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RNA TERMINUS CHEMISTRY POTENTIATES DECAY EVENTS THAT
TARGET HAC1 mRNA DURING THE UNFOLDED PROTEIN RESPONSE.
by
Patrick Cherry
B.A. Hendrix College, 2013
A thesis submitted to the
Graduate School of the
University of Colorado in partial fulfilment
of the requirements for the degree of
Doctor of Philosophy
Molecular Biology Program
2019
ii
This dissertation for the Doctor of Philosophy degree of
Patrick Cherry
has been approved for the
Molecular Biology Program
by
Sandy Martin, Chair
Richard Davis
Jeffrey Kieft
Michael McMurray
James Costello
Jay Hesselberth, Advisor
Date: May 17, 2019
iii
Cherry, Patrick (PhD, Molecular Biology Program)
RNA-terminus chemistry potentiates decay events that target HAC1 mRNA during splicing to
regulate the unfolded protein response.
Thesis directed by Associate Professor Jay R. Hesselberth.
ABSTRACT
RNA repair enzymes catalyze rejoining of an RNA molecule after cleavage of
phosphodiester linkages. RNA repair in budding yeast is catalyzed by two separate enzymes
that process tRNA exons during their splicing and HAC1 mRNA exons during activation of the
unfolded protein response (UPR). The RNA ligase Trl1 joins 2′,3′-cyclic phosphate and 5′-
hydroxyl RNA fragments, creating a new phosphodiester linkage with a 2′-phosphate at the
junction. The 2′-phosphate is subsequently removed by the 2′-phosphotransferase Tpt1, which
catalyzes phosphate transfer to NAD+, producing nicotinamide and a unique ADP-ribose
metabolite. I bypassed the essential functions of TRL1 and TPT1 in budding yeast by expressing
“pre-spliced”/intronless versions of the ten normally intron-containing tRNAs, indicating this
repair pathway does not have additional essential functions. Consistent with previous studies,
expression of intronless tRNAs failed to rescue the growth of cells with deletions in components
of the SEN complex, implying an additional essential role for the splicing endonuclease. Finally,
I optimized a PCR-based method to detect RNA 2′-phosphate modifications and show that they
are present on ligated HAC1 mRNA.
In the unfolded protein response, stress in the endoplasmic reticulum (ER) activates a large
transcriptional program to increase ER folding capacity. During the budding yeast UPR, Ire1
excises an intron from the HAC1 mRNA and the exon products of cleavage are ligated, and the
translated protein induces dozens of stress-response genes. The trl1∆ and tpt1∆ mutants
accumulate tRNA and HAC1 splicing intermediates indicative of specific RNA repair defects. As
iv
expected, failure to induce the unfolded protein response in trl1∆ cells grown with tunicamycin
is lethal owing to their inability to ligate HAC1 after its cleavage by Ire1. In contrast, tpt1∆ mutants
grow in the presence of tunicamycin despite reduced accumulation of spliced HAC1 mRNA. I
show that phosphorylation of two different HAC1 splicing intermediates is required for their
degradation by the 5′→3′ exonuclease Xrn1 to enact opposing effects on the UPR. I also found
that ligated but 2′-phosphorylated HAC1 mRNA is cleaved, yielding a decay intermediate with
both 5′- and 2′-phosphates at its 5′-end that inhibit 5′→3′ decay. These decay events expand
the scope of RNA-based regulation in the budding yeast UPR, and these RNA repair mutants
enable new studies of the role of RNA repair in cellular physiology.
The form and content of this abstract are approved. I recommend its publication.
Approved: Jay R. Hesselberth
v
DEDICATION
I dedicate this dissertation to my parents. Absolutely none of this would be possible
without your unconditional love and encouragement. I dedicate this work to my dad, who shares
with me the same sentiments of hard work and fairness, who supports me always, and who is
full of wise advice for the good times and the bad times. I dedicate this work to Mar, who came
into my life later than most parents, but who accepted instant parenthood and did an outstanding
job. Mar modeled values I now hold close, like equality in opportunity, acceptance, and honesty
(and continues to do so). This is for you.
vi
ACKNOWLEDGEMENTS
Firstly, I must acknowledge and thank my advisor, Jay Hesselberth, for his tireless
encouragement and positivity. Jay’s intense curiosity is contagious. Solely through
inquisitiveness have we discovered some of the most interesting behaviors of RNA. Jay
accepted me into the lab from a cold email I sent him the summer before starting graduate
school, and since then it has been a great fit. I thank Jay for his mentorship in my growth as a
scientist. Thank you, Jay.
I also thank my committee, Sandy Martin, Dick Davis, Michael McMurray, Jeff Kieft, and
Jim Costello. Thank you for taking the time to mentor me on the most perplexing problems in
my project, as well as for the encouragement to reach higher.
Many mentors have guided me to this point from being just a freshman in undergrad
looking for something more challenging than coursework to work on: Joy Sturtevant, my first
science mentor, introduced me to the amazing power of yeast genetics—in Candida albicans no
less—and to the workings of academic labs at the Louisiana State University Health Sciences
Center in New Orleans, LA. Michael Shiloh at University of Texas Southwestern Medical Center
in Dallas, TX thought me perseverance in the face of recalcitrant cloning problems. Andres Caro
at Hendrix College in Conway, AR taught both my Biochemistry I and II courses with Oscar-
worthy acting of “so, you are running from a bear,” as the recurring stimulus for epinephrine
release. Andrew Schurko, also at Hendrix, taught a fantastic genetics class with Nasonia
vitripennis wasps as genetic models and let my science presentations get into the weeds in
Advanced Genetics. Liz Gron, also at Hendrix, trained me in the methods and reasoning of
analytical chemistry, and got in some future grad school and career mentoring between lessons
and labs.
All members of the Hesselberth lab have helped me succeed in graduate school. Laura
White has been my partner-in-science for virtually all endeavors, from “paired science” (not
vii
typically recommended) to maintaining lab equipment to inviting speakers. I co-first-authored
my very first paper with Laura, and in doing so I discovered a special bond that comes from the
mutual appreciation of scientific talent and thought. Additionally, Sally Peach advanced so much
of this project before I took over and was a mentor and teacher of programming to me and many
others. I arrived in the lab my first year to Kerri York, Monica Ransom, and Suzi Brian, all of whom
taught me many yeast, nucleic acid, and protein manipulation techniques, along with some
graduate school survival mentoring. Mandy Richer’s balancing of outdoor excursions with lab
responsibilities inspired me to keep up my other hobbies while remaining dedicated to my thesis
project. Also, Mandy’s ongoing fight for fair treatment against the RNA Mafia is working;
evidence of this hangs outside Jay’s office door to this day. I appreciate Rachel Ancar’s presence
in the lab, her patient listening skills, her fearless approach to trying new things in science, and
her slightly absurd sense of humor. Chris Snowden, Jackie Pierce, Maggie Balas, Alexis
Zukowski are all “next door lab neighbors,” and so received the brunt of my questions about
methods and ideas that Hesselberth lab members were not familiar. For all of you, I am so
thankful.
Sometimes there is an easier way to get to the good, fun science in a project, and cloning-
by-phone can save time and frustration. Strains, samples, and knowledge have been dropped
into my life from some fantastic people, whom I would like to thank now. Roy Parker, Stewart
Shuman, Beate Schwer, Anita Hopper, Eric Phizicky, Jane Jackman, Erich Chapman, Jeff Moore,
Dave Barton, and Volker Thiel.
I am lucky to have so many mentors and PIs looking out for me at the University. Bob
Sclafani, Rytis Prekeris, David Engelke, Lori Sussel, Suja Jagannathan, Chad Pearson, and Tânia
Reis all lent me support and advice. Sabrena Heilman, Caitlin Moloney, and Michele Hwozdyk-
Parsons helped me manage my responsibilities to the Molecular Biology Program and the
viii
Graduate School. Thank you to the Molecular Biology Program, the Department of Biochemistry
and Molecular Genetics, The RNA Bioscience Initiative, and Victor W. & Earleen D. Bolie.
Among the ranks of the RNA Mafia are lab mates Rachel Ancar and Laura White, as well
as Maggie Balas, Ryan Sheridan, Erik Hartwick, Lena Steckelberg, Ben Akiyama, Marissa Rhule,
and Laura Hudish. I am thankful for my friends in Molecular Biology program: Molly Kingsley,
Jenn Rabe, Cassi Estrem, Sara Flubacher, Sara Espinosa, James Till, Ryan Sheridan, as well as
my graduate school cohort outside of the program: Mike Shaffer, Kyle Smith, Nicolle Witte, Tania
Eliseeva, Brooke Sinnen, Joe Bednarek, Alex Barret, Katie Mishall Barret, and Travis Nemkov.
Having friends outside of science is healthy, and even fun. My non-science friends
arguably enrich my life so much more than the science friends on my quest to be an instrument
of appreciation, and I share with them bonds formed in the struggles of climbing, biking,
camping, hiking, gaming, pop-culture-critiquing [drag], and occasionally free-soloing (sorry,
Mom and Dad). So I am thankful for Monty Prekeris, Jake Beck, Nick Goedecke, Jack Rugile,
Mike Gorodinsky, Alex Hart, Alex Imhof, Mark Rohr, Nick Turner, Fred Gravagna, Denise Meyer,
Evan Robertson, Ben Parefsky, Gina Loftus, Mike Slater, Camila Restrepo, and Lena Gerber.
I am so thankful to my peers who believed in me in undergrad: Cheryl Mathis, Joana Ortiz-
Baca, Johnny Tran, Ina Agee, Lauren Irby, Gabe Gonzales, Rima AbiSamra, Sloan Zimmerman,
and Olivia Urbanoitz. I thank Emily Winters for inspiring me to chase my dreams, one skype
session at a time. Thank you to my high school science buddies Justin Shapiro and Matt
McTernan for getting through grad school alongside me; science wouldn’t be as fun without your
A-day-ing around.
Thank you to my parents, for so much love, understanding, and encouragement.
The science and learning have been fantastic, but the memories and connections to my
favorite people in life are what make grind of graduate school feel worthwhile. Thank you all, so
much.
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TABLE OF CONTENTS
CHAPTER
I INTRODUCTION ............................................................................................................... 1
Endoribonucleases impose terminus chemistry ......................................................... 2
Intrinsic RNA cleavage .................................................................................... 5
Self-cleaving ribozymes .................................................................................. 6
Sen2 & Sen34 .................................................................................................. 7
Las1, Grc3, & 35S rRNA processing ............................................................... 9
Stress-induced tRNA cleavage ..................................................................... 11
Ribotoxins in interspecies conflict ................................................................ 12
RNA decay in co-translational mRNA surveillance ....................................... 14
Ire1, the “splicing” endoribonuclease of the unfolded protein response (UPR) ............................................................................................. 17
RNA terminus modification ....................................................................................... 19
Clp1 RNA 5′-kinase ....................................................................................... 20
Grc3 RNA 5′-kinase ...................................................................................... 20
CNP (cyclic nucleotide phosphodiesterase) ................................................. 21
RtcA (3′-terminal phosphate cyclase) ........................................................... 21
RNA ligases combine RNA terminus modification domains with a ligase domain. .. 22
Trl1, the ligase of fungi & plants .................................................................... 22
RtcB, the ligase of prokaryotes & animals .................................................... 23
Fidelity of RNA ligases .................................................................................. 25
RNA repair in cellular physiology .............................................................................. 26
The splicing of tRNAs .................................................................................... 26
The unfolded protein response (UPR) ........................................................... 28
x
RNA repair in bacterial stress responses and interspecies conflict .............. 35
Terminus chemistry and exonucleolytic RNA decay ................................................ 37
5′→3′ RNA decay enzymes require substrates be 5′-phosphorylated ........ 38
3′→5′ RNA degradation ................................................................................ 42
Kinase-Mediated Decay ............................................................................................ 44
KMD of tRNA introns ..................................................................................... 44
KMD of rRNA processing intermediates ....................................................... 46
KMD of 3′-fragments of no-go mRNA decay (NGD) ..................................... 47
KMD of T4 bacteriophage mRNA .................................................................. 47
Toward mutants of RNA repair ................................................................................. 48
II GENETIC BYPASS OF ESSENTIAL RNA REPAIR ENZYMES IN BUDDING YEAST ...... 50
Abstract ..................................................................................................................... 50
Introduction ............................................................................................................... 51
Materials & Methods ................................................................................................. 53
General Methods ........................................................................................... 53
Northern blotting ........................................................................................... 54
Detection of 2′-Phosphate linkages by RT-PCR .......................................... 54
Expression vector for intronless tRNAs ........................................................ 55
HAC1 epitope tagging and western blotting ................................................. 55
Results & Discussion ................................................................................................. 60
Genetic bypass of essential RNA repair genes in budding yeast ................. 60
RNA repair mutants have defects in translation ............................................ 63
RNA repair mutants accumulate intermediates and products of tRNA splicing .......................................................................................................... 68
RNA repair mutants have defects in unfolded protein response activation ....................................................................................................... 72
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Summary ................................................................................................................... 78
III MULTIPLE DECAY EVENTS TARGET HAC1 mRNA DURING SPLICING TO REGULATE
THE UNFOLDED PROTEIN RESPONSE ........................................................................ 80
Abstract ..................................................................................................................... 80
Introduction ............................................................................................................... 80
Materials & Methods ................................................................................................. 83
Cell culture and RNA preparation ................................................................. 83
RT-PCR/qPCR .............................................................................................. 83
Primer Extension ........................................................................................... 83
Northern blotting ........................................................................................... 84
Yeast strains and plasmids ........................................................................... 84
Results ...................................................................................................................... 89
RNA repair mutants have unique HAC1 mRNA processing defects. ............ 89
Kinase-mediated decay of cleaved HAC1 3′-exon competes with its ligation. .......................................................................................................... 92
Kinase-mediated decay of excised intron is required for HAC1 translation. ..................................................................................................... 96
Incompletely processed HAC1 mRNA is endonucleolytically cleaved and degraded .............................................................................................. 101
Discussion ............................................................................................................... 110
Summary ................................................................................................................. 115
IV FUTURE DIRECTIONS .................................................................................................. 117
Does kinase-mediated decay also regulate Xbp1 splicing in animals? .................. 117
Identify precise HAC1s secondary cleavage location and nuclease. ...................... 120
Applications of 2′-phosphorylated RNA to enhance stability in vivo ...................... 123
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Transcriptome-wide sequencing of products of RNA repair via enrichment of 2′-PO4
RNAs from tpt1∆ yeast ........................................................................................... 124
“Fungification” of metazoan cells to replace RtcB with Trl1 for marking repaired
transcripts with 2′-PO4. ........................................................................................... 126
REFERENCES ........................................................................................................................... 128
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LIST OF FIGURES
Figure 1.1: Mechanisms of endoribonuclease cleavage. ............................................................. 4
Figure 1.2: Model of RNA processing in tRNA splicing ................................................................ 8
Figure 1.3: Mechanisms of ligation. ............................................................................................ 24
Figure 1.4: Model of S. cerevisiae unfolded protein response activation ................................... 33
Figure 1.5: Three key mechanisms that couple HAC1 splicing to its translation. ...................... 34
Figure 1.6: Typical mRNA decay in the cytoplasm by Xrn1 and the exosome. ......................... 41
Figure 1.7: Examples of Kinase-Mediated Decay (KMD) ............................................................ 45
Figure 2.1: Genetic bypass of essential components of tRNA splicing with intronless tRNAs. . 61
Figure 2.2: Growth phenotype of RNA repair mutants. .............................................................. 64
Figure 2.3: tRNA processing phenotypes of RNA repair mutants. ............................................. 69
Figure 2.4: UPR-related phenotypes of RNA repair mutants. .................................................... 74
Figure 3.1. HAC1 mRNA processing defects in RNA repair and decay mutants. ...................... 90
Figure 3.2. Kinase-mediated decay of HAC1 3′-exon competes with its ligation. ..................... 93
Figure 3.3: Kinase-mediated decay of excised HAC1 intron is required to activate the unfolded
protein response. ............................................................................................................ 97
Figure 3.4: Incompletely processed HAC1s mRNA is cleaved and degraded. ........................ 102
Figure 3.5: A 5′- and 2′-phosphorylated HAC1 decay intermediate inhibits Xrn1. .................. 105
Figure 3.6: Kinetic analysis of HAC1 mRNA processing in cells lacking Tpt1. ........................ 108
Figure 3.7: Decay of HAC1 splicing intermediates regulates UPR activation, suppression, and
attenuation. ................................................................................................................... 111
Figure 4.1: Hypothesis of competition between decay and ligation in animals. ...................... 119
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LIST OF TABLES
Table 1.1: Exoribonucleases and their terminus requirements .................................................. 39
Table 2.1: Oligonucleotide sequences. ...................................................................................... 57
Table 2.2: Strain numbers and genotypes. ................................................................................. 58
Table 2.3: Intron-containing tRNA copy number and intron-dependent modifications in S.
cerevisiae. ....................................................................................................................... 67
Table 3.1: HAC1 processing intermediates ................................................................................ 85
Table 3.2: Oligonucleotide sequences ....................................................................................... 86
Table 3.3: Strain numbers and genotypes .................................................................................. 87
1
CHAPTER I
I INTRODUCTION
RNA is a versatile molecule in the cell, functioning as a messenger encoding a protein
(mRNA), a decoder of the genetic code (tRNA), a structural and catalytic component of the
spliceosome (snRNA) and the ribosome (rRNA), and a plethora of other known (snoRNA, ncRNA,
siRNA, etc.) and undiscovered functions. The lifecycle of RNA can be complex, with many
intervening processing events between birth and death (transcription and decay). Emerging
evidence suggests that some of these processing and decay pathways compete, and so this
dissertation seeks to address how processing and decay compete, and what effect these
competition(s) have on the cell.
To address the question of how processing and decay compete, as well as how this
competition affects cellular physiology, I focused on the unfolded protein response (the UPR), a
cell-autonomous, homeostatic signaling pathway that requires the cleavage, processing, and
ligation of an mRNA. To study the effects of RNA terminus modification and decay, I developed,
characterized, and deployed tools to add and subtract various terminus modification activity to
and from the cells, notably the 10x tRNA Block, a genetic bypass of the essential yeast
(Saccharomyces cerevisiae) tRNA ligase and 2′-phosphotransferase genes, as well as a
heterologous RNA ligase, and exonucleolytic decay pathways. In Chapter II, I demonstrate that
expression of intronless forms of the ten intron-containing tRNAs encoded in the yeast genome
is sufficient to bypass the essential genes TRL1 and TPT1. I remark that the 10x tRNA Block
construct failed to bypass any genes in the tRNA splicing endonuclease (SEN) complex,
indicating that they have an essential function beyond splicing tRNAs. Lastly, I introduce the
molecular and growth phenotype those RNA repair mutants have with respect to the UPR.
Chapter III further describes all my findings on how RNA ligation and decay interact at the stage
2
of 5′-phosphorylated products: having HAC1 3′-exon decay compete with ligation suppresses
the UPR in the absence of stimulus, and how kinase-mediated decay (KMD) of HAC1 intron is
required for robust induction of the UPR upon stimulus. In Chapter IV, I summarize these
findings, place them in context to show how they advance the field, and propose future directions
the project could take. Together, these findings support a generalized RNA decay mechanism,
kinase-mediated decay, and provide examples of how KMD and its competition with ligation
affect cellular physiology.
Endoribonucleases impose terminus chemistry
The termini of RNA are determined by their mode of synthesis: prior to modification, RNA
polymerases produce an RNA molecule with a 5′-triphosphate at the beginning, left behind from
the nucleotide triphosphate used to initiate synthesis, and with a 2′,3′-cis-diol at the end, left
behind by the last ribonucleotide triphosphate incorporated. However, many processing events
impinge on the RNA termini: in eukaryotes, RNA polymerase II (polII)-transcribed RNAs (mRNAs,
lncRNAs) undergo capping to add a 7-methylguanosine 5′→5′-triphosphate adduct (Shatkin
1976), effectively hiding the 5′-end from 5′→3′ exoribonucleases and marking the transcript as
a valid substrate for translation by the ribosome. Additionally, pol II transcripts are
polyadenylated at the polyadenylation site on the primary transcript, which Ysh1 (CPSF-73)
cleaves (Jenny et al. 1996; Ryan et al. 2004) and onto which Pap1 transfers non-templated
adenosine ribonucleotides (Lingner et al. 1991). After synthesis, many RNAs are post-
transcriptionally modified either by “trimming” (exonucleolytically) (see rRNA below) or by
cleavage (endonucleolytically) (see Sen2, Sen34, Las1 below).
The biochemistry of an endoribonuclease active site determines the RNA termini that are
produced from the incision: metal-ion-dependent endoribonucleases (e.g. RNase H, P, Z, MRP)
coordinate the scissile phosphodiester bond and a water molecule to yield “canonical” RNA
3
termini: a 5′-phosphate (5′-PO4) and a 2′,3′-cis-diol (Fig. 1.1A); in contrast, endoribonucleases
that do not coordinate a metal ion exploit the 2′-hydroxyl of the ribonucleotide to attack the
adjacent phosphodiester, yielding two RNA molecules, one with a 5′-hydroxyl (5′-OH) and the
other with a 2′,3′-cyclic phosphate (2′,3′-cP) (Fig. 1.1B). The consequence to the cell of which
variety of endoribonuclease catalyzes the cleavage is that metal-ion-dependent endonucleases
yield ‘clean ends,’ or termini of canonical chemistry that can enter directly into downstream
biochemical reactions, like ligation or exonucleolytic decay. In contrast, metal-ion-independent
endoribonucleases yield products with ‘dirty ends’ that need modification before they are
compatible for downstream reactions.
Metal-ion-dependent endoribonucleases generally employ aspartic acid residues in the
active site to coordinate two divalent metal ions (e.g. Mg2+) to stabilize additional negative charge
on the non-bridging oxygens of the phosphate group and to favor dissociation of water to form
a basic hydroxide that will serve as the nucleophile. By nucleophilic attack, the hydroxide forms
a bond with the phosphorous, releasing the leaving group, the 3′-oxygen of the nucleotide 5′ of
the phosphate (Yang 2011) (see electron-pushing curved arrows in Fig. 1.1A). This class of
endoribonucleases pertains little to this dissertation and is relegated to this brief mention in this
section.
Metal-ion-independent endoribonucleases (e.g. Sen2, Sen34, Rny1/Angiogenin, Las1,
PrrC, Ire1) use a variety of basic active site amino acids to catalyze cleavage. Generally, a basic
residue will deprotonate the 2′-hydroxyl (2′-OH) of a ribonucleotide, allowing the electronegative
2′-oxygen to perform nucleophilic attack on the neighboring phosphorous, forming a 2′,3′-cyclic
phosphate. Then, generally, an acidic residue will induce the 5′-oxygen of the downstream
ribonucleotide to serve as the leaving group, yielding a 5′-hydroxyl product (Fig. 1.1B). Metal-
ion independent endoribonucleases fulfill a plethora functions for the cell, which are reviewed
below.
4
Figure 1.1: Mechanisms of endoribonuclease cleavage.
A: Metal-ion-dependent endoribonucleases coordinate metal ions to prepare the scissile phosphodiester for nucleophilic attack by a hydroxide nucleophile obtained by abstracting a proton from a water molecule. Small curved arrows represent electron pushing; small dotted lines represent coordination or hydrogen bonding. The products of such cleavage are a 5′-fragment with a 2′,3′-cis-diol and a 3′-fragment with a 5′-phosphate.
B: Metal-ion-independent endoribonucleases use acidic (AH+) and basic (B-) amino acid residues to induce reactivity of the 2′-O, causing it to perform nucleophilic attack (curved arrows) on the scissile phosphate, resulting in a 5′-fragment with a 2′,3′-cyclic phosphate and a 3′-fragment with a 5′-hydroxyl.
5
Intrinsic RNA cleavage
Any phosphodiester bond in an RNA molecule is susceptible to intrinsic cleavage because
a 2′-hydroxyl is ‘intrinsically’ present adjacent to the phosphate backbone. Intrinsic cleavage—
the non-enzymatic, spontaneous cleavage of RNA—releases 5′-hydroxyl and 2′,3′-cyclic
phosphate products. Intrinsic cleavage is an ever-present threat to RNA in aqueous solution,
and its kinetics can be accelerated in the presence of heat, base, and divalent cations (e.g. Mg2+,
Mn2+, Zn2+, Pb2+). Heat increases the dissociation constant of water (KW), increasing the
concentration of hydroxide anions that deprotonate the 2′-OH, the first step of the cleavage
reaction. Alkaline pHs (pH ≥ 8) also increase hydroxide concentration, similarly increasing the
frequency of deprotonation of 2′-OH in RNA. Metal ions function as Brønsted–Lowry bases by
abstracting a proton from the 2′-OH of the ribose, yielding a more nucleophilic 2′-oxyanion
capable of attacking the adjacent phosphate (Brown et al. 1985). These factors are the
motivation for typically keeping RNA cold when resuspended in liquid solution, for why RNA
buffers are neutral-to-slightly-acidic in pH, and for why RNA storage buffers often contain EDTA.
Intrinsic cleavage occurs on a spectrum of three-dimensional conformations, with the
optimal being an “in line” conformation that facilitates an SN2-like transesterification reaction
mechanism, in which the attacking 2′-oxygen is 180° opposed to [“in line” with] the downstream
ribonucleotide’s 5′-oxygen, which is the leaving group in the reaction (Usher 1969; Soukup and
Breaker 1999). Single-stranded RNA is more susceptible to adopting the in-line conformation
than double-stranded RNA, meaning single-stranded RNA is more vulnerable to intrinsic
cleavage. Intrinsic cleavage is not sequence-specific, making it a useful step in kits and protocols
that require unbiased fragmentation of RNA for downstream applications (such as high-
throughput sequencing).
6
Self-cleaving ribozymes
Some of the most striking examples of the versatility of RNA are self-cleaving ribozymes.
Six known ribozymes carry out such catalysis, each yielding 5′-hydroxyl and 2′,3′-cyclic
phosphate products. The hepatitis delta virus (HDV) (Sharmeen et al. 1988) and glmS (Winkler et
al. 2004) ribozymes can cleave their own RNA chain, and the hammerhead (Prody et al. 1986),
hairpin (Buzayan et al. 1986), Varkud satellite (VS) virus (Saville and Collins 1990) and twister
ribozymes (Roth et al. 2014) can even ligate compatible RNA termini after cleavage. These self-
cleaving ribozymes catalyze cleavage using general acid-base proton transfer (Das and Piccirilli
2005), similar to the Brønsted–Lowry proton transfer at work in intrinsic RNA cleavage.
The most easily comprehensible example of self-cleaving ribozymes is the hairpin
ribozyme because its active site closely mimics that of RNase A. In the hairpin ribozyme, two
purine ribonucleotides act as a general base and acid, analogous to the two histidines in the
RNase A active site. Furthermore, a conserved adenine base stabilizes the excess negative
charge on the phosphate during the transition state of the cleavage reaction, closely mimicking
the role a lysine’s positively-charged ε-amine plays in the active site of RNase A. The ribozyme
catalyzes cleavage without the use of metal-ion co-factors in the active site but does use metal
ions to fold into a catalytically-active conformation (Nesbitt et al. 1997; Young et al. 1997; Hampel
and Cowan 1997). Cleavage proceeds via the pathways depicted in Fig. 1.1B, yielding 5′-OH
and 2′,3′-cP products.
The cleavage reaction mechanism depicted in Fig 1.1B is reversible, meaning that self-
cleaving ribozymes capable of positioning a 2′,3′-cyclic phosphate RNA and a 5′-hydroxyl RNA
in the active site can catalyze ligation of the non-canonical termini natively. In fact, the only
distinction between self-cleaving ribozymes that can or cannot re-ligate RNA is having base-
pairing interactions on both the 5′-side and the 3′-side of the incision event, providing the
necessary mechanism to position the termini for the reverse reaction, re-ligation. The
7
hammerhead, hairpin, and VS ribozymes are all capable of the ligation reaction, ultimately using
it to self-circularize (Buzayan et al. 1986; Canny et al. 2007; Jones et al. 2001).
Sen2 & Sen34
The yeast genome codes for an estimated 275 tRNAs (transfer RNAs), 61 of which contain
introns (Lowe and Eddy 1997; Chan and Lowe 2016). Among those, ten isodecoding tRNAs are
supplied exclusively by genes interrupted with introns (Chan and Lowe 2009). In 1977, Howard
Goodman, Maynard Olson, and Benjamin Hall discovered the first tRNA intron in the DNA
sequence encoding SUP4, a tRNATyr gene with a 14 base-pair (bp) intron downstream of the
anticodon (Goodman et al. 1977), and many additional tRNA introns have been found since
(Valenzuela et al. 1978; van Tol and Beier 1988). Soon after, Anita Hopper, Fred Banks, and Vicky
Evangelidis first published evidence that a nuclease was required for complete tRNA
biosynthesis (Hopper et al. 1978). The evolutionary origin and physiological purpose of tRNA
introns remain unknown, however their mechanism of removal is well-studied.
Cleaving introns out of tRNA transcripts is an essential function of the Splicing
EndoNuclease (SEN), a complex composed of four essential proteins (Dhungel and Hopper
2012). The primary transcripts of intron-containing tRNAs undergo primary nuclear export into
the cytoplasm, where they meet up with the SEN complex on the outer surface of the
mitochondria (in budding yeast (Peebles et al. 1983; Yoshihisa et al. 2007; Yoshihisa 2014)). The
catalytic subunits of the SEN complex, Sen2 and Sen34 (Trotta et al. 1997), recognize their
intron-containing tRNA substrates on a structural basis and cleave at the exon-intron junctions
(Fig. 1.2). Transfer RNA introns are typically small (14 to 106 nt) and share no sequence similarity,
even within the same organism (Belfort and Weiner 1997). The introns are recognized primarily
by structure by virtue of being located at the end of the anti-codon stem and not interrupting the
structure of the rest of the tRNA (Swerdlow and Guthrie 1984).
8
Figure 1.2: Model of RNA processing in tRNA splicing
Intron-encoding genes for tRNAs are transcribed by RNA polymerase III, trimmed at the 5′- and 3′-ends, and exported into the cytoplasm. The SEN complex cleaves the intron out of the tRNA, leaving two tRNA “halves.” Trl1 ligates the halves together, leaving a 2′-phosphate at the ligation junction. Tpt1 removes the 2′-phosphate left behind by Trl1. Depending on the tRNA, it may return to the nucleus for further processing, or remain in the cytoplasm. The intron cleaved out of the tRNA originates with a 5′-hydroxyl, an RNA terminus on which Xrn1 does not initiate decay. The RNA kinase domain of Trl1 phosphorylates the 5′-end of the intron, transforming it into a substrate of Xrn1 (Wu and Hopper 2014). The 5′-PO4 introns are robustly degraded.
9
Specifically, two conserved positions, called “cardinal positions” are always located the same
number of nucleotides away from the beginning and end of the intron, giving the SEN complex
the “measurements” for where to cut (Bufardeci et al. 1993; Di Nicola Negri et al. 1997).
The catalytic subunits of the SEN complex, Sen2 and Sen34, release the 5′- and 3′-tRNA
exons as 2′,3′-cP and 5′-OH termini, respectively, and the exons remain associated with each
other via extensive secondary structure. Similarly, the intron is released with 5′-OH and 2′,3′-cP
termini. Exoribonuclease 1 (Xrn1) degrades the discarded tRNA intron in the 5′→3′ direction (Wu
and Hopper 2014). The tRNA halves are ligated in healing and sealing reactions reviewed in the
“RNA repair in cellular physiology” section of this chapter. Upon ligation, some tRNAs return to
the nucleus via “tRNA retrograde nuclear import” for additional modifications (Shaheen and
Hopper 2005; Takano et al. 2005).
In addition to tRNAs, the SEN complex also cleaves the mRNA CBP1, a nuclear-encoded
gene for a mitochondrial protein (Tsuboi et al. 2015). Cytochrome b mRNA processing 1
promotes the stability and translation of the mitochondrial mRNA for cytochrome b (Krause et
al. 2004). It’s interesting that a non-tRNA substrate of the SEN complex would function at the
mitochondria given that the SEN complex localizes to the outer surface of the mitochondria in
yeast (Yoshihisa et al. 2003; 2007). However, CBP1 is not an essential gene and therefore likely
does not contribute to the essential function(s) of the SEN complex (Dieckmann et al. 1982).
Nevertheless, while tRNA splicing can be accomplished by moving the SEN complex to the
nucleus, nuclear SEN fails to complement deletion of mitochondrial SEN complex (Dhungel and
Hopper 2012), suggesting that SEN has yet another essential function in the cytoplasm of yeast.
Las1, Grc3, & 35S rRNA processing
Ribosomes are the ribonucleoprotein ribozymes that catalyze the mRNA-dependent
synthesis of proteins. Ribosomes have a complex synthesis pathway, coalescing four ribosomal
RNAs (rRNA) and approximately 80 proteins (Fromm et al. 2017). A major step of ribosome
10
assembly is the production and processing of the 35S pre-rRNA, synthesized as a tandem
transcript of the 18S, 5.8S, and 25S rRNAs, along with intervening and surrounding sequence
called Internal Transcribed Spacers (ITS) and External Transcribed Spacers (ETS) that are
removed through the pathway of pre-rRNA maturation (Woolford and Baserga 2013). This long
transcript was discovered in the 45S fraction of a sucrose gradient and called the “giant RNA”
(Scherrer 2003). Eukaryotic rRNA processing was first evinced in the 1960s, when Klaus
Scherrer, Jim Darnell, and Robert Perry demonstrated that 14C-uridine-labeled giant RNA rapidly
shifted to the 35S, and then 28S and 16S fractions (Scherrer and Darnell 1962; Scherrer et al.
1963; Perry 1962; Perry et al. 1964). They concluded that the “giant RNA” was the “pre-RNA”
that was transcribed and then processed into smaller RNAs. The field of pre-rRNA processing
has advanced, but to this day, some details of the precise molecular mechanisms of ribosomal
RNA processing remain undiscovered (Scherrer 2003).
The rRNA of budding yeast (S. cerevisiae), is encoded in the genome (as “rDNA”) at ~150
copies in a tandem repeat array located on chromosome 12 (Fernández-Pevida et al. 2015). RNA
polymerase III (polIII) transcribes the 5S rRNA, at which point Rex1/Rnh70 hydrolyzes 7 to 13
nucleotides off the 3′-terminus (van Hoof et al. 2000). In contrast to the former, simpler rRNA
maturation pathway, RNA polymerase I (polI) transcribes the 25S, 18S, and 5.8S rRNAs as a
tandem 35S rRNA transcript, which requires eight endonucleolytic cleavages (A0, A1, D, A2, A3,
B1L, C2, and B0) and eight exonucleolytic trimming steps (B1s, B2, C1′, C1, and four trimming steps
from 7S/L to 5.8S/L) to form rRNAs of the mature length (Woolford and Baserga 2013; Fernández-
Pevida et al. 2015). Despite 57 years passing between the first publication of pre-rRNA
processing and the date of this dissertation’s writing, the agents that catalyze three of the
endonucleolytic cleavages (A0, A1, and B1L) remain unidentified (Scherrer 2003). The
endonuclease that cleaves at C2, however, was identified as Las1 in 2015 (Gasse et al. 2015).
Internal Transcribed Spacer 2 (ITS2) lies between the 5.8S and 25S rRNAs of the 27SBS pre-
11
rRNA. Las1 cleaves ITS2 to produce the 7SS and 26S pre-rRNAs (Veldman et al. 1981). In a
downstream step, Rat1 (Xrn2) trims the 26S pre-rRNA to yield a mature-length 25S rRNA
(Geerlings et al. 2000). Las1 is a metal-ion-independent endoribonuclease (Gasse et al. 2015), a
topic that will be revisited in the “Kinase-Mediated Decay” section of this chapter.
Stress-induced tRNA cleavage
Cleavage of tRNAs at or near their anticodon for emergency regulation of translation is a
theme in biology. In both bacteria and eukaryotes, cellular stresses can result in the cleavage of
tRNAs in their anticodon loops to produce 3′ and 5′ halves called “tiRNAs.” The Escherichia coli
anticodon nuclease, PrrC, mounts an antiviral response against bacteriophage infection by
cleaving tRNALys in the anticodon loop, preventing the tRNA from participating in translation
(Levitz et al. 1990; Kaufmann 2000). In addition, colicin E5 causes cell death by translational
arrest by cleaving 3′ of the queuine base modification at the wobble position in the anticodon
loop in tyrosine, histidine, asparagine, and aspartic acid tRNAs (Ogawa et al. 1999; Masaki and
Ogawa 2002). Eukaryotes also cleave tRNAs during times of stress. The ciliate Tetrahymena and
the parasitic protozoan Giardia lamblia both cleave tRNAs under starvation conditions (Lee and
Collins 2005; Li et al. 2008), and budding yeast, plants, and humans cleave tRNAs in response
to oxidative stress specifically (Thompson et al. 2008; Thompson and Parker 2009b). However,
a major difference between prokaryotic and eukaryotic tRNA cleavage is that prokaryotes tend
to cleave a high proportion of susceptible tRNAs, whereas eukaryotic cleavage is ever only a
minority of cleavable tRNAs (Thompson and Parker 2009a).
The regulation of the stress-inducible endoribonucleases Rny1 in yeast and angiogenin in
humans occurs through remarkably similar pathways despite the nucleases sharing no
evolutionary history. Rny1 is a yeast vacuolar endoribonuclease belonging to the RNase T2 family
and serves to harden yeast against osmotic and heat-shock stresses (MacIntosh et al. 2001;
Thompson and Parker 2009b). Rny1 is normally trafficked through the yeast endomembrane
12
system and secreted, reducing its chances of encountering cytoplasmic RNA. However, upon
oxidative stress, Rny1 is released into the cytoplasm, where it cleaves tRNAs, causing
widespread translational pausing (Pelechano et al. 2015).
In a similar mode of regulation, human angiogenin, a member of the RNase A family, is a
secreted protein that is stored in the lysosome but also held in the nucleolus with its inhibitory
binding partner Rnh1. While the anticodon nuclease function of angiogenin is similar to that of
Rny1 from yeast, the two are not homologs; curiously, Rny1 has an ortholog in humans,
RNASET2, but it appears not to cleave tRNAs (Yamasaki et al. 2009). Angiogenin was first
discovered by purification from the HT-29 adenocarcinoma cell line (Fett et al. 1985) and was
shown to produce tiRNAs in in vivo (Yamasaki et al. 2009; Fu et al. 2009). Angiogenin is activated
by stress and translocates from the nucleolus to the cytoplasm where it dissociates from Rnh1
and cleaves tRNAs to produce tiRNAs. In addition to slowing translation by depleting available
tRNAs, the tiRNAs themselves may also promote apoptosis (Emara et al. 2010). Transfection of
5′-tiRNAs (but not corresponding 3′-tiRNAs) promotes stress granule formation and inhibition of
translation independent of eIF2α phosphorylation. 5′-tiRNAs can also inhibit translation initiation
by displacing eIF4G from mRNAs in vitro (Ivanov et al. 2011). Interestingly, tiRNAs can
competitively bind cytochrome c over the apoptosis pathway protein APAF1, protecting cells
from apoptosis during osmotic stress (Saikia et al. 2014).
Ribotoxins in interspecies conflict
Under intense resource competition, organisms can benefit from sabotaging a competing
species. Microbes especially compete for scarce resources at close quarters, making the
nutrient intake of one microbe a great loss for a competing microbe. So many organisms deploy
toxins at the expense of a competing or predatorial species. These toxins may take the form of
small molecules, peptides, or fully-functioning proteins that enzymatically catalyze some toxic
reaction. Ribotoxins are the class of toxic enzymes whose substrates are RNA. Ribotoxins have
13
a variety of RNA substrates, but typically target RNA molecules central to the metabolism of their
victim, such as ribosomal RNA or transfer RNA for their roles in protein synthesis. Cleavage of
these RNA targets leads to malfunction of translation machinery and, via an unknown
mechanism, cell death due to double-strand breaks of genomic DNA during the S phase of the
cell cycle (Klassen et al. 2004; Klassen and Meinhardt 2005).
Colicins are a group of secreted proteins belonging to plasmid-encoded toxin-antitoxin
systems that kill neighboring E. coli cells that do not harbor the col plasmid (James et al. 1996;
Cramer et al. 1999; Masaki and Ohta 1985). The toxic mechanism of these proteins varies, but a
family of colicins harm their victims via their nuclease activity. Colicin E3 cleaves the 3′ 49
nucleotides off the 16S rRNA in E. coli, removing a rRNA helix of the 30S ribosomal subunit
critical for its interaction with the 50S subunit and the A-site tRNA (Bowman et al. 1971a; 1971b;
Dahlberg and Dahlberg 1975). The colicin E3 cleavage event stops translation of protein,
ultimately leading to cell death. Colicin E5 and D inhibit translation in E. coli by specifically
cleaving tRNAs, leading to cell death (Tomita et al. 2000). The particular queuine post-
transcriptional modification to a guanine residue of the wobble base in the anticodon of some
tRNAs is sufficient for colicin E5 and D cleavage (Ogawa et al. 1999). However, colicin E5 and D
are capable of cleaving tRNAs lacking the queuine so long as they contain a specific anticodon
sequence (Ogawa et al. 2006).
There are also secreted ribotoxins deployed among eukaryotes. Kluyveromyces lactis
Zymocin and Pichia acaciae Killer (PaT) are secreted fungal defense toxins that penetrate S.
cerevisiae cells and, upon arrival in the cytoplasm, cleave the anticodon loop of tRNAGlu(GAA)
and tRNAGln(UUG), respectively. Consequent depletion of the specific tRNAs arrests yeast
growth by translational inhibition, cell cycle arrest, and DNA damage response activation (the
latter by an unknown mechanism) (Lu et al. 2005; Klassen et al. 2004; Klassen and Meinhardt
2005; Klassen et al. 2008).
14
Both Zymocin and Killer toxins represent examples of tRNA ribotoxins that rely on an RNA
base modification for targeting. Zymocin specifically recognizes the U34-wobble base
modification (5-methoxycarbonylmethyl-2-thiouridine, or mcm5s2U) of tRNAGlu(GAA) as carried
out by a modification pathway requiring tRNA methyltransferase Trm9. Consequently, trm9∆
cells and cells overexpressing tRNAGlu(GAA) to the point of saturation of Trm9 pathway enzyme
kinetics exhibit resistance to the Zymocin ribotoxin (Jablonowski et al. 2006). Pichia acaciae Killer
toxin (PaT) is a linear-plasmid encoded toxin-antitoxin system that confers toxin secretion again
competing yeast and simultaneous resistance to production of the toxin (Worsham and Bolen
1990; Chakravarty et al. 2014). PaT targeting also relies on the mcm5s2U modification, but this
time of tRNAGln(UUG) (Klassen et al. 2008). Furthermore, PaT can make two incisions in the
tRNAGln(UUG), releasing a dinucleotide the eliminates the possibility of recovery from the toxin
via RNA repair (Meineke et al. 2012).
RNA decay in co-translational mRNA surveillance
Messenger RNAs encode protein sequences that are translated into amino acid sequences
by the ribosome, and this process of translation is monitored for irregularities in the mRNA. The
coding quality of mRNA is controlled through a family of processes called co-translational mRNA
surveillance, an umbrella term for a series of processes that monitor ribosome progress along
the open reading frame to detect mRNAs that are defective in translation. Adaptor proteins
interact with the ribosome to detect aberrant mRNAs and direct them toward RNA decay
pathways. Principally, three types of co-translational mRNA surveillance are known: Nonsense-
Mediated Decay (NMD), Non-Stop Decay (NSD), and No-Go mRNA Decay (NGD) (Reviewed in
(Shoemaker and Green 2012)). Together, these processes ensure that a message accurately
encodes a peptide sequence.
No-go mRNA decay (NGD) is one of the co-translational mRNA surveillance processes and
detects stalled ribosomes on messages (Doma and Parker 2006). When ribosomes encounter
15
strong stalls in translation elongation, as caused by stem-loops, rare codons, or poly-positively-
charged amino acids, NGD is activated and the mRNA is cleaved, releasing 2′,3′-cP and 5′-OH
RNA products (Navickas et al. 2018). The cytoplasmic exosome degrades the 5′-product of
endonucleolytic cleavage via recruitment through Ski7, and Xrn1 degrades the 3′-product. No
specific mRNAs have been identified as being consistently subject to NGD, indicating that NGD
is likely a general-purpose quality control pathway that removes stochastically corrupted mRNAs
from the pool of translating messages.
While the nuclease of NGD is not known (and is speculated to be the ribosome itself), the
release-factor-like proteins Dom34 and Hbs1 recognize these aberrantly translating ribosomes
and remove the ribosome from the message in a stop-codon-independent manner. Dom34 and
Hbs1 appeared central in the discovery of no-go decay (Doma and Parker 2006), and the two
proteins were suspected of working as a ribosome recycling factors because of Dom34 is
paralogous to eukaryotic Release Factor 1 (eRF1), and Hbs1 to eRF3. Dom34 and Hbs1 were
later shown to form a heterodimer that mimics the conformation of a eukaryotic translation
termination complex (Chen et al. 2010; Becker et al. 2011) and even uses energy from the
hydrolysis of GTP to separate the 60S and 40S ribosomal subunits (Becker et al. 2011).
Conflicting reports indicate that Dom34 and Hbs1 are required for endonucleolytic cleavage of
the no-go mRNA (Passos et al. 2009) and some indicate no requirement (Ikeuchi and Inada 2016).
Either way, the factor that induces cleavage of the mRNA just upstream of the stalled ribosome
remains unidentified.
Non-Stop mRNA Decay (NSD) is rapid degradation of mRNAs that lack a translation
termination codon (Frischmeyer et al. 2002; van Hoof et al. 2002). Non-stop mRNAs can occur
naturally by alternative polyadenylation within the Open Reading Frame (ORF), by mutation, and
by endonuclease cleavage. NSD is activated by the ribosome reaching the 3′-terminus of the
mRNA, causing it to stall. Because the ribosome is stalled without a stop codon in the A-site,
16
the stop-codon-independent release factors Dom34 and Hbs1 recycle the ribosome, and Ski7
couples the process to rapid 3′→5′ degradation of the mRNA by the exosome (Horikawa et al.
2016). NSD is related to no-go decay by the need for stop-codon-independent ribosome release
factors, but NSD is different from other forms of exosomal degradation because the GTPase
domain of Ski7 is required for the process (van Hoof et al. 2002). Decapping followed by 5′→3′
decay, as carried out by Dcp1 and Xrn1, plays a minor role in removal of non-stop mRNAs from
circulation (Frischmeyer et al. 2002; Inada and Aiba 2005). Furthermore, the peptide resulting
from the non-stop mRNA is rapidly degraded by the ubiquitin-proteasome pathway via Ltn1 or
Not4 ubiquitinylating the peptide from the stalled ribosome (Wilson et al. 2007; Dimitrova et al.
2009; Bengtson and Joazeiro 2010).
Lastly, Nonsense-Mediated Decay (NSD) is the quality-control mechanism that degrades
mRNAs with aberrant stop codons. Such a quality control system was first suggested in 1979
when Regine Losson and Francois Lacroute introduced amber nonsense codons into the yeast
URA3 gene, causing destabilization of the URA3 mRNA, otherwise known for its legendary
stability (Losson and Lacroute 1979). Other causes of NMD include long 3′-untranslated regions
that alter the relationship of the poly(A) tail to the ORF (Muhlrad and Parker 1999a; Kebaara and
Atkin 2009), alternative translation start codons out of frame with the larger ORF that lead to
premature termination (Welch and Jacobson 1999), pre-mRNAs with introns that contain stop
codons (He et al. 1993; Sayani et al. 2008), mRNAs that contain frame shift sequences or
mutations (Belew et al. 2011), and mRNAs that contain upstream ORFs (Gaba et al. 2005; Guan
et al. 2006). These aberrant mRNAs become targets of Upf1, Upf2, and Upf3, which associate
with the translation termination complex and cause deadenylation-independent decapping or
shortening of the poly(A) tail, ultimately leading to Xrn1 degrading the mRNA (Muhlrad et al. 1994;
Cao and Parker 2003; Mitchell and Tollervey 2003; Baker and Parker 2004). Additionally, Upf1
17
alone can act to repress translation of an aberrant mRNA (Muhlrad and Parker 1999b) and can
target nonsense mRNAs to P-bodies (Sheth and Parker 2006).
Ire1, the “splicing” endoribonuclease of the unfolded protein response (UPR)
Ire1 (Inositol-Requiring Enzyme 1, alias: Ern1) (Nikawa and Yamashita 1992; Mori et al.
1993) is the transmembrane sensor protein that conveys the signal of unfolded proteins from
within the lumen of the endoplasmic reticulum (ER) to the cytoplasm (Walter and Ron 2011).
When unfolded proteins accumulate in the ER, the unfolded protein response (UPR) is triggered,
leading to corrective changes in gene expression that increase the size and protein-folding
capacity of the ER (covered in-depth in the “RNA repair in cellular physiology” section)
(Kozutsumi et al. 1988; Schuck et al. 2009). Ire1 conveys this signal via two enzymatic activities
present on its cytoplasmic domain: (i) it autophosphorylates other Ire1 molecules
(autophosphorylation in trans) upon oligomerization caused by accumulating unfolded proteins
in the ER lumen; and (ii) Ire1 activates its endoribonuclease domain, which is capable of cleaving
cytoplasmic RNAs in the vicinity of the ER membrane (Moore and Hollien 2012).
A structural examination of Ire1 explains much of its function in sensing unfolded proteins
and activating the UPR (Korennykh and Walter 2012). The primary/domain structure of Ire1 is
composed of an N-terminal sensing domain in the lumen of the ER, then a transmembrane helix,
then a cytoplasmic CDK2-like serine/threonine kinase, and finally a C-terminal ribonuclease
domain (Sidrauski and Walter 1997). A crystal structure of the luminal sensing domain revealed
abundant β-strands that coalesce to form a β-sheet platform (Credle et al. 2005) with the
surprising finding of a deep pocket with architectural similarity to the disordered-peptide-binding
pocket of major histocompatibility complex (MHC) (Achour et al. 1998; Olson et al. 2006). This
finding supports the model that the sensor domain of Ire1 can directly bind disordered/unfolded
proteins in the lumen of the ER to support oligomerization/activation (Credle et al. 2005; Gardner
and Walter 2011). In contrast, a competing model of activation of Ire1 is based on the
18
interpretation of a different crystal structure with an occluded groove on the β-sheet platform
(Zhou et al. 2006) to support a model in which unfolded proteins sequester BiP/Kar2 away from
direct Ire1 binding, allowing Ire1 the opportunity to dimerize and oligomerize (Bertolotti et al.
2000). However, unfolded proteins are likely the direct ligand of luminal domain of Ire1, as shown
by in vitro binding studies with hydrophobic and basic peptides, as well as in vivo
immunoprecipitation of Ire1 to a constitutively-unfolded protein (Gardner and Walter 2011).
What the conflicting models have in common is that the luminal sensor domain of Ire1 must
undergo a conformation change upon protein folding stress (independent of its ligand) in order
to self-associate and form oligomers on the surface of the ER (Gardner and Walter 2011).
Quantitatively, the amount of Ire1 oligomerization is directly proportional to RNase activity,
strongly implying that Ire1 self-association is the mechanism of RNase domain activation
(Korennykh et al. 2011a). The higher-order assembly of Ire1 stabilizes a region of the protein
called the helix-loop element (Korennykh et al. 2009), whose residues are critical for cleavage of
substrate RNA (Lee et al. 2008). Important to this interaction are the phosphoserine and
phosphothreonine residues formed from the specific autophosphorylation in-trans activity. The
phospho-amino acids form salt-bridges across monomers of the higher-order assembly, further
stabilizing the interaction. The interface of two Ire1 proteins (dimer) stabilizes the helix-loop
element, completing the active site of the RNase domain. This dimer structure of the RNase
domain is further supported by the 1:2 stoichiometry of binding: in vitro, one stem-loop RNA
substrate binds per two molecules of Ire1 (Korennykh et al. 2011b).
Across phyla, Ire1 has only three well-supported substrates that all function similarly as
transcription factors in vivo: HAC1 in yeast, bZIP60 in plants, and Xbp1 in animals. All three
mRNAs have stem-loop structures in common: a short stem with a seven- or eight-nucleotide
loop bearing the consensus sequence 5′-CNGNNGN-3′ (Gonzalez et al. 1999; Sidrauski and
Walter 1997; Yoshida et al. 2001). How these substrates of Ire1 function once they have been
19
cleaved is reviewed in the “Unfolded protein response” area of the “RNA repair in cellular
physiology” section below.
Ire1 has additional functions in the UPR beyond cleaving RNA for splicing. Hollien and
Weissman first identified ER-associated mRNAs that Ire1 could cleave (other than Xbp1) using a
microarray on total RNA isolated from Drosophila S2 cells (Hollien and Weissman 2006). A subset
of RNAs downregulated during protein folding stress returned to normal expression levels when
Ire1 was knocked down with RNAi, but not under Xbp1 knock-down. Northern blots showed the
RNAs were endonucleolytically cleaved. Later named Regulated Ire1-Dependent Decay (RIDD)
(Hollien et al. 2009), it was shown to be a general phenomenon of Ire1 across many phyla,
including mammals, plans, and fission yeast (Kimmig et al. 2012) (Tam et al. 2014; Guydosh et
al. 2017), but not budding yeast (Niwa et al. 2005). In mammals, RIDD cleaves RNAs that maintain
Xbp1-like stem-loop structural motifs near the ER membrane (Moore and Hollien 2015). In fission
yeast, where no UPR-associated transcription factor is present, Ire1 cleaves RNAs with some
degree of site-specificity, enacting RIDD on some substrates, but also increasing the stability of
other mRNA substrates (Kimmig et al. 2012). The lack of significant sequence identity (29%)
between the Ire1 protein of S. cerevisiae and S. pombe may confer the functional divergence (Li
et al. 2018), possibly explaining the differences in substrates of the endoribonuclease domain of
Ire1.
RNA terminus modification
Further processing steps or downstream functions sometimes require the modification of
the terminus of an RNA molecule. Cells often express discrete enzymes with RNA terminus
modification activity, like kinase, cyclic-phosphodiesterase, or cyclase activities, to assist in
processing RNA molecules for maturation or decay. 5′-termini can be phosphorylated
specifically by RNA 5′-kinases. Examples of RNA 5′-kinases include bacteriophage T4
20
polynucleotide kinase (PNK) (Wang et al. 2002) and the kinase domain of fungal or plant RNA
ligase (Trl1) (Wang and Shuman 2005; Wang et al. 2006). These kinases prepare the terminus of
the substrate RNA for ligation because the respective ligase will need a 5′-PO4 for transferring
an NTP onto the terminus, effectively “charging” the terminus as a high-energy 5′-Npp-RNA
intermediate for the ligation reaction. By changing the biochemistry of these RNA termini, the
RNA ligases/kinases expose the RNA to decay enzymes that specifically recognize the 5′-
phosphate of RNA (Wu and Hopper 2014). Notably, an absence of 2′- or 3′- direct kinases has
implications for the fidelity of ligation of RNA fragments, which is described in the next section
on RNA Ligases.
Clp1 RNA 5′-kinase
Clp1 (5′-RNA kinase) is an essential structural component of the yeast co-transcriptional
RNA processing complex (CLeavage and Polyadenylation factor 1a subunit) and a 5′-RNA kinase
in metazoans. Stefan Weitzer and Javier Martinez were motivated to identify the kinase that
converts 5′-OH synthetic siRNAs to the required 5′-PO4 terminus for incorporation into the RNA-
induced silencing complex (RISC) (Nykänen et al. 2001; Weitzer and Martinez 2007). The
homologous S. cerevisiae Clp1 was found not to have ATP-hydrolyzing activity, and so appears
to be a structural component of the cleavage and polyadenylation complex in yeast rather than
a catalytic subunit (Noble et al. 2007). Weitzer and Martinez also found that tRNA 5′- and 3′-
exons and introns are in vivo substrates of the kinase in animal cells. Clp1 is the candidate RNA
kinase for degradation of introns cleaved from spliced tRNAs in humans. In mice and humans,
mutations in Clp1 have been linked to neurodegenerative diseases, possibly caused by defects
in tRNA intron decay (Hanada et al. 2013; Karaca et al. 2014; Schaffer et al. 2014).
Grc3 RNA 5′-kinase
Grc3 (name etymology not known) is an essential 5′-RNA kinase in yeast first
bioinformatically identified as a triple-A ATP/GTPase by protein sequence homology (El-
21
Moghazy et al. 2000). Grc3 was identified in a microarray screen for non-coding RNA processing
and individually confirmed to participate in 27S pre-rRNA processing (a process that ultimately
produces mature 5.8S and 25S rRNAs) (Peng et al. 2003). Then Grc3 was demonstrated to be a
polynucleotide kinase associated with rRNA transcription centers, where it assists RNA
polymerase I terminate transcription (likely by the torpedo model) (Braglia et al. 2010). Another
study showed that an important function of Grc3 kinase is to phosphorylate the downstream
pre-rRNA fragment generated from Las1 cleavage, allowing the 5′-PO4-dependent Xrn2/Rat1
exonuclease to partially trim the pre-rRNA (Gasse et al. 2015).
CNP (cyclic nucleotide phosphodiesterase)
CNP (Cyclic Nucleotide Phosphodiesterase) is a 2′,3′-cyclic nucleotide phosphodiesterase
that catalyzes the irreversible hydrolysis of 3′-phosphodiester bonds in 2′,3′-cyclic nucleotides
to produce 2′-PO4/3′-OH termini (Sprinkle 1989). CNP is found extensively throughout cells of
the nervous system of mammals, making up ~4% of protein in myelinating glial cells (Vogel and
Thompson 1988), and is required in oligodendrocytes for survival of neuronal axons (Lappe-
Siefke et al. 2003). Surprisingly, only one in vivo substrate of CNP has been identified: Xbp1, the
mRNA of the unfolded protein response of animals (Unlu et al. 2018). Nonetheless, CNP is
frequently employed in biochemical manipulations of RNA termini in vitro and in vivo (Schwer et
al. 2008).
RtcA (3′-terminal phosphate cyclase)
RtcA (2′,3′-cyclase) uses ATP to cyclize 3′-phosphate RNA termini to 2′,3′-cyclic
phosphates. RtcA is widely distributed across the genomes and proteomes of eukaryotes and
prokaryotes (Genschik et al. 1997). 2′,3′-cyclase activity was first discovered in the lysate of
HeLa cells (Filipowicz et al. 1983) and in Xenopus cell nuclei (Laski et al. 1983). 2′,3′-cyclic
phosphate RNAs, like the kind that RtcA yields, are the substrates of the RNA ligases discussed
in the next section, conferring RtcA with a possible function of preparing substrates for ligases.
22
However, cyclic phosphates are stable once formed, so it is unclear what the physiological
substrates of RtcA could be across the domains of life. Potential physiological roles for RtcA
have been proposed after discovering additional products that RtcA is capable of generating,
including 5′-adenylated RNA (from 5′-PO4 substrates) and 5′-adenylated DNA in a nicked double
strand, owing to much overlap in the catalytic mechanism of RtcA with RNA/DNA ligases
(Chakravarty and Shuman 2011). Notably, yeast have no 2′,3′-cyclase activity in cell lysate (Billy
et al. 2000) and RtcA is not essential in E. coli (Genschik et al. 1998).
RNA ligases combine RNA terminus modification domains with a ligase domain.
Cells deploy a variety of strategies to “clean up”—or “heal”—the ends of RNA released
from cleavage events. Some of the RNA terminus modifications discussed in the previous
section resurface in multifunctional RNA ligases. The modification strategy used by the cell
largely depends on the RNA ligase expressed in the organism: fungi and plants use a “5′→3′”
mechanism of ligation, whereas animals, bacteria, and archaea use a “3′→5′” mechanism,
named so because of the terminus on which the phosphate becomes incorporated into the
phosphodiester backbone of the RNA molecule (Fig. 1.3) (Unlu et al. 2018).
Trl1, the ligase of fungi & plants
The tRNA Ligase in plants and fungi, Trl1 (alias: Rnl1, for RNA Ligase) was at first only
detected via its activity in yeast and in wheat germ extract (Knapp et al. 1978; Abelson 1979;
Konarska et al. 1981). In 1983, Greer et al. determined the enzymatic steps of the ligase from
enzyme isolated by activity purification (Greer et al. 1983), the same technique used a decade
earlier on T4 Bacteriophage RNA ligase (Silber et al. 1972). In 1986, Eric Phizicky, John Abelson
et al. showed that Trl1 can perform ligation in purified form on tRNA halves and an arbitrary
oligoribonucleotide substrates, implying that Trl1 is a general RNA ligase (Phizicky et al. 1986).
Trl1 has three separable domains (Fig. 1.3A): (i) a cyclic phosphodiesterase domain that
23
hydrolyzes the 2′,3′-cyclic phosphate to a 2′-phosphate/3′-hydroxyl; (ii) a RNA 5′-kinase
domain; and (iii) a 5′-adenylation domain that forms the RNA-5′-adenylate intermediate that
undergoes nucleophilic attack by the 3′-OH terminus to form a phosphodiester bond (Greer et
al. 1983; Schwartz et al. 1983; Xu et al. 1990; Apostol and Greer 1991). The termini-modifying
activities constitute “healing” of the RNA ends, and the adenylyl-transferase/ligase activity
represents “sealing” of the two molecules of RNA together. All three domains/functions of Trl1
are essential in yeast, but they can be expressed as separate proteins (Sawaya et al. 2003).
The completed ligation reaction heals the termini, and seals together the RNA molecules
with a phosphodiester bond, but a 2′-phosphate “scar” remains at the ligation junction due to
the cyclic phosphodiesterase specificity of Trl1. Plants and fungi have an enzyme, 2′-
PhosphoTransferase 1 (Tpt1) to remove the 2′-phosphate (Fig. 1.3A) (McCraith and Phizicky
1990). Tpt1 transfers the 2′-phosphate to a molecule of NAD+, displacing the nicotinamide and
yielding an unusual product, ADP-ribose 1″,2″-cyclic phosphate (Culver et al. 1993).
RtcB, the ligase of prokaryotes & animals
RtcB, RNA ligase present in archaea, bacteria, and animals, uses a different strategy to
ligate RNA. The locus of rtcB in the E. coli genome (in an operon alongside rtcA, an RNA 2′,3′-
terminal cyclase, and rtcR, a σ54 transcription factor) and its sequence conservation with other
bacterial ligases suggested it could be an RNA ligase (Galperin and Koonin 2004). RtcB and
HSPC117 were nearly simultaneously discovered as the RNA ligases responsible for tRNA
splicing in prokaryotes and animals, respectively (Tanaka and Shuman 2011; Popow et al. 2011).
Soon after, RtcB was expressed in yeast to show that it could complement deletion of the
essential gene TRL1 and that it also functioned as the ligase in the unfolded protein response
(Tanaka et al. 2011b). (The unfolded protein response, and the effect of RtcB on it, is discussed
in greater detail below and in Chapter III.) Furthermore, the biochemical activity of RtcB was
24
characterized as distinct from that of plant and fungal ligases, involving a “two-step,” 3′→5′
reaction mechanism (Zofallova et al. 2000; Tanaka et al. 2011a; Chakravarty et al. 2012).
Figure 1.3: Mechanisms of ligation.
A: In fungi and plants, the tRNA Ligase 1, Trl1 (also called RNA Ligase 1, Rnl1), is a multifunctional enzyme with distributive cyclic phosphodiesterase (CPDase), 5′-kinase, and ligase (involving transient 5′-andelylation) activity. The CPDase hydrolyzes the 2′,3′-cP to a 2′-PO4 and a 3′-OH, which serves as the nucleophile during ligation to displace the high-energy adenylate group. The ligation product has a 2′-phosphate “scar” (in blue, left over from the CPDase reaction), which is transferred to a molecule of NAD+, releasing nicotinamide (not depicted) and ADP-ribose 1″,2″-cyclic phosphate (depicted). The phosphate highlighted in red (5′) is incorporated into the phosphodiester backbone, hence the name “5′→3′ ligation.”
B: In animals, bacteria, and archaea, the enzyme RtcB (HSPC117) also works as a multifunctional enzyme with cyclic phosphodiesterase (CPDase) and ligase (with transient 3′-gyanelation) activity. The CPDase hydrolyzes the 2′,3′-cP to a 2′-OH and a 3′-PO4. The 5′-OH serves as the nucleophile during ligation to displace the high-energy guanylate group. The 3′-PO4 highlighted in red ultimately becomes incorporated into the phosphodiester backbone, giving it the name “3′→5′ ligation.”
25
RtcB/HSPC117 hydrolyzes the 2′,3′-cyclic phosphate to a 3′-phosphate, which it then
guanylates to form a 3′-ppG intermediate; finally, the 5′-OH, so-far unmodified throughout this
process, displaces the guanylate group, sealing the RNAs together with a phosphodiester bond
(Fig 1.3B).
Meanwhile, the metazoan ligase RtcB (alias: HSPC117), already known to carry out tRNA
ligation, was demonstrated to be a ligase essential for activation of the unfolded protein response
(UPR) in the nematode C. elegans, as shown via deletion mutants expressing a pre-spliced tRNA
construct (Kosmaczewski et al. 2014). In mammals, HSPC117 was identified using a synthetic
biology approach coupled with a siRNA screen (Lu et al. 2014) and with HeLa whole-cell extracts
coupled with RNAi knock-down of RtcB (and its partner protein, Archease) (Jurkin et al. 2014).
Fidelity of RNA ligases
The biochemistry of RNA termini encodes information about the fidelity of the molecule,
allowing ligases to enforce fidelity requirements on their substrates. Generating a seamless
ligation of two RNA ends is essential for producing tRNAs with correct anticodon loop structure
and preserving the coding potential of the 3′-exon of HAC1/Xbp1/bZIP60 in the unfolded protein
response. But this observation raises the question: how do ligases ensure fidelity? There are no
known terminal 2′- or 3′-RNA kinases, meaning that any RNA with a 2′,3′-cyclic phosphate, 2′-
phosphate, or 3′-phosphate was cleaved at that exact site, and so is a faithful substrate for
ligation. Supporting this idea are three examples of ligase specificities for such substrates, thus
enforcing fidelity of their ligation products. Firstly, RtcB, the ligase of prokaryotes and animals,
requires a 3′-PO4 to transfer the guanylate group in preparation for ligation. In the case where
the cyclic phosphate is hydrolyzed to a 2′-PO4, the RNA cyclase RtcA can restore the substrate
to a 2′,3′-cyclic phosphate, preserving the fidelity of the terminus. Secondly, in an interesting
twist, S. cerevisiae ligase, Trl1, requires a 2′-PO4 of its ligation substrates despite the 2′-PO4 not
being involved in the biochemical reaction itself (Schwer et al. 2004). Similarly, the RNA ligase of
26
the plant Arabidopsis thaliana requires a 2′-PO4 on its tRNA substrates for ligation (Wang et al.
2006). Thirdly, a [counter]example of a low-fidelity ligase is found in T4 RNA Ligase 1 (T4 Rnl1).
The previously discussed enforcement of fidelity via the 2′ or 3′ terminus of the RNA is not
present in bacteriophage T4 Rnl1, and consistent with that observation, the ligation products of
T4 Rnl1 occasionally lack nucleotides from what were the 3′-ends of its substrates (Schwer et
al. 2004). Thus, the RNA termini contain information about the fidelity of the molecule, and ligases
can read this information to enforce fidelity requirements on their substrates. In the next section,
the biochemistry of the 5′-terminus of RNA is explored for its effect on ligation and other
processes.
RNA repair in cellular physiology
RNA repair is distributed widely throughout biology. Therefore, it is not surprising that it
has been discovered in a variety of contexts, performing a variety of tasks for the cell, or
sometimes not for the cell, reviewed in the T4 bacteriophage RNA repair section below. RNA
repair ligates tRNA exons back together once their introns have been excised, and similarly, RNA
repair enables the unfolded protein response to activate by ligating the exons of an inducibly-
cleaved mRNA. Many stress responses use RNA repair to reverse the response, like after stress-
induced cleavage of tRNAs in prokaryotes or restoring ribosome function after MazF cleavage.
RNA repair is also a direct response during interspecies conflict with ribotoxins, examples of
which include the PnkP/Hen1 RNA repair and immunization system (Chan et al. 2009) and
defense against the Killer and Zymocin anti-fungal toxins (Klassen et al. 2008; Lu et al. 2005).
The splicing of tRNAs
The genomes of organisms encode some fraction of tRNA genes with introns, first
discovered as intervening sequence (IVS) in the SUP4 gene (Heinemann et al. 2010; Goodman
et al. 1977). The budding yeast genome contains an estimated 275 tRNA genes, 61 of which
27
contain introns (Lowe and Eddy 1997; Chan and Lowe 2016). Among those, ten isodecoding
tRNAs are supplied exclusively by genes interrupted with introns (Chan and Lowe 2009). The
splicing endonuclease complex (SEN) recognizes the structure of the intron-containing tRNA and
precisely cleaves at each side of the intron, releasing the intron and cleaved tRNA molecule.
Because the catalytic subunits of the SEN complex catalyze cleavage independent of metal ions,
the resulting tRNA fragments have 2′,3′-cyclic phosphate (2′,3′-cP) and a 5′-hydroxyl (5′-OH)
termini (Trotta et al. 1997).
In 1986, Eric Phizicky and John Abelson purified the ligase responsible for putting the two
halves of the tRNA together (Phizicky et al. 1986). Subsequent studies in John Abelson’s and
Chris Greer’s labs revealed the mechanistic steps tRNA Ligase (Trl1, alias: Rlg1 for RNA ligase
1) uses to ligate tRNAs (Greer et al. 1983). Trl1 performs RNA repair by preparing the termini for
ligation and then forming the phosphodiester bond in three successive steps (Figs 1.2 & 1.3A):
(i) the cyclic phosphodiesterase domain (CPDase) of Trl1 hydrolyzes the 2′,3′-cP to form a 2′-
phosphate, 3′-hydroxyl terminus; (ii) the RNA 5′-kinase domain phosphorylates the 5′-OH to
form a 5′-PO4; (iii) the ligase domain adenylates the 5′-PO4 to form a 5′-App intermediate, and
uses the 3′-OH to displace the adenylate, forming a ligated product with a 3′→5′ phosphodiester
bond and a residual 2′-phosphate at the junction of ligation.
The ligated tRNA is released from Trl1 with a residual 2′-PO4, which has to be removed for
the tRNA to undergo further modification and function in translation (Culver et al. 1997; Spinelli
et al. 1997). The 2′-phosphotransferase enzyme, Tpt1, specifically catalyzes the removal of the
2′-PO4 via a transesterification reaction of the phosphate onto a molecule of NAD+, yielding a
canonical 5′-3′-phosphodiester with an adjacent 2′-OH and an unusual product: ADP-ribose-
1″,2″-cyclic phosphate (McCraith and Phizicky 1990; 1991; Culver et al. 1993). Sherry Spinelli
and Eric Phizicky isolated the TPT1 gene, a highly-specific 2′-dephosphorylation enzyme, and
28
showed, using the conditional mutant tpt1-1, that spliced tRNAs are under-modified at
nucleotides near the splice site (Spinelli et al. 1997).
In humans, some tRNAs also require splicing by the conserved TSEN complex (tRNA
splicing endonuclease), with the catalytic subunits conservatively named TSEN2 and TSEN34
(Paushkin et al. 2004). However, in contrast to yeast, human tRNA splicing uses a different ligase
with a more direct mechanism (Abelson et al. 1998). In 2011 Johannes Popow and Javier
Martinez discovered the ligase, HSPC117 in humans, using an activity-guided purification
procedure on HeLa cell extracts, and they confirmed the requirement of the enzyme in cells via
RNAi knock-down (Popow et al. 2011). Popow et al. noted that the high degree of sequence
conservation between human HSCP117 and E. coli RtcB suggests conservation of the ligase
function(s), even between such distantly related organisms. Given that the RNA ligase in animals
RtcB, which does not yield ligation products with 2′-phoshphates (Chakravarty et al. 2012), is
able to activate the UPR (Lu et al. 2014), it is surprising that animal cells also express a 2′-
phosphotransferase, Trpt1. However, Trpt1 is dispensable in mice (Harding et al. 2008), calling
into question why this conserved enzyme remains present in an organism so evolutionarily
distant from the last common ancestor that used an RNA ligase that generated 2′-
phosphorylated products.
Once spliced out of the tRNAs, the introns are degraded by Xrn1, the primary means of
degrading these RNAs (Wu and Hopper 2014), but an intervening phosphorylation step would
be required to make the 5′-OH introns (as released by the Sen2 in the SEN complex) into a 5′-
PO4 substrate of Xrn1 (see kinase-mediated decay section).
The unfolded protein response (UPR)
The unfolded protein response (UPR) is a homeostatic intracellular signaling pathway,
conserved throughout eukaryotes, that signals to the nucleus during Endoplasmic Reticulum
(ER) stress and ultimately increases the protein folding capacity of the ER. Misfolding proteins,
29
protein processing overload, oxidative stress, calcium efflux, and viral infection each constitute
a stress to the ER and can cause UPR activation (Kozutsumi et al. 1988). Activation of the UPR
increases the protein folding capacity of the cell by upregulating expression of dozens of genes
like chaperones, membrane lipid biosynthesis genes, and ER-resident protein degradation
genes, ultimately resulting in an enlargement of the organelle and an increase of the ER’s protein
folding capacity (Schuck et al. 2009; Cox et al. 1997; Scheuner et al. 2001; Travers et al. 2000;
Casagrande et al. 2000).
As a homeostatic process, the UPR can be physiologically activated (i.e. adaptive/healthy)
in cells with high protein-processing requirements, like pancreatic β cells secreting insulin,
activated B-cells (plasma cells) secreting antibody, differentiating T-cells dividing rapidly, muscle
cells, and in active neurons synthesizing large amounts of protein (Okabayashi et al. 1985;
Reimold et al. 2000; Pramanik et al. 2018; Reimold et al. 2001; Clauss et al. 1993; Wang et al.
2010; Tan et al. 2018). However, UPR activation can be pathological (i.e. maladaptive/unhealthy)
in genetic diseases that cause protein misfolding (e.g. retinitis pigmentosa, cystic fibrosis) and
in type II diabetes (Ozcan et al. 2004). Additionally, and for the purposes of study, the UPR can
be activated by small molecules that put stress on the endoplasmic reticulum. Tunicamycin is a
potent inhibitor of N-linked glycosylation of proteins, thereby reducing protein stability in the ER
(Duksin and Mahoney 1982). Thapsigargin is often chosen for inducing the unfolded protein
response in mammalian cells, where it inhibits the sarcoplasmic Ca2+ ATPase, disrupting ionic
homeostasis in the ER (Rogers et al. 1995; DuRose et al. 2006). Dithiothreitol (DTT) can also be
used to induce the UPR by breaking the disulfide bonds formed in the ER, destabilizing those
proteins (Lee 1992).
While the UPR is present in all eukaryotes, different phyla possess different sets of
pathways of transmitting the stress signal, with the most conserved arm of being the Ire1
signaling pathway. Ire1 is the transmembrane sensor protein that conveys the signal of unfolded
30
proteins from within the lumen of the ER to the cytoplasm. Ire1 conveys this signal via two
enzymatic activities of its cytoplasmic domain: (i) it autophosphorylates other Ire1 molecules (in
trans) upon oligomerization caused by accumulating unfolded proteins; and (ii) Ire1 activates its
endoribonuclease domain, which can cleave cytoplasmic RNAs near the ER surface.
In budding yeast, Saccharomyces cerevisiae, Ire1 is the only known mechanism of
signaling ER stress to the nucleus. The discovery of the unprecedented mechanism of signaling
(and mRNA splicing) was demonstrated by a series of experiments published nearly
simultaneously by Peter Walter’s lab at UCSF and Kazutoshi Mori, first at UT Southwestern and
then at Kyoto University. Walter and Mori jointly received the Lasker Award in 2014 for their
discovery. But the idea that cells tailor their protein processing capacity on the basis of stress
was supported by initial observations of glucose-regulated proteins (GRPs) increasing in
expression not just under glucose starvation stress (Lee et al. 1983), but also under treatment
with amino acid analogues (Kelley and Schlesinger 1978). The connection between protein
metabolism and chaperones GRP78 (BiP, Kar2 in yeast) and GRP94 was inferred by expressing
influenza hemagglutinin gene alleles known to misfold and accumulate in the ER, which in turn
caused GRP78 and GRP94 expression to rise (Munro and Pelham 1986; Normington et al. 1989;
Kozutsumi et al. 1988).
Studies of the BiP gene of yeast, KAR2 (Normington et al. 1989; Rose et al. 1989;
Nicholson et al. 1990), set the stage for the discovery of Ire1 and Hac1 as signal transducers of
the unfolded protein response. A 22-bp conserved sequence in the promoter of KAR2 appeared
in the promoters of many other UPR-inducible genes and was sufficient to promote transcription
of a reporter gene (lacZ) upon induction of protein folding stress (Mori et al. 1992). Building on
that result, Mori then showed that the 22-bp element (named the UPRE for “UPR Response
Element”) is required for KAR2 induction under protein folding stress (Kohno et al. 1993). The
Walter and Mori labs cloned the UPRE upstream of a lacZ gene expressed in yeast, effectively
31
making a colorimetric screen for deletion mutants with defective UPR signaling. Using this
approach, Walter and Mori independently identified Ire1 (Inositol-requiring enzyme 1, alias: Ern1)
as a required component of the signaling pathway (Cox et al. 1993; Mori et al. 1993). Because
Mori also showed that Ire1 is a transmembrane protein with its N-terminus in the lumen and that
the cytoplasmic C-terminal kinase is required for signaling, Mori et al. hypothesized that Ire1
directly phosphorylated a latent transcription factor, causing it to activate, localize to the nucleus,
and drive expression of UPRE genes.
Consistent with that hypothesis, Mori’s group identified the transcription factor of the UPR,
HAC1 (aliases: Ern4 & Ire15), using a yeast one-hybrid screen that showed that Hac1 binds the
UPRE in vivo, activates transcription of UPRE genes, and is required for UPR signaling (Nojima
et al. 1994; Mori et al. 1996). Soon after, Peter Walter’s group used a synthetic overexpression
assay to determine that HAC1 is the transcription factor of the UPR, but that its expression is
regulated by a splicing event (Cox and Walter 1996).
The discovery that UPR signaling was transduced through splicing, not phosphorylation,
was compounded by the even more unexpected finding that yeast’s tRNA ligase was required
for splicing (and therefore signaling) (Cox and Walter 1996; Sidrauski et al. 1996). Walter and
colleagues found that Hac1 protein was only present in UPR-induced cells (as assayed by
immunofluorescence and western blot) and not in uninduced cells. This finding is inconsistent
with the typical mode of regulating a transcription factor’s activity, where the protein is present
at all times but inactive without stimulation. Northern blot analysis showed that the mRNA for
HAC1 decreased in length by about 250 nucleotides (nt) upon induction of the UPR in an Ire1-
dependent manner. Sequencing of the shortened HAC1 RNA species revealed that a 252 nt
“intervening sequence” (IVS) was missing from the shortened species (Cox and Walter 1996).
Furthermore, a random-mutagenesis genetic screen (sectoring screen) for mutants unable to
lose a KAR2 plasmid, indicating a defect in the UPR, uncovered an allele, rlg1-100, of the
32
essential tRNA ligase, TRL1 (alias RLG1) (Sidrauski et al. 1996). TRL1 is essential in yeast
because it ligates together tRNAs after the splicing endonuclease cleaves out the intron (Phizicky
et al. 1992), and so the rlg1-100 allele was discovered to be a separation-of-function mutant
allele, still enabling the yeast to ligate tRNA haves together, as required for viability, but not able
to ligate the halves of the HAC1 mRNA. Soon after, Walter’s group found Ire1 to be a site-specific
endoribonuclease that cleaves the HAC1 mRNA in a stress-dependent manner, providing a
mechanism for how information in the endoplasmic reticulum can be communicated to the
nucleus (Sidrauski and Walter 1997; Kawahara et al. 1998).
The aforementioned experiments revealed the only known signaling pathway entailing the
cytoplasmic splicing of an mRNA (Fig. 1.4): Normally bound by Kar2 under healthy conditions,
Ire1 instead binds unfolded or misprocessed proteins under conditions of ER stress, causing it
to oligomerize and activate the endoribonuclease domain (Gardner and Walter 2011; Lee et al.
2008; Korennykh et al. 2009). Ire1 recognizes HAC1 mRNA via its characteristic stem-loops,
which Ire1 cleaves to release the 5′-exon with a 2′,3′-cyclic phosphate terminus, the 3′-exon
with a 5′-hydroxyl terminus, and the intron with like termini (Gonzalez et al. 1999). The 5′- and
3′-exons are held together by extensive base-pairing, and so the termini are robustly ligated
together by Trl1 (as described previously). This spliced mRNA encodes a potent transcription
factor that localizes to the nucleus and upregulates dozens of genes, about 7% of the yeast
genome (Travers et al. 2000).
Hints that HAC1u mRNA processing was both necessary and sufficient for robust UPR
signaling came from expression of manipulated HAC1 ORFs and mRNAs, with the 5′-exon/UTR,
intron, and 3′-exon/UTR each responsible for important regulatory functions (Fig. 1.5).
33
Figure 1.4: Model of S. cerevisiae unfolded protein response activation
Unfolded proteins in the endoplasmic reticulum (ER) cause Ire1 to oligomerize and activate its cytoplasmic endoribonuclease domain, which site-specifically recognizes the cleavage site flanking the intron of HAC1 mRNA. Ire1 cleaves the intron out from the mRNA, at which point Trl1 ligates the two exons together, leaving a 2′-phosphate at the ligation junction due to its catalytic mechanism. Tpt1 removes the 2′-phosphate from the HAC1 mRNA, leaving the ligation event seamless. HAC1 mRNA is then translated to produce Hac1, a potent transcription factor that is trafficked to the nucleus to upregulate dozens of genes that increase protein folding capacity of the cell, thus relieving ER stress and re-establishing homeostasis. Notably, Hac1 promotes transcription of its own mRNA, making HAC1 splicing a positive feedback loop. Cleavage of HAC1 mRNA is reported to be the rate-limiting step in the pathway.
34
Figure 1.5: Three key mechanisms that couple HAC1 splicing to its translation.
HAC1 mRNA has two key features that ensure its translation is inhibited by the presence of the intron, and one feature that makes the signaling cascade quick to respond to ER stress. Firstly, HAC1 intron forms a strong base-pairing interaction with the 5′-untranslated region (5′-UTR) of the message, effectively preventing scanning ribosomes from reaching the open reading frame to begin translation (Rüegsegger et al. 2001). Secondly, in the event that a ribosome escapes inhibition and initiates translation on unspliced HAC1, a failsafe ORF encoded in the intron, in frame with the ORF of the 5′–exon, codes for a potent degron, resulting in rapid ubiquitination and degradation of the resulting protein (Di Santo et al. 2016). Thirdly and lastly, HAC1 3′-UTR folds into the bipartite element (BE) that anchors the mRNA to the cytoplasmic surface of the ER, near Ire1, on standby for rapid response to ER stress (Aragón et al. 2009).
35
HAC1u was found co-sedimenting with polyribosomes (Chapman and Walter 1997) despite there
being very little Hac1u (Unspliced Hac1) protein produced in unstressed cells (Cox and Walter
1996); this paradox was resolved by results indicating that the intron of HAC1u acts to prevent
scanning ribosomes from initiating translation on the unspliced mRNA (Rüegsegger et al. 2001).
In 2000, Kazutoshi Mori and Peter Walter both showed that swapping of the C-terminal domain
of Hac1 via the splicing of its mRNA was required for robust accumulation of the protein
transcription factor, but the C-terminal domain of Hac1s (spliced Hac1) was dispensable for
transcriptional activation (Mori et al. 2000). The Hac1 transcription factor upregulates
transcription of its own locus, HAC1, and that positive feedback loop is required for sustained
UPR activation (Ogawa and Mori 2004). HAC1u mRNA is tethered to the cytoplasmic surface of
the ER by the bipartite element (BE) in its 3′-UTR, a requirement for its rapid processing in the
event of protein folding stress (Aragón et al. 2009). And as recently as 2016, studies from David
Weinberg’s lab showed that the ORF of the intron of HAC1u in frame with that of the 5′-exon
encodes a potent degron, causing the protein resulting from spurious translation of HAC1u to be
rapidly degraded by the ubiquitin-proteasome pathway, as mediated by the ubiquitin ligase
DUH1 (alias: DAS1) (Di Santo et al. 2016). Importantly, both the translation initiation block and
the intron-encoded degron work together to maintain tight suppression of Hac1 protein
production when the cell is not experiencing ER stress, and thus not splicing HAC1.
RNA repair in bacterial stress responses and interspecies conflict
Microbes are subjected to quickly changing environmental conditions and to direct conflict
with microbes of other species. Two interesting cases of RNA breakage and repair occur in
microbes in response to stress and to interspecies conflict. In both instances, a situation of
broken RNA is rectified by repair. The RNA cleavage can be caused either by the microbe itself
or a competing microbe, but the repair is carried out to aid in recovery. Both examples support
the idea that RNA repair constitutes an adaptive function across many phyla.
36
E. coli under stress adapt their protein synthesis program by cleaving their rRNA and
mRNA, causing the ribosomes to favor translating a specialized subset of mRNAs. The
endoribonuclease MazF is part of a the mazEF toxin-antitoxin system (Aizenman et al. 1996;
Engelberg-Kulka et al. 2006) and was identified as the endoribonuclease that enacts this
translational program. MazF cleaves the 3′ 43 nucleotides off of the 16S rRNA, introducing
70S∆43 specialized ribosomes in the cell (Vesper et al. 2011). These ribosomes lack helix 45 and
the anti-Shine-Dalgarno sequence (aSD), both of which are essential for the translation of typical
mRNAs in E. coli (Shine and Dalgarno 1974). The specialized ribosomes preferentially translate
“leaderless” mRNAs that were similarly processed by MazF (Zhang et al. 2005), thus imposing
an alternative translation program.
Given the immense energetic costs of manufacturing a ribosome de novo, Temmel et al.
hypothesized that E. coli may ligate the rRNA back together to restore the ribosomes to their
pre-stress state once the stress passes. They identified RtcB as the ligase that reverses MazF
cleavage of rRNA to restore normal translation after stress (Temmel et al. 2016). Additionally, the
rtcB mRNA is itself processed by MazF, including it in the subpopulation of mRNAs selectively
translated during stress. This discovery provided a biological role for RtcB in E. coli and
demonstrated how RNA cleavage and ligation can be used to regulate an important bacterial
stress response.
PnkP/Hen1 RNA repair and immunization system is a cooperating 2′-O-methyltransferase
and RNA ligase system that repairs RNAs (usually tRNAs) damaged by ribotoxin endonucleases
and installs a methyl group at the 2′-position to block that oxygen from participating in cleavage
reactions in the future, effectively immunizing the RNA from cleavage again by the same
ribotoxin. Hen1, the methyltransferase, was first discovered in plants as a factor required for the
processing of miRNAs (3′-terminal 2′-O-methylation) (Kishi et al. 2005). Based on peptide
sequence similarity, homologous Hen1 proteins are also found in the genomes of various
37
bacteria, but their function in bacteria was unknown (Tkaczuk et al. 2006). Unexpectedly, the
Clostridium thermocellum operon coding for Hen1 contains a downstream gene possessing
kinase, phosphatase, and adenylyltransferase activities, hallmarks of an RNA ligase (Martins and
Shuman 2005). Chan and colleagues cloned and purified the gene from Anabaena variabilis and
showed that, in complex with the Pnkp ligase, the proteins carry out RNA repair on ribotoxin-
cleaved tRNAs and install a methyl group at the 2′-O position of the 5′-cleavage fragment prior
to ligating the fragments together (Chan et al. 2009).
Given that most ribotoxins use a metal-ion-independent mechanism to cleave their target
RNAs in their victims (Ogawa et al. 1999; Soelaiman et al. 2001; Morad et al. 1993; Nariya and
Inouye 2008; Tomita et al. 2000; Pedersen et al. 2003), the 2′-O-methylation is an effective
strategy to overcome the effects of the ribotoxins by blocking further cleavage. Quantitatively,
Wang et al. found a 31% decrease in cleavage susceptibility in vitro, a much-preferred outcome
over creating a futile cycle of cleavage and ligation in response to the ribotoxin (Chan et al. 2009).
In the Pnkp-Rnl-Hen1 complex, methylation occurs before ligation, and methylation may even
be conducive to the ligase component because methylated substrates are ligated 10-fold faster
(Wang et al. 2015). Furthermore, from a structural perspective, the methyltransferase active site
is situated between the 3′-phosphatase and the ligase active sites, maximizing the likelihood of
methylation of substrates prior to ligation.
Terminus chemistry and exonucleolytic RNA decay
The chemistry of the end of a molecule of RNA can have major impacts on which
exonucleases will engage with it and at what rates they will catalyze degradation. The three
known 5′→3′ exoribonucleases all require 5′-PO4 termini to catalyze decay. 3′→5′
exoribonucleases have a more heterogeneous set of requirements. These biochemical signals
sent from the RNA termini to the exonucleases operating on them represent a form of regulation
38
of their activity in decay and processing. Salient exoribonucleases are discussed here and
summarized in Table 1.1.
5′→3′ RNA decay enzymes require substrates be 5′-phosphorylated
All RNA is subject to decay, and cells require regulated RNA decay machinery to adapt to
their environment by changes in behavior through new signals sent from the nucleus. Messenger
RNAs are capped at their 5′-ends for protection from RNA degradation machinery in the cell
(Furuichi et al. 1977; Shimotohno et al. 1977). One of the primary decay factors against which
caps protect their RNAs is a 5′-phosphate dependent 5′→3′ exoribonuclease discovered by
Audrey Stevens by purifying it from yeast (Stevens 1980). Later named XRN1 (for
exoribonuclease 1) (Larimer and Stevens 1990), Xrn1 was shown to be abundant in cells
(Ghaemmaghami et al. 2003) and to processively degrade its substrates (Stevens 2001).
In budding yeast, the poly-adenylate tail (poly(A) tail) is a master regulator of mRNA
stability. Typically, RNA decay is initiated via deadenylation (Decker and Parker 1993) by the
Ccr4/Not complex (Tucker et al. 2001) or the Pan2/Pan3 complex (Boeck et al. 1996; Brown and
Sachs 1998). Deadenylation is a highly regulated process, with several RNA-binding proteins
known as actors in this regulation (Parker 2012). The principal regulator of deadenylation is the
Poly(A)-Binding protein, Pab1, which inhibits deadenylation by the Ccr4 complex when bound
to poly(A) tails (Caponigro and Parker 1995).
Once an RNA is deadenylated, the most common outcome is that the decapping complex
(Dcp1/Dcp2) hydrolyzes the 7-methylguanosine cap of the RNA to produce a 5′-phosphate,
exposing the 5′-terminus to degradation by Xrn1 (Hsu and Stevens 1993). Alternatively,
decapped RNAs can also be degraded in a 3′→5′ scheme by the cytoplasmic RNA exosome
(Anderson and Parker 1998). Comparatively, the 5′→3′ decay pathway, as executed by Xrn1,
appears to be more consequential to exponentially-growing cells.
39
Table 1.1: Exoribonucleases and their terminus requirements
Name Direction Species Active Termini
Inhibitory Termini
Reference
Xrn1/Xrn4 5′→3′ Eukaryotes 5′-PO4 5′-OH (Stevens 2001)
(Nagarajan et al. 2013)
Xrn2/Rat1 5′→3′ Eukaryotes 5′-PO4 5′-OH (Johnson 1997) (Geerlings et al. 2000)
Dxo1 5′→3′ Yeast 5′-PO4 5′-OH (Chang et al. 2012)
Dis3/Rrp44 3′→5′ Eukaryotes 2′,3′-OH,
2′,3′-cP N/A
(Meaux and van Hoof 2006)
Rrp6 3′→5′ Eukaryotes 2′,3′-OH 3′-PO4 (Burkard and Butler 2000)
Usb1 3′→5′ Yeast 2′,3′-OH
2′,3′-cP 3′-PO4 (Didychuk et al. 2017)
PNPase 3′→5′ Bacteria 2′,3′-OH 2′-, 3′-PO4 (Munir et al. 2018a)
40
Cells defective for cytoplasmic 5′→3′ RNA decay (e.g. xrn1∆, dcp2∆) exhibit slow growth or are
inviable (depending on genetic background), whereas yeast with mutations in 3′→5′ decay
factors (e.g. ski2∆, ski3∆) (discussed further below) exhibit a very modest growth phenotype
(Giaever et al. 2002). Interestingly, cells lacking any combination of both 5′→3′ decay and 3′→5′
decay are inviable (Anderson and Parker 1998), indicating that the decay pathways can
compensate for one another, leading to a synthetic lethal genetic interaction when ablated
simultaneously. A small proportion of the RNA in a yeast cells may decapped independent of
deadenylation (Muhlrad et al. 1995), and some RNAs may infrequently be subject to
endonucleolytic cleavage as a means of regulated decay (unpublished data, Y. Harigaya & R.
Parker) (Parker 2012), but the overarching trend is that decay is regulated primarily at the
deadenylation stage, followed by decapping, and then finally by exonucleolytic degradation (Fig.
1.6).
Beyond Xrn1, two alternative 5′-phosphate-dependent 5′→3′ decay pathways exist:
Rat1/Xrn2 and Dxo1. Rat1 (alias: Xrn2) is an essential nuclear exonuclease found by a screen for
mutants unable to export poly(A) RNA into the cytoplasm (giving it the name Ribonucleic Acid
Trafficking 1) (Amberg et al. 1992). Overexpression of wild-type Rat1 does not completely rescue
cells from the growth and RNA decay defects imparted by deleting Xrn1, but Rat1 mutants that
increase localization to the cytoplasm fully rescue the xrn1∆ phenotype (Poole and Stevens
1995). Conversely, Xrn1 tagged with a nuclear localization signal can complement the rat1-1
conditional mutant at non-permissive temperatures (Johnson 1997). Thus, Xrn1 and Rat1 are
functionally similar exoribonucleases that have become compartmentalized in eukaryotes.
41
Figure 1.6: Typical mRNA decay in the cytoplasm by Xrn1 and the exosome.
(1) The Ccr4/Not complex or Pan2/Pan3 complex deadenylate the poly(A) tail of the mRNA. (2) Dcp1 and Dcp2 are induced by to short poly(A) tail to de-cap the mRNA, producing a 5′-PO4 terminus. (3) Xrn1 is the primary decay factor in the cytoplasm, and it rapidly and processively degrades 5′-PO4 RNAs in the 5′→3′ direction. (4) The exosome is the secondary decay factor of the cytoplasm and is composed of a complex of structural and regulatory proteins, as well as the exoribonuclease Rrp44. The exosome degrades deadenylated mRNAs in the 3′→5′ direction. (Parker 2012)
42
A third enzyme was discovered to have distributive 5′→3′ exoribonuclease activity during
structural studies that were focused on decapping (Chang et al. 2012). Thus, Decapping
eXOnuclease 1 (Dxo1) was named for its combined nuclease activities on methylated and
unmethylated 7-methylguanosine caps and 5′-phosphorylated RNA. Dxo1 is structurally similar
to Rai1 (Rat1 Interacting protein 1), a nuclear decapping enzyme with affinity toward
unmethylated 7-methylguanosine caps, and Dom3Z, the mammalian homolog of Rai1 (Xue et al.
2000).
3′→5′ RNA degradation
A second RNA decay pathway executed by the cytoplasmic exosome in yeast can degrade
RNA from the opposite terminus, proceeding in a 3′→5′ direction (Fig. 1.6). A generalized 3′→5
decay pathway was proposed because mutants of components required for 5′→3′ decay are
viable and continue to degrade most RNAs, albeit at slower rates (Hsu and Stevens 1993;
Muhlrad et al. 1994; 1995; Beelman et al. 1996), and mutations of both the 5′→3′ and 3′→5′
pathways are synthetic lethal (Johnson and Kolodner 1995). The exosome was identified as a
multi-protein complex via an immunoprecipitation experiment performed on Rrp4 (Ribosomal
RNA Processing 4), a protein involved in 3′→5′ trimming of the 5.8S rRNA (Mitchell et al. 1996).
Mass spectrometry identified proteins associated with Rrp4, Rrp41 (Ski6), Rrp42, Rrp43, and
Rrp44 (Dis3), all required for 3′→5′ processing of the 5.8S rRNA, and the complex was dubbed
the exosome. In vitro characterization demonstrated that: Rrp4 is a distributive RNA hydrolase;
Rrp44 is a processive RNA hydrolase; and Ski6 (Rrp41) is a processive phosphorolytic enzyme
(Mitchell et al. 1997). Further structural studies revealed the exosome complex to be composed
of 10 key proteins: the exonuclease and endonuclease domain-containing Dis3 (Rrp44) protein,
homologous to E. coli RNase R (vacB), and a nine-member ring structure formed by six RNase
PH-like proteins (Rrp41, Rrp45, Rrp46, Rrp43, Mtr3, and Rrp42) and three small RNA-binding
43
proteins (Rrp4, Rrp40, and Csl4) analogous to bacterial PNPase (Mian 1997; Cheng et al. 1998;
Liu et al. 2006).
The activity of the exosome is largely determined by its association with regulatory
substrate recognition complexes (Araki et al. 2001). Beyond rRNA processing, subsequent
studies indicated that the exosome is present in the cytoplasm and functions as a general
pathway for degradation of mRNAs. In addition to Ski6, Anderson and Parker demonstrated that
Ski2, Ski8, and the adaptor protein Ski7 function as part of the cytoplasmic exosome, performing
both routine turnover of RNA, as well as degradation of aberrant mRNA, like in non-stop and no-
go mRNA decay (Anderson and Parker 1998; Wang et al. 2005). In contrast to the principal
enzymes of 5′→3′ decay, there is mixed evidence for terminus requirements for the RNA
exosome. Stacie Meaux and Ambro van Hoof showed, using self-cleaving ribozymes, that the
yeast cytoplasmic exosome degraded 2′,3′-cP just as well as 2′,3′-cis-diol terminated RNAs
(Meaux and van Hoof 2006). However, 3′-PO4 RNAs appear to be inhibitory to Rrp6, the catalytic
subunit of the nuclear exosome, similar to how DNA (with its 2′-H) may be inhibitory (Burkard
and Butler 2000). Usb1 (U Six Biogenesis 1), the 3′→5′ exoribonuclease in yeast essential for
U6 snRNA biogenesis, is also inhibited by 3′-PO4 (Didychuk et al. 2017). As Usb1 degrades the
pre-U6 snRNA, which starts off with a 2′,3′-cis-diol, Usb1 generates 2′,3′-cP with each
exonucleolytic step. Interestingly, Usb1 has a second activity of cyclic-phosphate hydrolysis to
a 3′-PO4, which inhibits further exonuclease activity, thereby preventing Usb1 from degrading
too much RNA off the 3′-end of U6 snRNA.
In addition to terminus chemistry requirements, a recent discovery provided an example
of a 3′→5′ exoribonuclease (phosphorylase) that is blocked by a non-canonical internal linkage.
Dissatisfied with the ability to resolve 2′-phosphorylated RNA 15-mers on polyacrylamide gels,
Stewart Shuman discovered that PNPase (PolyNucleotide Phosphorylase) from Mycobacterium
smegmatis is halted site-specifically by internal 2′-PO4 (Munir et al. 2018a).
44
Kinase-Mediated Decay
A major caveat of 5′→3′ RNA decay is that all three known exonucleases that degrade
5′→3′, Xrn1, Rat1, and Dxo1, require 5′-phosphates of their substrates. This phenomenon is
consequential because many metal-ion-independent RNA incision events produce 5′-hydroxyl
RNAs: Sen2/Sen34 cleavage of tRNAs (Knapp et al. 1979); Ire1 cleavage of HAC1 mRNA
(Gonzalez et al. 1999); Rny1 and angiogenin (Thompson and Parker 2009a); no-go mRNA decay
(Navickas et al. 2018); intrinsic cleavage of RNA. These products of cleavage are effectively
immune to 5′→3′ decay because the enzymes of 5′→3′ decay do not recognize the 5′–OH RNAs
as substrates. Any evidence of their 5′→3′ decay would necessarily imply a 5′-phosphorylation
event, as would be provided by a 5′-RNA kinase, hence Kinase-Mediated Decay (KMD).
Examples of KMD already exist: tRNA introns, pre-rRNA, no-go mRNA decay 3′-fragments, and
T4 bacteriophage mRNA.
KMD of tRNA introns
Introns from tRNAs, once cleaved out from transcripts, undergo kinase-mediated decay.
The catalytic subunits of the SEN complex, Sen2 and Sen34, catalyze metal-ion-independent
cleavage, and release 5′-OH and 2′,3′-cP products (Knapp et al. 1979). Anita Hopper uncovered
an intron retention defect for tRNAIle(UAU) in xrn1∆ yeast via an unbiased reverse-genetic screen,
indicating that the 5′-OH tRNA introns are converted to 5′-PO4 in order to be degraded by Xrn1
(Fig. 1.7A) (Wu et al. 2015), A follow-up paper showed that, in addition to tRNAIle(UAU), liberated
introns from tRNALeu(CAA), tRNALys(UUU), tRNATrp(CCA), and tRNAPro(UGG) accumulated in the
absence of Xrn1 (Wu and Hopper 2014). Furthermore, using the previously described conditional
mutants of the tRNA ligase rlg1-4 and rlg1-10 (Phizicky et al. 1992), Hopper and Wu showed that
ligase-mutant yeast grown at the non-permissive temperature also accumulated liberated tRNA
introns.
45
Figure 1.7: Examples of Kinase-Mediated Decay (KMD)
A: The splicing endonuclease (SEN) complex cleaves the intron out of the pre-tRNA, releasing it as a 5′-OH. The kinase domain of Trl1 phosphorylates the 5′-terminus, yielding a 5′-PO4, which is a substrate or Xrn1. (Wu et al. 2015)
B: Las1 cleaves at the C2 site in internal transcribed spacer 2 (ITS2) of the 27SB pre-rRNA, releasing a 26S pre-rRNA with a 5′-OH terminus. Grc3 RNA 5′-kinase phosphorylates the 5′-end, yielding a 5′-PO4 substrate of Rat1/Xrn2. (Gasse et al. 2015)
C: No-go mRNA decay (NGD) releases cleavage fragments with a 5′-OH by an unknown nuclease/mechanism. Trl1 phosphorylates the 5′-terminus of the 3′-cleavage fragment, yielding a 5′-PO4, which is a substrate or Xrn1. (Navickas et al. 2018)
D: During T4 phage infection of E. coli, the regulatory nuclease RegB cleaves the Shine-Dalgarno sequence of “early” RNAs during the transition to “late” gene expression. The cleaved mRNAs are released with 5′-OH termini and require phosphorylation by T4 polynucleotide kinase (PNK) for recognition and decay by host RNase E and RNase G. (Durand et al. 2012)
46
The experiment elegantly demonstrated that the tRNA introns have to be phosphorylated by the
5′-RNA kinase domain of yeast tRNA ligase (Trl1) prior to degradation by Xrn1, constituting an
example of kinase-mediated decay (Fig. 1.2).
KMD of rRNA processing intermediates
A second example of the kinase-mediated decay phenomenon was discovered at work
during ribosomal RNA (rRNA) processing. During the production of 25S and 5.8S rRNAs, their
tandem polI transcript has to be cleaved and trimmed to form rRNAs of the mature length. Lisa
Gasse and Ed Hurt identified the endoribonuclease that cleaves at the C2 position in ITS2 of the
27SBS pre-rRNA as Las1, a metal-ion-independent endonuclease. Once Las1 cleaves ITS2 at
the C2 site, yielding a 26S rRNA with a 5′-OH, the 5′–P exonuclease, Rat1, “trims” the 26S pre-
rRNA. Gasse et al. showed that the polynucleotide kinase Grc3 phosphorylates the 5′-terminus
of the 26S pre-rRNA to allow Rat1 to trim ribonucleotides off the 5′-end (Fig. 1.7B) (Gasse et al.
2015). This sequential generation of 5′-OH RNA, phosphorylation to a 5′-PO4, and
degradation/chew-back by a 5′-PO4 RNase (Rat1/Xrn2) constitutes kinase mediated decay as a
mechanism of rRNA maturation.
The factors that carry out this example of KMD are conserved in human pre-rRNA
processing. The LAS1L endonuclease cleaves both the C1′ site and 30 nt 3′ of the E′ site of the
ITS2 spacer in 45S pre-rRNA, producing the 6SS and 26S pre-rRNAs (Schillewaert et al. 2012).
Similar to Grc3, human Nol9 is the nucleolar RNA 5′-kinase (Heindl and Martinez 2010) and likely
phosphorylates the 5′-terminus of the 26S pre-rRNA, preparing it for trimming by XRN2, the
human 5′-phosohate-dependent 5′→3′ exoribonuclease homologous to Rat1 from budding
yeast. Thus, kinase-mediated decay is likely a conserved mechanism of pre-rRNA processing in
humans.
47
KMD of 3′-fragments of no-go mRNA decay (NGD)
The coding quality of mRNA is controlled through a family of processes called co-
translational mRNA surveillance, an umbrella term for a series of processes that monitor
ribosome progress along the open reading frame to detect mRNAs that are defective in
translation. Adaptor proteins interact with the ribosome to detect aberrant mRNAs and direct
them toward RNA decay pathways. No-go mRNA decay (NGD) is one of these co-translational
mRNA surveillance processes that detects and rectifies stalled ribosomes on messages (Doma
and Parker 2006). When ribosomes encounter strong stalls in translation elongation, such as
stem-loops, rare codons, poly-positively-charged amino acids, NGD is activated and the mRNA
is cleaved, releasing 5′ and 3′ RNA products. The cytoplasmic exosome degrades the 5′-product
of endonucleolytic cleavage, and Xrn1 degrades the 3′-product.
While the endoribonuclease that catalyzes the cleavage of no-go decay mRNAs remains
unknown, the cleavage site reliably occurs 5′ of the elements that causes the ribosome(s) to
stall. In a 2018 bioRχiv pre-print, Navickas, et al. demonstrate that the products of cleavage are
generated with 5′-OH termini (Navickas et al. 2018). With Xrn1 as the required decay factor for
degradation of 3′-NGD fragments (and Dxo1 to a much lesser degree), a 5′-phosphorylation step
would be a required intermediate step. Using the temperature-sensitive allele of Trl1 (rlg1-4),
Navickas, et al. showed that a phosphorylation event was indeed required prior to Xrn1-mediated
decay of the no-go decay product (Fig. 1.7C).
KMD of T4 bacteriophage mRNA
Bacteria have 3′→5′ and 5′→3′ decay programs for RNA, but these exonucleases typically
do not initiate decay (Belasco 2010). Instead, RNA decay typically starts with an endonucleolytic
cleavage event that generates substrates for exonucleases in the cell. One example of this decay
mode occurs via a cascading series of endonucleolytic cleavages that generate short substrates
for 3′→5′ exoribonucleases. To initiate decay, the bacterial RNA pyrophosphohydrolase (RppH)
48
converts the 5′-triphosphate terminus to a 5′-phosphate (Celesnik et al. 2007). Next, RNases E
and G recognize the 5′-PO4 terminus and cleave downstream, generating yet another (shorter)
5′-PO4 RNA that serves as a substrate for RNases E and G until the remaining RNA is sufficiently
short (Mackie 1998; Spickler et al. 2001). The fragmented RNA products of RNases E and G are
degraded by the bacterial 3′→5′ exoribonucleases RNase R, RNase II, and polynucleotide
phosphorylase (PNPase) (Andrade et al. 2009).
During infection of E. coil, T4 Bacteriophage must coordinate the expression of early and
late genes. The phage uses a virally-encoded endoribonuclease, RegB, to site-specifically cleave
at the Shine-Dalgarno sequence of early genes (Uzan et al. 1988; Sanson et al. 2000). RegB is a
metal-ion-independent endoribonuclease and so generates 5′-OH products of cleavage (Saïda
et al. 2003). However, with RNases E and G being 5′-phosphate-dependent decay factors, an
intervening phosphorylation step would be necessary for them to degrade early mRNAs on the
phage’s behalf. T4 phage supply a polynucleotide kinase (PNK) that phosphorylates these 5′-
OH termini and permits RNases E and G to degrade the early genes after RegB cleavage (Fig.
1.7D) (Durand et al. 2012). This RegB-directed KMD mechanism presents an interesting example
of KMD in prokaryotes, the only example known to date. The example is also remarkable
because of the cooperation the T4 phage exhibits by adapting its viral mRNA regulatory system
to the endonuclease and kinase of the general RNA decay machinery of the bacterium.
Toward mutants of RNA repair
To date, investigations of the role of RNA repair have occurred through temperature-
sensitive (conditional) mutants of the tRNA ligase, TRL1/RNL1, requiring shifting the yeast to the
non-permissive temperature for experiments in the absence of ligase functions (Phizicky et al.
1992), complicating the study of substrates of RNA repair. Furthermore, conditional mutants of
49
the 2′-phosphotransferase, TPT1, obscure detection of the products of ligation, as they are
marked by a residual 2′-phosphate at the ligation junction (Spinelli et al. 1997).
An outright genetic deletion of the genes TRL1 or TPT1 is not possible because they are
essential, but those genotypes would be useful for studying the substrates and products of RNA
repair, as well as any additional contributions the enzymes make to RNA processing, such as
kinase mediated decay. In Chapter II, I present my studies on performing a genetic bypass of
the essential genes TRL1 and TPT1 using a construct to express “pre-spliced” tRNAs, showing
that the only essential function of Trl1 and Tpt1 is to splice intron-containing tRNAs. I go on to
use the genetic bypass, called the “10x tRNA Block,” in Chapter III, to describe how RNA repair
regulates the unfolded protein response (UPR) downstream of Ire1-mediated cleavage, using
KMD to effectively suppress the UPR under normal conditions and to facilitate activation of the
UPR upon endoplasmic reticulum stress. In Chapter IV, I propose using the findings of these
studies to discover substrates and products of RNA repair in both fungi and metazoans and
stabilize RNAs with 5′-terminal 2′-phosphates.
50
CHAPTER II
II GENETIC BYPASS OF ESSENTIAL RNA REPAIR ENZYMES IN BUDDING YEAST1
Abstract
RNA repair enzymes catalyze rejoining of an RNA molecule after cleavage of
phosphodiester linkages. RNA repair in budding yeast is catalyzed by two separate enzymes
that process tRNA exons during their splicing and HAC1 mRNA exons during activation of the
unfolded protein response. The RNA ligase Trl1 joins 2′,3′-cyclic phosphate and 5′-hydroxyl
RNA fragments, creating a new phosphodiester linkage with a 2′-phosphate at the junction. The
2′-phosphate is subsequently removed by the 2′-phosphotransferase Tpt1, which catalyzes
phosphate transfer to NAD+, producing nicotinamide and a unique ADP-ribose metabolite. I
bypassed the essential functions of TRL1 and TPT1 in budding yeast by expressing “pre-
spliced”/intronless versions of the ten normally intron-containing tRNAs, indicating this repair
pathway does not have additional essential functions. Consistent with previous studies,
expression of intronless tRNAs failed to rescue the growth of cells with deletions in components
of the SEN complex, implying an additional essential role for the splicing endonuclease. The
trl1∆ and tpt1∆ mutants accumulate tRNA and HAC1 splicing intermediates indicative of specific
RNA repair defects and are hypersensitive to drugs that inhibit translation. As expected, failure
to induce the unfolded protein response in trl1∆ cells grown with tunicamycin is lethal owing to
their inability to ligate HAC1 after its cleavage by Ire1. In contrast, tpt1∆ mutants grow in the
presence of tunicamycin despite reduced accumulation of spliced HAC1 mRNA. Finally, I
optimized a PCR-based method to detect RNA 2′-phosphate modifications and show that they
1Published with permission under a Creative Commons License (Attribution-NonCommercial 4.0 International) from RNA: PD Cherry, LK White, K York, and JR Hesselberth. Genetic bypass of essential RNA repair enzymes in budding yeast. 2018. 24: 313-323. doi: 10.1261/rna.061788.117
51
are present on ligated HAC1 mRNA. These RNA repair mutants enable new studies of the role
of RNA repair in cellular physiology.
Introduction
RNA repair enzymes catalyze rejoining of an RNA molecule after cleavage of
phosphodiester linkages. RNA repair is carried out by enzymes that prepare the RNA termini for
ligation, ligate the termini, and “clean-up” the ligated product when required. RNA repair systems
are present in all domains of life and catalyze rejoining of the 5′-hydroxyl and 2′,3′-cyclic
phosphate products of cleavage. RNA repair catalyzes the maturation of endogenous tRNAs and
mRNAs (Schwer et al. 2004), and can also counteract the action of endonucleases during inter-
organismal conflicts (Burroughs and Aravind 2016; Amitsur et al. 1987; Nandakumar et al. 2008).
E. coli RtcB can repair ribosomal RNA cleaved during stress by the endonuclease MazF, thereby
reversing ribosomal heterogeneity and restoring translational activity to MazF-processed
ribosomes (Temmel et al. 2016). The Pnkp–Hen1 RNA repair complex, which is present in more
than 250 bacterial species, combines the enzymatic activities of the bacteriophage T4 RNA
repair system with the Hen1 methyltransferase, which installs a 2′-O-methyl group to “immunize”
repaired RNA molecules against future cleavage by the same ribotoxin (Wang et al. 2015). In
light of these and other recent developments in our understanding of the functional and genetic
diversity of RNA ligases (Burroughs and Aravind 2016; Kosmaczewski et al. 2015), I developed
a genetic bypass of the Saccharomyces cerevisiae RNA repair system, enabling direct study of
the roles of RNA repair proteins in budding yeast.
In budding yeast, S. cerevisiae, after tRNA intron excision, ligation of the exons is carried
out in successive steps (Phizicky et al. 1986). First, the splicing endonuclease (SEN) complex
cleaves the intron at two sites, creating two exon products, one with a 2′,3′-cyclic phosphate
terminus and one with a 5′-hydroxyl terminus, and an intron product with 5′-hydroxyl and 2′,3′-
52
cyclic phosphate termini that is degraded (Wu and Hopper 2014). In a second step, the two
exons are ligated by the RNA ligase Trl1. Trl1 joins substrates with 2′,3′-cyclic phosphate and
5′-hydroxyl termini via: (i) conversion of the 2′,3′-cyclic phosphate to a 2′-PO4/3′-hydroxyl; (ii)
phosphorylation of the 5′-OH; (iii) adenylylation of the 5′-PO4 and iv) nucleophilic attack on the
adenylate by the 3′-OH, producing AMP and a new 5′→3′ phosphodiester linkage with a 2′-PO4
(Greer et al. 1983). The 2′-PO4 left at the ligation junction is removed by the NAD+-dependent 2′-
phosphotransferase Tpt1, creating a canonical 5′→3′ phosphodiester linkage (McCraith and
Phizicky 1990; Culver et al. 1993) (Fig. 2.1A).
Trl1 has a second known role in the cell: activating the unfolded protein response (UPR).
Trl1 ligates the two exons of the HAC1 mRNA after cleavage by Ire1, activating the UPR
(Gonzalez et al. 1999). Ire1 excises an intron from the HAC1 pre-mRNA, and Trl1 subsequently
ligates the HAC1 exons together, enabling its translation into a transcription factor that localizes
to the nucleus and drives transcription of hundreds of stress response genes (Sidrauski et al.
1996). Yeast cells that lack Trl1 and Tpt1 and that express RNA repair enzymes from T4
bacteriophage are viable, but they exhibit low-fidelity HAC1 mRNA cleavage and ligation,
suggesting the 2′- PO4/3′-hydroxyl terminus produced by the cyclic phosphodiesterase domain
of Trl1 directs precise ligation (Schwer et al. 2004). Trpt1, the mammalian 2′-
phosphotransferase, was shown to be dispensable for UPR activation in mammals (Harding et
al. 2008). However, subsequent studies showed that the HSPC117/RtcB RNA ligase—which
does not create 2′-PO4 ligation products (Chakravarty et al. 2012)—activates the mammalian
UPR (Lu et al. 2014), explaining why Trpt1 2′-phosphotransferase activity is dispensable
(Harding et al. 2008). The role of Tpt1 in budding yeast in the UPR has not been previously
explored.
53
Using a genetic bypass strategy, RNA repair was previously shown to be essential for
growth only for fulfilling the need of C. elegans (Kosmaczewski et al. 2014) and trypanosomes
(Lopes et al. 2016) to splice tRNAs. Using a similar strategy, I designed and tested a genetic
bypass for deletion of the essential RNA repair enzymes Trl1 and Tpt1 in budding yeast and
show that rescued trl1∆ and tpt1∆ cells (the “RNA repair mutants”) have unique phenotypes for
both tRNA splicing and HAC1 mRNA splicing during the unfolded protein response.
Materials & Methods
General Methods
Saccharomyces cerevisiae W303 strains were cultured in YPD and synthetic “drop-out”
media (Sherman 2002) for experimental cultures and for plasmid selection, respectively. Single-
copy URA3 SEN plasmids were created by recombining SEN-containing genomic fragments
from the yeast tiling collection (Jones et al. 2008) into Advanced Gateway plasmids (Jones et al.
2008) via Gateway LR reactions (Life Technologies). Plasmid counterselection was performed on
synthetic complete solid media containing 5-fluoroorotic acid (FOA, US Biologicals) (1 mg/mL).
Yeast transformations were performed using a Lithium Acetate PEG-3350 Sheared Salmon
Sperm protocol (Gietz and Schiestl 2007). Genotypes of yeast were confirmed by PCR using
forward and reverse primers flanking the disrupted locus and outward-facing primers within the
disruption cassette. Success of FOA plasmid counterselections was confirmed using RT-PCR
for the shuffled gene in DNased total RNA isolated from shuffled strains and positive controls.
Experiments with tunicamycin (Sigma) treatment were performed at final concentrations of
0.08 µg/mL in solid media and at 2.5 µg/mL in liquid media, with negative controls using DMSO
(solvent). Cultures were incubated at 30°C unless otherwise indicated. UPR inductions ±
tunicamycin were carried out for 2 hours during exponential phase growth. For growth assays,
cells were grown in YPD overnight and liquid cultures were normalized to OD600 = 0.2 before plating
54
tenfold serial dilutions on indicated media. For translation inhibition assays, dilutions were
spotted onto YPD agar supplemented with 2 µg/mL anisomycin (Sigma), 100 ng/mL hygromycin
B (Invivogen), or 100 ng/mL cycloheximide (Sigma).
Northern blotting
Oligonucleotide probes (IDT) were designed to hybridize with RNA species to be analyzed
(Table 2.1). Oligonucleotide probes were 5′-radiolabeled with [γ-32P]-ATP (Perkin Elmer) using
T4 PNK (Enzymatics), and excess unincorporated label was removed with G-25 Sephadex (GE)
spin columns. Probes were heated to 100°C and diluted into 10 mL ULTRAhyb Oligo
hybridization buffer (Life Technologies). Total RNA (normalized to SCR1 loading) was
electrophoresed, along with RiboRuler Low Range RNA Ladder (Thermo) and ss10 ssDNA ladder
(Simplex Sciences) on pre-cast 10% polyacrylamide urea TBE Novex gels (Life Technologies) at
160 V for 90 minutes. Gels were stained with SybrGold Nucleic Acid Stain (Life Technologies),
imaged, and electrotransferred with a Genie electroblotter (Idea Scientific) onto Hybond N+ nylon
membranes (GE) at 19 V for 60 minutes submerged in 1x TBE. RNA was crosslinked to the
membranes with 0.12 joules of 254 nm UV light (Stratalinker 1800, Stratagene). Nylon
membranes were blocked with 10 mL ULTRAhyb Oligo for 1 hour at 42°C and hybridized with
probe at 42°C for 18 hours. Blots were washed at 42°C for 30 min each with buffer (2X SSC,
0.1% SDS) and were developed by phosphorimaging (GE, Molecular Dynamics).
Detection of 2′-Phosphate linkages by RT-PCR
A reverse primer that anneals directly downstream of the 2′-phosphate site (Table 2.1,
HAC1-R0 2-P) was designed so that the first deoxyribonucleotide incorporated by the reverse
transcriptase is complementary to the 2′-phosphorylated ribonucleotide. Total RNA (2 µg) was
DNased (Turbo DNase Kit, Ambion), treated ± with calf intestinal phosphatase (NEB), and acid-
phenol extracted. DNased and ±CIP-ed total RNA was denatured at 65°C for 5 minutes with 10
pmol of the reverse primer and then transferred to ice. The concentration of dNTPs in reverse
55
transcription reactions was reduced from 500 µM to 1 µM to inhibit reverse transcriptase
(SuperScript III, Life Technologies, lot #1826824) at sites of 2′-phosphorylation (Fig. 4E,
schematic). PCR was performed with the same reverse primer and a forward primer that anneals
to the 5′-exon of HAC1; 28 cycles were performed on low dNTP reactions and 22 cycles were
performed on RT reactions with normal dNTP concentrations. Amplified DNA was
electrophoresed in a 2% agarose TBE 1X SybrGold (Life Technologies) gel and imaged.
Expression vector for intronless tRNAs
I designed a sequence to express ten pre-spliced tRNAs using the SUP4 promoter and
RPR1 terminator, respectively (Good and Engelke 1994). The full sequence is available at
Addgene, record 70125. Two restriction sites at the 5′- and 3′-ends of the sequence for SacI
and ClaI are underlined. Ten tRNA genes are encoded with an intervening spacer (“CTTTGT”)
derived from a dicistronic tRNA (Engelke et al. 1985). From 5′ to 3′, the tRNAs are: Phe(GAA),
Leu(CAA), Lys(UUU), Ser(GCU), Ile(UAU), Trp(CCA), Tyr(GUA), Pro(UGG), Ser(CGA) and
Leu(UAG). The pol III promoter contains sequence elements within the SUP4 Tyr(GUA) tRNA
(Good and Engelke 1994), and thus tRNATyr(GUA) occurs twice in the construct. The sequence
was synthesized as a gBlock from IDT and recombined into pDONR221 in a Gateway BP reaction
(Life Technologies). The product plasmid (pDONR221-10X-tRNA) was recombined by a Gateway
LR reaction into pAG424-ccdB (created by removing the GPD promoter from pAG424-GPD-
ccdB (Alberti et al. 2007), which has the TRP1 selectable marker and a 2µ origin for high-copy
number per cell (Chan et al. 2013). The pDONR221-10X-tRNA, pAG424-ccdB (empty vector) and
pAG424-10X-tRNA plasmids are available from Addgene (plasmids 70125, 70124, and 70123).
HAC1 epitope tagging and western blotting
Oligonucleotide probes HAC1-pML104-plus and HAC1-pML104-minus (IDT) (Table 1) form
an insert that was ligated into the Cas9 and sgRNA yeast expression plasmid pML104 (Addgene
entry 67638) (Laughery et al. 2015). Yeast strains were simultaneously transformed with pML104-
56
HAC1-ct and FLAG donor DNA with homology to the HAC1 gene for homology-directed repair.
Co-transformed yeast were selected on uracil drop-out media (U.S. Biologicals) and confirmed
to have an in-frame FLAG epitope tag by Sanger sequencing. Cell lysates were analyzed by
immunoblotting using an anti-FLAG M2 antibody (Sigma) and secondary anti-mouse HRP-
conjugated antibodies (ThermoFisher Scientific) with Enhanced Chemiluminescence detection
(Promega). Loading controls included Ponceau staining of the membrane and anti-GAPDH
antibody (UBP-Bio).
57
Table 2.1: Oligonucleotide sequences.
Oligonucleotide Name Oligonucleotide sequence (5′→3′)
tRNA Ile UAU 5-Exon N Probe TATAAGCACGAAGCTCTAACCACTGAGCTACACGAGC
tRNA Ile UAU Intron N Probe CGTTGCTTTTAAAGGCCTGTTTGAAAGGTCTTTGGCACAGAAACTT
Phe-GAA-5-Exon-Probe CTTCAGTCTGGCGCTCTCCCAACTGAGCTAAATCCGC
Leu-CAA-5-Exon-Probe CTTGAATCAGGCGCCTTAGACCGCTCGGCCAAACAACC
Pro-UGG-5-Exon-Probe CCCAAAGCGAGAATCATACCACTAGACCACACGCCC
Tyr-GUA-5-Exon-Probe TTACAGTCTTGCGCCTTAAACCAACTTGGCTACCGAGAG
HAC1-F RT-PCR ACCTGCCGTAGACAACAACAAT
HAC1-R RT-PCR AAAACCCACCAACAGCGATAAT
HAC1-F 2-P RT-PCR ATGGGAGCTGCAGATGTTTAAG
HAC1-R0 2-P GAATTCAAACCTGACTGCGCTT
TPT1-F RT-PCR TGTTCAGGTCGCTCAATAATGT
TPT1-R RT-PCR TCTTTTCGAGCGGTATGTTTCT
TRL1-F2 RT-PCR GTGGCAGAATATTGCGATGA
TRL1-R2 RT-PCR ATCCTCCAAGGTGTTCGATG
KAR2-QPCR-F AAGACAAGCCACCAAGGATG
KAR2-QPCR-R AGTGGCTTGGACTTCGAAAA
PGK1-QPCR-F TCTTAGGTGGTGCCAAAGGTT
PGK1-QPCR-R GCCTTGTCGAAGATGGAGTC
HAC1-pML104-plus GATCTTCATGAAGACAATCGCAAGGTTTTAGAGCTAG
HAC1-pML104-minus CTAGCTCTAAAACCTTGCGATTGTCTTCATGAA
58
Table 2.2: Strain numbers and genotypes.
All strains are background W303 (MATa {leu2-3,112 trp1-1 can1-100 ura3-1 ade2-1 his3-11,15}).
Strain ID Genotype Source
YJH829 tpt1∆::LEU2 (TPT1 CEN ARS URA3) B. Schwer
YJH830 tpt1∆::LEU2 (TPT1 CEN ARS URA3) (pAG424-ccdB)
YJH832 tpt1∆::LEU2 (TPT1 CEN ARS URA3) (pAG424-10x-tRNA)
YJH834 tpt1∆::LEU2 (pAG424-10x-tRNA)
YJH681 trl1∆::KanMX (TRL1 CEN URA3) B. Schwer
YJH708 trl1∆::KanMX (TRL1 CEN URA3) (pAG424)
YJH709 trl1∆::KanMX (TRL1 CEN URA3) (pAG424-10x-tRNA)
YJH835 trl1∆::KanMX (pAG424-10x-tRNA)
YJH836 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3)
YJH837 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3) (SEN2 LEU2 2µ)
YJH838 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3) (SEN15 LEU2 2µ)
YJH839 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3) (SEN34 LEU2 2µ)
YJH840 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3) (SEN54 LEU2 2µ)
YJH841 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3)
YJH842 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3) (SEN2 LEU2 2µ)
YJH843 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3) (SEN15 LEU2 2µ)
YJH844 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3) (SEN34 LEU2 2µ)
YJH845 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3) (SEN54 LEU2 2µ)
YJH846 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3)
YJH847 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3) (SEN2 LEU2 2µ)
YJH848 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3) (SEN15 LEU2 2µ)
YJH849 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3) (SEN34 LEU2 2µ)
YJH850 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3) (SEN54 LEU2 2µ)
59
YJH851 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3)
YJH852 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3) (SEN2 LEU2 2µ)
YJH853 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3) (SEN15 LEU2 2µ)
YJH854 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3) (SEN34 LEU2 2µ)
YJH855 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3) (SEN54 LEU2 2µ)
YJH856 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3) (pAG424-ccdB)
YJH857 sen2∆::KanMX (SEN2 tiling block CEN ARS URA3) (pAG424-10x-tRNA)
YJH858 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3) (pAG424-ccdB)
YJH859 sen15∆::KanMX (SEN15 tiling block CEN ARS URA3) (pAG424-10x-tRNA)
YJH860 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3) (pAG424-ccdB)
YJH861 sen34∆::KanMX (SEN34 tiling block CEN ARS URA3) (pAG424-10x-tRNA)
YJH862 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3) (pAG424-ccdB)
YJH863 sen54∆::KanMX (SEN54 tiling block CEN ARS URA3) (pAG424-10x-tRNA)
YJH864 HAC1-C-terminal-3x-FLAG
YJH865 trl1∆::KanMX HAC1-C-terminal-3x-FLAG (pAG424-10x-tRNA)
YJH866 tpt1∆::LEU2 HAC1-C-terminal-3x-FLAG (pAG424-10x-tRNA)
60
Results & Discussion
Genetic bypass of essential RNA repair genes in budding yeast
Ten S. cerevisiae tRNA isodecoders are encoded with introns (Chan and Lowe 2009),
which must be accurately processed for cells to faithfully translate messenger RNA (Hopper
2013). I adapted a strategy first documented in C. elegans (Kosmaczewski et al. 2014) to express
these ten tRNAs in “pre-spliced” form (Fig. 1B, the 10x-tRNA plasmid) and found that expression
of these intronless tRNAs rescues the growth of cells with deletions in the essential genes TRL1
and TPT1 (Fig. 2.1C, 2.2A). This result is consistent with previous findings that TPT1 is essential
only in the context of the generation of 2′-phosphorylated tRNAs by Trl1 (Schwer et al. 2004)
and that a growth defect caused by TRL1 knockdown in the trypanosome Trypanosoma brucei
is rescued by expressing intronless tRNATyr (Lopes et al. 2016).
The SEN complex cleaves introns from pre-tRNAs (Fig. 2.1A), and each component of the
heterotetrameric endonuclease (Sen2, Sen15, Sen34 and Sen54) is essential for growth (Trotta
et al. 1997). I created cells with single genomic deletions in each of the SEN2, SEN15, SEN34,
and SEN54 genes and found that in each case cells were only viable when complemented by
plasmid-mediated expression of the cognate SEN gene (Fig. 2.1D). I expressed the ten pre-
spliced tRNAs in cells containing deletions of each of the SEN genes and found that pre-spliced
tRNAs failed to rescue deletion of any component of the SEN complex (Fig. 2.1C), consistent
with this complex having an essential function beyond splicing of pre-tRNA (Dhungel and Hopper
2012). The gene encoding the only other known substrate of the SEN complex, CBP1, is non-
essential (Tsuboi et al. 2015), suggesting that the SEN complex has another essential function,
possibly to process unknown RNA substrates.
61
Figure 2.1: Genetic bypass of essential components of tRNA splicing with intronless
tRNAs.
A. Functions of tRNA splicing enzymes. Introns from pre-tRNAs are removed by the SEN complex, which contains the Sen2 and Sen34 endonucleases that cleave the 5′ and 3′ splice sites, respectively, as well as Sen15 and Sen54. Following cleavage, the tRNA exons are ligated by the multifunctional RNA ligase Trl1, producing a ligated tRNA with a 2′-PO4 at the splice
junction. Tpt1 removes the 2′-PO4 in an NAD++-dependent reaction, producing ADP-ribose-
1″,2″-cyclic phosphate and nicotinamide.
62
Figure 2.1 (continued): Genetic bypass of essential components of tRNA splicing with
intronless tRNAs.
B. Schematic of a plasmid encoding the ten S. cerevisiae tRNAs in intronless form. The tRNAs are expressed from a high-copy 2µ TRP1 plasmid containing a SUP4 promoter and terminated by the RPR1 terminator.
C. Genetic bypass of RNA repair mutants by expression of intronless tRNAs (“10x-tRNA” plasmid). Strains expressing a URA3 covering plasmid were transformed with an empty vector (TRP1 2µ) or a high-copy plasmid encoding intronless tRNAs (B). URA+ TRP+ colonies (top row) were selected and struck on FOA media (bottom row), which selects against cells with the covering plasmid, to assess intronless tRNA-mediated bypass. Plates were photographed after 5 to 7 days of incubation at 30°C. Intronless tRNAs complement deletion of TRL1 and TPT1 but do not rescue deletion of SEN components (bottom right).
D. Cells with genomic deletions of SEN genes (SEN2, SEN15, SEN34, SEN54) and a single-copy URA3 plasmid expressing the deleted gene were individually transformed with high-copy LEU2 plasmids containing the genomic locus of each of the SEN genes. LEU+ colonies were plated on –LEU media (top), and FOA media (bottom) to select against the covering plasmid. Only those plasmids that contain the cognate genomic locus of the deleted SEN gene were able to rescue growth on FOA (bottom).
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RNA repair mutants have defects in translation
Genetically-bypassed trl1∆ and tpt1∆ cells share common growth phenotypes that may
reflect translational defects in the absence of RNA repair or incomplete rescue by intronless
tRNAs. I found that rescued trl1∆ and tpt1∆ cells grow slowly at 30°C and fail to grow at 37°C
on rich media (Fig. 2.2B). These growth defects could be due to differences in the abundance or
functionality of pre-spliced tRNAs or due to the accumulation of endogenous intermediates of
tRNA splicing. Alterations in tRNA pools and post-transcriptional modifications of specific tRNA
species have been implicated in regulating translational dynamics in both prokaryotes and
eukaryotes (Kirchner and Ignatova 2015), providing a possible explanation for the temperature
sensitivity. Rescued trl1∆ cells should accumulate unligated pre-tRNA half-molecules that have
been cut by the SEN complex, whereas tpt1∆ cells should accumulate spliced and ligated pre-
tRNAs that retain a 2′-PO4 at the ligation junction (Fig. 2.1A). Accumulation of these molecular
species could have distinct effects on translation. However, both RNA repair mutants exhibit
broad sensitivity to sub-lethal doses of translational inhibitors (Fig. 2.2C).
It is possible that the pre-spliced tRNAs used to rescue trl1∆ and tpt1∆ mutants are
suboptimal due to defects in their expression levels, amino acid charging, or post-transcriptional
modification. S. cerevisiae tRNAs have an average of 13 post-transcriptional modifications per
tRNA species (Phizicky and Hopper 2010), and the diverse set of modifications added to tRNAs
can impact maintenance of reading frame, translational fidelity, and tRNA stability (Hopper 2013).
The rapid tRNA decay pathway (RTD) comprises a quality control mechanism that degrades
aberrant tRNA molecules via the 5′→3′ exonucleases Rat1 and Xrn1. Structural stability is a
major determinant of RTD substrates: both the introduction of destabilizing mutations in tRNA
sequences and the deletion of modification enzymes that act to enhance tRNA stability cause
temperature sensitivity in budding yeast.
64
Figure 2.2: Growth phenotype of RNA repair mutants.
Laura K White performed the experiments displayed in this figure.
A. RNA repair mutants rescued by pre-spliced tRNAs expressed from the (10x-tRNA TRP1 2µ) plasmid grow on -TRP and FOA media, but not on -URA media. Plates were photographed after 3 days of growth at 30°C.
B. Growth of RNA repair mutants is temperature-sensitive. Cells were serially diluted, spotted on
YPD media and incubated at the indicated temperatures. Wild-type and complemented trl1∆
(TRL1) and tpt1∆ (TPT1) cells grow at all temperatures, whereas bypassed trl1∆ and tpt1∆ cells fail to grow at 37°C.
65
Figure 2.2 (continued): Growth phenotype of RNA repair mutants.
C. Sensitivity of RNA repair mutants to translational inhibitors. Cells were serially diluted and spotted on YPD media or YPD media supplemented with sub-lethal doses of the translational inhibitors anisomycin, hygromycin B and cycloheximide at the concentrations indicated. Wild-type cells are viable under all conditions, whereas RNA repair mutants fail to grow in the presence of inhibitors of translation.
D. Deletion of XRN1 does not suppress the temperature sensitive phenotype of trl1∆ rescued with pre-spliced tRNAs. All plates were photographed after 3 to 4 days of incubation at 30°C or 37°C.
66
However, growth at 37°C can be rescued by compensatory mutations that restore tRNA
structural stability or by deletion of the Xrn1 exonuclease (Whipple et al. 2011). Based on these
prior observations, I tested whether RTD is responsible for temperature sensitivity by deleting
XRN1 in a rescued trl1∆ mutant and found that the double deletion, trl1∆ xrn1∆, grows no better
at 37°C than the single trl1∆ mutant (Fig. 2.2D), suggesting that the temperature sensitivity of
RNA repair mutants is not a consequence of RTD.
Whereas our results suggest that pre-spliced tRNAs are not rapidly degraded due to
structural instability, these intronless tRNA species may still be hypomodified or otherwise
suboptimal. Previous studies of tRNA introns identified two types of post-transcriptional
modifications that are only added at the pre-tRNA stage, when an intron is still present. Site-
specific introduction of 5-methylcytidine (m5C) or pseudouridine in the anticodon or at position
40 is dependent on the presence of the intron, and these intron-dependent modifications occur
in four of the ten intron-containing tRNAs in S. cerevisiae (Grosjean et al. 1997) (Table 2.3). In
particular, incorporation of m5C at the wobble position in the pre-tRNALeu(CAA) is catalyzed by
the tRNA-specific methyltransferase Trm4, and disruption of TRM4 causes sensitivity to the
antibiotic paromomycin, an aminoglycoside that interferes with translational fidelity (Wu et al.
1998). This observation is consistent with our finding that RNA repair mutants rescued with pre-
spliced tRNAs fail to grow on rich media containing low doses of the translational inhibitors
anisomycin, hygromycin B, or cycloheximide (Fig. 2.2C). Thus, this phenotype could be
indicative of hypomodification of one or more of the intronless tRNAs from the construct.
Alternatively, it is possible that bypass of trl1∆ and tpt1∆ with intronless tRNAs provides a
complete rescue of tRNA splicing but causes reduced fitness due to an additional but
nonessential role of RNA repair related to translation.
67
Table 2.3: Intron-containing tRNA copy number and intron-dependent modifications in S.
cerevisiae.
Laura K White produced this table for our 2018 publication in RNA.
tRNA Species Copy Number Intron Length (nt) Intron-Dependent Base Modifications
tRNA-Ser(CGA) 1 19
tRNA-Ile(UAU) 2 60 Pseudouridine, positions 34 & 36
tRNA-Leu(UAG) 3 19
tRNA-Ser(GCU) 4 19
tRNA-Trp(CCA) 6 34
tRNA-Lys(UUU) 7 23
tRNA-Tyr(GUA) 8 14 Pseudouridine, position 35
tRNA-Leu(CAA) 10 32 5-methylcytidine, position 35
tRNA-Phe(GAA) 10 18-19 5-methylcytidine, position 40
tRNA-Pro(UGG) 10 30-33
Genomic copy number of tRNAs listed are reproduced here, reorganized, from (Chan and Lowe 2009); intron-dependent tRNA modifications are from (Grosjean et al. 1997).
68
RNA repair mutants accumulate intermediates and products of tRNA splicing
To evaluate the ability of cells lacking RNA repair to produce mature tRNAs, I analyzed the
processing of tRNAIle(UAU) by northern blot using total RNA from rescued trl1∆ and tpt1∆ cells,
unshuffled control strains, wild-type, and xrn1∆ mutants. Wild-type, trl1∆ (TRL1), and tpt1∆
(TPT1) cells have expected tRNA processing intermediates, including primary transcript (145 nt),
pre-tRNA with 5′- and 3′-processing by RNaseP, Rex1, RNase Z, and Lph1 (136 nt), mature
tRNA at 76 nt, excised intron at 60 nt, and 5′-exon at 38 nt (Fig. 3 A, B). Total RNA from trl1∆
mutants complemented with intronless tRNAs contains a band of the same size as mature tRNA
from wild-type controls, and this tRNA must arise from the intronless tRNA construct because
these cells cannot ligate exons processed from genomic intron-containing tRNAs (Fig. 2.3A, lane
4). Cells that contain the pre-spliced tRNA plasmid exhibit high molecular weight species to
which the northern probe hybridizes, which are likely tandem tRNA transcripts that are not
completely processed. This result is consistent with the construct functioning as a single
transcriptional unit, owing to its design with a single SUP4 pol III promoter and RPR1 pol III
terminator; additionally, the intronless tRNA genes may also be transcribed individually via
internal promoters (Galli et al. 1981).
The S. cerevisiae genome contains tRNA genes at varying copy number (Table 2.3) (Chan
and Lowe 2009), raising the possibility that rescue by a high-copy plasmid expressing intronless
tRNAs could alter the levels of specific tRNA isodecoders. To assess the ability of the genetic
bypass to produce quantities of tRNAs similar to those found in wild-type cells, I performed
northern blotting with probes hybridizing to the 5′-exons of tRNAs with varying genomic copy
number.
69
Figure 2.3: tRNA processing phenotypes of RNA repair mutants.
Diagrams of tRNA primary structure, annotations, and probe locations are depicted next to each northern blot. SCR1 loading control blots are below and were the basis of loading for each lane.
A. Northern blot using probe that hybridizes to the 5′-exon of tRNAIle(UAU) identifies pre-processed tRNA intermediates derived from primary transcripts of intronless tRNAs (lanes 4, 5, and 7, bracket annotation), as well as intermediates arising from the intron-containing endogenous
tRNAs (at ~150 nt). Each strain produces mature tRNAs (~76 nt). The trl1∆ mutant (lane 4) is unable to ligate tRNAIle(UAU) exons arising from chromosomal copies of the gene, and thus mature tRNAs in these cells occur from processing of the intronless tRNA transcript. A band at ~32 nt in lanes 4 and 7 is likely a product of SEN cleavage of chromosomally-encoded intron-containing tRNAs that are not re-ligated.
70
Figure 2.3 (continued): tRNA processing phenotypes of RNA repair mutants.
B. Northern blot with probe to tRNAIle(UAU) intron shows increased accumulation of the intron in
trl1∆ mutants (lanes 4 and 7), as well as in xrn1∆ and in double deletion trl1∆ xrn1∆ mutants (lanes 6 and 7, respectively). A band at ~90 nt putatively represents 5′-exon/intron (question mark). Densitometry quantifications of intron signal relative to wild-type (lane 1) and normalized to SCR1 signal are displayed below lane numbers.
C. Northern blot with probe to the 5′-exon of tRNATyr(GUA), a tRNA with a copy number of 8 in the budding yeast genome, reveals mature tRNA bands at approximately the same intensity across wild-type (lane 1) and repair mutant strains (lanes 4 & 5).
D. Northern blot with probe to the 5′-exon of tRNAPhe(GAA), a tRNA with a copy number of 10 in the genome, exhibits approximately equal density of mature tRNA band intensity in wild-type (lane 1) and deletions in RNA repair genes (lanes 4 & 5).
E. Northern blot with probe to the 5′-exon of tRNALeu(CAA), a tRNA with a copy number of 10 in
the genome, shows a decrease in mature tRNA band intensity in the trl1∆ mutant (lane 4), and
also in the tpt1∆ mutant (lane 5), as compared to wild-type and covered strains (lanes 1 through 3).
F. Northern blot with probe to the 5′-exon of tRNAPro(UGG), a tRNA with a copy number of 10 in
the genome, reveals a decrease in mature tRNA band intensity in both trl1∆ and tpt1∆ RNA repair mutants (lanes 4 & 5, respectively). All lanes show a doublet of bands, with the upper band being consistent in size with mature tRNAPro(UGG), and the lower band is annotated with an asterisk.
Lane 5 displays a strong tpt1∆-dependent band, annotated with a question mark.
71
In the case of low copy number, tRNAIle(UAU) is encoded at two copies per genome, and the
intronless tRNA plasmid produces quantities of mature tRNAIle(UAU) similar to wild-type cells
(Fig. 2.3A, lane 1 versus 4 & 5). In contrast, the isodecoders tRNATyr(GUA), tRNALeu(CAA),
tRNAPhe(GAA), and tRNAPro(UGG) are encoded in the genome at relatively high copy number (8,
10, 10, and 10 copies, respectively) (Table 2.3). The intronless tRNA plasmid produces
tRNATyr(GUA) and tRNAPhe(GAA) at abundances similar to wild-type (Fig. 2.3C & 2.3D, lane 1
versus 4 & 5), but the plasmid fails to produce equivalent amounts of tRNALeu(CAA) and
tRNAPro(UGG) (Fig. 2.3E & 2.3F, lanes 1 versus 4 & 5). These results show that there is not a
monotonic relationship between copy number in the genome and the ability of the intronless
tRNA plasmid to produce wild-type quantities of each tRNA. However, these results do not
comment on the modification status of intronless tRNAs. Furthermore, the discrepancy in
quantities of tRNALeu(CAA) and tRNAPro(UGG) produced by the intronless tRNA plasmid could
contribute to the slow growth phenotype of the RNA repair mutants (Fig. 2.2).
The northern blot for tRNAPro(UGG) (Fig. 2.3F) reveals several additional features of interest
that appear to be unique to this tRNA isodecoder. Doublet bands of spliced tRNAPro(UGG), which
I also detected in all strains tested, have been previously observed by denaturing polyacrylamide
gel (Winey et al. 1986). The upper of these two bands has a length consistent with the 75 nt
mature tRNAPro(UGG) (Chan and Lowe 2009). The lower band, approximately 10 nucleotides
shorter, cannot be explained as an intron-containing tRNA half or processing intermediate, as
the intron for tRNAPro(UGG) is 31 nt and a processing intermediate that contained the intron and
both exons would migrate at ~103 nt. Because these intermediates accumulate in the RNA ligase
mutant—and ligase is required for intron splicing—the pre-spliced tRNA construct is also
competent to produce both species (i.e., the putative processing intermediate marked with an
asterisk and mature tRNA). In addition, both tpt1∆ (TPT1) and tpt1∆ cells (Fig. 2.3F, lanes 3 and
72
5) contain a third tRNAPro(UGG) band (indicated by a question mark) that accumulates to high
levels in tpt1∆ cells and is not seen in other genotypes, which may correspond to an additional
tRNA species resolved via 2D PAGE analysis of tRNA from a tpt1 conditional mutant (Spinelli et
al. 1997). It is possible that this third band is derived from endogenous spliced tRNAPro(UGG) that
retains a 2′-phosphate at the splice junction in the absence of TPT1; however, the precise
identity of these three species of tRNAPro(UGG) remains unresolved.
In addition to analysis of mature tRNA production, I further investigated the fate of the
introns from endogenous tRNA genes in the context of the RNA repair mutants. Northern blot
analysis of the tRNAIle(UAU) intron recapitulated the finding that decay of excised tRNA introns
requires both Xrn1 and Trl1(Wu and Hopper 2014). Bypassed trl1∆ cells show an increase in
levels of tRNA intron (47-fold compared to wild-type, normalized to SCR1 levels), as do xrn1∆
cells (6.3-fold) and double mutant trl1∆ xrn1∆ cells (22-fold) (Fig. 2.3B, lanes 4, 6, and 7). These
observations are in line with kinase-mediated decay of cleaved tRNA introns, in which the 5′-
RNA kinase activity of Trl1 (Fig. 2.1A) is required to phosphorylate the 5′-hydroxyl intron
products of SEN cleavage to enable their 5′→3′ exonucleolytic decay by Xrn1, which specifically
degrades 5′-phosphorylated RNA substrates (Stevens 2001).
RNA repair mutants have defects in unfolded protein response activation
Induction of the unfolded protein response (UPR) requires the ligation of two exons after
intron excision from the HAC1 mRNA (Gonzalez et al. 1999). I tested cells lacking RNA repair for
their ability to activate the UPR. Bypassed trl1∆ mutants fail to grow on media containing the
UPR-inducing drug tunicamycin (Fig. 2.4A), confirming that Trl1 is required for UPR activation in
response to protein folding stress (Sidrauski et al. 1996). In contrast, tpt1∆ mutants grow equally
well in the presence and absence of tunicamycin (Fig. 2.4A), suggesting that the 2′-PO4
remaining on HAC1 mRNA after ligation in tpt1∆ cells does not interfere with its translation or
73
that partial HAC1 translation is sufficient for UPR activation (see below). Cells with Hac1-FLAG
(in wild-type, trl1∆, and tpt1∆ backgrounds) were also spotted onto tunicamycin containing
media and had similar growth to cells with untagged Hac1, confirming that Hac1-FLAG can
function in the UPR (Fig. 2.4A).
I corroborated the growth assay by analyzing HAC1 splicing in trl1∆ and tpt1∆ cells using RT-
PCR (Fig. 2.4B). The wild-type strain catalyzes cleavage and ligation of HAC1 upon UPR
stimulation with tunicamycin, whereas the trl1∆ mutant exhibits no detectable spliced HAC1 and
a reduction in unspliced HAC1. The decrease in unspliced HAC1 observed in lanes 4 and 6 (Fig.
2.4B) as compared to their tunicamycin-null control lanes can be explained by two facts: first,
both strains express Ire1, the endonuclease responsible for cleaving HAC1 mRNA prior to
ligation; second, the Hac1 protein is a transcription factor that activates transcription of the
HAC1 gene, creating a positive feedback loop to sustain UPR activation (Ogawa and Mori 2004).
When Ire1 cleaves HAC1 mRNA, but Trl1 is unavailable for ligation, Hac1 protein is not translated
and cells fail to activate the positive feedback loop. Then, without the newly-transcribed HAC1
mRNA, the remaining pool of HAC1 is efficiently cleaved so that the cDNAs synthesized do not
contain both PCR priming sites and thus fail to amplify in this PCR assay. These simultaneous
deficits could lead to a decrease in unspliced HAC1 in RNA repair mutants treated with
tunicamycin as compared to their null-treatment controls.
74
Figure 2.4: UPR-related phenotypes of RNA repair mutants.
A. Growth assay of RNA repair mutants on UPR-inducing media. Yeast cells (wild-type, trl1∆,
and tpt1∆) were serially diluted and spotted onto rich media (YPD) and tunicamycin-containing media (80 ng/mL tunicamycin) to induce the UPR. Plates were imaged after 3 days of growth at
30°C. Wild-type and tpt1∆ cell growth is unaffected by tunicamycin, whereas trl1∆ cells fail to grow on media containing tunicamycin. Serial dilution growth assays for cells of the same genetic background as above, but with C-terminal FLAG tags on Hac1, are shown below.
75
Figure 2.4 (continued): UPR-related phenotypes of RNA repair mutants.
B. Analysis of HAC1 splicing in RNA repair mutants. Total RNA from untreated and tunicamycin-
treated wild-type, trl1∆, and tpt1∆ cells was analyzed by RT-PCR using primers specific for HAC1, producing products at 499 bp (unspliced HAC1) and 247 bp (spliced HAC1). A no-template (NT) control is shown in lane 7. The proportion of spliced HAC1 upon tunicamycin
treatment increases in wild-type (lanes 1 and 2) and tpt1∆ cells (lanes 5 and 6), and spliced HAC1 is visible in RNA from both cells (lanes 2 and 6, 247 bp). Spliced HAC1 is undetectable in
trl1∆ cells (lane 4) upon tunicamycin treatment (lanes 3 and 4), owing to the inability of trl1∆ cells to ligate HAC1 exons. Asterisk marks an unknown PCR product dependent on tunicamycin treatment. Cells with C-terminal FLAG tags of Hac1 were also analyzed for splicing in the same manner (below) with a no-template (NT) control in lane 9. Reactions lacking reverse transcriptase (RT-) were negative for amplification (data not shown).
C. Hac1 protein levels in RNA repair mutants. Whole cell lysates were prepared from wild-type,
trl1∆, and tpt1∆ cells expressing C-terminal Hac1-FLAG and grown in the presence and absence of tunicamycin. Lysates were analyzed by SDS-PAGE and nitrocellulose transfer followed by Ponceau S staining and cross-reaction with anti-FLAG and anti-GAPDH antibodies. Scale to the left is nominal molecular mass of a protein ladder (kDa); the expected mass of Hac1-FLAG is 31 kDa. Hac1-FLAG is detected in wild-type cells upon tunicamycin addition but is undetectable in
trl1∆ and tpt1∆ cells.
D. Induction of the UPR-responsive KAR2 gene in RNA repair mutants. Amounts of KAR2 mRNA
(normalized to PGK1 mRNA abundance) were measured by RT-qPCR in wild-type, trl1∆, and
tpt1∆ cells in the presence and absence of tunicamycin. Error bars are 95% confidence intervals, n=3. Relative abundance of KAR2 mRNA increased 20-fold in wild-type cells treated with
tunicamycin, whereas the corresponding levels of KAR2 did not increase in trl1∆ and increased
1.4-fold in tpt1∆ cells.
E. Detection of ligated and 2′-phosphorylated HAC1 mRNA. Total RNA was treated with calf intestinal phosphatase (CIP) to remove 2′-phosphates (diagram) and reverse-transcribed using HAC1-specific primer under high (500 µM) concentrations of dNTPs. The cDNA products were PCR amplified, yielding products for unspliced (456 bp) and spliced (204 bp) HAC1 mRNA. Using high dNTP concentrations, I find that splicing of HAC1 in wild-type cells increases upon tunicamycin treatment (compare lanes 1 and 2 versus 3 and 4), similar to B, but is unaffected by CIP treatment (compare lanes 1 versus 2, and 3 versus 4). Likewise, I find that spliced HAC1
mRNA in tpt1∆ cells increases in response to tunicamycin (compare lanes 9 and 10 versus lanes 11 and 12), albeit to a lesser extent that wild-type cells and is unaffected by CIP treatment (compare lanes 9 versus 10 and lanes 11 versus 12). An asterisk marks an unknown PCR product dependent on tunicamycin treatment.
F. Detection of ligated and 2′-phosphorylated HAC1 mRNA. Using low (1 µM) dNTP concentrations, I find that spliced HAC1 is preferentially amplified in wild-type cells over unspliced HAC1 (lanes 1 through 4). The abundance of spliced HAC1 mRNA from wild-type cells increases in response to tunicamycin but is unaffected by CIP treatment (compare lanes 1 versus
2, and 3 versus 4). RT-PCR analysis of HAC1 mRNA from tpt1∆ cells reverse transcribed under low dNTP concentrations shows both unspliced and spliced forms of HAC1 mRNA, and spliced HAC1 mRNA increases in response to tunicamycin treatment (compare lanes 10 versus 12).
However, in contrast to wild-type, amplification of spliced HAC1 mRNA from tpt1∆ is strongly
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Figure 2.4 (continued): UPR-related phenotypes of RNA repair mutants.
dependent on prior treatment with CIP. In the absence of tunicamycin and CIP treatment, spliced HAC1 mRNA is undetectable, whereas treatment with CIP enables reverse transcription (compare lane 9 to lane 11; see panel with enhanced contrast to the right). Similarly, the
abundance of spliced HAC1 mRNA from tpt1∆ cells increases in response to tunicamycin, and its abundance is further increased upon CIP treatment (compare lane 11 versus 12).
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I found that tpt1∆ cells accumulate spliced HAC1, but to a lesser extent than wild-type,
suggesting that the 2′-phosphorylated products of ligation interfere with processing or stability
of the HAC1 mRNA (Fig. 2.4B, compare lanes 2 versus 6). Cells with Hac1-FLAG spliced HAC1
mRNA with the same pattern as their untagged controls (Fig. 2.4B). An immunoblot for Hac1-
FLAG (Fig. 2.4C) showed that wild-type cells showed no detectable Hac1 until the UPR was
induced with tunicamycin, but Hac1-FLAG could not be detected in lysate from trl1∆ or tpt1∆
cells.
In addition, trl1∆ cells failed to increase expression of the chaperone KAR2 (Fig. 2.4D), a
representative UPR-responsive gene strongly induced by Hac1 transcription factor protein (Mori
et al. 1992; Nikawa et al. 1996) indicating that trl1∆ cells fail to induce UPR-responsive genes.
Consistent with reduced spliced HAC1 mRNA accumulation (Fig. 2.4B), tpt1∆ mutants also
show reduced expression of KAR2 (Fig. 2.4D). However, this reduced degree of UPR induction,
as observed by HAC1 mRNA splicing and KAR2 expression, is nonetheless sufficient for growth
of tpt1∆ cells in the presence of tunicamycin (Fig. 2.4A). Despite accumulation of spliced HAC1
mRNA (Fig. 2.4B), accumulation of KAR2 mRNA (Fig. 2.4D) and growth on tunicamycin (Fig.
2.4A), I was unable to detect Hac1-FLAG by immunoblot from tpt1∆ cells. (Fig. 2.4C). It is
possible that the 2′-PO4 allows only partial translation of the spliced HAC1 mRNA 5′-exon,
leading to production of Hac1 N-terminal bZIP domain and low level UPR activation, but
precluding translation of the C-terminal FLAG epitope. Consistent with this possibility, translation
of unspliced HAC1 mRNA is sufficient to restore growth on tunicamycin-containing media in the
absence of the Duh1 ubiquitin ligase (Di Santo et al. 2016).
I found that tpt1∆ mutants accumulate less spliced HAC1, less Hac1 protein, and less
KAR2 mRNA than wild-type yeast despite having a functional RNA ligase. To determine whether
ligated HAC1 mRNA retains a 2′-PO4 at the ligation junction in tpt1∆ cells, I adapted a reverse-
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transcriptase-based method from previous reports that show 2′-phosphates inhibit reverse
transcription (Dhungel and Hopper 2012; Schutz et al. 2010). I used calf intestine phosphatase
(CIP) treatment, which removes 2′-phosphates from RNA (Fig. 2.4E schematic) (McCraith and
Phizicky 1990), and RT-PCR to test whether 2′-phosphates were present in ligated HAC1 mRNA.
Using low concentrations of dNTPs (1 µM) (Fig. 2.4F) and a reverse transcription primer adjacent
to the expected site of 2′-phosphorylation, CIP treatment substantially enhanced detection of
spliced HAC1 mRNA in tpt1∆ cells, but levels of amplified HAC1 in RNA from wild-type cells
were unaffected, indicating that a 2′-PO4 remains at the ligation junction in the HAC1 mRNA
from tpt1∆ cells. Reverse transcription reactions with typical dNTP concentrations (500 µM) (Fig.
2.4E) restored amplification of unspliced HAC1 mRNA. I surmise that reduced dNTP
concentrations lower the processivity of reverse transcriptase, favoring synthesis of shorter
cDNA substrates. The presence of spliced HAC1 mRNA in tpt1∆ cells in the absence of CIP
treatment (Fig. 2.4F, lane 11) could indicate that the assay is not quantitatively sensitive to sites
of 2′-phosphorylation or that not all molecules of ligated HAC1 mRNA retain 2′-phosphates
despite the absence of Tpt1. In any case, 2′-phosphorylated RNA reduces cDNA synthesis under
these conditions, enabling their detection via this RT-PCR strategy.
Summary
I showed the one essential function of RNA repair in budding yeast is catalyzing the ligation
of tRNA halves resulting from splicing; however, the reduced-growth phenotypes of RNA repair
mutants caused by various translational inhibitors, a UPR-inducing drug, and elevated
temperature suggest either that the pre-spliced tRNAs do not function at the same
efficiency/abundance as endogenously-encoded tRNAs—or that RNA repair is generally helpful
to cells, albeit not essential. I demonstrated that “pre-spliced” tRNA genes are transcribed
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and/or processed to a size consistent with wild-type mature tRNA. Furthermore, I provided
evidence of dysfunction of the UPR in both the ligase mutant and the 2′-phosphotransferase
mutant, suggesting that 2′-phosphorylated HAC1 mRNA contributes to UPR induction
dynamics. Lastly, I showed that RNA retaining 2′-PO4 residues is detectable when the enzyme
responsible for removing them is deleted.
These new genetic reagents enable studies to identify other targets of RNA repair. For
example, cells lacking the Tpt1 2′-phosphotransferase accumulate 2′-phosphates at sites of
RNA ligation, enabling their possible identification by methods to identify 2′-O-modifications by
RNA-seq (Birkedal et al. 2015) or by affinity purification by tagging with mutant Tpt1 enzyme and
biotin-NAD (Steiger et al. 2005). These approaches address the diversity of RNA repair in biology
and are the subject of further study.
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CHAPTER III
III MULTIPLE DECAY EVENTS TARGET HAC1 mRNA DURING SPLICING TO
REGULATE THE UNFOLDED PROTEIN RESPONSE2
Abstract
In the unfolded protein response (UPR), stress in the endoplasmic reticulum (ER) activates
a large transcriptional program to increase ER folding capacity. During the budding yeast UPR,
Ire1 excises an intron from the HAC1 mRNA and the exon products of cleavage are ligated, and
the translated protein induces dozens of stress-response genes. Using cells with mutations in
RNA repair and decay enzymes, we show that phosphorylation of two different HAC1 splicing
intermediates is required for their degradation by the 5′→3′ exonuclease Xrn1 to enact opposing
effects on the UPR. We also found that ligated but 2′-phosphorylated HAC1 mRNA is cleaved,
yielding a decay intermediate with both 5′- and 2′-phosphates at its 5′-end that inhibit 5′→3′
decay and suggesting that Ire1 degrades incompletely processed HAC1. These decay events
expand the scope of RNA-based regulation in the budding yeast UPR and have implications for
the control of the metazoan UPR.
Introduction
During the unfolded protein response (UPR), protein folding stress in the lumen of the
endoplasmic reticulum leads to oligomerization of the transmembrane kinase/endoribonuclease
Ire1 and the processing of a cytoplasmic mRNA to yield splicing intermediates with 2′,3′-cyclic
phosphate (PO4) and 5′-hydroxyl (OH) termini (Gonzalez et al. 1999). In budding yeast, excision
2 Published with permission under a Creative Commons License (Attribution 4.0 International (CC BY 4.0)) from eLife: PD Cherry, SE Peach, and JR Hesselberth. Multiple decay events target HAC1 mRNA during splicing to regulate the unfolded protein response. 2019. doi: 10.7554/eLife.42262
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of an intron from the HAC1u mRNA (“u” denoting the unspliced mRNA) by Ire1 is followed by
exon ligation by the multifunctional Trl1 RNA ligase (Sidrauski et al. 1996) involving 5′-
phosphorylation of the 5′-OH product, adenylylation of the 5′-PO4, and resolution of the 2′,3′-
cyclic PO4 to a 2′-PO4/ 3′-OH. The newly produced 3′-OH serves as the nucleophile to attack
the 5′-adenylate intermediate, yielding a ligated mRNA with an internal 2′-PO4. The 2′-PO4 is
assumed to be removed in a separate reaction by the 2′-phosphotransferase, Tpt1, in a NAD+ -
dependent reaction (Culver et al. 1997). The spliced mRNA, called HAC1s mRNA (“s” denoting
spliced mRNA) (Li et al. 2018), is translated into a transcription factor that activates the
expression of dozens of stress-response genes to mitigate protein-folding stress (Ron and
Walter 2007). In addition, Hac1 activates its own promoter in a positive feedback loop that
generates more HAC1u and permits sustained UPR activation (Ogawa and Mori 2004) (Fig. 3.1A).
Control of this positive feedback loop ensures UPR suppression during normal growth and
rapid activation upon stress exposure. To facilitate the control of UPR activation, HAC1u contains
cis- regulatory elements that suppress unintended translation and promote rapid processing. If
a ribosome initiates on HAC1u , translation through the 5′-exon/intron junction yields a truncated
protein with an intron-encoded C-terminal peptide “degron” that targets it for ubiquitylation and
degradation (Di Santo et al. 2016). A stem-loop in its 3′-untranslated region (the “3′-BE”) tethers
HAC1u to the ER membrane, ensuring rapid Ire1-mediated cleavage following ER stress (Aragón
et al. 2009). Finally, a long-range base-pairing interaction between the 5′-UTR and intron
prevents ribosome initiation to suppress translation of HAC1u mRNA (Chapman and Walter 1997;
Di Santo et al. 2016).
Previous work found unexpected roles for RNA decay and repair enzymes acting on HAC1
mRNA in the budding yeast unfolded protein response. Ire1 is a metal-ion-independent
endonuclease that produces RNA cleavage products with 5′-OH termini (Gonzalez et al. 1999).
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In cells lacking the cytoplasmic 5′→3′ exonuclease Xrn1, HAC1 splicing intermediates
accumulate with 5′-PO4 termini, indicating that an RNA 5′-kinase phosphorylates HAC1
processing intermediates and that not all HAC1 splicing intermediates are productively ligated
(Harigaya and Parker 2012; Peach et al. 2015). In addition to its role in HAC1 exon ligation, Trl1
is required to relieve translational attenuation of HAC1s by an unknown mechanism (Mori et al.
2010). In cells expressing the T4 bacteriophage RNA repair enzymes PNK and RNL in lieu of
TRL1, ligated HAC1 molecules contained single nucleotide deletions from the 3′-terminus of the
5′-exon, indicating that a 3′→5′ exonucleolytic activity acts on the cleaved 5′-exon (Schwer et
al. 2004) and nuclear 3′→5′ decay of HAC1u liberates the 3′-BE, tuning the activation potential
of the UPR (Sarkar et al. 2018).
Recent studies showed that RNA decay also plays a role in the UPR in other organisms.
During UPR activation in the fission yeast, Ire1 incises specific mRNAs to promote their
stabilization or degradation (Kimmig et al. 2012; Guydosh et al. 2017). This mode of Ire1 cleavage
is similar to the metazoan Regulated Ire1-Dependent Decay (RIDD) pathway wherein Ire1 incises
some ER-localized mRNAs and the cleavage products are degraded by Xrn1 and the
cytoplasmic exosome (Hollien and Weissman 2006).
Here, we used budding yeast with mutations in RNA repair and decay enzymes to show
that HAC1 splicing intermediates are processed at multiple steps prior to ligation, limiting the
impact of spurious Ire1 activation and unintentional HAC1 cleavage. Our studies also show that
incompletely spliced HAC1s mRNA is targeted for degradation, which may be used to attenuate
the UPR.
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Materials & Methods
Cell culture and RNA preparation
Single colonies were inoculated in drop-out media supplemented with relevant amino
acids and incubated at 30°C overnight with rotation. Cultures were diluted to an OD600 of 0.2 in
yeast-extract, peptone, dextrose (YPD) media, and UPR induction was carried out when yeast
were growing at mid-log phase with a 2-hour treatment (unless otherwise indicated) with
tunicamycin (final concentration of 2.5 μg/mL, Sigma-Aldrich) or DMSO mock treatment. Cells
were harvested by centrifugation, and total RNA was isolated by hot acid phenol extraction. For
RT-PCR and RT-qPCR experiments, total RNA was treated with TURBO DNase (2 U, Ambion)
to degrade contaminating genomic DNA.
RT-PCR/qPCR
DNase-treated RNA was reverse transcribed with 200 U of SuperScript III reverse
transcriptase (Invitrogen) using a gene-specific reverse primer (Table 3.2). Products analyzed on
a 1.5% agarose TBE gel, stained with 1x GelRed (Sigma) and imaged with a Bio Rad GelDoc.
Densitometry was performed with Bio-Rad Image Analysis software and splicing quantifications
were computed and visualized in R using ggplot2 and cowplot R Packages. Quantitative PCR
(qPCR) for KAR2 was also performed on cDNA as generated above and assayed for KAR2 and
PGK1 using Sso Advanced Universal SYBR Green Supermix (Bio Rad) and cycled on a Bio Rad
C1000 384-well thermal cycler and plate reader. Output Ct values were analyzed in Microsoft
Excel and plotted in R using ggplot2 and cowplot R Packages.
Primer Extension
Primers specific for HAC1 mRNA and U6 snRNA were PAGE-purified and ethanol
precipitated. Oligonucleotide primers were 5′-end-labeled with PNK (Enzymatics) and γ-3 2P-
ATP (Perkin Elmer) and purified with Sephadex G-25 spin columns (GE Healthcare resin, Thermo
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empty columns). Radiolabeled primers and total RNA (15 μg) were heated to 65°C for 5 minutes
and cooled to 42°C. SuperScript III reverse transcriptase (200 U, Invitrogen) was added and
reverse transcription reactions were run with a final concentration of 500 μM dNTPs. Primers
were extended for 30 minutes at 42°C, 15 minutes at 45°C, and 15 minutes at 50°C. SuperScript
III RT was inactivated by heating for 20 minutes at 75°C. RNA was destroyed with in 10 mM
NaOH at 90°C for 3 minutes and neutralized with HCl. Formamide loading dye was added and
products were run on an 8% acrylamide TBE 7M Urea gel. Gels were dried (Bio Rad) and
exposed on a phosphor-imager screen and imaged on a Typhoon 9400 (GE Healthcare).
Northern blotting
Total RNA (3 μg) was electrophoresed on 6% acrylamide TBE 7M urea gels and transferred
to nylon membrane (Hybond N+, GE) by electroblotting. Membranes were UV-crosslinked (254
nm, 120 mJ dose), blocked in ULTRAhyb-Oligo Buffer (Ambion), and incubated with 5′-32P-
labeled oligonucleotide probes (Table 3.2) in ULTRAhyb-Oligo at 42°C for 18 hours. Membranes
were washed with 2X SSC/0.5% SDS washing buffer two time for 30 minutes each, exposed on
a phosphor-imager storage screen, and imaged on a Typhoon 9400 (GE Healthcare).
Membranes were stripped of original probe with 3 washes in stripping buffer (2% SDS) at 80°C
for 30 minutes per wash. Membranes were re-blocked and probed a second time for the loading
control, SCR1 (Table 3.2).
Yeast strains and plasmids
Yeast strains and sources used in this study are listed in Table 3.3.
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Table 3.1: HAC1 processing intermediates
Name Size (nt) Visual Summary Description
HAC1u 1450* Full-length, genomic HAC1 transcript
HAC1s 1198* Spliced HAC1; intron removed
HAC1 5′-exon 728 Everything 5′ of the intron
Cleaved 5′-exon ~678 Fragment of 5′-Exon missing ~50 nt off its 3′-end
HAC1 intron 252 Liberated intron (alone)
circularized intron ~500 Circularized intron, visible in wild-type & RtcB cells
HAC1 3′-exon 474* Everything 3′ of the intron
Cleaved 3′-exon ~524* 3′-Exon with ~50 nt of 5′-Exon on its 5′-end
5′-exon + intron 980 5′-exon + Intron
Intron + 3′-exon 726* Intron + 3′-exon
*Size does not include poly(A) tail.
Sizes of HAC1 processing intermediates are predicted from strand-specific RNA sequencing data (Levin et al. 2010) mapped to the sacCer1 genome.
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Table 3.2: Oligonucleotide sequences
Oligonucleotide Name Oligonucleotide sequence (5′→3′)
HAC1-F RT-PCR ACCTGCCGTAGACAACAACAAT
HAC1-R RT-PCR AAAACCCACCAACAGCGATAAT
KAR2 qPCR F AAGACAAGCCACCAAGGATG
KAR2 qPCR R AGTGGCTTGGACTTCGAAAA
PGK1 qPCR F TCTTAGGTGGTGCCAAAGGTT
PGK1 qPCR R GCCTTGTCGAAGATGGAGTC
HAC1 5′-exon probe 1 AAGTCTCTTGGTCCGACGCGGAATCGCGCA
HAC1 5′-exon probe 2 CTGGATTACGCCAATTGTCAAGATCAATTG
HAC1 intron probe 1 AACCGGCTCCTCCCCCATCAGAGAACCACGA
HAC1 intron probe 2 GGACAGTACAAGCAAGCCGTCCATTTCTTAGT
HAC1 3′-exon probe (primer
extension and northern)
ACCGGAGACAGAACAGTAGAAACCACTAAGCG
KAR2 probe ACCGTAGGCAATGGCGGCTGCGGTTGGTTC
SCR1 probe (oRP100) GTCTAGCCGCGAGGAAGG
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Table 3.3: Strain numbers and genotypes
All strains are background W303 (MATa {leu2-3,112 trp1-1 can1-100 ura3-1 ade2-1 his3-11,15}).
Strain ID Genotype Source
YJH682 “wild-type” (CRY1) Mingxia Huang
YJH632 xrn1∆::HygMX
YJH745 ski2∆::NatMX
YJH898 dxo1∆::KanMX
YJH867 xrn1∆::HygMX dxo1∆::KanMX
YJH829 tpt1∆::LEU2 (TPT1 CEN ARS URA3) Schwer et al., 2004 PNAS
YJH830 tpt1∆::LEU2 (TPT1 CEN ARS URA3) (pAG424-ccdB)
YJH832 tpt1∆::LEU2 (TPT1 CEN ARS URA3) (pAG424-10x-tRNA)
YJH834 tpt1∆::LEU2 (pAG424-10x-tRNA)
YJH980 tpt1∆::LEU2 (pAG424-10x-tRNA) (pAG413-NPr-TPT1)
YJH891 tpt1∆::LEU2 (pAG424-10x-tRNA) (pAG413-NPr-tpt1-R138A)
YJH902 tpt1∆::LEU2 xrn1∆::HygMX (pAG424-10x-tRNA)
YJH901 tpt1∆::LEU2 ski2∆::NatMX (pAG424-10x-tRNA)
YJH681 trl1∆::KanMX (pRS416-TRL1) Schwer et al., 2004 PNAS
YJH708 trl1∆::KanMX (pRS416-TRL1) (pAG424)
YJH709 trl1∆::KanMX (pRS416-TRL1) (pAG424-10x-tRNA)
YJH835 trl1∆::KanMX (pAG424-10x-tRNA)
YJH887 trl1∆::KanMX (pAG424-10x-tRNA) (pRS413-TRL1)
YJH811 trl1∆::KanMX (pAG424-10x-tRNA) (pRS413-trl1-D425N)
YJH812 trl1∆::KanMX xrn1∆::HygMX (pAG424-10x-tRNA) (pRS413-trl1-D425N)
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YJH912 trl1∆::KanMX (pAG424-10x-tRNA) (pRS413-trl1-K114A)
YJH913 trl1∆::KanMX (pAG424-10x-tRNA) (pRS413-trl1-K114A-D425N)
YJH808 trl1∆::KanMX (pRS423-TPI-RtcB)
YJH809 trl1∆::KanMX xrn1∆::HygMX (pRS423-TPI-RtcB)
YJH899 trl1∆::KanMX xrn1∆::HygMX (pAG424-10x-tRNA)
YJH900 trl1∆::KanMX ski2∆::NatMX (pAG424-10x-tRNA)
YJH903 trl1∆::KanMX hac1∆::HygMX (pAG424-10x-tRNA) (pAG413-GPD-HAC1u)
YJH904 trl1∆::KanMX hac1∆::HygMX (pAG424-10x-tRNA) (pAG413-GPD-HAC1s)
YJH920 hac1∆::NatMX (pAG413-GPD-HAC1u)
YJH921 hac1∆::NatMX (pAG413-GPD-HAC1s)
YJH923 tpt1∆::LEU2 hac1∆::NatMX (pAG424-10x-tRNA) (pAG413-GPD-HAC1u)
YJH924 tpt1∆::LEU2 hac1∆::NatMX (pAG424-10x-tRNA) (pAG413-GPD-HAC1s)
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Results
RNA repair mutants have unique HAC1 mRNA processing defects.
I recently showed that the functions of the essential RNA repair enzymes Trl1 and Tpt1 in
budding yeast can be genetically bypassed by the expression of intronless tRNAs, which are
able to support translation in trl1∆ and tpt1∆ cells (Cherry et al. 2018). Because Trl1 is required
for HAC1s ligation and subsequent UPR activation (Sidrauski et al. 1996), trl1∆ cells are unable
to grow on media containing tunicamycin (Fig. 3.1B). In contrast, the general growth defect of
tpt1∆ cells is unaffected by tunicamycin (Cherry et al. 2018) (Fig. 3.1B), indicating that these cells
can activate the UPR. Combination of trl1∆ and tpt1∆ with mutations in 5′→3′ and 3′→5′ decay
factors xrn1∆ or ski2∆ led to more pronounced growth defects than the single deletions, but
removal of these decay factors did not affect the growth deficit of trl1∆ or tpt1∆ cells on
tunicamycin (Fig. 3.1B).
Given the multiple enzymatic roles of Trl1 and Tpt1 during RNA repair, I sought to
understand how the loss of these enzymes affected HAC1 mRNA splicing. I visualized HAC1
splicing intermediates by northern blotting with probes for the HAC1 3′-exon and intron (Fig.
3.1C, D) and found robust cleavage and ligation of HAC1u in wild-type cells in the presence of
tunicamycin, leading to high levels of HAC1s . As expected, trl1∆ cells lacking RNA ligase activity
did not produce HAC1s upon tunicamycin treatment. However, cleaved 3′-exon and intron
accumulated upon tunicamycin treatment in trl1∆ cells (Fig. 3.1C, D), indicating a defect in 3′-
exon decay. Cleaved HAC1 3′-exon often appears as a smear of products between ~450 nt and
~575 nt (Fig. 1C); I attribute this size heterogeneity to differences in poly(A) tail presence or
length, as the 5′-ends of these products occur uniformly at one site (Fig. 5A).
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Figure 3.1: HAC1 mRNA processing defects in RNA repair and decay mutants.
A. Schematic of the budding yeast unfolded protein response. ER stress activates Ire1 (green),
which excises an intron (thin line) from HAC1u mRNA. The 5′ (black) and 3′ (grey) exons are
ligated by Trl1 yielding spliced HAC1 (HAC1s ) with a 2′-phosphate at the new ligation junction,
which is subsequently removed by the 2′-phosphotransferase Tpt1. HAC1s mRNA is translated into a transcription factor (Hac1, purple) that upregulates the HAC1 gene itself (a positive feedback loop), as well as several chaperones and heat shock proteins that resolve the stress.
B. Yeast cells with mutations in RNA repair and decay factors were serially diluted (5-fold) and spotted onto agar media (YPD and YPD containing tunicamycin (Tm; 0.16 μg/mL)), grown at 30°C for 2 days, and photographed. The “10x tRNA” plasmid encodes for 10 intronless tRNAs that bypass the lethality of trl1∆ and tpt1∆ (Cherry et al., 2018). The top panels depict cells with deletions of the RNA ligase TRL1, and the bottom panels depict growth of cells deletions of the 2′-phosphotransferase TPT1.
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Figure 3.1 (continued): HAC1 mRNA processing defects in RNA repair and decay mutants.
C. HAC1 processing in RNA repair and decay mutants (3′-exon probe). HAC1u cleavage and ligation were analyzed in mutants of TRL1 RNA ligase and TPT1 2′-phosphotransferase by denaturing acrylamide gel northern blotting using a probe to the HAC1 3′-exon. Diagrams of
HAC1u, HAC1s, and HAC1 splicing intermediates are drawn next to predominant bands (see
Table 1 for descriptions and sizes of all annotations). HAC1u is cleaved and ligated to produce
HAC1s in wild-type cells (lanes 1 & 2). Intron/3′-exon and 3′-exon splicing intermediates
accumulate in trl1∆ cells, but HAC1s is not produced (lanes 3 & 4). In tpt1∆ cells, a small amount
of cleaved 3′-exon is present in the absence of tunicamycin (lane 5), whereas HAC1s and 3′-exon accumulate upon tunicamycin induction (lane 6). Cells lacking xrn1∆ grown in the absence
of tunicamycin produce HAC1s and Intron/3′-exon and 3′-exon splicing intermediates (lane 7),
and tunicamycin addition causes an increase in production of HAC1s (lane 8). The blot was stripped and reprobed using a probe for SCR1 as a loading control.
D. HAC1 processing in RNA repair and decay mutants (intron probe). Linear intron (252 nt) is
excised from HAC1u upon tunicamycin treatment (lanes 1 & 2) and linear intron is excised and accumulates in trl1∆ cells in the presence and absence of treatment (lanes 3 & 4). Excised intron is present a low level in tpt1∆ cells (lanes 5 & 6), whereas xrn1∆ cells accumulate high levels of full-length intron and shorter, intron-derived decay intermediates (lanes 7 & 8). A star denotes excised and circularized intron, which migrates at ~500 nt.
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The 2′-phosphotransferase Tpt1 is essential in budding yeast to remove 2′-phosphate
groups from ligated tRNAs (Culver et al. 1997), but its role in HAC1 mRNA processing during the
UPR has not been defined. Although the growth of tpt1∆ cells is unaffected by tunicamycin (Fig.
3.1B), specific perturbations of HAC1 processing in tpt1∆ cells indicate that residual 2′-
phosphate groups on HAC1 mRNA cause defects in cleavage and ligation. Whereas tunicamycin
treatment led to cleavage of HAC1u and production of HAC1s in tpt1∆ cells, the levels of HAC1s
(Fig. 3.1C) and excised intron (Fig. 3.1D) are significantly lower than in wild-type cells. In
addition, despite the fact that tpt1∆ cells have functional RNA ligase, cleaved 3′-exon
accumulated to high levels upon tunicamycin treatment (Fig. 3.1C).
Kinase-mediated decay of cleaved HAC1 3′-exon competes with its ligation.
To further investigate the 3′-exon decay defect, I examined splicing of HAC1 in xrn1∆ cells.
I found that HAC1s accumulated in the absence of tunicamycin (Figs. 3.1C, 3.2A, C, D, and E).
This promiscuous processing was surprising given that HAC1s is undetectable in wild-type cells
under normal growth conditions, and it suggested that Xrn1 somehow limits production of
HAC1s. In xrn1∆ cells, 3′-exon accumulated to modest levels in both the absence and presence
of tunicamycin (Fig. 3.2A), whereas in trl1∆ cells, HAC1 3′-exon accumulated to higher levels
(Fig. 3.2A, 3.2B). Moreover, the abundance of 3′-exon was similar in trl1∆ and trl1∆ xrn1∆ cells
(Fig. 3.2A), indicating that Xrn1 requires Trl1 for 3′-exon degradation. Previous work showed that
Trl1 5′-kinase activity is required for the Xrn1-mediated degradation of excised tRNA introns in
budding yeast (Wu and Hopper 2014), and I considered whether this pathway also degraded
HAC1 3′-exon. Indeed, expression of a kinase-inactive version of Trl1 (Trl1-D425N) (Wang et al.
2006) did not restore Xrn1-mediated decay of the 3′-exon (Fig. 3.2B), affirming that Trl1 ligase
5′-kinase activity is required for Xrn1-mediated suppression of HAC1 splicing.
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Figure 3.2: Kinase-mediated decay of HAC1 3′-exon competes with its ligation.
A. Decay of cleaved HAC1 3′-exon requires Trl1. A northern blot for HAC1 3′-exon reveals that xrn1∆ (lanes 3 & 4) and trl1∆ (lanes 5 & 6) are mutations sufficient to cause 3′-exon to accumulate, compared to wild-type. In xrn1∆ cells, the accumulations appears tunicamycin-treatment independent, whereas in trl1∆ cells, the accumulation increases upon treatment. The accumulation is also present in xrn1∆ trl1∆ cells (lanes 7 & 8).
B. Decay of cleaved HAC1 3′-exon requires the catalytic activity of Trl1 5′-kinase. We expressed a kinase-inactive missense mutant of Trl1 (Wang et al., 2006), trl1- D425N, to assess the contribution of RNA 5′-kinase activity to decay of HAC1 intermediates. Expression of trl1-D425N (lanes 5 & 6) caused similar accumulation of 3′-exon as in the trl1∆ deletion, and xrn1∆ trl1- D425N cells have levels of 3′-exon similar to trl1- D425N alone (compare lanes 6 & 8), indicating that Trl1 5′-kinase activity is required for Xrn1-mediated decay.
C. 5′-kinase and ligase domains contribute to the abundance of liberated 3′-exon. We expressed a ligase-inactive missense mutant of Trl1 (Sawaya et al. 2003), trl1-K114A, to assess the contribution of ligation activity to levels of HAC1 intermediates. Expression of trl1-K114A (lanes 5 & 6) lead to a moderate accumulation of 3′-exon. The double missense mutant, trl1-K114A-D425N (lanes 7 & 8), has 3′-exon accumulation much like that of trl1-D425N, albeit stronger.
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Figure 3.2 (continued): Kinase-mediated decay of HAC1 3′-exon competes with its ligation.
D. Dxo1 activity does not affect HAC1 3′-exon abundance or promiscuous ligation. Northern blot analysis of 3′-exon shows that dxo1∆ cells phenocopy wild-type cells (2 & 6), whereas and xrn1∆ and dxo1∆ xrn1∆ cells accumulate similar levels of 3′-exon, indicating that Dxo1 does not contribute substantially to 3′-exon abundance.
E. Cells lacking Trl1 and expressing E. coli RtcB promiscuously splice HAC1s. Northern blot
analysis for HAC1 5′-exon shows that trl1∆ (RtcB) cells promiscuously splice HAC1s , similar to xrn1∆ cells (compare lanes 5 and 7 to lane 3), and the defects in HAC1 splicing in trl1∆ (RtcB) cells are unaffected by xrn1∆ (compare lanes 7 & 8 to 5 & 6).
F. RT-PCR assay to measure HAC1 splicing shows promiscuous splicing of HAC1s in RtcB xrn1∆ cells. Similar to E, here an endpoint RT-PCR assay using primers that flank the intron or splice
junction assesses and semi-quantifies HAC1 splicing. Tunicamycin induces HAC1s production
in wild-type cells, (lanes 1 & 2) but HAC1s is detected in in trl1∆ (RtcB) and xrn1∆ cells under normal growth conditions (without tunicamycin, lanes 3 & 5). An asterisk marks an unidentified PCR product.
G. Model for kinase-mediated decay of cleaved HAC1 3′-exon. Ire1 cleavage produces a 3′-exon with a 5′-OH that is phosphorylated by Trl1 5′-kinase. The 5′-phosphorylated product is then adenylated and ligated to the HAC1 5′-exon by Trl1 or degraded by the 5′-phosphate-dependent 5′→3′ exonuclease Xrn1. In xrn1∆ cells (top), the lack of robust 5′→3′ decay favors ligation, leading to promiscuous splicing under normal growth conditions. In trl1∆ cells expressing RtcB (bottom), RtcB directly ligates the 5′-OH products of Ire1 cleavage, and the lack of Trl1 5′-kinase activity renders Xrn1 decay irrelevant, causing promiscuous production of
HAC1s.
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I also tested whether the ligase activity of Trl1 affected HAC1 3′-exon abundance using an
adenylyl-transferase/ligase defective allele (Trl1-K114A) (Sawaya et al. 2003) and found that
additional 3′-exon accumulates compared to wild-type (Fig. 2C), indicating that ligation also
contributes to processing of free 3′-exon.
I also examined the accumulation of 3′-exon in cells lacking Dxo1, a distributive, 5′-
phosphate-dependent 5′→3′ exonuclease (Chang et al. 2012) and found that HAC1 3′-exon
accumulation was unaffected in dxo1∆ cells. In addition, the levels of 3′-exon were similar in
xrn1∆ and dxo1∆ xrn1∆ cells (Fig. 3.2C), indicating that Xrn1 is the primary factor responsible for
5′→3′ decay of the 3′-exon.
Together these data indicate that ligation and Xrn1-mediated 5′→3′ decay compete for
the 5′-phosphorylated 3′-exon splicing intermediate (Fig. 3.2F, top). Examination of HAC1
splicing in trl1∆ cells expressing the E. coli RtcB RNA ligase (Tanaka et al. 2011b) provided
additional evidence of a competition between ligation and decay. RtcB catalyzes ligation of 2′,3′-
cyclic PO4 and 5′-OH RNA termini via a unique mechanism involving nucleophilic attack of the
5′-OH on a 3′-guanylate intermediate; accordingly, RtcB does not have 5′-kinase activity
(Chakravarty et al. 2012). I found that upon tunicamycin treatment, HAC1s was produced in trl1∆
(RtcB) cells (Fig. 3.2D), as shown previously (Tanaka et al. 2011b). However, under normal
growth conditions, trl1∆ (RtcB) cells also promiscuously spliced HAC1s at levels similar to xrn1∆
cells (Figs. 3.2D & E). I propose that because HAC1 ligation by RtcB does not involve a 5′-
phosphate intermediate, Xrn1 is unable to degrade the 5′-hydroxyl exon product of Ire1
cleavage, tipping the balance toward ligation and producing HAC1s under normal growth
conditions (Fig. 2F, bottom). Thus Xrn1-mediated decay of HAC1 3′-exon appears to counteract
a low rate of background Ire1 cleavage to ensure the UPR is only activated when legitimately
stressed.
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Kinase-mediated decay of excised intron is required for HAC1 translation.
In several instances, cells with mutations in repair and decay factors can splice HAC1 but
fail to grow on media containing tunicamycin (Fig. 3.3A), indicating that HAC1s production is not
sufficient to activate the UPR. I assayed expression of KAR2, an ER chaperone and direct target
of the Hac1 transcription factor (Kohno et al. 1993) and found that all repair and decay mutants
express significantly less KAR2 mRNA upon tunicamycin treatment (Fig. 3.3B), consistent with
another layer of UPR regulation downstream of HAC1s production. Excised HAC1 intron
accumulates upon tunicamycin treatment in trl1∆, xrn1∆, and trl1∆ xrn1∆ cells (Fig. 3.3C).
Moreover, the intron decay products that accumulate in these cells are indicative of kinase-
mediated decay: in xrn1∆ cells that lack Xrn1 but have 5′-kinase activity, intron products appear
with a few distinct, smaller products below the full-length 252 nt excised intron (Fig. 3.3C). In
contrast, excised intron accumulated as a uniform, ~250 nt product in trl1∆ cells that lack 5′-
kinase activity (Fig. 3.3C), independent of XRN1 status. Moreover, production of shorter decay
products (like those present in xrn1∆ cells) was dependent on Trl1 5′-kinase catalytic activity
(Fig. 3.3D). Interestingly, expression of the adenylyl-transferase-dead/ligase-dead allele, trl1-
K114A, also led to accumulation of some free HAC1 intron (Fig. 3I), potentially indicating a role
for ligation in the processing of liberated HAC1 intron. Together, these data show that excised
HAC1 intron is a substrate for kinase-mediated decay with strict dependence on a 5′-
phosphorylation step to promote 5′→3′ decay.
A single deletion of Dxo1 had no effect on HAC1 intron degradation (Fig. 3.3E); however,
the sizes of smaller decay products in xrn1∆ dxo1∆ cells were subtly different than in xrn1∆ cells
(Fig. 3.3E), indicating that Dxo1 and other exonucleases partially degrade excised HAC1 intron,
but only when it accumulates in xrn1∆ cells.
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Figure 3.3: Kinase-mediated decay of excised HAC1 intron is required to activate the
unfolded protein response.
A. A serial dilution (5-fold) yeast growth assay on rich media (YPD) and tunicamycin-containing media (+Tm) compares the growth of xrn1∆ , trl1∆ (10x tRNA), and trl1∆ (RtcB) cells to resist
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Figure 3.3 (continued): Kinase-mediated decay of excised HAC1 intron is required to
activate the unfolded protein response.
protein-folding stress. Growth of wild-type cells is unaffected by tunicamycin, whereas growth of xrn1∆ cells is partially inhibited by tunicamycin. Cells that lack ligase (trl1∆), and cells expressing E. coli RtcB RNA ligase in lieu of TRL1 (Tanaka et al., 2011) both fail to grow on media containing tunicamycin, and this growth defect is not affected by xrn1∆.
B. Expression of a UPR-responsive gene is compromised in RNA repair and decay mutants. RT-qPCR of mRNA for KAR2 (BiP), a direct target of Hac1 (Kohno et al., 1993), performed on total RNA from the indicated genotypes shows that wild-type cells induce KAR2 expression by 16-fold upon tunicamycin treatment. (Error bars are 95% confidence interval, n=3; comparison bars represent p < 0.01, Student’s t-test). trl1∆ cells show an insignificant increase in UPR induction, whereas tpt1∆ cells have elevated KAR2 levels in the absence of tunicamycin, which does not change significantly after tunicamycin treatment. trl1∆ (RtcB) and xrn1∆ cells have a modest increase in expression, but not to the same degree as wild-type (p < 0.01).
C. Excised HAC1 intron is stabilized in xrn1∆ and trl1∆ cells. Northern blot analysis using a probe to HAC1 intron reveals that excised intron (252 nt) and partially-degraded intron intermediates accumulate in xrn1∆ cells. Ligase-delete cells (trl1∆ ) also accumulate intron as a uniformly sized 252 nt product. In xrn1∆ and trl1∆ cells, intron accumulates in the absence tunicamycin.
D. Catalytic activity of Trl1 5′-kinase is required for 5′→3′ decay of excised HAC1 intron. Northern blot analysis using a probe to HAC1 intron shows that a missense mutation in the 5′-kinase domain of Trl1 (trl1-D 425N) phenocopies the HAC1 accumulation of trl1∆ cells (lanes 4 & 6, also C).
E. The distributive 5′→3′ exonuclease Dxo1 can partially degrade HAC1 intron. Northern blot analysis for HAC1 intron on total RNA from dxo1∆ and dxo1∆ xrn1∆ cells shows that Dxo1 can partially degrade HAC1 intron when it accumulates in xrn1∆ cells (compare lanes 4, 6 & 8). F. A slow-migrating intron species accumulates in trl1∆ cells expressing RtcB. Northern blot analysis of RNA from wild-type and trl1∆ cells shows that wild-type cells accumulate linear, partially degraded intron (lanes 1-4), whereas trl1∆ (RtcB), and trl1∆ xrn1∆ (RtcB) cells accumulate a slower-migrating species (~500 nt; lanes 5-8).
G. Cells expressing RtcB accumulate circular HAC1 intron. To test whether the slower-migrating band was circularized, total RNA was treated with RNase R and analyzed by northern blot. The slower-migrating species is largely protected from degradation, indicating it is a circle. Linear HAC1 intron and SCR1 (bottom) are degraded upon RNase R treatment (lanes 2, 4, 6 & 8). A panel of enhanced contrast shows that the slower migrating species in wild-type cells are circular, excised HAC1 introns, resistant to RNase R. Circular intron only occurs in samples from cells expressing a ligase.
H. The cytoplasmic exosome degrades HAC1 intron when 5′→3′ decay is disabled. Northern blot analysis using a HAC1 intron probe on total RNA from ski2∆ (a component of the cytoplasmic exosome) cells showed that ski2∆ cells accumulate excised HAC1 intron (lane 4) at levels similar to wild-type (lane 3). In trl1∆ ski2∆ cells, a lack of both kinase-mediated decay (trl1∆) and 3′→5′ decay (ski2∆) causes accumulation of excised intron relative to trl1∆ cells (compare lanes 6 & 8).
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Figure 3.3 (continued): Kinase-mediated decay of excised HAC1 intron is required to
activate the unfolded protein response.
I. Catalytic activity of Trl1 ligase domain contributes to processing of excised HAC1 intron. We expressed a ligase-inactive Trl1 allele, trl1- K114A, as in Fig. 3.2C. Expression of trl1-K114A (lanes 5 & 6) leads to a modest accumulation of intron, though not to the same extent as in the kinase-inactivated mutant (trl1-D425N) (lanes 3 & 4) . The double missense mutant, trl1-K114A-D425N (lanes 7 & 8), exhibits intron accumulation similar that of trl1-D425N.
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Consistent with this notion, we found that the cytoplasmic exosome also contributes to HAC1
intron turnover (Fig. 3.3H), but this mode of decay is unlikely to regulate the UPR as the growth
of ski2∆ cells is unaffected by tunicamycin (Fig. 3.1B).
In trl1∆ (RtcB) cells, excised HAC1 intron accumulates as a circle, evinced by its altered
mobility and resistance to Xrn1-mediated decay in vivo (Fig. 3.3F), and its resistance to RNase
R degradation in vitro (Fig. 3.3G). Robust circularization of the HAC1 intron by RtcB in trl1∆ cells
is facilitated by 5′-OH and 2′,3′-cyclic PO4 termini created by Ire1 cleavage and the absence of
Trl1 end modification activities that could otherwise produce termini incompatible with RtcB
ligation (5′-PO4 or adenylylate; and 2′-PO4/3′-OH). It is noteworthy that circularized intron
accumulates to high levels in the absence of tunicamycin (Fig. 3.3F & 3.G), indicating that Ire1
catalyzes a low level of intron excision (and 3′-exon excision (Fig. 3.1C)) from HAC1u during
normal growth, leading to the accumulation of stable, circularized introns in the presence of
RtcB.
The HAC1 intron and 5′-UTR form an extensive base-pairing interaction that inhibits
ribosome initiation (Chapman and Walter 1997; Di Santo et al. 2016). Thus, together these data
evoke a model in which kinase-mediated decay of the excised HAC1 intron is required for HAC1s
translation, and a failure to degrade HAC1 intron—even when HAC1s is produced—prevents
HAC1s translation and subsequent expression of stress-responsive genes. Despite their ability
to make HAC1s, xrn1∆ and trl1∆ (RtcB) cells have growth defects on media containing
tunicamycin (Fig. 3.3A) and the relative severity of their defects parallels the accumulation of
excised intron in these cells. Cells lacking Xrn1 have a modest growth defect and accumulate
linear intron, which can be degraded by other exonucleases (Fig 3.3C-E, G). In contrast, trl1∆
(RtcB) cells have a severe growth defect and accumulate high levels of a stable, circularized
intron that is immune to exonucleolytic decay (Fig. 3.3F & G). I believe these findings resolve the
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mystery of the previously identified “second function” of Trl1 required for UPR activation (Mori
et al. 2010), namely that—in addition to its ligase activity—Trl1 initiates kinase-mediated decay
of the excised HAC1 intron, relieving its repressive effect on HAC1s and activating translation.
Incompletely processed HAC1 mRNA is endonucleolytically cleaved and degraded
In addition to the canonical 5′-exon product of 5′-splice site cleavage, a second product
uniquely accumulates in tpt1∆ and tpt1∆ ski2∆ cells that is ~50 nt shorter than full-length 5′-exon
(Fig. 3.4A). Expression of a catalytically inactive form of Tpt1 (Tpt1-R138A, (Sawaya et al. 2005))
in tpt1∆ cells failed to rescue this defect (Fig. 3.4B), affirming that the catalytic activity of Tpt1 is
required to prevent accumulation of the shorter 5′-exon fragment. A corresponding elongated
3′-exon fragment accumulates in tpt1∆, and more intensely in tpt1∆ xrn1∆ cells, indicating it is
degraded by Xrn1 (Fig. 3.4C-E). The elongated 3′-exon is specifically detected using a northern
probe with a sequence complementary to the distal 3′-end of the 5′-exon (Fig. 3.4C & D),
indicating that a portion of the 5′-exon is responsible for the increased size of this decay
intermediate. Moreover, a fragment of similar size hybridizes to a probe for the 3′-exon,
suggesting that the sequence derived from 5′-exon is linked to the 3′-exon (Fig. 3.4E); together
these data indicate that HAC1s is cleaved upstream of the 2′-phosphorylated ligation junction in
tpt1∆ cells, and these products are degraded by both Xrn1 and the cytoplasmic exosome.
To determine whether HAC1s sequence is sufficient for cleavage, I expressed plasmid-
encoded HAC1u and HAC1s in hac1∆ cells to identify HAC1 splicing intermediates. I performed
the analysis on trl1∆ cells, reasoning that if HAC1s sequence were sufficient to cause cleavage,
we would expect an accumulation of 3′-exon in cells unable to ligate or degrade the products
by kinase-mediated decay. Tunicamycin treatment of trl1∆ hac1∆ cells expressing HAC1u caused
Ire1-mediated cleavage and accumulation of cleaved 3′-exon (Fig. 3.4F, lane 2).
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Figure 3.4: Incompletely processed HAC1s mRNA is cleaved and degraded.
A. A shortened form of 5′-exon accumulates in tpt1∆ cells and is degraded by the cytoplasmic exosome. Northern blot analysis of tpt1∆ and tpt1∆ ski2∆ using a probe to 5′-exon identified a shortened form of liberated 5′-exon, about 50 nt smaller than full-length (red arrowheads). A region of enhanced contrast shows the specific accumulation of this product in tpt1∆ and tpt1∆
ski2∆ cells. The bottom diagram depicts the relative positions of probes 1 and 2 on the HAC1 5′-exon; a red arrowhead marks the putative site of cleavage.
B. Production of shortened 5′-exon requires the catalytic activity of Tpt1. Northern blot analysis of tpt1∆cells expressing either a plasmid-encoded wild-type copy of TPT1 or a catalytically-inactive missense mutant (tpt1-R138A) (Sawaya et al. 2005) using a probe for 5′-exon. Production of the cleaved 5′-exon seen in tpt1∆ cells (lane 4, red arrow marks the band) is rescued by plasmid-mediated expression of wild-type TPT1 (lane 6) but not by Tpt1-R138A (lane
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Figure 3.4 (continued): Incompletely processed HAC1s mRNA is cleaved and degraded.
8, red arrow marks the band), confirming that Tpt1 catalytic activity is required to prevent HAC1s secondary cleavage.
C. The shortened 5′-exon is missing a portion of its 3′-end. A northern blot was probed with 5′-exon probe 2, which hybridizes to the 3′-most 30 nt of HAC1 5′-exon (see diagram in A). Compared to the blot in A, the shortened 5′-exon species is absent (lanes 4 and 8) and instead the probe detects smaller bands consistent with the length of the elongated 3′-exon (see region of enhanced contrast).
D. A lengthened form of 3′-exon accumulates in tpt1∆cells. Total RNA from tpt1∆ and xrn1∆ cells was analyzed by northern blot with 5′-exon probe 2, a probe that anneals to the 3′-most 30 nt of HAC1 5′-exon. D and E share the same band interpretation key, so dashed lines have been drawn across to illustrate the different positions of typical 3′-exon (474 nt) and elongated 3′-exon (524 nt).
E. The elongated 3′-exon from tpt1∆ cells co-migrates and co-hybridizes with 5′-exon probe 2 (see D). Stripping the blot from D and re-hybridizing it with HAC1 3′-exon probe detects the same, elongated band(s) as in the 5′-exon probe 2 blot (lane 8), as well as HAC1 3′-exon bands of typical length.
F. Expression of “pre-spliced” HAC1s is not sufficient to promote cleavage. Total RNA from trl1∆
hac1∆ cells expressing HAC1u and HAC1s from a plasmid was analyzed by northern blot with a
3′-exon probe. Full length HAC1u expressed from this construct is cleaved upon tunicamycin
addition (lane 2). Expression of “pre-spliced” HAC1s yields a single product with no decay
intermediates (lanes 3 and 4), indicating that “pre-spliced” HAC1s—which was not produced by ligation and therefore lacks a 2′-phosphate—is not sufficient to recapitulate the cleavage found in tpt1∆ cells.
G. Expression of “pre-spliced” HAC1s is not sufficient to promote secondary cleavage. Total RNA from hac1∆ and tpt1∆ hac1∆ cells expressing HAC1u and “pre-spliced” HAC1s from a plasmid was analyzed by northern blot with a 3′-exon probe. In hac1∆ cells, full length HAC1u expressed from this construct is cleaved and ligated upon tunicamycin addition (lane 2). Expression of “pre-spliced” HAC1s yields a single product with no processing intermediates (lanes 3 and 4), indicating that “pre-spliced” HAC1s is not further processed. The HAC1u construct expressed in cells of genotype tpt1∆ hac1∆ (10x tRNA) gets cleaved upon tunicamycin treatment (lane 6), yielding a secondary cleavage fragment. Expression of pre-spliced HAC1s in tpt1∆ hac1∆ cells does not produce any additional processing fragments, indicating that 2′-phosphorylated products from HAC1u are required for secondary cleavage in tpt1∆ cells.
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However, I found that full length HAC1s was the only product produced in the trl1∆ hac1∆ cells
in the presence and absence of tunicamycin (Fig. 3.4F). Furthermore, expression of HAC1s in a
tpt1∆ background failed to produce additional fragments (Fig. 3.4G), whereas expression of
HAC1u is sufficient in the tpt1∆ background to produce free HAC1 3′-exon, consistent with its
secondary cleavage. Together, these results (Fig. 3.4F & 3.4G) indicate that the HAC1s transcript
is not sufficient to recapitulate the secondary cleavage, and that HAC1u must be cleaved and
ligated to produce the 2′-phosphorylated HAC1s secondary cleavage substrate.
To further characterize this secondary cleavage product, I determined the 5′ end of the
elongated 3′-exon. I analyzed the 5′-ends of cleaved 3′-exon by primer extension and found that
cleaved 3′-exon was barely detectable in wild-type cells upon tunicamycin addition, whereas 3′-
exon accumulated in trl1∆ cells due to a lack of kinase-mediated decay (Fig. 3.5A, lanes 2 and
8). In tpt1∆ cells treated with tunicamycin, two products accumulated upon tunicamycin addition:
a product consistent with canonical length 3′-exon and a small amount of elongated 3′-exon
(Fig. 3.5A, lane 4). The elongated product accumulated in tpt1∆ xrn1∆ cells, again indicating it is
degraded by Xrn1 (Fig. 3.5A, lane 6). To test this prediction (summarized in Fig. 3.5B), I measured
the susceptibility of 3′-exon fragments to treatment in vitro with recombinant Xrn1 (rXrn1/TEX).
As expected, fragments from xrn1∆ cells were degraded by rXrn1 (Fig. 3.5C), establishing that
they have 5′-PO4 termini, whereas 3′-exon fragments from trl1∆ cells were resistant to rXrn1
degradation (Fig. 3.5C), indicating that they have 5′-OH termini.
The instructive findings came from examining 3′-exon accumulation in tpt1∆ cells. The 3′-
exon fragment of canonical length that accumulates in tpt1∆ cells was resistant to rXrn1
treatment (Fig. 3.5C, lanes 11 and 12), while the elongated fragment of 3′-exon is susceptible to
rXrn1 treatment (Fig. 3.5C, lanes 15 and 16).
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Figure 3.5: A 5′- and 2′-phosphorylated HAC1 decay intermediate inhibits Xrn1.
A. Primer extension analysis of HAC1 3′-exon cleavage products. Primer extension using a probe for 3′-exon was performed on total RNA from wild-type, tpt1∆ (10x tRNA) , tpt1∆ xrn1∆
(10x tRNA) , and trl1∆ (10x tRNA) cells treated with or without tunicamycin. A loading control for U6 snRNA is depicted in the bottom panel. The extension product of cleaved 3′-splice site is 174 nt, found in wild-type cells treated with tunicamycin (lane 2) and is present in untreated trl1∆ c ells but increases upon tunicamycin treatment (lanes 7 and 8). An extension product from tpt1∆
cells accumulates upon tunicamycin treatment (lanes 3 and 4) and co-migrates with the product from wild-type (lane 2) and trl1∆ (lane 8) cells. In addition, a faint elongated product at ~225 nt is present in tpt1∆ cells (lane 4). This elongated product accumulates to higher levels in tpt1∆ xrn1∆ cells treated with tunicamycin (lane 6).
B. Model of HAC1 3′-exon primer extension product lengths. A 5′-radiolabeled (yellow star) primer (HAC1 3′-exon probe, see Table 2) anneals to HAC1 3′-exon mRNA and primes cDNA synthesis by a reverse transcriptase. The reverse transcriptase stops synthesizing cDNA when it runs out of RNA template at the 5′-terminus of the 3′-exon. The model shows three situations as observed in A: 1) canonical 3′-exon cleavage fragments (lower right), as observed in trl1∆ cells; 2) extended HAC1 3′-exon (above reaction arrow), as caused by secondary cleavage of
HAC1s, observed in tpt1∆ xrn1∆ (10x tRNA) cells; and 3′-extended HAC1 3′-exon, on which Xrn1(cellular or in vitro) initiated decay, but failed to proceed past the 2′-P (beneath arrow).
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Figure 3.5 (continued): A 5′- and 2′-phosphorylated HAC1 decay intermediate inhibits Xrn1.
C. Susceptibility of 3′-exon cleavage products to in vitro Xrn1 degradation. Total RNA from A
was treated with recombinant Xrn1 (rXrn1) and analyzed by primer extension for the 3′-exon. The loading control performed on U6 snRNA is depicted in the bottom panel. The 3′-exon from trl1∆ cells accumulates upon tunicamycin treatment (compare lanes 1 & 2 to 3 & 4) but the 3′-exon is resistant to rXrn1 (lanes 3 and 4) because it lacks a 5′-phosphate due to lack of Trl1 5′-kinase activity in these cells. In contrast, the 3′-exon products from xrn1∆ cells, which have Trl1 5′-kinase and thus 5′-phosphates, are degraded by rXrn1 (lanes 6 and 8). The 3′-exon product in tpt1∆ cells is resistant to rXrn1 treatment (compare lanes 9 to 10, and 11 to 12), despite the fact these cells have both Trl1 and Xrn1. The elongated 3′-exon product that accumulates in tpt1∆
xrn1∆ cells is partially degraded by rXrn1 (compare lane 15 to 16) and the decay intermediate co-migrates with cleaved 3′-exon.
D. Model depicting cleavage and decay of 2′-phosphorylated HAC1s. HAC1s is cleaved (likely by Ire1) ~50 nt upstream of the 2′-phosphorylated ligation junction, creating a 3′-product with ~50 nt of sequence of the 5′-exon (black) and an internal 2′-phosphate. Xrn1 initiates decay at the 5′-terminus, degrading the 5′-exon portion (black) up to the site of 2′-phosphorylation. The product of partial decay contains a 5′- and 2′-phosphate at its first position, which inhibits further degradation by Xrn1.
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These observations indicate that 3′ product of secondary cleavage of HAC1s is a substrate of
Xrn1 in vivo and in vitro, raising the possibility that it may also be a substrate of kinase-mediated
decay, depending on the chemistry of the endoribonuclease that generates the secondary
cleavage. I propose these products are created via two steps: (i) HAC1s is cleaved ~50 nt
upstream of the 2′-phosphorylated ligation junction; (ii) Xrn1 partially degrades the intermediate
fragment to the site of 2′-phosphorylation, which inhibits further degradation (Fig. 3.5D). Under
this model, the 3′-exon fragment that accumulates in tpt1∆ cells has both 5′-PO4 and 2′-PO4
moieties at its first position, which inhibits Xrn1-mediated decay in vivo and in vitro (Fig. 3.1C;
3.4D; 3.5A & C).
The accumulation of HAC1 decay intermediates in tpt1∆ cells over time further supports a
model of HAC1s cleavage by Ire1. In tpt1∆ cells, the accumulation of secondary cleavage product
coincides with the increase in production of HAC1s at 20 minutes (Fig. 3.6). In tpt1∆ cells, cleaved
HAC1 3′-exon is present at low levels at steady state and accumulates over the course of two
hours upon tunicamycin treatment (Fig. 3.6A). HAC1s is also generated in tpt1∆ cells, but at
significantly reduced levels compared to wild-type (Fig. 3.6A). It is also notable that tpt1∆ xrn1∆
cells (Fig. 3.6B) and tpt1∆ ski2∆ cells (Fig. 3.6D) accumulate more spliced HAC1s than tpt1∆ cells
(Fig. 3.6E), suggesting that some HAC1s molecules or splicing intermediates in tpt1∆ cells are
degraded, possibly because they contain 2′-PO4 moieties. At all time points, tpt1∆ cells contain
more free 3′-exon than HAC1s, a ratio opposite to wild-type cells (Fig. 3.6A), indicating that that
3′-exon cleaved from HAC1u accumulates as a result of partial decay of 5′- and 2′-
phosphorylated HAC1s. The augmented accumulation of 3′-exon in tpt1∆ cells is also observed
in the primer extension analysis (Fig. 3.5A).
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Figure 3.6: Kinetic analysis of HAC1 mRNA processing in cells lacking Tpt1.
A. Northern blot analysis of a time course of tunicamycin treatment (0 to 120 minutes) in tpt1∆
(10x tRNA) cells showing the dynamics of HAC1u splicing using a probe for 3′-exon. The SCR1
loading control is shown below the panel. RNA from wild-type cells treated for 0 and 120 minutes
tunicamycin was loaded in lanes 7 and 8. HAC1s accumulates slowly over 120 minutes and its
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Figure 3.6 (continued): Kinetic analysis of HAC1 mRNA processing in cells lacking Tpt1.
abundance at the end of the time-course is 35-fold lower than wild-type (lanes 6 and 8). A 3′-exon cleavage product (~475 nt) is present at time 0 and accumulates over time; its increase is
coincident with the appearance of HAC1s (20-minute time point) and its abundance at 120
minutes exceeds the level of HAC1s . A 3′-exon cleavage product in wild-type cells (~500 nt) is apparent at 120 minutes (lane 8).
B. Northern blot analysis of HAC1 splicing in tpt1∆ xrn1∆ (10x tRNA) cells using a probe for 3′-exon. Conditions are the same as A. HAC1s increases at 20 minutes and its final level (lane 6) is
higher than HAC1s in tpt1∆ cells (A, lane 6), approaching wild-type levels (lane 8). Levels of the product of 5′-splice site cleavage (intron/3′-exon) accumulate over 120 minutes. The 3′-exon product (~475 nt) and cleaved 3′-exon (~500 nt) begin accumulating at 20 minutes, coincident
with increased levels of HAC1s . A 3′-exon cleavage product in wild-type cells (~500 nt) is apparent at 120 minutes (lane 8).
C. Northern blot analysis of HAC1 splicing in tpt1∆ (10x tRNA) cells using a probe for 5′-exon.
Conditions are the same as A. HAC1s increases at 20 minutes up to final level (lane 6) (A, lane 6), approaching wild-type levels. Canonical 5′-exon (~725 nt) and secondarily-cleaved 5′-exon (~675 nt; red arrowheads) begin accumulating at 20 minutes, coincident with increased levels of
HAC1s.
D. Northern blot analysis of HAC1 splicing in tpt1∆ ski2∆ (10x tRNA) cells using a probe for 5′-exon. Deletion of SKI2 in the context of tpt1∆ increases the abundance of nearly all intermediates relative to tpt1∆ alone (A). Shortened HAC1 5′-exon is marked with red arrowheads. The product of 3′-splice site cleavage (5′-exon/intron) is present at time 0 and accumulates over 120 minutes,
whereas HAC1s accumulates beginning at 20 minutes.
E. Northern blot analysis of HAC1 splicing using a probe for 5′-exon to compare the beginning and end time points of time course experiments performed on tpt1∆ , tpt1∆ xrn1∆, and tpt1∆ ski2∆. Results from densitometry of the spliced and unspliced bands are written beneath the blot. Less HAC1 splicing takes place in tpt1∆ cells compared to wild-type, and disabling decay factors ski2∆ and xrn1∆ in the tpt1∆ mutant lead to increased HAC1 splicing, compared to tpt1∆ alone.
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Xrn1’s in vitro ability to only partially degrade elongated HAC1 3′-exon from tpt1∆ xrn1∆ cells,
and inability to degrade canonical 3′-exon from tpt1∆ cells, indicates that the accumulation of
free HAC1 3′-exon is likely caused primarily by blocked 5′→3′ degradation.
Discussion
Many different regulatory events impinge on HAC1 mRNA to control its localization and
processing. It has been assumed that cleavage of HAC1u by Ire1 is the rate-limiting step for UPR
activation. Counter to this view, we found that decay of HAC1 splicing intermediates is required
for both UPR activation and suppression. I found several examples wherein “kinase-mediated
decay” (KMD) degrades HAC1 splicing intermediates containing 5′-OH termini by sequential 5′-
phosphorylation and 5′-phosphate-dependent 5′→3′ exonucleolytic degradation (Fig. 3.7).
I propose that after 3′-splice site cleavage by Ire1, the Trl1 5′-kinase domain associates
with and phosphorylates the 5′-OH of the 3′-exon product (Fig. 3.7). Dissociation of the Trl1
kinase active site from the 5′-PO4 product then enables a competition between re-association
of Trl1 (now its adenylyltransferase/ligase domain) to catalyze ligation—or Xrn1 to catalyze
degradation. In some circumstances, this balance is tipped to favor ligation even in the absence
of overt UPR stimulation: a lack of decay in xrn1∆ cells favors ligation, whereas the lack of 5′-
kinase activity in trl1∆ (RtcB) cells renders Xrn1 decay irrelevant (Fig. 3.2F). Xrn1 is abundant in
budding yeast (Ghaemmaghami et al. 2003), which may efficiently suppress the UPR under
normal conditions by degrading spuriously cleaved HAC1 3′-exon intermediates. Regulation of
the ligation step of HAC1s splicing makes intuitive sense because it is the last opportunity to act
during splicing; once ligated, HAC1s is again protected by a 7-methylguanosine cap and a poly-
(A) tail.
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Figure 3.7: Decay of HAC1 splicing intermediates regulates UPR activation, suppression,
and attenuation.
Activation of Ire1 by ER stress activates its endoribonuclease activity, leading to cleavage of the
5′- and 3′-splice sites of HAC1u. The cleaved 3′-exon (grey) is phosphorylated by the 5′-kinase activity of Trl1, permitting either its ligation or decay by Xrn1. The 5′-exon (black) and intron (thin line) form an extensive base-pairing interaction (grey squiggle). Kinase-mediated decay of the
intron is required to activate the translation of HAC1s . Ligated but 2′-phosphorylated HAC1s may also be cleaved by Ire1 upstream of the ligation junction, and the cleavage products are degraded by Xrn1 and the cytoplasmic exosome. In cells lacking Tpt1, a unique KMD intermediate accumulates with 5′- and 2′-phosphates, which inhibit Xrn1-mediated degradation.
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Spurious HAC1 splicing was previously reported in trl1∆ cells expressing the tRNA ligase from
Arabidopsis thaliana (Mori et al. 2010). It is noteworthy that there are mechanistic differences
between the tRNA ligases of A. thaliana and another yeast (K. lactis) (Remus and Shuman 2014),
suggesting that these differences could impact the balance between kinase-mediated decay and
ligation in budding yeast expressing plant RNA ligase.
I also found that excised HAC1 intron is a substrate of kinase-mediated decay (Fig. 3.3).
Indeed, we believe that phosphorylation of HAC1 intron by Trl1 to promote kinase-mediated
decay is the previously proposed “second role” of Trl1 ligase in activating HAC1 translation
independent of ligation (Mori et al. 2010). In this previous study, excised and circularized HAC1
intron was found to remain associated with HAC1s, inhibiting translation. Kinase-mediated decay
of excised intron therefore likely relieves the long-range base-pairing interaction that prevents
HAC1s translation (Cox et al. 1997; Rüegsegger et al. 2001; Di Santo et al. 2016), explaining how
HAC1s can accumulate without concomitant UPR activation. This second layer of control over
HAC1s translation by KMD adds another failsafe mechanism to prevent its translation and
unintentional UPR activation.
Previous examples of kinase-mediated decay of bacterial mRNAs (Durand et al. 2012),
eukaryotic tRNA introns (Wu and Hopper 2014), ribosomal RNA processing intermediates (Gasse
et al. 2015), and no-go mRNA decay cleavage products (Navickas et al. 2018) suggest that this
mode of decay may be widespread. Coupling of RNA 5′-kinase and 5′→3′ exonucleolytic decay
activities in the context of kinase-mediated decay may regulate the UPR in other organisms.
Splicing of Xbp-1 mRNA in metazoans (the functional homolog of HAC1) is catalyzed by Ire1-
mediated removal of an intron and ligation by RtcB RNA ligase (Kosmaczewski et al. 2014).
Fundamental differences in the chemistry of RNA ligation between fungal and metazoan ligases
suggest that Xbp-1 may be subject to a biochemically distinct mode of regulation. Because RtcB
depends on 5′-OH and 3′-PO4 termini for catalysis (Chakravarty et al. 2012), activities that
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remodel 5′-OH RNA termini could divert Ire1-generated 5′-OH splicing intermediates from
productive ligation. To that point, cyclic nucleotide phosphodiesterase (CNP) and RtcA (a 2′,3′-
RNA cyclase) were shown to “tune” the UPR in metazoans by competing with RtcB/HSCP117
for ligation substrates (Unlu et al. 2018). Specifically, CNP hydrolyzes the 2′,3′-cyclic phosphate
compatible RtcB to a 2′-PO4 incompatible with RtcB, decreasing ligation of Xbp1. Conversely,
RtcA, which converts 2′-PO4 RNA to 2′,3′-cyclic phosphate, makes the terminus compatible
with RtcB, thus enhancing Xbp1 splicing.
Additionally, an RNA 5′-kinase (e.g., Clp1) may phosphorylate the 3′-exon product of Xbp-
1 cleavage, simultaneously inhibiting ligation by RtcB and promoting its degradation by a 5′-
phosphate-dependent exoribonuclease to limit UPR activation. In this vein, it is noteworthy that
kinase-inactivating mutations in Clp1 cause neurodevelopmental defects and neuronal
dysfunction in humans, mice, and zebrafish (Hanada et al. 2013; Karaca et al. 2014; Schaffer et
al. 2014), possibly due to chronic UPR activation in neural tissues (Clayton and Popko 2016).
Xrn1 degrades mRNA fragments generated during metazoan Regulated Ire1-dependent
Degradation (RIDD) (Hollien and Weissman 2006); however, it is not known how these 5′-OH
cleavage products of Ire1 are phosphorylated for Xrn1-mediated decay. The RNA 5′-kinase
Clp1(Weitzer and Martinez 2007) and polynucleotide kinase Nol9 (Heindl and Martinez 2010) are
candidates for this activity, though neither are known to phosphorylate mRNA decay
intermediates.
While 5′→3′ decay plays a major role in UPR regulation, I found little evidence for UPR
regulation by 3′→5′ decay activity. As shown previously (Schwer et al., 2004), the exposed 3′-
end of cleaved HAC1 5′-exon is a substrate for 3′→5′ decay (Fig. 3.4) . But while excised HAC1
intron is stabilized in ski2∆ cells lacking cytosolic 3′→5′ decay (Fig. 3.3H), their growth is
unaffected by tunicamycin (Fig. 3.1B) , indicating that 3′→5′ decay of the excised intron does
not contribute substantially to intron-mediated HAC1s repression.
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I also found evidence that Ire1 cleaves incompletely processed HAC1s mRNA, which is
ligated but contains an internal 2′-PO4 moiety. Cleavage of HAC1s leads to 5′- and 3′-fragments
that are degraded; however, the 3′-fragment is only partially degraded by kinase-mediated
decay, producing a 5′- and 2′-phosphorylated molecule that cannot be degraded by Xrn1.
Consistent with these findings, a recent study also showed that an RNA with an internal 2′-
phosphate group is protected from 3′→5′ decay by E. coli PNPase in vitro (Munir et al. 2018b);
those and our results together indicate that site-specific installation of a 2′-PO4 is an effective
strategy to protect an RNA from complete exonucleolytic degradation in vivo or in vitro.
Decay intermediates produced from incompletely processed, 2′-phosphorylated HAC1s
have not been previously observed and suggest a plausible regulatory role for Tpt1 in regulating
HAC1s fate. I have yet to determine the impact of 2′-phosphorylation on HAC1s translation, but
given that tpt1∆ cells grow on tunicamycin (Fig. 3.1B & (Cherry et al. 2018)) and activate KAR2
gene expression (Fig. 3.3B), it is likely that some Hac1 protein is produced from 2′-PO4 HAC1s
mRNA. It also remains to be determined how and why incompletely processed HAC1s is cleaved.
Insofar as the HAC1s cleavage substrate is initially produced by tunicamycin-dependent Ire1
cleavage and ligation, we conjecture that Ire1 incises ligated, 2′-phosphorylated HAC1s
upstream of the original ligation junction, yielding smaller 5′-exon and larger 3′-exon fragments.
Formally, I cannot currently rule out the possibility that another endonuclease catalyzes
secondary HAC1s cleavage; however, Ire1 is the only endoribonuclease known to site-
specifically incise HAC1 mRNA. I note that ire1∆ mutants are unable to initiate processing of
HAC1u and therefore do not make HAC1s in the first place (Sidrauski and Walter 1997),
precluding direct analysis of HAC1s cleavage in ire1∆ mutants.
I showed that the 2′-PO4 is required for cleavage, as expression of “pre-spliced” HAC1s
does not lead to cleavage (Fig. 3.4F & G). It is possible that the presence of a 2′-PO4 in the
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context of a composite Ire1 splice site (i.e., formed from two halves of the original 5′- and 3′-
splice sites (Sidrauski and Walter 1997; Hooks and Griffiths-Jones 2011)) on HAC1s is recognized
by Ire1, but because a 2′-OH is the nucleophile for transesterification by metal-independent Ire1
(Gonzalez et al. 1999), the 2′-PO4 inhibits the chemical step of incision. This model also provides
a plausible mechanism to explain how Ire1 incises a neighboring, non-canonical site. We
propose that a failure of Ire1 to release 2′-phosphorylated HAC1s would enable the active site of
a nearby Ire1 molecule—in the context of its activated, oligomeric form (Korennykh et al. 2009)—
to catalyze site-specific incision at the second, upstream site. My results also raise the question
of why Ire1 would cut incompletely processed HAC1s. Here, cleavage of incompletely processed
(ligated, but 2′-phosphorylated) HAC1s could be a means to inactivate HAC1s after prolonged
stimulation to attenuate the UPR (Chawla et al. 2011; Rubio et al. 2011).
Summary
In the unfolded protein response (UPR), protein-folding stress in the endoplasmic reticulum
(ER) activates an extensive transcriptional program to increase ER folding capacity. During the
budding yeast UPR, the trans-ER-membrane kinase-endoribonuclease Ire1 excises an intron
from the HAC1 mRNA and the exon cleavage products are ligated and translated to a
transcription factor that induces dozens of stress-response genes. HAC1 cleavage by Ire1 is
thought to be the rate limiting step of its processing. Using cells with mutations in RNA repair
and decay enzymes, I showed that phosphorylation of two different HAC1 splicing intermediates
by Trl1 RNA 5′-kinase is required for their degradation by the 5′→3′ exonuclease Xrn1 to enact
opposing effects on the UPR. Kinase-mediated decay (KMD) of cleaved HAC1 3′-exon competes
with its ligation to limit productive splicing and suppress the UPR, whereas KMD of the excised
intron activates HAC1 translation, likely by relieving an inhibitory base-pairing interaction
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between the intron and 5′-untranslated region. We also found that ligated but 2′-phosphorylated
HAC1 mRNA is endonucleolytically cleaved, yielding a decay intermediate with both 5′- and 2′-
phosphates at its 5′-end that inhibit 5′→3′ decay and suggesting that Ire1 initiates the
degradation of incompletely processed HAC1s to proofread ligation or attenuate the UPR. These
multiple decay events expand the scope of RNA-based regulation in the budding yeast UPR and
may have implications for the control of the metazoan UPR by mRNA processing.
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CHAPTER IV
IV FUTURE DIRECTIONS
This work illustrates four contributions to the field: (i) the only essential functions of Trl1
and Tpt1 are tRNA splicing; (ii); the tRNA splicing endonuclease (SEN) has an additional essential
function beyond tRNA splicing; (iii) kinase-mediated decay (KMD) occurs on two substrates,
HAC1 intron to relieve unfolded protein response repression and on HAC1 3′-exon to suppress
spurious UPR activation; and (iv) 2′-phosphorylated RNA is stabilized from Xrn1 degradation.
Many unanswered questions remain, addressed in detail below. Kinase-mediated decay occurs
on 5′-OH substrates in yeast, and likely other organisms that use a Trl1/Tpt1 mechanism of
ligation. But what about in organisms, like humans, that use a RtcB mechanism? Also, beyond
tRNAs, does RtcB serve an additional essential function? Can this Trl1/Tpt1 genetic bypass in
yeast be applied to identify the products of RNA repair, transcriptome-wide? And can the 2′-PO4
stalling of Xrn1 degradation be put to use to stabilize RNAs of our choice? What utility could
these 2′-PO4 stabilized RNAs have?
Does kinase-mediated decay also regulate Xbp1 splicing in animals?
In Chapter III of this dissertation (Cherry et al. 2019) I unveiled the first examples of kinase-
mediated decay (KMD) on mRNA. For both the 3′-exon and intron of HAC1 mRNA, the
endoribonucleolytic cleavage releases them as 5′-hydroxyl fragments that can be 5′-
phosphorlated and subsequently degraded by Xrn1. In addition to these being the first examples
of KMD among mRNAs, they also regulate/tune activation of the unfolded protein response, and
without them UPR signaling functions improperly. In this section, I lay out how KMD may regulate
the metazoan UPR.
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Animals express an RNA ligase with a different biochemical mechanism than that of yeast,
but that difference could still permit a competition between decay and ligation. In fungi and
animals, Ire1 (IRE1α) functions as a metal-ion-independent endoribonuclease, releasing
products with 5′-OH termini. Mammalian RtcB/HSPC117 can then take those 5′-OH termini and
use them directly in ligation (Fig. 1.3). Because RtcB guanylates the 3′-terminus of the other
ligation substrate, RtcB biochemically requires a 5′-OH RNA to ligate. Therefore, an RNA 5′-
kinase could interfere with the ligation reaction by phosphorylating the 5′-terminus before RtcB
encounters it (Fig. 4.1). Humans have two RNA 5′-kinases that could participate in this
competition: Clp1 and Nol9 (Weitzer and Martinez 2007; Heindl and Martinez 2010). Both
proteins catalyze the ATP-dependent phosphorylation of the 5′-terminus of RNA with little
sequence specificity. Competition experiments could be carried out in the trl1∆ (RtcB) yeast
backgrounds already on hand, but that would be rather indirect because it would test
competition for substrate between enzymes from two different organisms expressed in yet
another organism. Furthermore, there is the potential for a synthetic lethal interaction of
expression, which may occur if the chosen 5′-kinase vastly outcompetes RtcB (by activity or
sheer protein abundance) for cleaved tRNAs, which RtcB is ligating together to keep the cells
alive. That result itself would be interesting and informative about the relative activity of the two
enzymes, but it would be an indirect assessment of competition.
A more human-specific experiment would be to knock-down and overexpress Clp1 and
Nol9 RNA 5′-kinases and test for a difference in Xbp1 splicing without and with UPR stimulation
(with thapsigargin or tunicamycin). Clp1 is an important structural protein in the RNA polymerase
II cleavage and polyadenylation complex (Minvielle-Sebastia et al. 1997; de Vries et al. 2000),
which may necessitate relegating Clp1 exclusively to the nucleus with a Clp1-nuclear-
localization-sequence construct that would limits its contact with cytoplasmic Xbp1 mRNA.
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Figure 4.1: Hypothesis of competition between decay and ligation in animals.
At left, the model depicts the 3′-exon of HAC1 released from Ire1 cleavage with a 5′-hydroxyl terminus. The multifunctional Trl1 phosphorylates the 5′-terminus. As a 5′-phosphate, the 3′-exon can either be ligated (Trl1) or be degraded (Xrn1). On the right, a proposed model of competition in animals shows the 3′-exon of Xbp1 immediately after cleavage by Ire1α with a 5′-OH. At this step, the exon can either be ligated (RtcB/HSPC117) or be phosphorylated (Clp1 kinase). The ligation pathway would commit the 3′-exon to activating the UPR via Xbp1 translation, whereas the 5′-PO4 3′-exon would be degraded by Xrn1.
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To test the disease relevance of Clp1 and UPR signaling, one of the disease-linked missense
mutations (clp1-R140H or -G419A) (Karaca et al. 2014) could be expressed to test for spurious
Xbp1 splicing, as would be expected from compromised kinase activity if decay and ligation
compete.
However, there are several reasons that KMD-based regulation of the UPR is likely not
occurring in animals. Firstly, compartmentalization of RNA 5′-kinases may deprive the cytoplasm
of robust kinase activity. Clp1 is ~85% localized to the nucleus (Schaffer et al. 2014); Nol9 is a
nucleolar-localized protein that participates in ribosomal RNA maturation (Heindl and Martinez
2010). Secondly, RtcB is an efficient ligase, and competing with its rapid catalysis would require
a sufficiently rapid 5′-kinase. While human RtcB/HSPC117 has not been directly tested for its
ligation prowess, E. coli RtcB, which operates by the same biochemical mechanism, is a fast
ligase, especially for RNA termini held in proximity to one another (Tanaka and Shuman 2011).
Thirdly animals have two other signaling mechanisms for rectifying ER stress, ATF6 and PERK,
which can compensate for signaling (to some degree) and thus limit the evolutionary pressure to
develop intricate or nuanced mechanisms of regulation on the Ire1/Xbp1 arm of the response.
Furthermore, regulation of the UPR in animals via KMD on the intron of Xbp1 is unlikely because
the intron has not been shown to participate in regulation once cleaved out of the mRNA. (In
contrast, the “dual function” of Trl1 in yeast is to start KMD of HAC1 intron (Mori et al. 2010).) In
fact, the secondary structure of Xbp1 favors ejection of the intron upon cleavage at the exon-
intron junctions, zipping up to position the 5′- and 3′-splice sites in close proximity for rapid
ligation (Peschek et al. 2015).
Identify precise HAC1s secondary cleavage location and nuclease.
Products of secondary cleavage of HAC1s are observed in tpt1∆ cells. The 5′-fragment,
shortened by the cleavage, accumulates in tpt1∆ ski2∆ cells (Fig. 3.4A); the 3′-fragment,
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lengthened by the cleavage, accumulates in tpt1∆ xrn1∆ cells (Fig. 3.4D & E). As the project
currently stands, it is not clear how these secondary cleavage fragments are generated or where
precisely the secondary cleavage occurs on the HAC1s mRNA. Below, I state two hypotheses
about the mechanism of secondary cleavage, and I propose two genetic experiments to test the
hypotheses. I also propose two methods to precisely identify the site of secondary cleavage of
HAC1s.
Given that expression of “pre-spliced” HAC1s from a plasmid is not sufficient for secondary
cleavage of the transcript, the 2′-PO4 appears to be a necessary feature of the HAC1s mRNA
from tpt1∆ cells for secondary cleavage. Thus, all experiments on this topic will have to occur
on tunicamycin-treated tpt1∆ cells so that 2′-PO4 HAC1s mRNA can be generated. Two
hypotheses I have for the generation of this fragment are: (1) Ire1 cleaves a different site on 2′-
PO4 HAC1s; or (2) no-go mRNA decay occurs because 2′-PO4 in an open reading frame can stall
ribosomes during translation elongation.
Testing whether Ire1 is cleaving HAC1s is complicated by the requirement for Ire1 to
generate 2′-PO4 HAC1s in tpt1∆ cells. Therefore, I would generate a degron-tagged Ire1 strain
(Johnston et al. 1995; Sheridan and Bentley 2016) in the tpt1∆ background so that Ire1 could be
stabilized in the presence of methotrexate to produce 2′-PO4 HAC1s, and then depleted by
removal of methotrexate from the media. With a starting amount of HAC1s in the cells, northern
blotting RNA harvested in a time course after degradation of Ire1 would show whether any
additional secondary cleavage products accumulate after removal of Ire1. A failure to accumulate
more secondary cleavage product would indicate that Ire1 is required for the secondary
cleavage. Given that Ire1 is an endonuclease already known to cleave HAC1, the finding would
debut a curious function of Ire1, where it cleaves a product (at a non-canonical site) that it
generated, meaning that it can antagonize the same signaling pathway that it activates. Such a
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function is reminiscent of a quality-control pathway, where Ire1 would recognize defective
products and mark them for decay.
Alternatively, secondary cleavage HAC1s products could be caused by no-go mRNA
decay, wherein an elongating ribosome(s) stalls upon encountering the 2′-PO4 in the open
reading frame. It is unclear how much Hac1 protein is produced in tpt1∆ cells because the
epitope tag I chose to use in Chapter II was a C-terminal tag, and therefore would be synthesized
less than an N-terminal tag on an mRNA that causes stalls in translation. If no-go decay is
responsible for the secondary cleavage of HAC1, a simple genetic deletion of DOM34, the
release-factor-like protein that detects and rescues stalled ribosomes, would prevent the
secondary cleavage, and so the fragment would become undetectable on northern blots or
primer extension assays. A complementary approach would be to express a mutant HAC1
mRNA with no start codons and so would test whether translation is required for generating the
secondary cleavage.
The precise 5′-end of the extended 3′–fragment generated from secondary cleavage of
HAC1s remains undetermined. Northern blot and primer extension results suggest that the
secondary cleavage of HAC1s occurs ~40 nt upstream of the original ligated site, generating the
observed products. Simultaneously, northern results indicate that sequence from the
downstream region of the 5′-exon goes missing from the shortened form of 5′-exon, consistent
with cleavage at an alternative site. I propose to perform 5′-end capture, PCR amplification, and
Sanger sequencing of extended 3′-exon to determine precisely where the secondary cleavage
site is. A similar process could be done with 3′-end-capture of the shortened 5′-exon.
Alternatively, the sequence of the extended 3′-exon could be read of by dideoxy-sequencing the
RNA with ddNTPs spiked into primer extension reactions. Identifying the precise location of
secondary cleavage could provide insight into the nuclease that cleaves at that site and would
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provide helpful information for performing in vitro experiments to recapitulate the 2′-phosphate-
dependent cleavage of HAC1 RNA.
Applications of 2′-phosphorylated RNA to enhance stability in vivo
Typical cellular RNAs resist 5′→3′ decay via the regulated process of capping and
decapping (Parker 2012). Typically, 5′-PO4 RNAs are destined for decay by the abundant and
processive exonuclease Xrn1. However, in the course of studying the effect of tpt1∆ on the
unfolded protein response, I found via northern blot that an elongated form of HAC1 3′-exon
was present in cells at the expense of a shortened 5′-exon (Fig. 3.4). Curiously, the stabilized
3′-exon accumulated at its typical length in tpt1∆ cells, but tpt1∆ xrn1∆ caused the band to shift
upward, suggesting additional sequence at the 5′-end, ruling out the previously hypothesized
reason for variability of the band: poly(A) tail length changes. Primer extension analysis revealed
that only those two products are recovered from cells, consistent with a processive decay factor.
Recombinant Xrn1 (rXrn1) incubated with RNA samples in vitro degraded extended 3′-exon to
the canonical 3′-exon site, where the 2′-PO4 resides in tpt1∆ cells. Thus, I concluded that internal
2′-phosphorylation of a substrate of Xrn1 is necessary for the stall in decay I observe (though it
may not be sufficient). Furthermore, I found that 3′-exon from tpt1∆ cells that has already been
partially degraded by cellular Xrn1 was resistant to further decay by rXrn1 in vitro, indicating that
a 5′-PO4, 2′-PO4 nucleotide at the beginning of an RNA molecule can prevent Xrn1 degradation.
Structures that stabilize RNAs from 5′→3′ decay are sought after by scientists and
biological agents alike. Scientists seek stable RNAs to maintain high levels of gene expression
in experiments and to ensure long half-lives of RNA-based drugs delivered to patients. Viruses
steal 5′-caps and fold their RNA genomes into three-dimensional structures to evade
degradation by 5′→3′ exoribonucleases (Plotch et al. 1979; 1981; Chapman et al. 2014). While
the 2′-PO4 left behind on the HAC1s mRNA (and on tRNAs) in tpt1∆ cells may not be
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physiological, the modification nonetheless represents a means of stabilizing an RNA of one’s
choice. Much like the Xrn1-resistant RNA structures, 2′-PO4 could be chemically or enzymatically
installed on an RNA of one’s choice, either terminally or internally. A 5′-terminal 2′-PO4 would
stabilize the RNA in its entirety, whereas an internal 2′-PO4 would permit decay of the RNA up
to a point. Furthermore, the 5′,2′-phosphorylated RNA would be stable even in cells with
functioning Tpt1/Trpt1 activity because the 2′-phosphotransferase activity of Tpt1/Trpt1 appears
restricted to internal 2′-sites, whereas terminal 2′-PO4 would likely be resistant to Tpt1/Trpt1
activity (Zillmann et al. 1992; Culver et al. 1997).
The functions of RNAs stabilized with 5′-PO4, 2′-PO4 termini would likely be restricted to
non-coding roles in the cell, with the one exception of open reading frames translated with the
help of an Internal Ribosome Entry Site (IRES) (Jang et al. 1988). Because translation initiation
largely relies on 5′-caps and poly(A) tails (Dever et al. 2016), a 2′-PO4 stabilized RNA would likely
need another means of recruiting the translational machinery to the ORF. However, the plethora
of non-coding functions of RNAs (e.g. long non-coding RNAs, antisense RNAs, RNAi) would all
be compatible with the 2′-PO4 stabilization technique.
Transcriptome-wide sequencing of products of RNA repair via enrichment of 2′-PO4 RNAs
from tpt1∆ yeast
What RNAs get cleaved and put back together? The field of RNA repair is interested in
identifying the substrates and/or products of RNA repair, especially in eukaryotes. While the
mechanism of repair (i.e. the enzymes) are well studied, their substrates are less so.
Endonucleolytic cleavage of mRNA has a broad range of nucleases and substrates, only a
handful of which are known to be repaired. Studies that captured and sequenced the specific
termini of endoribonucleolytic cleavage (2′,3′-cP capture and 5′-OH capture) (Schutz et al. 2010;
Cooper et al. 2014; Peach et al. 2015) have detected some promising leads, but as many
125
examples show, not all cleaved RNAs are destined to be ligated back together. Currently, the
limited number of known RNA repair substrates include bacterial rRNA, tRNA, and HAC1/Xbp1
mRNA. But is RNA repair a broader phenomenon in biology? The tpt1∆ (10x tRNA) yeast may be
able to help us find out. If capturing the substrates of repair (e.g. products of cleavage) haven’t
answered this question, the next logical step is to capture products of ligation.
Cells with a Trl1/Rnl1 ligation system that are ablated for Tpt1/Trpt1 activity accumulate
products of ligation with a stable 2′-PO4 at the ligation junction. This 2′-PO4 is a site-specific
mark of ligation, and so provides evidence that RNA repair took place on that molecule at that
site. Enrichment of these RNAs could be accomplished by exploiting a characterized mutation
of Tpt1 that could transfer a biotin group to 2′-PO4 sites. Tpt1 uses a 2-step mechanism to
remove 2′-PO4: (i) use NAD+ to attach an ADP-ribose molecule to the 2′-PO4, displacing the
nicotinamide group, and then (ii) use the neighboring 2″-OH to displace the RNA away from the
ADP-ribose. The mutant Tpt1-K69A-R71S has a compromised second step, meaning that the
mutant enzyme will quickly transfer an ADP-ribose group onto a 2′-PO4, but then slowly remove
the ADP-ribose group (Steiger et al. 2005). This difference in rates would generate a population
of marked RNAs toward the beginning of incubation with the mutant Tpt1 in vitro. If, instead of
NAD+, 6-Biotin-17-NAD+ were given as a substrate for Tpt1-K69A-R71S in vitro reaction, the
RNAs would be marked with an affinity-resin compatible tag. To pull down the biotinylated RNAs,
Tpt1 would need to be inactivated, and the labeled RNA would be affinity-purified from the rest
of the RNA. After binding to the resin, either the RNA could be reverse transcribed into a cDNA
library, or the RNA could be fragmented followed by the library preparation, or the RNA even
eluted with wild-type Tpt1, which would complete the reaction and release the RNAs with a
seamless 2′-OH at the ligation junction. Enrichment of RNAs via Tpt1-assisted affinity-
purification could identify which RNAs undergo repair, but the method may only yield limited
information about the specific site of repair.
126
An alternative approach, in light of the Xrn1-stall observation, would be to digest the RNA
up to sites of 2′-PO4 in vitro, and then to perform 5′-end capture and sequencing on the RNAs
that survive incubation with Xrn1. The benefit of this method would be that the site of repair
could be identified at single-nucleotide resolution, as is indicated by the rXrn1 digestion and
primer extension results on 2′-PO4 HAC1 3′-exon (Fig. 3.5). However, Xrn1 can be halted by
elements other than 2′-PO4, so the assay could be made specific to 2′-PO4 by performing paired
Tpt1± pre-treatments in vitro on the RNA samples to be analyzed by subsequent rXrn1
degradation and library preparation. Site-specific information would not only identify the RNAs
on which repair occurs but would also identify the sequence motif at which the repair occurs,
obtaining data that could aid in determining which possible RNAs are regulated in a concerted
manner based on primary or secondary structure. These classes of recognition motifs could then
be associated with RNA cleavage or repair programs that activate under specific circumstances,
ultimately establishing the paradigm of regulation by RNA breakage and reunion.
“Fungification” of metazoan cells to replace RtcB with Trl1 for marking repaired transcripts
with 2′-PO4.
The capture and sequencing of 2′-PO4 RNAs could be extended to organisms that naturally
use a different, mutually-exclusive mechanism of ligating RNA (i.e. RtcB/HPSC117). Recently,
Trl1 was used to replace RtcB in mouse embryonic stem cells (Unlu et al. 2018). However, the
Trpt1 gene (Tpt1 homolog), previously shown to be dispensable in mouse cells (Harding et al.
2008), became essential upon switching the ligase to Trl1, consistent with Trpt1 complementing
tpt1∆ in yeast (Hu et al. 2003) and with mammalian cells using RtcB (Popow et al. 2011), a ligase
that does not yield 2′-phosphorylated products. That fact means that, after “yeast-ifying” the
RNA ligase of mammals, a “pre-spliced” tRNA bypass mechanism could be used to bypass the
essential function of Trpt1, allowing Trpt1 to be deleted and permitting the cell to accumulate
127
2′-PO4 products of ligation. As described previously, these could then be captured and
sequenced to determine how widespread RNA repair is, and on what RNAs it typically acts.
128
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