGCAAAGCTGGGCGTTT
EMBOSS_001
118 -AG-------ACA-TTCCCCGCTTCCC---ACT----CCAAAGTCCGCCA
||
||| |||..|
||.|
|||
|||.||..|.|||
EMBOSS_001
193 CAGGTCGCGCACACTTCTGC---TCGCGGGACTGCAACCACAGAACCCCA
EMBOSS_001
152 AGAAG-------------------------CGTATCCCGCT--------G
|.|||
|||
||||
|
EMBOSS_001
240 ATAAGAGCGGGAGTGCTGCGCGACAGTGAACGT----CGCTTGTCGATGG
EMBOSS_001
169 AGCGGCG-----------TGGCGCG--GGGGCGTCATCCGTCAGCTCCCT
.||.|||
||.|||| ||||.| |.|.||
||.
EMBOSS_001
286 TGCAGCGCTGGAACCTGCTGCCGCGTTGGGGTG-CTTGCG-------CCC
P2
EMBOSS_001
206 CTAGTTACGCA--GGCAG--TGC---GTGTC----------CGCGC---A
|..|||..||| ||||| |||
|..||
||.||
|
EMBOSS_001
328 CATGTTTGGCAGCGGCAGCCTGCACTGAATCAGCAATGGAACGAGCGAGA
CR-V
EMBOSS_001
236 CCAACC-------------------------ACAC---GGGGCTCATTCT
.|.|||
||||
||||||.|.|||
EMBOSS_001
378 GCGACCGTCCTTCATGGACGGCGACTCGTGAACACAATGGGGCTTACTCT
EMBOSS_001
258 --CAGCGCGGCTGTT---270 Hm
|||.|
||.|
EMBOSS_001
428 GGCAGTG----TGCTCCCT
442 Cm

5 Hm
50 Cm
34 Hm
98 Cm
78 Hm
144 Cm
117 Hm
192 Cm
151 Hm
239 Cm
168 Hm
285 Cm
205 Hm
327 Cm
235 Hm
377 Cm
257 Hm
427 Cm

Figure 4.4: Sequence alignment of C. merolae RNase MRP RNA with Homo sapiens RNase MRP RNA. The
alignment highlights conserved regions that serve as a basis for predicting structural motifs within the RNase MRP
RNA. The sequence similarity between the RNase MRP RNAs of these two organisms is 40.1%. Conserved regions
within Domain 1, including CR-I, CR-IV, and CR-V, are evident. The P4 helix is formed by pairing elements from
the CR-I and CR-V regions.

64

Figure 4.5: Predicted secondary structure of C. merolae RNase MRP RNA. The structure was generated based
on sequence alignments with RNase MRP RNAs from Drosophila melanogaster, Saccharomyces cerevisiae, and
Chlamydomonas reinhardtii. Conserved regions, including the P1, P2, and P3 helices, as well as the CR-I, CR-IV,
and CR-V regions, are highlighted. The P4 helix, formed by the pairing of elements from the CR-I and CR-V
regions, is also depicted.

65

4.3.2 Identification of Protein Constituents of the RNase MRP Complex
Having predicted the structure of C. merolae RNase MRP RNA, I sought a more complete
picture of the C. merolae MRP particle by identifying the corresponding protein constituents of
RNase MRP. I used PSI-BLAST and reciprocal BLAST searches to identify homologs of the
known protein subunits of RNase MRP from various organisms. I predicted the protein
constituents of the C. merolae RNase MRP complex based on what has been found in
Saccharomyces cerevisiae, Homo sapiens, Chlamydomonas reinhardtii, and Drosophila
melanogaster as shown below (Table 4.1).
Table 4.1: Predicted Protein Constituents of RNase MRP in Cyanidioschyzon merolae Identified Through
Comparative Genomic Analysis.

S.
cerevisiae
MRP
Proteins
POP1
POP3
POP4
POP5
POP6
POP7
POP8
SNM1
RMP1
RPP1/p30
RPR2

S.
cerevisiae
Protein
Length
(aa)
875
195
279
173
158
140
133
198
201
293
144

C.merolae
Gene
CMR143C
CMG096C
CMO008C
CMM152C
CMI187C

Best Hit
(BH)
+
+
+
+
+

aa = Amino acids

C.merolae
Protein
Length
(aa)
920
282
194
298
163

+ = Present

Query
Cover
(QC) %
23
46
54
67
70

% ID
30
26
25
26
20

- = Absent

E-value
2e-08
3e-08
0.018
2e-09
3e-18

Reciprocal
Best Hit
(RBH)
+
+
+
+
+

e = 10^

In comparing the RNase MRP protein constituents between Saccharomyces cerevisiae and
Cyanidioschyzon merolae, I observed that only five of the eleven proteins in S. cerevisiae have
homologs in C. merolae. These homologous proteins vary in their degree of conservation based
66

on amino acid length, query coverage (QC), E-value, and percentage identity. POP1, although
showing similar amino acid lengths (920 in C. merolae vs. 875 in S. cerevisiae), POP1 has
relatively low query coverage (23%), an E-value of 2 × 10⁻⁸, and a percentage identity of 30%.
This suggests that while the overall length is comparable, the protein is less conserved in terms
of sequence similarity, indicating functional divergence. POP4 has a query coverage of 46%, an
E-value of 3 × 10⁻⁸, and a percentage identity of 26%. With nearly identical lengths (282 vs. 279
amino acids), this suggests moderate conservation, but the lower percentage identity indicates
some sequence variability. POP5, with 54% query coverage, an E-value of 0.018, and a
percentage identity of 25%, shows moderate conservation, though the length of the protein in C.
merolae (194) is slightly longer than in S. cerevisiae (173). The higher E-value and lower
identity indicate that POP5 may have diverged more than others. RPP1, showing 67% query
coverage, an E-value of 2 × 10⁻⁹, and a percentage identity of 26%, is highly conserved in terms
of length (298 vs. 293), with relatively higher conservation overall compared to POP5. RPR2 is
the most conserved protein of the five, with 70% query coverage, the lowest E-value (3 × 10⁻¹⁸),
and a percentage identity of 20%. Despite the lower identity percentage, the close similarity in
length (163 vs. 144) and the high query coverage indicate that RPR2 retains a highly conserved
structure. In conclusion, RPR2 and RPP1 are the most conserved based on query coverage, Evalue, and amino acid length, with RPR2 showing particularly strong structural conservation.
POP1, despite its similar length, has the lowest conservation based on sequence similarity and
query coverage, making it the least conserved of the group.

67

4.4. Discussion
I was surprised to observe the presence of stably-expressed, non-coding RNAs in the introns of
C. merolae, and particularly their apparent accumulation in response to heat stress. To
investigate their function, I sought to investigate the identity of that from the gene CMK142T.
The comparative genomic and computational analyses carried out in this study reveal the
conservation of RNase MRP RNA and proteins in C. merolae. By aligning the predicted
secondary structure of C. merolae RNase MRP RNA with those from other well-characterized
organisms, including Saccharomyces cerevisiae, Homo sapiens, Drosophila melanogaster, and
Chlamydomonas reinhardtii, I observed that key structural motifs, particularly those within
Domain 1, are conserved across these species (Figure 4.1 to Figure 4.4).
The sequence alignments of C. merolae RNase MRP RNA with those of Drosophila
melanogaster, Saccharomyces cerevisiae, and Chlamydomonas reinhardtii, is evidence that
certain structural motifs and conserved regions are maintained across these diverse species. The
alignment results consistently demonstrate the preservation of key elements, such as the P1, P2,
and P3 helices, as well as the CR-I, CR-IV, and CR-V regions, which are integral to the RNase
MRP RNA's function. These conserved motifs not only highlight the evolutionary significance of
these regions but also provide insights into the structural stability and functional integrity of the
RNase MRP complex across different organisms.
The consistent observation of these conserved regions across multiple species justifies the
prediction of the secondary structure of C. merolae RNase MRP RNA (Figure 4.5). The
predicted structure, grounded in both computational models and comparative genomic data, aims
to reflect the conserved structural motifs identified in these alignments. This suggests that the
68

catalytic core and the RNA processing function of RNase MRP have been maintained through
evolution, which is indicative of their essential role in cellular processes.
However, the identification of protein constituents reveals a notable divergence in the
complexity of the RNase MRP complex in C. merolae. While the well-studied RNase MRP in S.
cerevisiae comprises 11 protein subunits (Piccinelli et al., 2005; Rosenblad et al., 2006; Lopez et
al. 2009), the complex in C. merolae appears to be composed of only five (5) proteins. This
reduction in protein constituents may reflect adaptations specific to C. merolae, possibly due to
its minimalistic genome and the specialized environment in which it thrives. The retention of the
core proteins POP1, POP4, POP5, RPP1, and RPR2 highlights their likely important roles in the
assembly and function of the RNase MRP complex in this red alga.
These findings underscore the balance between conservation and adaptation within the RNase
MRP complex, emphasizing the potential for functional diversity even among conserved
molecular machines. Further experimental validation of these computational predictions is
necessary to elucidate the precise roles of these protein subunits and to understand how the
reduced complexity of RNase MRP in C. merolae affects its functionality.

69

Chapter 5 - Mutational analysis of the RNA component of C. merolae RNase
MRP Reveals a Shift in the Stoichiometry of the Two Forms of 5.8S rRNA

70

5.1 Introduction
I conducted a mutational analysis of the RNA component of C. merolae RNase MRP to elucidate
its function and impact on rRNA processing. To achieve this, I employed two distinct
methodologies: direct gene replacement and plasmid shuffling. In the former, I directly replaced
the WT MRP with a mutant MRP via homologous recombination while the latter involved
generating an MRP deletion strain covered by a counter-selectable plasmid with WT MRP. Upon
transforming a second plasmid with a mutant MRP, the WT can be “shuffled” out by counterselection.
In Saccharomyces cerevisiae, the RNase MRP RNA, encoded by the essential gene NME1, had
been extensively mutagenized to provide insights into its structure and functions. Researchers
employed a gene shuffle technique to create yeast strains expressing 26 independent mutations in
the NME1 gene. These strains were characterized based on their growth at various temperatures,
their ability to utilize different carbon sources, the stability of RNase MRP RNA, and the
efficiency of 5.8S rRNA processing (Shadel et al., 2000). This detailed analysis revealed that 11
mutations were lethal, six exhibited temperature-sensitive lethality, and several mutants
displayed a preference for non-fermentable carbon sources (Table 5.1). Importantly, the severity
of growth defects in these mutants correlated directly with the extent of disruption in 5.8S rRNA
processing (Figure 5.1), thereby identifying the essential regions of RNase MRP RNA for its
nuclear function. My investigation in C. merolae mirrors the approach used in yeast, offering a
perspective on the conserved roles of RNase MRP across species. Specifically, I tried three
mutations but was able to examine the effects of two mutations (highlighted in green in Table
5.1) of the RNA component of C. merolae RNase MRP on the stoichiometry of the two forms of
5.8S rRNA and assessed the overall stability of RNase MRP RNA. By juxtaposing our results
71

with those obtained in yeast, we aim to demonstrate that C. merolae MRP has a similar function
to yeast MRP in regulating the balance of the two forms of 5.8S rRNA.
Table 5.1. Summary of NME1 Mutagenesis Data
1

2

3

The table summarizes the mutagenesis data for individual NME1 mutants, with each mutant listed on the left. The
specific base changes are identified, with numbering corresponding to the position within the NME1 transcript
(Schmitt and Clayton, 1992). 1Growth phenotypes are scored on a scale from 0 to 4, where 0 indicates no growth
and 4 represents wild-type growth. When two numbers are provided, they reflect growth on dextrose alone or
dextrose and glycerol. 2 The ratios of 5.8S rRNA were measured from ethidium bromide-stained gels and represent
steady-state levels. 3 RNA stability is also quantified on a scale from 0 to 4, with 0 indicating no detectable RNA
and 4 corresponding to wild-type RNA levels (Shadel et al., 2000).

72

Figure 5.1. NME1 mutation results in altered 5.8S rRNA isoform ratio. (top) present Northern blot analyses for
the indicated NME1 mutants, using an NME1 probe to detect MRP RNA and an SCR1 probe as a control for RNA
loading levels. (bottom) display ethidium bromide-stained agarose gels, highlighting the 5.8S rRNA profiles for
each mutant. For temperature-sensitive mutants, RNA was harvested after shifting to the non-permissive
temperature. Inviable mutants were maintained with a wild-type copy of the NME1 gene (Shadel et al.,2000).

5.2 Materials and Methods.
5.2.1 Construction of Plasmid for Mutagenesis
To generate a plasmid for mutagenesis, I employed a systematic approach involving sequential
restriction digests, gel electrophoresis, oligonucleotide duplex formation, ligation, and cloning.
This procedure was executed in two distinct phases (Figures 5.2 and 5.3).

73

Figure 5.2. Modification of PCR2.1 with Multiple Cloning Sites. Arrows indicate the directionality of the
workflow, starting from the bottom of the figure.

74

To obtain a backbone vector for subsequent procedures, a restriction digest was performed. The
reaction mixture included 3,000 ng of the pCR2.1 plasmid, 1X CutSmart Buffer (New England
Biolabs), and the restriction enzymes EcoRI and NotI, totaling 30 µL. This mixture was
incubated at 37 C for 60 minutes. The backbone fragment of interest from the digested pCR2.1
was purified using the E.Z.N.A. Gel Extraction Kit (Omega Bio-tek).
To prepare the oligonucleotide duplex, 50 pmoles of each oligo, oSDR2744 and oSDR2745 (see
Table 5.2), were combined from 5 µL of 10 µM solutions of each oligo. The mixture was
supplemented with 0.5 µL of 1 M KCl and subjected to a heat denaturation step at 65 C for 5
minutes. Following heating, the mixture was allowed to cool gradually to room temperature
for 20 minutes. The resulting oligonucleotide duplex, with a concentration of 5 µM, was then
diluted 1:100 in water to achieve a final concentration of 50 nM, equivalent to 50 fmol/µL.
To introduce a customized multiple cloning site (MCS) into the prepared backbone vector, a
ligation reaction was conducted. The reaction mixture comprised 1X T4 DNA Ligase Buffer
(New England Biolabs), 1X T4 DNA Ligase, 73 ng of the pCR2.1 backbone fragment, and 50
fmol of the MCS duplex in a total volume of 10 µL. The mixture was incubated at room
temperature for 15 minutes. Subsequently, half of the ligation reaction was added to 50 µL of
DH5α competent E. coli cells that had been thawed on ice. The cells were then incubated on ice
for 30 minutes, heat-shocked at 42 C for 45 seconds, and returned to ice for 2 minutes. The cells
were then mixed with 150 µL of LB medium, plated onto LB-agar plates containing
carbenicillin, and incubated overnight at 37 C.
Following incubation, four colonies were selected for inoculation into LB medium containing
100 µg/mL ampicillin for overnight growth in a shaker at 300 rpm and 37 C. Plasmid DNA was

75

isolated from each bacterial culture using the E.Z.N.A. Plasmid DNA Mini Kit I (Omega Biotek). To verify the presence of the insert, I performed an RE digest using PmeI and AgeI.

Figure 5.3. Introduction of CMK142 Region into the Modified pCR 2.1 Vector for Mutagenesis. Arrows indicate
the directionality of the workflow, starting from the bottom of the figure.

76

To generate the CMK142 insert DNA, PCR amplification was conducted using Q5 High-Fidelity
DNA Polymerase (New England Biolabs). The reaction mixture included 1X Q5 Reaction
Buffer, 200 µM dNTPs, 0.5 µM each of primers oSDR2307 and FUB94 (Table 5.2), 5 ng of
Cyanidioschyzon merolae wild-type genomic DNA as the template, and 1 U of Q5 High-Fidelity
DNA Polymerase, with a final volume of 50 µL. Thermocycling conditions were as follows: an
initial denaturation at 98 C for 30 seconds, followed by 35 cycles of denaturation at 98 C for 5
seconds, annealing at 66 C for 20 seconds, and extension at 72 C for 30 seconds, with a final
extension at 72 C for 2 minutes.
Restriction digests of both the purified insert DNA (200ng) and the modified pCR2.1 vector
(designated PSR1126, 2 µg) were performed with PmeI-HF and AgeI-HF, and purified using the
E.Z.N.A. Gel Extraction Kit (Omega Bio-tek). The K142 insert was ligated into the vector
and confirmed by RE digest using Pme1 and Age1.
5.2.2 Construction of Knockout Plasmid
To generate a knockout plasmid via molecular cloning to knock out the endogenous CMK142T
in an attempt to establish a plasmid shuffling system in C. merolae, I employed a systematic
approach involving sequential restriction digests, gel electrophoresis, ligation, and cloning. This
procedure was executed in two distinct phases (Figures 5.4 and 5.5).

77

Figure 5.4. Construction of PSR1113 Vector with a Sulfadiazine Marker. Arrows indicate the directionality of
the workflow, starting from the bottom of the figure.

78

To obtain a backbone vector for subsequent steps, a restriction digest was carried out, the
reaction mixture comprised 1X CutSmart Buffer (New England Biolabs), 2,000 ng of PSR887,
SphI-HF, in a final volume of 30 µL. The digestion was incubated at 37 C for 1 hour. This was
done to remove the URA5.3 marker in PSR887.
To generate the sulfadiazine marker, PCR amplification was conducted using Q5 High-Fidelity
DNA Polymerase (New England Biolabs) as described in section 5.2.1 using 0.5 µM each of
primers oSDR2743 and oSDR2744 (Table 5.2), and 5 ng of PSR1052 as the template.
Thermocycling conditions were the same as described in 5.2.1, annealing at 65 C for 20 seconds,
and extension at 72 C for 39 seconds. Dephosphorylation of the restriction-digested pSR887
backbone plasmid was performed in a reaction mixture that contained 1X Shrimp Alkaline
Phosphatase Buffer (Promega), 405 ng of linearized pSR887, and 1 U of Shrimp Alkaline
Phosphatase in a total volume of 50 µl that was incubated at 37 C for 15 min followed by
inactivation of the enzyme at 65 C for 15 min. The insert DNA was ligated into the SAP-treated
PSR887 vector, transformed 50 µL of DH5α competent E. coli cells as described in section 5.2.1,
and confirmed by RE digest using BamH1 and Spe1.

79

Figure 5.5. Introducing CMK142 Homology Arms into PSR1113. Arrows indicate the directionality of the
workflow, starting from the bottom of the figure.

80

For the preparation of the LIC vector, 500 ng of vector DNA; PSR1113 (250 fmol) was digested
with either 0.5 µL of PacI in a 10 µL reaction containing 1X CutSmart Buffer at 37 C for 3
hours, or with 0.5 µL of SwaI in a 10 µL reaction containing 1X NEB Buffer 3.1 at 25 C for 3
hours. Following digestion, the reaction was heat-inactivated at 65 C for 20 minutes. Half of the
digested product was treated with T4 DNA polymerase (NEB) for 30 minutes at 22 C in a
mixture containing 0.5 µL of 10X CutSmart Buffer, 0.5 µL of 100 mM DTT, 0.5 µL of 50 mM
dCTP (for PacI) or dGTP (for SwaI), 0.4 µL of T4 DNA polymerase, and 3.1 µL of water. The
enzyme was then inactivated by heating at 75 C for 20 minutes. PCR amplification was
conducted using Q5 High-Fidelity DNA Polymerase (New England Biolabs) to generate the
insert DNA. The reaction mixture included 1X Q5 Reaction Buffer, 200 µM dNTPs, 0.5 µM of
each set of primers oSDR2646/oSDR2638(PacI) and oSDR2647 and oSDR2648 (SwaI) (Table
5.2), 1 ng of C. merolae Wild-type genomic DNA as the template, and 1 U of Q5 High-Fidelity
DNA Polymerase, in a final volume of 50 µL. Thermocycling conditions were the same as
described in section 5.2.1, annealing at 65 C for 20 seconds, and extension at 72 C for 30
seconds (PacI) and 10 seconds (Swa1).
For the LIC PCR preparation, a 250 fmol aliquot of the PCR product was treated with T4 DNA
polymerase under the same conditions as the vector digestion.
For the LIC reaction, 1 µL of the treated vector (25 ng, 10 fmol) was combined with 1 µL of the
treated PCR product (25 fmol) and incubated at room temperature for 5 minutes, transformed 50
µL of DH5α competent E. coli cells as described in section 5.2.1 and confirmed by RE digest
using BamH1 and Spe1.

81

5.2.3 Construction of Transient and Integrable Plasmids
The construction of transient and integrable plasmids was achieved through the generation of
three distinct plasmids, each serving a specific purpose. Two transient plasmids were developed,
each incorporating a different selectable marker - URA5.3 and Chloramphenicol Acetyl
Transferase (CAT) - to facilitate the plasmid shuffling technique. Additionally, one integrable
plasmid was constructed to carry the targeted mutations required for the direct replacement
method. This process was executed in three distinct phases.
For Phase I and II (Figures 5.6 and 5.7 respectively), preparation of the Pac1 LIC vector was
performed as described in Section 5.2.2. PCR amplification was conducted using Q5 HighFidelity DNA Polymerase (New England Biolabs), with 0.5 µM of each primer (oSDR2645 and
oSDR2637; Table 5.2) and 1 ng of Cyanidioschyzon merolae wild-type genomic DNA as the
template, as outlined in Section 5.2.1.
The thermocycling conditions were the same as described in Section 5.2.1, except for the
annealing temperature of 68 C for 20 seconds and an extension step at 72 C for 56 seconds.
The LIC reaction was then performed as outlined in Section 5.2.2. The LIC product was used to
transform DH5α competent E. coli cells as detailed in Section 5.2.1 and confirmed by RE digest
using XbaI for the URA plasmid and BamH1 for the CAT plasmid.

82

Figure 5.6 Introducing CMK142 region into the PacI site of PSR887. Arrows indicate the directionality of the
workflow, starting from the bottom of the figure.

83

Figure 5.7 Introducing CMK142 region into the PacI site of PSR886. Arrows indicate the directionality of the
workflow, starting from the bottom of the figure.

84

Figure 5.8 Introducing a second homology arm into the SwaI site of PSR1119. Arrows indicate the
directionality of the workflow, starting from the bottom of the figure.

Swa1 preparation of the LIC vector (500 ng of vector DNA; PSR1119 250 fmol from phase II)
was performed as described in Section 5.2.2. PCR amplification was conducted using Q5 High85

Fidelity DNA Polymerase (New England Biolabs), with 0.5 µM of each primer (oSDR2647 and
oSDR2648; Table 5.2) and 1 ng of Cyanidioschyzon merolae wild-type genomic DNA as the
template, as outlined in Section 5.2.1. The thermocycling conditions were the same as described
in Section 5.2.1, except for the annealing temperature of 68 C for 20 seconds and an extension
step at 72 C for 56 seconds. The LIC reaction was then performed as outlined in Section 5.2.2.
The LIC product was used to transform DH5α competent E. coli cells as detailed in Section 5.2.1
and confirmed by RE Digest using XbaI for the URA plasmid and BamH1 for CAT.
5.2.4 Making of Mutant Plasmids
For the mutational analysis of the RNase MRP RNA component in Cyanidioschyzon merolae, I
employed the "Around-the-Horn" site-directed mutagenesis technique. The primers for each set
of mutations were phosphorylated using T4 polynucleotide kinase (PNK) in a reaction mix
containing 10X Kinase buffer, MgSO₄, ATP, and PNK, followed by incubation at 37 C for 45
minutes. The reaction mixture contained 1X Q5 Reaction Buffer, 200 µM dNTPs, 0.5 µM of
each set of primers oSDR2649/oSDR2650, oSDR2651/oSDR2652 and oSDR2653/oSDR2654
(Table 5.2) for ∆343 – 356, ∆372 - 405 and G162A respectively, 5 ng of PSR1127 as the
template, and 1 U of Q5 High-Fidelity DNA Polymerase, in a final volume of 50 µL.
Thermocycling conditions were as follows: an initial denaturation at 98 C for 30 seconds,
followed by 35 cycles of denaturation at 98 C for 5 seconds, annealing at 63 C for 20 seconds,
and extension at 72 C for 2.17 minutes, with a final extension at 72 C for 2 minutes. The PCR
product was treated with DpnI to digest the template plasmid, and the desired PCR product was
purified with the E.Z.N.A. Cycle Pure Kit (Omega Bio-tek). A ligation reaction was conducted.
The reaction mixture comprised 1X T4 DNA Ligase Buffer (New England Biolabs), 1X T4
DNA Ligase (Blunt), and 91 ng of DpnI-treated PCR Product in a total volume of 10 µL. The
86

mixture was incubated at room temperature for 15 minutes, transformed 50 µL of DH5α
competent E. coli cells, and inoculated on an LB medium as described in section 5.2.1.
Whether or not my plasmid contained the mutations was initially determined by performing a
colony PCR on both the wild-type (PSR1127) and the mutants, assessing whether the size of the
PCR products corresponded to the expected mutant size or the size of the unedited PSR1127
plasmid. PCR was performed using Taq DNA Polymerase with Standard Taq Buffer (New
England Biolabs) and a DNA Engine Dyad Peltier Thermal Cycler (Bio-Rad Laboratories). Each
reaction mixture contained 1X Standard Taq Reaction Buffer, 200 µM dNTPs, 0.5 µM of each
primer (FuB 146 and FuB147 in Table 5.2), 2ng of template DNA (either wild-type PSR1127 or
mutants), 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 at 58 C for 20 sec, and extension at 68 C for 1 min, followed by a final extension
at 68 C for 5 min. The PCR products were run alongside a 100 bp DNA Ladder (New England
Biolabs) on a 2% agarose gel containing ethidium bromide at 150 V for 60 minutes and
visualized using a Gel Doc imaging system. After initial confirmation of successful mutations,
samples were sent to sequence at the Eurofins SimpleSeq Facility using the primer oSDR1445
(Table 5.2).

87

5.2.5 Inserting Mutated Regions from PSR1127 into Integrable Plasmids
Following sequencing and validation of the mutations in PSR1127, the resulting mutated region
of interest was excised and inserted into integrable plasmids for transformation into C. merolae.
Each restriction digest reaction mixture contained 1X CutSmart Buffer (New England Biolabs),
3,000 ng of plasmid DNA (either mutated PSR1127/PSR1127b or PSR1124), and PmeI-HF and
AgeI-HF in a total volume of 30 µl. The reactions were incubated at 37 C for 60 minutes. After
digestion, the 7,239-bp backbone from PSR1124 and the respective insert from the digested
mutant plasmids (636-bp from PSR1127 and 669-bp from PSR1127b) were purified using the
E.Z.N.A. Gel Extraction Kit (Omega Bio-tek). Ligation reaction and transformation of DH5α
competent E. coli cells was performed as described in section 5.2.1 using 80 ng of vector DNA,
and 68 ng of the inserts in a total volume of 10 µL. BamHI RE digest was used to verify
plasmids (Figures 5.9 and 5.10).

88

∆372 - 405

Figure 5.9 Introducing a ∆372 - 405 mutant into PSR1124 to generate PSR1128. Arrows indicate the
directionality of the workflow, starting from the bottom of the figure.

89

G162A

Figure 5.10 Introducing a G162A mutant into PSR1124 to generate PSR1130. Arrows indicate the directionality
of the workflow, starting from the bottom of the figure.

To generate the original linear DNA vector for the transformation of Cyanidioschyzon merolae,
PCR was carried out using Q5 High-Fidelity DNA Polymerase (New England Biolabs) and a
DNA Engine Dyad Peltier Thermal Cycler (Bio-Rad Laboratories). Each PCR mixture contained
1X Q5 Reaction Buffer, 200 µM dNTPs, 0.5 µM of each primer (oSDR2647 and oSDR2645), 5
ng of plasmid template DNA (pSR1128; figure 5.11 or pSR1130; figure 5.12), and 1 U of Q5
High-Fidelity DNA Polymerase in a total volume of 50 µL.

90

The thermocycling conditions were as follows: initial denaturation at 98 C for 30 seconds,
followed by 35 cycles of denaturation at 98 C for 5 seconds, annealing at 72 C for 20 seconds,
and extension at 72 C for 1 minute and 45 seconds. A final extension step was performed at 72 C
for 2 minutes. A portion of the PCR product was run alongside a 1 kb DNA Ladder (New
England Biolabs) on a 0.7% agarose gel containing ethidium bromide and visualized using a Gel
Doc imaging system. The remaining PCR products were purified using the E.Z.N.A. Cycle Pure
Kit (Omega Bio-tek), and concentrations were measured using a NanoDrop spectrophotometer.

Figure 5.11 Linear transformation product amplified from PSR1128.

Figure 5.12 Linear transformation product amplified from PSR1130.

91

Table 5.2. Oligonucleotides used in the construction and sequencing of Plasmid Shuffling or Direct
Replacement vectors. Primers oSDR2637, oSDR2638, oSDR2307, oSDR1445, FUB94, Fub146 and FUB147 were
designed by Dr. Martha Stark.

Oligonucleotides

Directionality

Sequence (5′ to 3′)

oSDR2643

Forward

GCATGC CGACGAGAACGTATAAGGAGTG

oSDR2644

Reverse

GCATGC ACACTTTTTGCCTGCACAAGTT

oSDR2645

Reverse

ATG TTA AGT GGA TTA C
GCAGGTTTTGAACGAGGTGG

oSDR2646

Forward

AGTTGAAGTATGTTACTCGTCTGTGTTTCTCTCGTGG

oSDR2647

Forward

CTTATCTCAATATTTGCAGGAGAAGACACCGCTACG

oSDR2648

Reverse

AACAATCACCAATTTGGTCATGGTGTGGTGAAACGC

oSDR2649

Forward

TCAGCAATGGAACGAGCG

oSDR2650

Reverse

CCGCTGCCAAACATGG

oSDR2651

Forward

GTGAACACAATGGGGCTTAC

oSDR2652

Reverse

GCTCGTTCCATTGCTGATTC

oSDR2653

Forward

AAGTGTCGGTGGTAAGCAAAG

oSDR2654

Reverse

TCTGACCTACTGTCCGG

oSDR2637

Forward

AGTTGA AGT ATGTTACACTAGAGACGCTTCCGTGAC

oSDR2638

Reverse

ATGTTAAGTGGATTACACCCGATTACCTTGCGTCA

oSDR2744

Forward

AATTCGTTTAAACGATATCACCGGTGC

oSDR2745

Reverse

GGCCGCACCGGTGATATCGTTTAAACG

oSDR2307

Forward

TCGGACGTGGTTAGTTGACG

oSDR1445

Reverse

GTAATACGACTCACTATAGGGCG

FUB94

Reverse

GCGATCCTGAATCTGGTCAA

FUB146

Forward

AATAAGAGCGGGAGTGCTG

FUB 147

Reverse

CCAGAGTAAGCCCCATTGTG

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5.2.6 Transformation of C. merolae
The protocols for cultivating and transforming Cyanidioschyzon merolae were adapted from
Kobayashi et al. (2010). The standard growth conditions for C. merolae included using either
liquid MA2G media [40 mM (NH₄)₂SO₄, 8 mM KH₂PO₄, 4 mM MgSO₄, 1 mM CaCl₂, 184 µM
H₃BO₃, 100 µM FeCl₃, 80 µM Na₂EDTA, 36 µM MnCl₂, 6.4 µM Na₂MoO₄, 3.08 µM ZnCl₂, 1.2
µM CuCl₂, 0.68 µM CoCl₂, 50 mM glycerol] or solid 0.75X MA2G media [with all components
reduced by 25% except for glycerol, and 4.62 mg/mL Gelzan (Caisson Labs) as a gelling agent]
for plating. Cultures were incubated at 42C with 2% CO₂ under continuous illumination at 90
µmol photons·m⁻²·s⁻¹.
Three days before transformation, wild-type C. merolae cells were diluted in MA2G media to
achieve an actively dividing culture with an OD₇₅₀ < 3.0 by the day before transformation. On the
day before transformation, the cells were further diluted to an OD₇₅₀ = 0.25-0.40, allowing the
culture to reach a target OD₇₅₀ = 0.8-1.0 within one day. In preparation, two plates containing
solid 0.75X MA2G media, supplemented with 250 µg/mL chloramphenicol, were poured and left
to dry overnight at room temperature, then stored upside down in a plastic sleeve at 4 C.
On the day of transformation, 40 mL of cells were centrifuged at 2,000 × g for 10 minutes,
washed in 1 mL of warm MA-I buffer [20 mM (NH₄)₂SO₄, 2 mM MgSO₄, 92 µM H₃BO₃, 18 µM
MnCl₂, 3.2 µM Na₂MoO₄, 1.54 µM ZnCl₂, 0.6 µM CuCl₂, 0.34 µM CoCl₂], centrifuged at 2,000
× g for 12 seconds, and resuspended in a total volume of 200 µL of warm MA-I buffer. Each 25µL aliquot of this cell suspension contained approximately 5.00-6.25 × 10⁶ cells. For each
transformation, 25 µL of cells were mixed with 100 µL of MA-I buffer containing the nucleic
acids to be delivered and 60 µg of sonicated salmon sperm DNA, followed by the addition of
125 µL of 60% w/v PEG 4000 in MA-I buffer. After the immediate addition of 1 mL of warm
93

MA2G, the entire transformation reaction was diluted to a final volume of 50 mL of warm
MA2G in a graduated glass cylinder.
Three transformations were performed: the first served as a negative control with no DNA, the
second contained 5 µg of PSR1128, and the third included 5 µg of PSR1130. One day posttransformation, plates were spotted with 15-µL aliquots of 20% v/v cornstarch in MA2GC
(MA2G conditioned media). The cells were centrifuged at 2,000 × g for 10 minutes, resuspended
in 300 µL of MA2GC, subjected to a series of serial dilutions, and 10-µL aliquots of cells were
dispensed onto each cornstarch spot. The plates were incubated under standard conditions until
colonies formed.
For cells transformed with the sulfadiazine (Sd) resistance marker both transient and integrated,
the selection process began one day post-transformation by culturing the cells in liquid MA2G
media containing 5 µg/mL and 7.5 µg/mL Sd. Following a 10-day incubation, when a noticeable
difference between the transformed cells and the control was observed, the Sd concentration was
increased to 10 µg/mL to enhance selection pressure. Four to six days after this increase, a clear
distinction between the control and transformed cells was evident. Subsequently, the transformed
cells were plated on 0.75X MA2G agar plates and incubated under standard growth conditions
until colonies formed.
For CAT transient transformations, to maintain the transformed plasmid under constant selection
pressure, I initiated the process 24 hours post-transformation by centrifuging the cells at 2,000 ×
g for 10 minutes. The cell pellet was resuspended in 1 mL of MA2G media supplemented with
150 µg/mL chloramphenicol (Cp). The resuspended cells were transferred to a 6-well plate
containing 6 mL of the same media and incubated under standard conditions. To maintain
selective pressure, 150 µg/mL chloramphenicol was added directly to the cultures every three
94

days, without media replacement. By day 6, both the control and experimental cultures appeared
yellow-green, indicating stress; however, by day 9, the experimental culture had transitioned to a
dark green color, while the control culture remained yellow-green.
5.2.7 Analysis of Colonies
Colonies were picked and inoculated into 16 µL of MA2G medium in a 96-well plate for highresolution screening by PCR using Taq polymerase. The colony PCR was performed using Taq
DNA Polymerase with Standard Taq Buffer (New England Biolabs) and a DNA Engine Dyad
Peltier Thermal Cycler (Bio-Rad Laboratories). Each reaction mixture contained 1X Standard
Taq Reaction Buffer, 200 µM dNTPs, 0.5 µM of each primer (oSDR2607/ oSDR2637 and
oSDR2259/ oSDR2645 in Table 5.3), 1 µl of template DNA (either wild-type 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 at 56 C for 20 sec, and extension at 68 C for 1 min and 31 sec, followed
by a final extension at 68 C for 5 min. The PCR products were run alongside a 100 bp DNA
Ladder (New England Biolabs) on a 0.7% agarose gel containing ethidium bromide at 150 V for
1 hour and visualized using a Gel Doc imaging system. The remainder of the resuspended
colonies were transferred to 200 µL of MA2G medium supplemented with chloramphenicol. To
minimize evaporation, 100 µL of water was added between the wells surrounding the cell
suspensions, and the plate was wrapped in grafting tape. The cultures were incubated for
approximately three days until small green cell clumps formed at the bottom of the wells.
Positive cultures were subsequently transferred to 2 mL of MA2G medium with chloramphenicol
in a 24-well plate and grown for an additional four days. Once the cultures appeared green,
genomic DNA was isolated from 0.25 mL of the culture using Quick Edward's Genomic DNA
95

preparation method (1-2 OD units of cells were harvested by centrifugation. The cell pellet was
resuspended in 200 µL of Edwards Buffer and vortexed for 5 seconds; a reduced volume of 100
µL was used for smaller pellets. Isopropanol (200 µL) was then added, mixed by inversion, and
centrifuged at 13,000 x g for 5 minutes. The supernatant was decanted, and the pellet was airdried for less than 2 minutes by inverting the tube on a paper towel. The DNA pellet was
resuspended in 100 µL of water, or 50 µL for smaller pellets, using a pipette. Insoluble material
was removed by centrifugation at 13,000 x g for 1 minute. The DNA solution was diluted 1:10,
and 1 µL was used for PCR; performed as described earlier, with a wild-type control included).
Table 5.3. Primers Used to Verify Successful Genomic Integration.
Oligonucleotides

Directionality

Sequence (5′ to 3′)

oSDR1831

Reverse

AACCTAACTCTCCGTCGCTA

oSDR2259

Forward

AGTTGAAGTATGTTACAAGCTCACAGGCAAAACAG

oSDR2263

Forward

TTCTAAATCGCTTCCAGCCG

oSDR2264

Reverse

GTTTTTGCCTGGTCCCTACA

OSDR2607

Forward

GTTCGGCATCCTGAACCTGA

5.2.8 Northern Blot Analysis
Northern blot analysis was performed as described earlier in section 2.1.2 to 2.1.4.12.

96

5.3. Results
5.3.1 Plasmid Construction
To investigate the function of RNase MRP in 5.8S rRNA processing, I conducted mutational
analysis on its RNA component. The aim was to assess whether mutations that disrupt the
stoichiometry of the two forms of 5.8S rRNA in S. cerevisiae also do so in C. merolae, thereby
confirming its conserved role in 5.8S rRNA biogenesis. For these experiments, I generated the
plasmids described in the materials and method section with their respective diagram of the
cloning steps. I initiated the construction of plasmids for mutagenesis by inserting an
oligonucleotide duplex, which harbored two multiple cloning sites (MCS), into the PCR 2.1
backbone. The pCR 2.1 vector was initially linearized by digestion with EcoRI and NotI (Figure
5.13) to create compatible ends for the MCS insertion, facilitating downstream cloning steps.

97

B

A

pSR1126 + Pme1 + Age1
1

2

3

4

10.0 kbp
10.0 kbp
3.0 kbp
3.0 kbp

0.5 kbp

0.5 kbp

Figure 5.13. Restriction digest analysis of pCR2.1 and pSR1126. A 0.7% agarose gel stained with ethidium
bromide displaying the results of restriction digests: (a) pCR2.1 digested with EcoRI and NotI, producing the
expected 3,886 bp backbone fragment along with a ~500 bp fragment that was subsequently discarded; and (b)
pSR1126 digested with PmeI and AgeI, resulting in the desired 3,913 bp product. The size markers used were the 1
kb DNA ladder (Ladder 1) and the 100 bp DNA ladder (Ladder 2) for reference.

The DNA insert was derived from CMK142 and ligated into the PmeI and AgeI multiple cloning
sites of pSR1126, resulting in the construction of pSR1127. The successful integration of the
insert was confirmed by restriction digest analysis of pSR1127 using PmeI and AgeI (Figure
5.14).
98

B
A

10.0 kbp

1.5 kbp

3.0 kbp

*
0.5 kbp

0.5 kbp
0.1 kbp

C

pSR1127 + Pme1 + Age1
1

2

3

4

5

6

10.0 kbp
3.0 kbp

0.5 kbp

Figure 5.14. Restriction Digest Analysis of Insert DNA and pSR1127. A 0.7% agarose gel stained with ethidium
bromide displaying (a) Insert DNA of expected size of 990 bp and the results of restriction digests: (b) Insert DNA
digested with PmeI and AgeI, producing the expected 670 bp fragment denoted with * (c) pSR1127 digested with
PmeI and AgeI with 1,2,3,4,5,6 displaying the desired 3,913 bp and 670 bp products. The size markers used were
the 1 kb DNA ladder or the 100 bp DNA ladder for reference.

99

To construct a plasmid containing the sulfadiazine resistance marker, pSR887, which originally
carried a URA5.3 marker, was digested with SphI and XhoI to excise the URA5.3 marker,
leaving the backbone for subsequent cloning. The sulfadiazine marker insert was generated from
pSR1052, and ligation was then performed to join the backbone, and the sulfadiazine marker
insert; pSR1113. The successful integration of the insert was confirmed by restriction digest
analysis of pSR1113 using two sets of restriction enzymes (BamH1/XhoI and BamH1/Spe1) to
also confirm insert directionality (Figure 5.15).

C
A

B

10.0 kbp
3.0 kbp

10.0 kbp
1.5 kbp
3.0 kbp

*

0.5 kbp

0.5 kbp

Insert DNA
Plasmid Backbone
Figure 5.15. Construction and restriction digest analysis of pSR1113. A 0.7% agarose gel stained with ethidium
bromide displaying (a) Insert DNA of expected size of 1, 913 bp and the results of restriction digests: (b) pSR887
digested with SphI and XhoI, producing the expected 2,978 bp backbone fragment (c) pSR1113 digested with
BamHI and XhoI displaying the expected 4,590 bp and 302 bp bands and BamH1 and Spe1 displaying the desired
2,867 bp and 2,025 bp products. The size markers used were the 1 kb DNA ladder or the 100 bp DNA ladder for
reference.

100

The knockout plasmid, pSR1117, designed for the disruption of the endogenous CMK142 gene
to establish a plasmid shuffling system in Cyanidioschyzon merolae, was constructed through
ligation-independent cloning (LIC). DNA fragments derived from CMK142 were inserted into
the SwaI and PacI sites of the pSR1113 vector. The insertion into the SwaI site was validated by
restriction digest analysis using BamHI and XhoI (Figure 5.16).

pSR1117A + BamH1 + Xho1

B

1

2

3

4

5

A

1.5 kbp

10.0 kbp

0.5 kbp

3.0 kbp

0.1 kbp
0.5 kbp
Swa1 Insert DNA
pSR1113 + Swa1 Insert = pSR1117A

Figure 5.16. Insertion of CMK142-derived DNA fragment into the SwaI site of the pSR1113 vector. A 0.7%
agarose gel stained with ethidium bromide displaying (a) Insert DNA of expected size of 505 bp and (b) The insert
cloned into the Swa1 site of pSR1113 and digested with BamHI and XhoI, producing the expected 4,590 bp and 755
bp fragments. The size markers used were the 1 kb DNA ladder and the 100 bp DNA ladder for reference.

101

The plasmid pSR1117A, containing the validated SwaI insert, was subsequently used as a
template for the insertion of the 3’ homology arm derived from the CMK142 gene into the PacI
site, resulting in the construction of the final knockout plasmid, pSR1117. Successful integration
of the fragment at the PacI site was confirmed by restriction digest analysis using BamHI and
XhoI (Figure 5.17).

pSR1117 + BamH1 + Xho1

B

1

2

3

4

5

6

A
10.0 kbp
3.0 kbp
2.0 kbp
10.0 kbp
3.0 kbp

1.0 kbp

0.5 kbp
0.5 kbp

pSR1117A + Pac1 Insert = pSR1117

Pac1 Insert DNA

Figure 5.17. Construction and Restriction Digest Analysis of pSR1117. A 0.7% agarose gel stained with
ethidium bromide displaying (a) Insert DNA of expected size of 1,037 bp and (b) The insert cloned into the Pac1
site of pSR1117A to generate pSR1117. The final plasmid is digested with BamHI and XhoI, with samples 1 to 6
producing the expected 3,019 bp, 2,010 bp, 745 bp, and 368 bp fragments. The size marker used was the 1 kb DNA
ladder for reference.

102

To generate two shuttle plasmids with distinct selectable markers (URA5.3 and CAT), insert
DNA was generated from CMK142 containing our MRP RNA locus and cloned into the PacI
sites of pSR887 (harboring the URA5.3 selectable marker) and pSR886 (harboring the CAT
selectable marker) Figure 5.18.

pSR1119
+ BamH1

pSR1114 + Xba1

A

B

1

2

3

4

1

2

3

C

10.0 kbp
3.0 kbp

10.0 kbp

10.0 kbp

3.0 kbp

3.0 kbp

0.5 kbp

1.0 kbp
1.0 kbp

Pac1 Insert DNA

pSR1114

pSR1119

Figure 5.18. Construction and Restriction Digest of two shuffle plasmids with distinct selectable markers. A
0.7% agarose gel stained with ethidium bromide displaying (a) Insert DNA of expected size of 2,776 bp that was
cloned into Pac1 sites of pSR887 and pSR886 via ligation independent cloning to generate pSR1114 and pSR1119
respectively (b) Restriction digest of pSR1114 with XbaI, producing the expected 7,067 bp and 1,472 bp fragments
and (c) Restriction digest of pSR1119 with BamHI displaying the expected 4,966 bp and 2,427 bp fragments. The
size marker used was the 1 kb DNA ladder for reference.

To generate an integrable CAT amplicon capable of incorporating mutant MRP and replacing the
endogenous MRP at the CMK142 locus, I constructed a CAT integrable plasmid. This was
achieved by inserting a 5' homology arm into the SwaI site of pSR1119 using ligationindependent cloning (Figure 5.19).
103

pSR1124 + BamH1

B

A

1

2

3

4

5

6

10.0 kbp
1.5 kbp

3.0 kbp

0.5 kbp
0.5 kbp

0.1 kbp

pSR1119 + Swa1 Insert = pSR1124

Swa1 Insert DNA

Figure 5.19. Construction and Restriction Digest of pSR1124. A 0.7% agarose gel stained with ethidium bromide
displaying (a) Insert DNA of expected size of 505 bp that was cloned into Swa1 sites of pSR1119 to generate
pSR1124 (b) Restriction digest of pSR1124 with BamHI, displaying the expected 4,966 bp and 2,932 bp fragments.
The size markers used were the 1 kb DNA ladder and the 100 bp DNA ladder for reference.

To generate mutant plasmids using "Around-the-Horn" site-directed mutagenesis, as described in
Materials and Methods section 5.1.4, phosphorylated primers were employed in a PCR reaction
using pSR1127 as the template to introduce the desired mutations. Figures 5.20 and 5.21 display
the mutation sites on the predicted RNase MRP RNA secondary structure in Cyanidioschyzon
merolae and the corresponding PCR results from the mutagenesis, respectively.

104

Figure 5.20: Predicted secondary structure of RNase MRP RNA in Cyanidioschyzon merolae with mutation sites
highlighted. The mutation sites are marked in green, with the corresponding equivalent sites in S. cerevisiae (Sc)
indicated.

105

10.0 kbp

3.0 kbp

0.5 kbp

Figure 5.21: PCR results from site-directed mutagenesis of RNase MRP RNA in Cyanidioschyzon merolae. The
resulting amplicons are 4,558 bp for the ∆343–356 deletion, 4,538 bp for the ∆372–405 deletion, and 4,572 bp for
the G162A point mutation. The 1 kb DNA ladder is used as a size marker.

The mutagenized PCR products were digested with DpnI and ligated, as detailed in the Materials
and Methods section. To preliminarily verify successful mutagenesis before sequencing, PCR
amplification was performed. This allowed for an initial assessment of differences in product
size between the mutagenized plasmids, using the non-mutagenized plasmid as a control. The
results are shown in Figure 5.22 with ∆372–405 deletions (1,2,3,4,5) yielding the expected 157
bp band.
106

.

∆343-356_D2
1

2

3 4

5

∆372-405_P19
1

2

3 4

5

1.5 kbp

0.5 kbp

0.1 kbp

Figure 5.22: PCR analysis of mutagenized plasmids compared to a non-mutagenized control, displayed on a
2% agarose gel stained with ethidium bromide. The control plasmid produced the expected 191 bp band, while
the ∆372–405 deletions (1,2,3,4,5) yielded the expected 157 bp band. Unexpected variable bands of 191 bp (2,5) and
~250 bp (3,4) were observed for the ∆343–356 deletion, instead of the anticipated 177 bp band. A 100 bp DNA
ladder was used as a size reference .

Samples of the ∆372–405 deletion and the G162A point mutants, were sent for sequencing.
Repeated attempts to achieve successful mutagenesis of the ∆343–356 deletion were
unsuccessful.
Following sequencing confirmation of successful mutagenesis, the plasmids were digested, and
the mutated regions were excised (Figure 5.23), as described in the Materials and Methods
section. The corresponding region from pSR1124 was also excised and replaced with the
mutated fragments through ligation, resulting in the construction of pSR1128 (∆372–405
107

deletion) and pSR1130 (G162A mutation). The successful integration of the insert was
confirmed by restriction digest analysis using PmeI and AgeI (Figure 5.24).

10.0 kbp

3.0 kbp

0.5 kbp

Figure 5.23 Restriction enzyme digestion and ligation process for constructing pSR1128 and pSR1130. A
0.7% agarose gel stained with ethidium bromide shows; digested pSR1124 yielding a backbone of interest fragment
of 7,231 bp and inserts of interest measuring 636 bp and 669 bp from ∆372–405 deletion and G162A mutation
respectively. The 1 kb DNA ladder is used as a size marker.

108

pSR1128 (∆372-405)

pSR1130 (G162A)

+ BamH1
A

1

2

3

4

5

+ BamH1
B

6

10.0 kbp

10.0 kbp

3.0 kbp

3.0 kbp

0.5 kbp

0.5 kbp

1

2

3

4

5

6

pSR1130

pSR1128

Figure 5.24. Restriction Digest of pSR1128 and pSR1130. A 0.7% agarose gel stained with ethidium bromide
shows (a) Restriction digest of pSR1128 with BamHI, displaying the expected 4,932 bp and 2,932 bp fragments. (b)
Restriction digest of pSR1130 with BamHI, displaying the expected 4,954 bp and 2,943 bp fragments. The 1 kb
DNA ladder is used as a size marker.

After confirmation of the correct mutant constructs with the expected sizes for each mutation
(pSR1128 displaying the expected 4,932 bp, and 2,932 bp fragments and pSR1130 the expected
4,954 bp and 2,943 bp fragments). Mutants were transformed into C. merolae.

109

A

B

10.0 kbp

10.0 kbp

3.0 kbp

3.0 kbp

0.5 kbp

0.5 kbp
pSR1130

pSR1128

Figure 5.25: Amplification of linear DNA from plasmids used for the transformation of Cyanidioschyzon
merolae. A 0.7% agarose gel stained with ethidium bromide displays (a) the 4,986 bp PCR product from pSR1128
used for the transformation of C. merolae, and (b) the 5,018 bp PCR product from pSR1130 used for C. merolae
transformation. A 1 kb DNA ladder was used as a size reference.

After 12 days post-transformation, multiple chloramphenicol-resistant colonies were observed.
Twelve colonies picked from the pSR1128 (∆372 – 405) transformants were screened for
homologous recombination using colony PCR (Figure 5.26) with each transformant tested with
two different sets of primers making a total of 24 screens. The colony PCR analysis identified
several promising candidates, two of which were further analyzed by extracting genomic DNA
and subjecting them to additional PCR tests.
110

∆372 – 405 (pSR1128)
1

2

3

4

5 6

7

8

9

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24

10.0 kbp
3.0 kbp

0.5 kbp

Figure 5.26 Initial colony PCR screening for genomic integration of pSR1128 (∆372–405). A 0.7% agarose gel
stained with ethidium bromide shows colony PCR results for 24 C. merolae colonies, tested with two primer sets.
The expected bands of 1,318 bp and 995 bp, indicating successful genomic insertion, are displayed. A 1 kb DNA
ladder was used as a size marker for reference.

This subsequent test was to confirm successful recombination and integration into the CMK142
locus and wild-type C. merolae genomic DNA was used as a control (Figure 5.27). The
successful transformants' clear resistance to chloramphenicol confirmed the functionality of the
APCC promoter, demonstrating its ability to drive the expression of the CAT gene.

111

∆372 – 405
(pSR1128)
1

2

10.0 kbp

3.0 kbp

0.5 kbp

Figure 5.27. Final PCR confirming the successful genomic integration of pSR1128 (∆372–405) into CMK142
locus. A 0.7% agarose gel containing ethidium bromide showing the PCR results for two promising ∆372–405 C.
merolae candidates (1 and 2) showing the expected 5,609 bp band for successful integration while the wild-type
(WT) sample acting as a negative control displays the expected 3,738 bp band. A 1 kb DNA ladder was used as a
size reference.

Similarly, for pSR1130 (G162A), multiple chloramphenicol-resistant colonies were observed
after 12 days post-transformation. Seven colonies picked from the pSR1130 (G162A)
transformants were screened for homologous recombination using colony PCR (Figure 5.28)
with each transformant tested with two different sets of primers making a total of 14 screens. The
colony PCR analysis identified several promising candidates, three of which were further
analyzed by extracting genomic DNA and subjecting them to additional PCR tests.

112

G162A (pSR1130)
1

2

3

4

5

6

7

8

9

10

11

12

13

14

10.0 kbp
3.0 kbp

0.5 kbp

Figure 5.28 Initial colony PCR screening for genomic integration of pSR1130 (G162A). A 0.7% agarose gel
stained with ethidium bromide shows colony PCR results for 14 C. merolae colonies, tested with two primer sets.
The expected bands of 1,318 bp and 995 bp, indicating successful genomic insertion are displayed. A 1 kb DNA
ladder was used as a size marker for reference.

Although some bands corresponding to nonspecific amplification were present, the expected
bands were still observed, indicating successful recombination and integration into the CMK142
locus, with wild-type C. merolae genomic DNA used as a control (Figure 5.29). The clear
chloramphenicol resistance observed in the successful transformants further confirmed the
functionality of the APCC promoter, demonstrating its ability to drive CAT gene expression.

113

G162A
(pSR1130)
1

2

3

10.0 kbp

3.0 kbp

0.5 kbp

Figure 5.29. Final PCR confirming the successful genomic integration of pSR1130 (G162A) into CMK142
locus. A 0.7% agarose gel containing ethidium bromide showing the PCR results for three promising G162A C.
merolae candidates (1, 2, and 3) showing the expected 5,641 bp band for successful integration while the wild-type
(WT) sample acting as a negative control displays the expected 3,738 bp band. A 1 kb DNA ladder was used as a
size reference.

5.3.2 Northern Blot Analysis
Following successful recombination and integration of the mutant RNase MRP RNA, I
conducted northern blot analysis to evaluate potential alterations in the stoichiometry of the two
forms of 5.8S rRNA. Figure 5.30 presents the results of this analysis.

114

A

Intron
RNase MRP

1.5% Formaldehyde Agarose
∆372 – 405 (P19)

Wild Type Cm
42C

B

42C

C

57C

57C

5.8SrRNA

5.8SrRNA
5SrRNA

5SrRNA
8% 7M Urea Denaturing PAGE

8% 7M Urea Denaturing PAGE

G162A (P8)

∆372 – 405 (P19)
42C

42C

57C

57C

E

D

5.8SrRNA
5.8SrRNA

5SrRNA
5SrRNA
Figure 5.30 Changes in the stoichiometry of the two forms of 5.8S rRNA following RNase MRP RNA
mutations in Cyanidioschyzon merolae. (a) Total RNA from the ∆372–405 and G162A mutant strains was
analyzed for RNase MRP expression using a 1.5% formaldehyde agarose gel (b) Total RNA from wild-type cells
analyzed on an 8% PAGE gel, stained with ethidium bromide to visualize the two forms of 5.8S rRNA. (c-d) Total
RNA from the ∆372–405 mutant strain and (e) from the G162A mutant strain were analyzed using 8% PAGE gels
and ethidium bromide staining to observe the 5.8S rRNA forms. The probes used were oSDR2487 = 5.8S and
oSDR2479 = RNase MRP.

115

The northern blot analysis results reveal that deletion of the P19 region (∆372–405) in C.
merolae's RNase MRP RNA leads to a notable shift in the stoichiometry of the two forms of
5.8S rRNA (Figures 5.30 c and d). The shift observed between Figure 2.7 and Figures 30c and
30d highlights a change in the relative abundance of the two forms of 5.8S rRNA after mutating
RNase MRP. In Figure 2.7, the short form of 5.8S rRNA is predominant, with the long form
being less visible. However, following the mutation of RNase MRP, the two forms appear in
equal amounts, as shown in Figures 30c and 30d. This suggests that the mutation affects the
processing of 5.8S rRNA, leading to altered stoichiometry between the long and short forms.
However, the same change in stoichiometry between the long and short forms of 5.8S rRNA is
observed at both 42 C and 57 C (Figures 5.30 c and d), suggesting that temperature does not play
a role in this shift. The equal amounts of both forms after RNase MRP mutation are consistent
across different temperature conditions, indicating that the observed change in stoichiometry is
driven by the mutation rather than by heat stress.
In contrast, the G162A mutant strain exhibited no discernible effect on 5.8S rRNA processing,
indicating that mutating this region does not impair RNase MRP functionality in C. merolae.

116

5.3.3 Establishing Plasmid Shuffling System in C. merolae
Even though the direct replacement via homologous recombination worked successfully (section
5.3.2 and Figures 5.30c and d) I attempted to use the plasmid shuffling system used in S.
cerevisiae to see if that gives the same result in C. merolae and 8as an attempt to establish this
system in C. merolae. The plasmid shuffling system involves transiently introducing a second
copy of the CMK142 gene on a plasmid containing a selectable marker, URA5.3. This step
enables the knockout of the endogenous CMK142 gene through homologous recombination
using a different selectable marker, Sulfadiazine. After successfully disrupting the native gene,
the plasmid-encoded CMK142 copy is transiently replaced or "shuffled out" with a mutant
version of CMK142 carried on another plasmid with a distinct selectable marker, CAT. This
transient process ensures selective pressure throughout each phase, allowing for the replacement
of the wild-type gene with the desired mutant.
To achieve this, I transiently transformed Cyanidioschyzon merolae T1 strains with a plasmid
containing a copy of the CMK142 gene (pSR1114), harboring the URA5.3 selectable marker. The
URA5.3 gene encodes orotidine-5'-phosphate decarboxylase, an essential enzyme in the de novo
pyrimidine (uracil) synthesis pathway. Since the T1 strains are auxotrophic for uracil, they rely
on external uracil for survival, allowing cells transformed with the plasmid to survive in uracildeficient conditions. After confirming the successful transformation, a linear DNA product was
amplified from pSR1117 (Figure 5.31) to facilitate the replacement of the endogenous CMK142
gene via homologous recombination.

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10.0 kbp

3.0 kbp

Figure 5.31 Amplification of linear DNA from pSR1117 used for the transformation of Cyanidioschyzon
merolae. A 0.7% agarose gel stained with ethidium bromide displays the expected 3,329 bp PCR product from
pSR1117 used for the transformation of C. merolae. A 1 kb DNA ladder was used as a size reference.

The integration was first selected using sulfadiazine, ensuring that only cells transformed with
the sulfadiazine-resistant marker survived in the presence of the sulfadiazine (Figure 5.32 and
Figure 5.33).

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Day 1 after recovery

A

Day 10 after recovery
B

nc = negative control

Int = Integrand

Figure 5.32. Selection of transformed cells with sulfadiazine. (a-b) Transformed C. merolae cells were cultured in
the presence of 5 µg/mL and 7.5 µg/mL sulfadiazine from day one post-transformation until day 10. Control groups
were included for reference, demonstrating the selective growth of cells carrying the sulfadiazine-resistant marker.
nc= negative control, and Int = Integrand.

After 10 days of selection, the concentration of sulfadiazine was increased to 10 µg/mL for all
cultures (Figure 5.33). This increase in sulfadiazine concentration heightened the stringency of
the selection process, favoring cells with the successful integration of the selectable marker. As a

119

result, non-transformed cells that persisted at the lower concentrations were effectively
eliminated, enriching the population of transformed cells.

d

Day 4 after increasing S concentration to
10ug/ml

A

B

d

Day 6 after increasing S concentration to
10ug/ml
Figure 5.33 Growth of transformed C. merolae cells under increasing sulfadiazine concentration. Transformed
C. merolae cells were grown in the presence of 10 µg/mL sulfadiazine from day 10 post-transformation, following
initial selection with 5 µg/mL and 7.5 µg/mL. (a and b) display 4 days and 6 days after increasing sulfadiazine
concentration respectively. Control groups were included for reference, demonstrating the selective growth of cells
carrying the sulfadiazine-resistant marker. nc= negative control, and Int = Integrand.

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After day 6 of stringent selection, cells were grown on an MA2G media plate for single colonies
as described in the materials and methods section. Four colonies were screened with two
different sets of primers making a total of 8 screens (Figure 5.34). The colony PCR analysis
identified promising candidates, two of which were further analyzed by extracting genomic DNA
and subjecting them to additional PCR tests.

pSR1117 Colony Screening
1

2

3

4

5

6

7

8

10.0 kbp
3.0 kbp

0.5 kbp

Figure 5.34 Initial colony PCR screening for genomic integration of pSR1117. A 0.7% agarose gel stained with
ethidium bromide shows colony PCR results for 8 C. merolae colonies, tested with two primer sets (1,3,5,7 sets –
oSDR2637 and oSDR2607) and (2,4,6,8 sets - oSDR2259 and oSDR2645). The expected bands of 1,358 bp and 979
bp, indicating successful genomic insertion, are displayed. A 1 kb DNA ladder was used as a size marker for
reference.

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This follow-up experiment aimed to confirm successful recombination and integration into the
CMK142 locus, using wild-type C. merolae genomic DNA as a control (Figure 5.35).

gDNA.
pSR1117
1

2

10.0 kbp

3.0 kbp

0.5 kbp

Figure 5.35. PCR confirming genomic integration of pSR1117. A 0.7% agarose gel containing ethidium bromide
showing the PCR results for two promising CMK142 knockout candidates (1 and 2) showing the expected 2,822 bp
band for wild-type (WT) genomic DNA sample and 3,329 bp band for integrands. A 1 kb DNA ladder was used as a
size reference.

Cells were cultured and transiently transformed with the pSR1128 (∆372–405) strain carrying
the CAT gene. By day 9 post-transformation, a distinct difference between the control and the
transformed cells was observed, and by day 12, the control cells had died while the transformants
appeared dark green (Figure 5.36).

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Int

nc

Day 3 post-transformation
(150ug/ml)

Day 6

(150ug/ml)

Day 9

(150ug/ml)

Day 11

Day 12

Figure 5.36. Chloramphenicol resistance in transformed C. merolae cells. Cells were transiently transformed
with the pSR1128 (∆372–405) strain harboring the CAT gene, with 150 µg/mL of chloramphenicol added every
three days. By day 9 post-transformation, clear differences between control and transformed cells were visible. By
day 12, control cells had died, while transformants exhibited a dark green coloration, confirming chloramphenicol
resistance driven by the APCC promoter. nc = negative control Int = Integrands.

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A final PCR analysis was performed to definitively confirm the knockout of CMK142 and assess
whether the prior transformation with sulfadiazine successfully disrupted CMK142 or if the
plasmid was merely integrated into the genome without knockout. This test ensured that the
targeted homologous recombination event effectively removed the endogenous CMK142 gene,
validating the transformation process.

(pSR1117)
1

2

3

4

10.0 kbp
3.0 kbp

0.5 kbp

Figure 5.37. Final PCR analysis revealed unsuccessful integration of pSR1117 into the CMK142 locus. A 0.7%
agarose gel stained with ethidium bromide shows PCR results for four CMK142 knockout candidates (lanes 1-4),
each displaying an unexpected band above 10,000 bp. The wild-type (WT) sample, serving as a negative control,
exhibits the expected 3,738 bp band. A 1 kb DNA ladder was used as a molecular size reference.

The appearance of an unexpected 10,000 bp band, rather than the expected 4,167 bp, indicates
that the CMK142 gene was not successfully knocked out during the transformation. The ability
of the cells to grow in the presence of sulfadiazine suggests that while the selection marker was
integrated into the genome, it did not disrupt the CMK142 locus as intended. Subsequent
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screenings of additional colonies yielded similar unsuccessful results. These findings suggest
that, although integration occurred elsewhere in the genome, the desired knockout of CMK142
was unsuccessful, necessitating further optimization of the plasmid shuffling technique approach
in C. merolae.
5.4. Discussion
The Northern blot analysis results presented in this chapter revealed that the deletion of the P19
region (Δ372–405) in the RNA component of Cyanidioschyzon merolae RNase MRP results in a
significant alteration in the stoichiometry of the two forms of 5.8S rRNA (Figure 5.30c and d).
This observation underscores the role of RNase MRP in the biogenesis of 5.8S rRNA, aligning
with its established function in Saccharomyces cerevisiae and Drosophila melanogaster. In these
model organisms, mutations in RNase MRP lead to a shift in the ratio between the long and short
forms of 5.8S rRNA (5.8S_L and 5.8S_S), typically maintained in a 10:1 ratio (long: short)
(Schmitt and Clayton 1993; Schneider et al., 2010; Shadel et al., 2000). In S. cerevisiae, RNase
MRP cleaves precursor rRNA at the A3 site, facilitating the generation of the shorter form of
5.8S rRNA (5.8S_S). The shift in stoichiometry observed in the C. merolae Δ372–405 mutant
suggests that this region of RNase MRP RNA is required for similar rRNA processing, pointing
to a conserved functional role of RNase MRP across eukaryotes.
Interestingly, in contrast to the P19 region deletion, the G162A mutant strain did not exhibit any
noticeable effect on 5.8S rRNA processing (Figure 5.30e). This finding indicates that the G162A
mutation does not disrupt RNase MRP functionality in C. merolae, as the stoichiometry of 5.8S
rRNA remains unchanged. This observation mirrors studies in S. cerevisiae, where mutations in
non-essential regions of RNase MRP RNA do not necessarily impair rRNA processing (Shadel
et al., 2000; Figure 5.1 and Table 5.1). The lack of an effect in the G162A mutant suggests that
125

this region is not essential for the RNA-protein interactions or catalytic activity necessary for
5.8S rRNA processing. This also implies that certain regions of RNase MRP RNA are
functionally redundant or dispensable, providing a more nuanced understanding of the structural
and functional dynamics of RNase MRP.
An array of mutations in the RNA component of RNase MRP, each exhibiting distinct
phenotypes, will be instrumental in future studies aimed at identifying the protein constituents of
the MRP complex and elucidating the mechanisms that regulate its cellular localization and
enzymatic function. By dissecting how different mutations impact RNase MRP activity,
researchers can better understand the RNA-protein interactions and structural features required
for the enzyme's role in rRNA processing.
These findings further support the hypothesis that the fundamental role of RNase MRP in 5.8S
rRNA maturation is evolutionarily conserved across eukaryotes. As seen in other organisms, C.
merolae depends on RNase MRP for precise regulation of 5.8S rRNA forms, with specific
regions, such as the P19 region, playing an essential role in this process. The G162A mutation's
lack of impact on 5.8S rRNA processing highlights that not all regions of the RNase MRP RNA
are equally required, adding valuable insights into the structure-function relationship within the
MRP complex.

126

Chapter 6 – General Conclusion and Remarks

127

I successfully clarified the role of RNase MRP in C. merolae, highlighting its conserved function
in the processing of ribosomal RNA (rRNA), particularly in the generation of the two forms of
5.8S rRNA. This work has provided significant insights into the function and adaptation of
RNase MRP in C. merolae. It was confirmed that C. merolae possesses two distinct forms of
5.8S rRNA in a 10:1 ratio (small: large), highlighting the role of RNase MRP in regulating rRNA
stoichiometry, which is essential for ribosome biogenesis. Additionally, the study demonstrated
that heat stress-induced intronic accumulation within the CMK142T gene does not affect RNase
MRP’s catalytic activity or the stoichiometry of 5.8S rRNA forms. This finding indicates that C.
merolae maintains rRNA processing efficiency despite environmental stress.
Also, despite a reduction in pre-rRNA levels under heat stress, the unaltered levels of mature 28S
and 18S rRNAs in C. merolae suggest a sophisticated regulatory mechanism that ensures
ribosome function is maintained even under extreme conditions. One possibility is that mature
rRNAs, once formed, are highly stable and resistant to degradation, allowing their levels to
remain constant even when rDNA transcription is inhibited (Grünberger et al., 2023).
Additionally, C. merolae may possess an efficient rRNA processing machinery that maximizes
the conversion of available pre-rRNA into mature rRNAs, compensating for the reduced
precursor synthesis. Moreover, the organism may regulate rRNA turnover, slowing down the
degradation of mature rRNAs during stress to preserve essential ribosomal components. These
adaptive mechanisms, which may include specialized pathways that protect and stabilize rRNA,
highlight the resilience of C. merolae and its ability to maintain cellular functions in the face of
environmental challenges. This contrasts with other organisms, where a short heat shock inhibits
pre-rRNA transcription and processing into mature rRNAs in mammals (Ghosha and Jacob
1996), heat stress inhibits rDNA transcription in animal cells (Ghosha and Jacob 1996; Coccia et
128

al.,2017), and In Arabidopsis thaliana, where heat stress disturbs nucleolar structure, inhibits
pre-rRNA processing, and provokes imbalanced ribosome profiles leading to undetectable
precursors of 18S, 5.8S, and 25S RNAs (Darriere et al., 2022), underscoring the evolutionary
diversity in stress response strategies.
I also explored the impact of specific mutations on RNase MRP function and the results show
that deletion of the P19 region (∆372–405) in the RNase MRP RNA component causes a shift in
the stoichiometry of the two forms of 5.8S rRNA, confirming RNase MRP's role in this process.
In contrast, the G162A mutation did not affect 5.8S rRNA processing, indicating that this
specific mutation does not interfere with the enzyme's functionality. These findings support the
hypothesis of evolutionary conservation of RNase MRP’s role in rRNA processing, aligning with
what has been observed in other organisms, such as yeast and humans (Piccinelli et al., 2005,
Schmitt and Clayton 1993, Rosenblad et al., 2006; Lopez et al. 2009, Goldfarb et al., 2017).
Furthermore, computational analyses revealed that RNase MRP RNA in C. merolae retains
conserved structural regions similar to those in other eukaryotes, supporting the evolutionary
conservation of its function. The RNase MRP complex in C. merolae comprises a reduced set of
five protein constituents compared to eleven in Saccharomyces cerevisiae, reflecting an
evolutionary adaptation to its unique cellular context.
The significance of this research lies in its contribution to understanding the fundamental role of
RNase MRP in eukaryotic biology. Studying C. merolae, a red alga with a streamlined genome,
provides valuable insights into how core cellular mechanisms function in simpler organisms, free
from the genetic redundancies often seen in more complex systems. The evolutionary
conservation of RNase MRP suggests that findings from this study could have broad biological
implications, extending to more complex organisms, including humans where RNase MRP
129

mutations in humans are linked to genetic disorders such as cartilage-hair hypoplasia (CHH)
(Hirose et al.,2006) a condition characterized by skeletal abnormalities, immunodeficiency, and
increased cancer susceptibility. Understanding the role of RNase MRP in C. merolae again helps
to deepen our knowledge of how similar mutations may affect their function in humans.
Looking forward, this study opens several new avenues for research. Further exploration of
RNase MRP's molecular mechanisms, especially how specific regions of the enzyme interact
with its protein components, will provide a deeper understanding of its function. Comparative
studies between C. merolae and other organisms can help to delineate conserved and divergent
aspects of RNase MRP across species. Also, structural studies using techniques like cryoelectron microscopy could offer high-resolution insights into the structure-function relationship
of RNase MRP in C. merolae, advancing our understanding of this complex. Additionally,
investigating the molecular mechanisms by which C. merolae maintains rRNA processing and
ribosome function under heat stress could uncover novel regulatory pathways and stress response
strategies. By addressing these areas, future research can build on the findings of this thesis to
further elucidate the roles and adaptations of RNase MRP in eukaryotes, contributing to a more
comprehensive understanding of ribosome biogenesis and cellular stress responses.
In conclusion, my research has significantly advanced our understanding of RNase MRP in C.
merolae, reinforcing the enzyme’s complex role in rRNA processing across species. The
evolutionary conservation of RNase MRP underscores its importance in cellular processes and
highlights its potential as a target for therapeutic intervention in diseases associated with rRNA
processing dysfunction.

130

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