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Regulation of Metabolic Stress-Induced snoRNAs Regulation of Metabolic Stress-Induced snoRNAs

Benjamin Steel Scruggs Washington University in St. Louis

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WASHINGTON UNIVERSITY IN ST. LOUIS

Division of Biology and Biomedical Sciences

Molecular Cell Biology

Dissertation Examination Committee Jean Schaffer, Chairperson

Peter Crawford Eric Galburt

Kathleen Hall Rob Mitra Daniel Ory

Heather True

Regulation of Metabolic Stress-Induced snoRNAs

by

Benjamin Steel Scruggs

A dissertation presented to the Graduate School of Arts and Sciences

of Washington University in partial fulfillment of the

requirements for the degree of Doctor of Philosophy

December 2012

Saint Louis, Missouri

ii

Table of Contents

List of Figures and Tables............................................................................................. v

List of Abbreviations.................................................................................................... vii

Acknowledgements....................................................................................................... x

Abstract....................................................................................................................... xii

 Chapter 1: Introduction to Lipotoxicity

Type 2 diabetes is a growing worldwide health concern ..............................................1

Lipotoxicity is a hallmark of metabolic disease .............................................................1

Mechanisms linking lipid overload to cell death............................................................2

Elucidating molecular mechanisms of lipotoxicity .........................................................3

Conserved functions of snoRNAs.................................................................................4

Palmitate treatment induces rpL13a snoRNA accumulation in CHO cells ...................6

Confirmation of the role for rpL13a snoRNAs in lipotoxicity ........................................7

Biogenesis of box C/D and H/ACA snoRNPs ...............................................................7

Summary.....................................................................................................................10

Chapter 2: rpL13a snoRNAs function as non-canonical box C/D snoRNAs

Introduction .................................................................................................................15

Results ........................................................................................................................16

In vivo expression of rpL13a snoRNAs ...................................................................16

In vivo rpL13a snoRNA knockdown reduces oxidative damage .............................17

rpL13a guided rRNA modifications are unaltered during lipotoxicity.......................17

rpL13a snoRNAs accumulate in the cytoplasm during lipotoxicity ..........................18

iii

Conclusions ................................................................................................................19

Figures ........................................................................................................................22

Chapter 3: SmD3 regulates intronic snoRNA biogenesis

Introduction .................................................................................................................30

Results ........................................................................................................................31

SmD3 haploinsufficiency confers resistance to palmitate-induced cell death .........31

Targeted knockdown of SmD3 recapitulates 6H2 phenotype .................................32

SmD3 disruption protects cells from generalized oxidative stress induction...........33

SmD3 regulates intronic non-coding RNA expression ............................................33

SmD3 knockdown perturbs snRNP biogenesis.......................................................35

6H2 cells display normal splicing efficiency ............................................................36

SmD3 knockdown does not alter alternative splicing ..............................................37

SmD3 controls intron lariat abundance ...................................................................38

Conclusions ................................................................................................................40

Figures ........................................................................................................................42

Chapter 4: Superoxide triggers cytosolic snoRNA accumulation

Introduction .................................................................................................................58

Results ........................................................................................................................59

Palmitate-induced superoxide precedes cytosolic snoRNA accumulation..............59

Direct superoxide induction causes rapid cytosolic snoRNA accumulation ............60

Superoxide levels correlate with cytosolic snoRNA accumulation ..........................61

iv

Conclusions ................................................................................................................62

Figures ........................................................................................................................65

Chapter 5: Discussion..................................................................................................73

Cytosolic functions of snoRNAs..................................................................................75

Regulation of snoRNA processing..............................................................................76

Regulation of snoRNA localization..............................................................................78

Summary.....................................................................................................................79

 

Chapter 6: Materials and Methods ..............................................................................80

References ....................................................................................................................97

v

List of Figures and Tables

Chapter 1

Figure 1.1 Genetic screen to isolate palmitate resistant CHO mutants..........................12

Figure 1.2 Secondary structure of box C/D and box H/ACA snoRNAs ..........................13

Figure 1.3 rpL13a-encoded box C/D snoRNAs are induced in palmitate treated CHO

cells ................................................................................................................................14

Chapter 2

Figure 2.1 In vivo expression of rpL13a snoRNAs .........................................................22

Figure 2.2 ASO knockdown of rpL13a snoRNA in vivo ..................................................23

Figure 2.3 rpL13a snoRNAs are required for oxidative stress in vivo ............................24

Figure 2.4 In vivo rpL13a snoRNA knockdown reduces oxidative damage ...................25

Figure 2.5 U32a, U33, and U35a contain antisense homology to rRNA ........................26

Figure 2.6 2’-O-methylation is unaffected in 6F2 cells ...................................................27

Figure 2.7 rpL13a snoRNAs accumulate in the cytoplasm under lipotoxic conditions ...28

Figure 2.8 rpL13a snoRNAs accumulate in the cytoplasm under lipotoxic conditions ...29

Chapter 3

Figure 3.1 6H2 cells are resistant to palmitate-induced cell death .................................42

Figure 3.2 6H2 cells are haploinsufficient for SmD3 ......................................................43

Figure 3.3 Targeted knockdown of SmD3 confers palmitate-resistance ........................45

Figure 3.4 SmD3 disruption protects cells from palmitate-induced and generalized

oxidative stress induction ...............................................................................................46

Figure 3.5 Intronic non-coding RNA expression is disrupted in 6H2 cells......................48

Figure 3.6 SmD3 knockdown disrupts U4 and U5 snRNPs ...........................................50

vi

Figure 3.7 Host gene expression is normal in 6H2 cells ................................................52

Figure 3.8 Alternative splicing is normal in 6H2 cells .....................................................54

Figure 3.9 SnoRNA-containing intron lariats are decreased in 6H2 cells.......................55

Figure 3.10 Role for SmD3 in snoRNA production.........................................................57

Chapter 4

Figure 4.1 Palmitate induces superoxide .......................................................................65

Figure 4.2 Doxorubicin is not a potent H2O2 inducer ......................................................67

Figure 4.3 Doxorubicin induces rapid cytosolic rpL13a snoRNA accumulation .............68

Figure 4.4 Manipulation of superoxide alters cytosolic rpL13a snoRNA accumulation ..70

Figure 4.5 Superoxide is linked to cytoplasmic snoRNA accumulation..........................72

Chapter 5

Table 5.1 Primer/oligo sequences ..................................................................................92

vii

Abbreviations

7-keto 7-ketocholesterol

ASO anti-sense oligo

BSA bovine serum albumin

CHO Chinese hamster ovary

CM-H2DCFDA 5-(and-6)-chloromethyl-2’,7’-dichlorodihydrofluorescein

diacetate, acetyl ester

CYT cytosolic

DETC diethyldithiocarbamate

DHE dihydroethidium

DPI diphenylene iodium

eEF1A-1 eukaryotic elongation factor 1A-1

FA fatty acid

FFA free fatty acids

gadd7 growth arrested DNA-damage inducible gene 7

GFP green fluorescent protein

H2O2 hydrogen peroxide

KD knockdown

LNA locked nucleic acid

LPS lipopolysaccharide

MCLA 2-methyl-6-(4-methoxyphenyl)-3,7-dihydroimidazol[1,2-

A]pyrazin-3-one, hydrochloride

mRNA messenger RNA

viii

miRNA microRNA

MnTBAP manganese(III) tetrakis(4-benzoic acid)porphyrin

NADPH nicotinamide adenine dinucleotide phosphate

ncRNA non-coding RNA

NUC nuclear

O2- superoxide

Palm palmitate

PBS phosphate buffered saline

PI propidium iodide

PAGE polyacrylamide gel electrophoresis

PKC protein kinase C

qPCR quantitiative polymerase chain reaction

RACE rapid amplification of cDNA ends

rpL13a ribosomal protein L13a

RNPs ribonucleoproteins

ROS reactive oxygen species

rRNA ribosomal RNA

scaRNA small Cajal body RNA

shRNA short hairpin RNA

siRNA small interfering RNA

SMN survival of motor neurons

snRNA small nuclear RNA

snRNP small nuclear ribonucleoprotein

ix

snoRNA small nucleolar RNA

snoRNP small nucleolar ribonucleoprotein

snrpD3 small nuclear ribonucleoprotein D3

SOD superoxide dismutase

triol 3β,5α,6β-cholestantriol

tRNA transfer RNA

WT wild-type

x

Acknowledgements

I would like to thank my thesis advisor, Dr. Jean Schaffer, for her immeasurable support

over the years. This work would not be possible without her encouragement and

guidance. I greatly appreciate her willingness to always let me test a new hypothesis,

and I am grateful for her help in making me a better scientist.

I would like to thank Dr. Peter Crawford, Dr. Kathleen Hall, Dr. Daniel Ory, Dr. Rob

Mitra, Dr. Heather True, and Dr. Eric Galburt for serving on my thesis committee. Their

support and suggestions have proved invaluable over the years. Much of this work can

be attributed directly to their guidance. I would especially like to thank Dr. Peter

Crawford for serving as my committee chair and many helpful conversations. I would

also like to thank Dr. Kathleen Hall for her encouragement in pursuing the field of RNA. I

would like to acknowledge NHLBI for funding through the Cardiovascular Training

Grant.

My graduate experience would not be the same without immense help from all

members of the Schaffer and Ory labs, both past and present. I would specifically like to

thank Dr. Dan Ory for his perspectives and many helpful suggestions. Thank you to

Katrina Brandis, George Caputa, Mia Henderson, Sarah Jinn, Josh Langmade, Jiyeon

Lee, Melissa Li, and Linda Zhao for so many helpful conversations about snoRNAs. I

especially would like to thank Carlos Michel for encouraging the lab to study snoRNAs

and helping me with my project. I must also thank Dr. Chris Holley for so many helpful

conversations about snoRNAs and ROS. It’s been a pleasure to share lab space, ideas,

xi

and experiments with you. Thank you to Sarah Gale and Dave Scherrer for all that you

do for the lab – it is a much better place with you both as a part of it.

To my fellow grad students Aggie Bielska, Rita Brookheart, and Maria Praggastis, thank

you for many great conversations and time spent in the lab. I would also like to

acknowledge Dr. Phyllis Hanson and Charlie Jackson for their collaboration on

exosomal ncRNA experiments.

Thank you to my parents for their steadfast support. They have always encouraged me

to pursue any aspirations, and I am extremely grateful.

Finally, I would like to thank my wife, Susan Scruggs, for her endless support and

encouragement. Thank you for understanding that one more hour for an experiment

always turns into two. Thank you for listening to me ramble about science. This has

been a much more enjoyable journey with you by my side.

xii

ABSTRACT OF THE DISSERTATION

Regulation of metabolic stress-induced snoRNAs

by

Benjamin Steel Scruggs

Doctor of Philosophy in Biology and Biomedical Sciences (Molecular Cell Biology)

Washington University in St. Louis, 2012

Professor Jean E. Schaffer, Chairperson

Accumulation of excess lipid in non-adipose tissues is associated with oxidative

stress and organ dysfunction and plays an important role in diabetic complications.

While a number of stress responses have been implicated, the precise molecular

mechanisms linking lipid accumulation and cellular dysfunction are not fully understood.

To elucidate molecular events critical for lipotoxicity, we used retroviral promoter trap

mutagenesis to generate mutant Chinese hamster ovary cell lines resistant to lipotoxic

and oxidative stress. This approach uncovered a previously unsuspected role for small

nucleolar RNAs (snoRNAs) as critical mediators of lipotoxic cell death.

Herein we show that under lipotoxic conditions, intronic snoRNAs in the rpL13a

gene accumulate in the cytosol during metabolic stress, suggesting that these non-

coding RNAs function non-canonically to target cytosolic RNAs. Moreover, we

demonstrate that the rpL13a snoRNAs play a critical role in vivo in amplification of

reactive oxygen species and downstream oxidative stress-mediated tissue injury.

Study of independent mutants from our genetic screen not only identified a role

for snoRNAs in lipotoxicity, but also provided new insights into the cellular machinery for

xiii

production of these non-coding RNAs. We demonstrate that the spliceosomal protein

SmD3 plays a critical role in expression of intronic non-coding RNAs, including the

rpL13a snoRNAs, by maintaining the abundance of snoRNA-containing intron lariats

from which they are processed. Our findings indicate that this function may involve

effects of SmD3 on small nuclear RNAs (snRNAs) U4 and U5.

Finally, we demonstrate a link between superoxide induction and cytosolic

snoRNA accumulation. Under lipotoxic conditions, superoxide induction precedes

cytosolic snoRNA accumulation. Other chemical inducers of superoxide also cause the

rpL13a snoRNAs to localize to the cytosol, and manipulation of superoxide dismutase

demonstrates that superoxide levels directly correlate with cytosolic snoRNA

expression.

Together, our studies identify an important role for snoRNAs in metabolic stress

responses. The rpL13a snoRNAs are essential for cell death in response to lipid

overload through functions that are distinct from their canonical role in ribosomal RNA

modification in the nucleolus. Moreover, these non-coding RNAs respond to chemical

inducers of reactive oxygen species and thereby may contribute more broadly to

disease pathogenesis that involves oxidative stress.

 

1

CHAPTER 1

Introduction to Lipotoxicity

Type 2 diabetes is a growing worldwide health concern

Metabolic syndrome, obesity, and type 2 diabetes are growing in prevalence at

an astounding rate in the United States and around the world. Occurrence of type 2

diabetes is increasing in both adult and juvenile populations. In 2011, the Centers for

Disease Control estimated that 26 million Americans have type 2 diabetes and 79

million have some form of metabolic syndrome, and those numbers are expected to

double or triple by 2050 (National Diabetes Fact sheet 2011, online). Type 2 diabetics

have increased risks for heart disease, stroke, hypertension, nephropathy, neuropathy,

and retinopathy. In the United States, diabetes costs $174 billion annually, including

$116 billion in direct medical expenses. Therefore, it is becoming increasingly important

to understand the underlying causes of complications associated with diabetes.

Lipotoxicity is a hallmark of metabolic disease

Diabetes complications are likely related to the systemic metabolic abnormalities

in this disease. While it is well appreciated that hyperglycemia contributes to diabetes

pathogenesis, elevations in serum triglycerides and free fatty acids also play an

important role in the pathogenesis of diabetic complications. Under physiological

conditions, mammalian adipose cells internalize and store large quantities of lipid.

However, under pathophysiological conditions, accumulation of fatty acids in non-

adipose tissues causes cell dysfunction and cell death that lead to impaired organ

2

function (112). This phenomenon, known as lipotoxicity, contributes to the pathogenesis

of heart failure, renal dysfunction, steatohepatitis, and progressive pancreatic

insufficiency (3, 46, 50, 96, 97).

Mechanisms linking lipid overload to cell death

In vitro models in which the media of cultured cells is supplemented with excess

fatty acid have been used to probe metabolic and signaling pathways involved in the

cellular response to lipid overload. In a time- and dose-dependent manner, long-chain

saturated fatty acids induce apoptosis in a variety of cell types (11, 18, 61, 66, 124), and

this response is enhanced by high glucose (20). Although lipid overload in non-adipose

cells is initially buffered by cytoprotective triglyceride stores (60, 63), when the limited

capacity for neutral lipid storage in non-adipose cells is exceeded, excess saturated

fatty acids initiate several cellular stress response pathways. Fatty acid-induced

endoplasmic reticulum stress can result in reactive oxygen species (ROS) generation

(99). Independently, oxidative stress is induced in a variety of cell types through

activation of NADPH oxidase, mitochondrial dysfunction due to remodeling of organelle

membranes, and excessive cycles of oxidative phosphorylation (45, 82, 101). NADPH

oxidase participates in palmitate-induced superoxide production (31) and is likely to

spark a cascade of ROS production as ROS are rapidly interconverted (34).

Dismutation of superoxide produces hydrogen peroxide. Subsequently, hydrogen

peroxide will produce highly reactive hydroxyl radicals in the presence of metal ions

through Fenton or Harber-Weiss reactions. Hydrogen peroxide can also be converted to

hypochlorous acid in the presence of myeloperoxidase. Superoxide can also react with

3

nitric oxide to form another highly reactive molecule, peroxynitrite. Excessive ROS lead

to detrimental consequences including protein carbonylation, lipid peroxidation, and

DNA damage (14, 17, 76). Administration of antioxidants to cultured cells and animal

models of lipotoxicity mitigates against lipotoxic cell death (8, 9, 56, 61), suggesting a

central role for oxidative stress in lipotoxicity.

Elucidating molecular mechanisms of lipotoxicity

Despite identification of stress pathways involved in lipotoxicity, the precise

molecular responses following lipid overload have yet to be elucidated. To identify

genes critical for the cellular lipotoxic response, we performed a genetic screen in

Chinese hamster ovary (CHO) cells, with mutagenesis by transduction with ROSAβgeo

retrovirus at low multiplicity of infection to achieve, on average, one insertion per ten

genomes. Although the integrated provirus contains a cDNA cassette for a β-

galactosidase-neomycin phosphotransferase fusion protein, it lacks its own promoter,

and thus its transcript is expressed only if the retrovirus inserts downstream of an active

promoter and splice donor site. Mutagenized cells that survived a round of neomycin

selection were then treated for 48 h in media supplemented with a lipotoxic

concentration of palmitate (500 µM) to model pathophysiological states. Under these

conditions, wild type (WT) cells were killed, but mutant cells, each with a single

disrupted gene critical for lipotoxicity, survived.

This genetic screen led to the identification of many unpredicted genes as

essential for the lipotoxic response. The eukaryotic elongation factor (eEF) 1A-1 was

shown to have a role in ROS and ER stress induced death (7). The non-coding RNA

4

gadd7 was shown to be induced by lipotoxic stress in a ROS-dependent fashion and

drive ROS propagation following palmitate treatment, demonstrating that gadd7

functions as a feed-forward regulator of lipid-induced and ROS-induced cell death (9). In

another mutant cell line (6F2), the promoter trap disrupted the locus for ribosomal

protein L13a (rpL13a) (72). Studies of this mutant revealed that the portions of this gene

essential for lipotoxicity are three highly conserved small nucleolar RNAs (snoRNAs)

embedded within the rpL13a introns, rather than the protein-coding exonic sequences.

These findings suggest a previously unsuspected role for snoRNAs in the regulation of

metabolic stress in mammalian cells.

Conserved functions of snoRNAs

SnoRNAs are typically nucleolar-localized RNAs that guide the modification of

other non-coding RNAs (ncRNAs). Eukaryotic cells contain more than 200 unique

snoRNAs comprised of two families—the box C/D snoRNAs and box H/ACA snoRNAs

(57). The vast majority of snoRNAs transiently base pair with complementary target

RNA to guide either 2’-O-methylation (C/D snoRNAs) or pseudouridylation (H/ACA

snoRNAs) of target RNA (69). These modifications are the most common covalent

modifications found in ribosomal RNA (rRNA) and are found in key regions of rRNA

including the peptidyl transferase center and the mRNA-decoding center. Both types of

modifications are essential for ribosome function, and the importance of these

modifications is emphasized by the evolutionary conservation of modification locations.

Other snoRNA modification targets include snRNAs in eukaryotes, transfer RNAs in

archaea, and spliced leader RNAs in trypanosomes (16, 19, 111). Spliceosome function

5

also depends on the modification of snRNAs by C/D and H/ACA snoRNAs. Modification

of snRNAs takes place in Cajal bodies providing evidence for extranucleolar roles for

snoRNAs (16). The precise function of modified nucleotides in rRNAs and snRNAs

remains unknown, but they are hypothesized to have a critical function, since complete

loss of modification causes lethality and specific mutations can cause disease (15).  

Both snoRNA families are structurally and functionally conserved from humans to

Archaea (77, 104, 105). Box C/D snoRNAs contain the conserved sequence motifs

UGAUGA (C box) and CUGA (D box) near the 5’ and 3’ ends of the snoRNA,

respectively. The two boxes are separated by ~60nt and internal C’ and D’ boxes are

often present in that region. Base-pairing to target RNAs typically takes place in the

region directly upstream of the D box or D’ box, involving a sequence known as the

antisense element (Figure 1.2) Box C/D snoRNAs are associated with four box C/D

snoRNP proteins, fibrillarin, Nop56, Nop58, and 15.5K/NHPX (55, 74, 93, 107, 123,

127). Fibrillarin catalyzes the 2′-O-methyl transfer at the site determined by the box C/D

snoRNA guide (118)

Similarly, H/ACA snoRNAs are characterized by a conserved secondary

structure. Each H/ACA snoRNA is defined by a hairpin–hinge–hairpin–tail structure with

two short conserved sequences called boxes H and ACA. One or both hairpins have an

internal loop with two short sequences that are complementary to the rRNA substrate.

These pseudouridylation pockets enable anti-sense base-pairing with target RNAs in

the region where they direct modifications (Figure 1.2). The box H/ACA snoRNAs form

snoRNP complexes with dyskerin, Nhp2, Nop10, and Gar1 proteins (4, 29, 35, 65, 120).

The enzyme responsible for uridine-to-pseudouridine isomerization is dyskerin (39).  

 

6

Palmitate treatment induces rpL13a snoRNA accumulation in CHO cells.

Identification of the snoRNA encoding rpL13a locus through the genetic screen

conducted in the Schaffer Lab suggested snoRNAs have a role in lipotoxicity. All

mammalian loci for rpL13a contain four highly conserved intronic box C/D snoRNAs that

are processed during splicing of the rpL13a pre-mRNA transcript (Figure 1.3A) (75).

These snoRNAs, U32a, U33, U34, and U35a, are encoded within introns 2, 4, 5 and 6,

respectively, and range in size from 61-82 nucleotides. U32a, U33, U34, and U35a are

highly conserved across species.

Lipotoxic conditions induce the pre-mRNA expression of many genes including

rpL13a. Consistent with up-regulation of the pre-mRNA, rpL13a snoRNAs also

accumulate under stress conditions as assessed by RNase protection assays with 32P-

labeled probes specific for the snoRNA sequences (Figure 1.3B & C). In comparison,

levels of the microRNA miR-16 do not change under these conditions. In WT CHO cells,

U32a, U33, and U35a snoRNAs are expressed at low levels under basal conditions and

increased following palmitate treatment, whereas U34 is not detected under basal or

lipotoxic conditions. In mutant 6F2 cells, induction of U32a, U33, and U35a is markedly

attenuated, consistent with disruption of the rpL13a locus (Figure 1.3B & C). Palmitate

induction of U32a, U33, and U35a snoRNAs is also observed in WT C2C12 murine

myoblasts, which demonstrate similar sensitivity to lipotoxic conditions (72). Moreover,

these snoRNAs are induced by saturated fatty acids known to cause lipotoxicity

(myristic, palmitic, and stearic acids), but not by unsaturated palmitoleic acid and oleic

acid, which are well tolerated by cells. These findings suggest a conserved role for

snoRNAs as mediators of lipotoxicity.

7

Confirmation of the role for rpL13a snoRNAs in lipotoxicity.

Demonstration of rpL13a snoRNAs as mediators of metabolic stress was

confirmed in two ways (72). First, stable cell lines generated in the mutant 6F2

background showed that snoRNA sequences within the rpL13a genomic locus are

required for complementation of 6F2 cells. Mutant cells transfected with a plasmid

containing 4.3 kb from the murine rpL13a genomic locus, including all eight exons and

intervening introns, restored palmitate-induced snoRNA expression, palmitate-induced

ROS, and palmitate-induced cell death. By contrast, when mutant cells were transfected

with a similar construct in which all four snoRNAs were removed, but promoter and

exon-intron structure was otherwise intact, complementation was lost. In an alternate

approach to genetic confirmation, simultaneous knock-down of U32a, U33, and U35a in

wild-type murine myoblasts by nucleofection with phosphorothioate-modified anti-sense

oligos (ASOs) rendered these cells resistant to palmitate-induced ROS and cell death.

Since loss of a single snoRNA alone did not recapitulate lipotoxicity resistance, these

data support a model in which the three snoRNAs function in concert to promote

palmitate-induced oxidative stress.

Biogenesis of box C/D and H/ACA snoRNPs

The ability of the rpL13a locus to regulate oxidative stress is dependent on

expression of the encoded snoRNAs, which are presumed to form small nucleolar

ribonucleoproteins (snoRNPs). Biogenesis of box C/D and H/ACA snoRNPs is a highly

regulated, multi-step process initiated during the transcription of a snoRNA host gene.

8

In vertebrates, the vast majority of box C/D and H/ACA snoRNAs are encoded within

introns of pre-mRNAs (25, 52, 58, 108, 109). All host introns for vertebrate snoRNAs

encode only a single intronic snoRNA. During splicing, snoRNAs are excised from the

pre-mRNA as part of the intron lariat. Mammalian pre-mRNA intron lariats are rapidly

debranched and degraded after splicing (83). 5’ to 3’ and 3’ to 5’ exonucleases recycle

the linearized intron lariat (79). Active recruitment of snoRNP proteins to the nascent

snoRNA during synthesis and/or splicing of the host pre-mRNA is required for efficient

intronic snoRNP production. The associated snoRNP proteins define the termini of the

snoRNA by protecting them from the processing exonucleases (13, 51, 108, 122).

Following trimming of the flanking intron, the mature snoRNP is released.

Biosynthesis of functional box C/D snoRNPs requires the ordered recruitment of

snoRNP proteins 15.5K/NHPX, Nop56, Nop58, and fibrillarin. To establish a scaffold for

the 15.5K/NHPX snoRNP protein, a functional Kink-turn structure must be formed by

the nascent snoRNA. The 5’ and 3’ box C and D terminal regions of the snoRNA form

this Kink-turn by folding into a stem-internal loop-stem structure (54, 123). Conserved

nucleotides in the C and D box motifs form non-canonical G-A, A-G, and U-U base pairs

establishing the Kink-turn and docking site for the 15.5K/NHPX protein. Binding of

15.5K/NHPX induces a sharp bend in the phosphodiester backbone of the two

contiguous RNA stems of the Kink-turn (102, 103, 115, 119). This conformational

change provides the structural requirements for the subsequent binding of Nop58,

Nop56, and two copies of fibrillarin (12). One fibrillarin and Nop58 bind to the box D and

C sequences in the upper stem of the Kink-turn. Nop56 and another copy of fibrillarin

bind to the internal C’ and D’ boxes.

9

Position within the intron is also critical for efficient processing of human box C/D

snoRNAs. Most box C/D snoRNAs are located about 80-90 nucleotides upstream of the

3’ splice site (38). An optimal distance of ~50 nucleotides between the snoRNA coding

region and the branch point of the host intron is required for efficient snoRNA

processing. Increasing or decreasing the length between the snoRNA and the branch

point significantly compromises snoRNA accumulation (37). The relationship between

snoRNA location and the branch site suggests there is a synergy between intronic

snoRNA processing and pre-mRNA splicing. Consistent with a splicing-depending

mechanism for box C/D snoRNA processing, 15.5K/NHPX is recruited to box C/D

intronic snoRNAs at the C1 splicing complex stage. A general splicing factor, intron

binding protein 160 (IBP160), provides a molecular link between the spliceosome and

the nascent box C/D snoRNA (36). IBP160 has putative helicase activity and is thought

to trigger box C/D snoRNP assembly by interacting with the U2 spliceosomal small

nuclear RNP (snRNP) or other splicing factors associated with the branch-point region

in the C1 splicing complex.  

Box H/ACA snoRNPs also require ordered recruitment of snoRNP proteins.

However, in contrast to box C/D snoRNAs, human H/ACA snoRNAs have no

preferential intronic location relative to the 5’ or 3’ splice sites of the host introns (88,

92). Human box H/ACA snoRNAs are commonly found in introns of longer than average

length. H/ACA snoRNAs are also processed efficiently from artificial host pre-mRNAs

regardless of position within the intron suggesting that box H/ACA snoRNAs are

processed independent of relative splice site location (88).

10

Many class specific factors regulate the biogenesis of box C/D snoRNAs and box

H/ACA snoRNAs, but the excision of all intronic snoRNPs requires splicing. The

relationship between pre-mRNA splicing and snoRNA biogenesis suggests the

spliceosomal machinery is critical for snoRNA expression. The spliceosome is a

complex RNA machine comprised of its own set of ncRNAs, the small nuclear RNAs

(snRNAs), and over 100 proteins (117). Each snRNA is bound to a unique set of

proteins to form a small nuclear ribonucleoprotein (snRNP). In addition to snRNP-

specific proteins, each of the major snRNAs (U1, U2, U4, and U5) is bound to a

common set of seven proteins (SmB/B’, SmD1, SmD2, SmD3, SmE, SmF, and SmG).

These proteins form a seven-membered ring around the short, highly conserved,

uridine-rich sequence on each snRNA called the Sm site (1, 47, 100). Assembly of Sm

cores onto snRNAs is regulated by the SMN complex (22, 71, 84). Following assembly

of the Sm protein heptameric ring, the 5’ cap of the snRNA is hypermethylated to form

the 2,2,7-trimethyl guanosine cap, and the mature snRNP is then imported into the

nucleus by snurportin and importin β for final association with snRNP-specific proteins

and utilization in mRNA splicing (23, 24, 42, 70, 86).

Summary

Herein, we describe work that provides new understanding of the function of the

rpL13a snoRNAs in cells and in a rodent model of acute oxidative stress, identifies a

role for the spliceosomal protein, SmD3, in regulating the biogenesis of intronic ncRNAs

including the rpL13a snoRNAs, and uncovers a link between superoxide production and

11

induction of the rpL13a snoRNAs. Together, our studies provide important new insights

into the production and function of snoRNAs during metabolic stress.

12

Figure 1.1  

   Figure 1.1 Genetic screen to isolate palmitate resistant CHO mutants. Wild type

(WT) CHO cells were transduced with the ROSAβgeo retrovirus, leading to integration

of the provirus containing a splice acceptor, promoterless β-galactosidase-neomycin

resistance cassette, and polyadenylation sequences. Promoter-trapping and gene

disruption at the site of integration was selected for by growth in neomycin (NEO), and

palmitate-resistant mutants were selected by growth in media with 500 µM palmitate

(palm) for 48 h.

����� ������ ��� � ��������� �������� ���

��������� ���� ������� �� ��

����

13

Figure 1.2

Figure 1.2 Secondary structure of box C/D and box H/ACA snoRNAs. A) Box C/D

snoRNAs (green) are characterized by box C and D motifs. Sequences upstream of box

D and D’ base pair (blue) with target RNAs (red) to guide the 2’-O-methylation of target

RNA. B) Box H/ACA snoRNAs are characterized a hairpin–hinge–hairpin–tail structure

and box H and ACA motifs. Pseudouridylation pockets allow for base pairing with target

RNAs to guide pseudouridylation modifications.

A B

14

Figure 1.3

Figure 1.3 rpL13a-encoded box C/D snoRNAs are induced in palmitate treated

CHO cells. (A)  rpL13a gene organization showing location of ROSAβgeo promoter trap

insertion and locations of intronic U32a, U33, U34, and U35a snoRNAs. (B, C) WT and

6F2 cells were untreated (UT) or supplemented with palmitate for 48 h. (B) Small RNA

was harvested and used in RNase protection assay with 32P-labeled hamster rpL13a

snoRNA probes or miR-16 probe as control. (C) Autoradiograms from RNase protection

experiments as in (B) were quantified by densitometry. All data expressed as mean ±

SE for 3 independent experiments. * p < 0.01 for palmitate treated vs. untreated.

U32a

U33

U34

U35a

miR-16

palmitate - + - + WT 6F2

A

B C

Michel CI et al. 2011 Cell Metab

rpL13a genomicU32a U33 U34 U35a

ROSA�geo

snoR

NA

: miR

-16

(rel

uni

ts)

UT Palm UT Palm0

2

4

6

8U32aU33U35a

WT 6F2

*

15

CHAPTER TWO

rpL13a snoRNAs function as non-canonical box C/D snoRNAs

INTRODUCTION

Box C/D snoRNAs U32a, U33, and U35a are critical for the propagation of ROS

following lipid overload in cultured cells (72). The mechanisms through which rpL13a

snoRNAs recognize metabolic stress and regulate the cellular stress response remain

unclear. Typically, box C/D snoRNAs serve as guides for ribonucleoprotein particles

during 2’-O-methylation of rRNA. These modifications occur within the nucleolus, where

the vast majority of snoRNAs reside. Extranucleolar functions for snoRNAs have also

been described. SnoRNAs are known to traffic to Cajal bodies where they can serve to

guide the modification of snRNAs (16). Recent studies have suggested a number of

additional non-canonical roles for snoRNAs. A brain specific snoRNA, HBII-52, has

been shown to interact with a pre-mRNA in the nucleoplasm and play a role in

alternative splicing (49). Biochemical and computational analyses from multiple groups

show snoRNAs can be processed into miRNAs in a Dicer dependent fashion,

suggesting snoRNAs may localize to the cytoplasm where processing and function

would take place (21, 78, 90). Additional snoRNAs have been identified or

computationally predicted to lack rRNA or snRNA antisense homology (43). These

“orphan” snoRNAs could serve as guides for currently unidentified target RNAs in

unknown locations.

16

Here, we extend the initial finding that rpL13a snoRNAs are critical mediators of

metabolic stress by examining their role in vivo. We demonstrate that U32a, U33, and

U35a contribute to oxidative stress and oxidative damage in a mouse model of

inflammation and oxidative stress. Furthermore, we broaden our understanding of

snoRNA function in the metabolic stress pathway in several ways. By examining

predicted rpL13a modification sites in 6F2 cells, we show that methylation of these

rRNA nucleotides is not affected by altered levels of the snoRNAs during lipotoxicity. By

characterizing the localization of rpL13a snoRNAs in wild type cells, we show that they

accumulate in the cytoplasm during lipotoxicity. This novel localization for intron-

encoded snoRNAs provides insight into the potential mechanism through which rpL13a

snoRNAs regulate oxidative stress.

RESULTS

In vivo expression of rpL13a snoRNAs. Our studies of the rpL13a snoRNAs show

that these molecules function in the cellular response to lipotoxic and oxidative stress.

To extend these findings to an in vivo model of oxidative stress, we examined the

expression of rpL13a snoRNAs in a well-established model of lipopolysaccharide (LPS)-

mediated liver injury that is characterized by inflammation, steatosis, and oxidative

stress (5). Compared to saline-injected control mice, liver tissue from LPS-treated mice

showed significant up-regulation of cytosolic U32a, U33, and U35a snoRNAs (Figure

2.1), demonstrating that the rpL13a snoRNAs are induced in vivo in response to

metabolic stress.

17

In vivo rpL13a snoRNA knockdown reduces oxidative damage. Based on these

findings, we analyzed the effects of loss-of-function of the rpL13a snoRNAs in the LPS-

mediated liver injury model. To achieve specific knockdown of the snoRNAs in vivo,

prior to LPS injection, mice were treated with three serial intraperitoneal injections of

antisense locked nucleic acid oligonucleotides directed against each of the three

snoRNAs or directed against GFP as a control. Antisense oligonucleotides directed

against the snoRNAs achieved 72, 84, and 74% knockdown of U32a, U33, and U35a,

respectively, in liver tissue following LPS injection without diminishing LPS-induced

inflammation (Figure 2.2). Knockdown of the snoRNAs mitigated LPS-induced oxidative

stress in the liver as demonstrated by dihydroethidium staining for superoxide and

oxidative damage to liver tissue proteins and lipids (Figures 2.3 and 2.4). These findings

indicate that rpL13a snoRNAs are required in vivo for propagation of oxidative stress.

rpL13a guided rRNA modifications are unaltered during lipotoxicity. Canonical box

C/D snoRNAs participate in ribonucleoproteins that localize to nucleoli. In S. cerevisiae

and in X. laevis, box C/D snoRNAs serve as guides that target 2’-O-methylation of

rRNAs with which they share short stretches of antisense homology (53). Although they

lack some sequence features of canonical 2’-O-methylation guide snoRNAs (internal

box C’ sequence not well-conserved, U33 lacks box D’, rRNA complementarity not

upstream of box D in U35a), U32a, U33, and U35a each contain 10-12 nucleotide

stretches of complementarity to rRNA sites of 2’-O-methylation (Figure 2.5), suggesting

a potential role as guide RNAs for 2’-O-methylation of G1328 in 18S and A1511 in 28S

(U32a), U1326 in 18S (U33), and C4506 in 28S (U35a) rRNAs (75). We reasoned that if

the mechanism of action of snoRNAs U32a, U33, and U35a in lipotoxic and oxidative

18

stress involved 2’-O-methylation of these rRNAs, modifications of these rRNA sites

should be diminished in 6F2 compared to WT cells under metabolic stress conditions

when the snoRNAs are induced in WT cells. However, primer extension studies showed

no differences in the extent of modification of these rRNA sites between WT and 6F2

cells under basal or palmitate-treated conditions (Figure 2.6). These data indicate that

under basal and lipotoxic conditions, either residual expression of U32a, U33, and U35a

in 6F2 cells is sufficient to support these modifications of rRNAs, or this function is

subserved by other molecules in eukaryotic cells. Furthermore, at a point in the lipotoxic

response at which absence of snoRNA induction is readily apparent and functionally

correlates with resistance to lipotoxicity in 6F2 cells, there is no corresponding change

in 2’-O-methylation of rRNAs.

rpL13a snoRNAs accumulate in the cytoplasm during lipotoxicity. We

hypothesized that if U32a, U33, and U35a were involved in functions other than

modification of ribosomal RNAs, then under lipotoxic stress conditions, they may have a

subcellular distribution distinct from canonical box C/D snoRNAs, which co-localize with

nascent rRNAs in the nucleolus. Following palmitate treatment of C2C12 cells, we

isolated nuclear and cytosolic RNAs by sequential detergent extraction and quantified

U32a, U33, and U35a by qRT-PCR. With palmitate treatment U32a, U33, and U35a

increase in the cytoplasm, whereas levels of these snoRNAs remain unchanged in the

nucleus (Figure 2.7A and B). Accumulation of rpL13a snoRNAs in the cytosol under

lipotoxic conditions was confirmed by fluorescence in situ hybridization. As expected,

anti-sense probe for snoRNA U3 demonstrated strong nucleolar localization, and this

19

was unaffected by lipotoxic stress (Figure 2.8A). Staining for the rpL13a snoRNAs was

performed in cells nucleofected with control ASO (GFP) or with ASO targeting each of

the rpL13a snoRNAs to ascertain the specificity of signal. Consistent with data from

RNase protection and qPCR assays, in control nucleofected cells expression of U32a,

U33, and U35a was low under normal growth conditions and increased under lipotoxic

conditions (Figure 2.8B, GFP-nucleofected panels). Prominent staining for each of

these snoRNAs was observed in the cytoplasm, but not nucleoli. The probe for U32a

also stains non-nucleolar regions of the nucleus. Cytoplasmic staining for the rpL13a

snoRNAs under lipotoxic conditions was markedly diminished when the snoRNAs were

depleted by specific ASOs that target each snoRNA (Figure 2.8B, U32a, U33, and U35a

ASO-nucleofected panels). Cytoplasmic staining for U32a, U33, and U35a was also

distinct from the nuclear pattern observed under lipotoxic conditions using a probe

specific for intron 1 (Figure 2.8A), indicating that the cytosolic distributions of the rpL13a

snoRNAs do not simply reflect localization of the pre-mRNA. The nuclear staining for

U32a resembled the staining for intron 1 and was not diminished with ASOs that target

U32a, suggesting that this represents detection of the pre-mRNA, which is not targeted

by U32a ASOs. Together our biochemical and in situ hybridization data support a model

in which U32a, U33, and U35a snoRNAs act in non-canonical roles in the cytoplasm

during lipotoxic stress.

CONCLUSIONS

SnoRNAs U32a, U33, and U35a are critical for the cellular response to metabolic

stress. In vivo knockdown of these three snoRNAs further illustrates their importance to

the pathophysiology of metabolic stress. The LPS-induced model of hepatic tissue injury

20

confirms that the rpL13a snoRNAs are induced in the setting of oxidative stress in vivo.

Moreover, our in vivo knockdown data show that these snoRNAs are required for

amplification of oxidative stress and for propagation of oxidative stress-mediated

damage to proteins and lipids. Because LPS has pleiotropic effects in vivo and because

ASO knockdown in vivo is limited by liver toxicity, it is not surprising that in vivo

knockdown of these snoRNAs provided only a partial reduction in the oxidative stress

response and did not blunt the release of serum transaminases into the circulation (not

shown). In future studies, genetic approaches to loss of function and/or blunting of

additional effector pathways (e.g., inflammatory signaling) may be required to block liver

injury entirely. Nonetheless, our in vivo knockdown studies show that the rpL13a

snoRNAs are required for the full induction of tissue oxidative stress in the murine LPS

model, thus providing in vivo evidence for a role of snoRNAs in metabolic stress.

We show here in a stable mutant cell line that decreased basal expression of

U32a, U33, and U35a snoRNAs and loss of palmitate induction of these snoRNAs is not

associated with changes in 2’-O-methylation of predicted rRNA targets, yet is

associated with resistance to lipotoxicity. While it is possible that these snoRNAs

contain more than one functional guide sequence to target multiple substrates including

rRNAs (13, 110), their accumulation in the cytosol during lipotoxicity suggests a non-

nucleolar function for these snoRNAs. Movement of snoRNAs to the cytosol is not

without precedent, since the independently transcribed box C/D snoRNA, U8, has been

shown to be exported from the nucleus (121). Here, we provide the first direct evidence

that intronic snoRNAs can also localize to the cytosol. Our data are most consistent with

a model in which the rpL13a snoRNAs function in lipotoxicity and oxidative stress

21

response pathways as non-canonical box C/D snoRNAs. Furthermore our findings

suggest that the rpL13a snoRNAs affect targets in the cytoplasm.

22

Figure 2.1

Figure 2.1 In vivo expression of rpL13a snoRNAs. (A) Mice were injected

intraperitoneally with lipopolysaccharide (LPS), or equivalent volume of phosphate-

buffered saline (PBS) as control, and liver tissue was harvested 12 h later. Cytosolic

RNA isolated from livers was used for quantification of rpL13a snoRNAs (relative to

36B4) or iNOS and Cox2 (relative to actin). Graph shows mean ± SE from a

representative experiment with N = 3 (PBS) and N = 4 (LPS) animals per group. * p <

0.05 for LPS vs. PBS.

U32a U33 U35a 0

2

4

6

targ

et:c

ontro

l RN

A(re

l uni

ts)

PBSLPS

**

*

0

500

1000

1500

2000

iNOS

*

0

10

20

30

40

50

Cox2

*

23

Figure 2.2

Figure 2.2 ASO knockdown of rpL13a snoRNAs in vivo. Mice were pretreated with

three serial doses of ASOs targeting rpL13a snoRNAs or GFP as control prior to LPS

injection and analysis of liver tissue. snoRNA, iNOS, and Cox2 expression in liver

cytosol was quantified as in Figure 2.3. n = 4 to 6 per group.

* p < 0.05 for GFP vs. snoRNA knockdown.

U32a U33 U35a 0.0

0.5

1.0

1.5

targ

et:c

ontro

l RN

A(re

l uni

ts)

gfpLNA + LPSsnoLNA + LPS

* * *0.0

0.5

1.0

1.5

iNOS0.0

0.5

1.0

1.5

2.0

Cox2

*

24

Figure 2.3

Figure 2.3 rpL13a snoRNAs are required for oxidative stress in vivo. Mice were

pretreated with three serial doses of ASOs targeting rpL13a snoRNAs or GFP as control

prior to LPS injection and analysis of liver tissue. Representative images show frozen

sections of liver tissue stained with DHE and parallel sections in which staining was

performed in the presence of pegylated superoxide dismutase (SOD) as control. Scale

bar, 100 µm. Graph shows quantification of fluorescence intensity. n = 4 to 6 per group.

* p < 0.05 for GFP vs. snoRNA knockdown.

DHE

DHE+

SOD

GFP LNA snoRNA LNA

0.0

0.5

1.0

1.5

DHE

Mea

n Fl

uore

scen

ce(re

l uni

ts)

*

sno LNAgfp LNA

25

Figure 2.4

Figure 2.4 In vivo rpL13a snoRNA knockdown reduces oxidative damage. Mice

were pretreated with three serial doses of ASOs targeting rpL13a snoRNAs or GFP as

control prior to LPS injection and analysis of liver tissue. Quantification of (A) protein

carbonylation by western blotting and (B) tissue oxysterols (7-ketocholesterol, 7-keto;

3β,5α,6β-cholestantriol, triol) 24 h following LPS. n = 4 to 6 per group. * p < 0.05 for

GFP vs. snoRNA knockdown.

0

200

400

600

800

1000

7-ketong

/mg

prot

ein

*

0.0

0.5

1.0

1.5

carbonyls

prot

ein

carb

onyl

s:ac

tin

(rel u

nits

)

*

0

20

40

60

80

triol

*

gfp LNAsno LNA

oxysterols

carbonyls

actin

gfp LNA sno LNAA B

26

Figure 2.5

Figure 2.5 U32a, U33, and U35a contain antisense homology to rRNA. Comparison

of rpL13a snoRNA sequences from mouse (Mm), hamster (Cg), and human (Hs)

species. C, D, and D’ box sequences are indicated in boxed bolded text. Regions with

antisense homology are underlined. Conserved and non-conserved nucleotides are

displayed in lower and upper case letters, respectively.

U32a

Mm GAgtcCAtgatgagcaacaCtcaccatctttcgtttgagtctcacgAcTGtgagatcaa-cccatgcaccgctctgagacTCCg AAgtcAGtgatgagcaacaAtcaccatctttcgtttgagtctcacgAcTGtgagatcaa-cccatgcaccgctctgagacTTHs AGgtcAGtgatgagcaacaTtcaccatctttcgtttgagtctcacgGcCAtgagatcaaCcccatgcaccgctctgagacCT

U33

Mm cAgcTTGtgatgagA-c-AtctcccactCATGttcgagttGcTcGacTatgagaTGactcTacatgcactaccatctgaggcTGTCg cAgcCTAtgatgaAG-cGAtctcccactGGTGttcgagttTcCcAacTatgagaCAactcTacatgcactaccatctgaggcTGGHs cGgcCGGtgatgagAAc-TtctcccactCACAttcgagttTcCcGacCatgagaTGactcCacatgcactaccatctgaggcCAC

U34

Mm cgtcTGtgatgttcTgcTaTtacctacattgtttgaGcctcatgaaaAcCCcactGgctgagacgcCg cgtcTGtgatgttcTgcTaTtacctacattgtttgaGcctcatgaaaAcGCcactAgctgagacgcHs cgtcCAtgatgttcCgcAaCtacctacattgtttgaTcctcatgaaaGcAGcactGgctgagacgc

U35a

Mm ggcaCatgatgtTCttatTctcacgatggtctTcggatgCcAcAgTTAGgGCaGtgCcgaTaatgccaAAggctAagctgatgccagGa Cg ggcaGatgatgtTTttTtTctcacgatggtctTcggatgCcAcCgATTGg-CaGtgCcgaTaatgccaCAggctCagctgatgccagTaHs ggcaGatgatgtCCttat-ctcacgatggtctGcggatgTcCcTgT---gGGaAtgGcgaCaatgccaATggctTagctgatgccagGa

From Michel et al. 2011

box C box D' box D

27

Figure 2.6

Figure 2.6 2’-O-methylation is unaffected in 6F2 cells. WT and 6F2 cells were

treated with 500 µM palmitate and total RNA analyzed for pseudouridylation or 2’-O-

methylation nucleotide modification of predicted sites using reverse transcriptase primer

extension. Autoradiograms show primer extension assays and parallel sequencing for

detection of (A) U32a and U33 target sites on 18S rRNA (G1328 and U1326,

respectively); (B) U32a target site on 28S rRNA (A1511); (C) U35a target site on 28S

rRNA (C5406); (D) unrelated snoRNA target sites on 18S rRNA as controls (snoRNAs

Z17a & Z17b target U121; snoRNAs U45a & U45c target A159). For each panel, arrows

point to bases in rRNA (numbered according to human rRNA sequence) that are

modified and corresponding DNA sequence is shown in the left-most four lanes.

ddNTP: ACGT [dNTP] H L H L H L H L

palmitate - - + + - - + +

G1328U1326,

A159

U121

A

A1511 C4506

ddNTP: ACGT [dNTP] H L H L H L H L

palmitate - - + + - - + +

CB

ddNTP: ACGT [dNTP] H L H L H L H L

palmitate - - + + - - + +

ddNTP: ACGT [dNTP] H L H L H L H L

palmitate - - + + - - + +

D

From Michel et al. 2011

WT 6F2 WT 6F2 WT 6F2 6F2WT

28

Figure 2.7

Figure 2.7 rpL13a snoRNAs accumulate in the cytoplasm under lipotoxic

conditions. (A, B) C2C12 cells were untreated (UT) or treated with palmitate for 24 h.

Cells were separated into cytosolic (CYT) and nuclear (NUC) fractions by sequential

detergent solubilization. (A) Fractions were analyzed by western blotting for a cytosolic

marker, hsp90, and for a nuclear marker, lamin B1. (B) Total RNA was prepared from

the fractions and analyzed for rpL13a snoRNA abundance relative to 36B4 by qRT-

PCR. Graphs show mean ± SE from a representative experiment (n = 3). * p < 0.05 for

palmitate treated vs. UT.

NUCU32a U33 U35a

0

1

2

3

4 UTpalmitate

snoR

NA

: 36B

4 R

NA

CYTU32a U33 U35a

0

1

2

3

4

snoR

NA

: 36B

4 R

NA

A B

**

*

Hsp90

Lamin B1

CYTNUC

29

Figure 2.8

Figure 2.8 rpL13a snoRNAs accumulate in the cytoplasm under lipotoxic

conditions. (A, B) C2C12 cells were analyzed by in situ hybridization under basal

conditions (UT) and following 24 h treatment with palmitate (PALM) using specific

snoRNA or control probes (red). Nuclei were stained with SYTOX Green. (A) Cells were

probed with U3 antisense probe for known nucleolar snoRNA, control U3 sense probe,

and control rpL13a intron 1 antisense probe. (B) Control GFP ASO-nucleofected and

specific snoRNA ASO-nucleofected cells were examined by in situ hybridization with

antisense probes for U32a, U33, and U35a. Bars, 10 µm.

UT

PALM

U3 sense

U3 antisense

rpl13a intronantisenseprobe

U33 antisense probeGFP U33

U35a antisense probeGFP U35a

UT

PALM

ASOU32a antisense probe

GFP U32a

A

B

30

CHAPTER THREE

SmD3 regulates intronic snoRNA biogenesis

INTRODUCTION

Most vertebrate snoRNAs are encoded within introns and co-transcribed with

their host genes (109). Intronic box C/D snoRNP protein assembly initiates during the

C1 complex stage of splicing (37), with subsequent intron lariat formation at the C2

complex stage of splicing followed by debranching and exonucleolytic trimming of the

mature snoRNP (51, 80, 85). The observations that rpL13a snoRNAs rapidly

accumulate in the cytosol during metabolic stress and are required for lipotoxic cell

death suggest that cytoplasmic RNAs may be their primary targets and that efficient

processing of these intronic elements is important for the lipotoxic response. However,

the precise molecular mechanisms through which snoRNAs are induced and regulated

during lipotoxicity remain to be elucidated.

Our genetic screen led to isolation of a second, independent mutant cell line

harboring a disruption in an RNA-related gene. This novel mutant cell line is

haploinsufficient for SmD3, a core component of the spliceosome. We demonstrate that

SmD3 participates in the lipotoxic response through regulation of intron lariat

abundance and biogenesis of intron-encoded rpL13a snoRNAs. We also provide

evidence linking the expression of SmD3 to the levels of critical snRNA components of

the spliceosome and generalized production of intronic non-coding RNAs (ncRNAs).

Our results extend the known function for SmD3 in splicing to a specific role within

individual snRNPs essential for the biogenesis of intronic ncRNAs.

31

RESULTS

SmD3 haploinsufficiency confers resistance to palmitate-induced cell death. The

6H2 mutant cell line was isolated from our genetic screen for genes that are critical for

the lipotoxic response. To quantify the degree of palmitate resistance in mutant 6H2,

WT and 6H2 cells were treated with palmitate for 48 h, and cell death was quantified by

propidium iodide (PI) staining and flow cytometric analysis. Compared to WT cells, 6H2

cells were significantly protected from palmitate-induced death (Figure 3.1A). By

contrast, treatment of WT and 6H2 cells with the general apoptosis inducers

camptothecin, staurosporine, and actinomycin D revealed no differences in sensitivity.

Therefore, palmitate-resistant 6H2 cells are not generally resistant to cell death.

The promoter trap mutagenesis facilitated identification of the disrupted gene

because of the unique fusion transcript produced upon a single productive integration.

To confirm the presence of a single retroviral integration, Southern blot analysis of 6H2

genomic DNA was performed, probing for the ROSAβgeo sequence (Figure 3.2A). The

presence of a single hybridizing band in DNA digested with multiple different restriction

enzymes is consistent with a single retroviral integration. To identify the disrupted gene

in the mutant 6H2 cells, mRNA was isolated and used for 5’ rapid amplification of cDNA

ends (5’ RACE). Unique sequence from the RACE product was analyzed using NCBI

BLAST, which revealed that the site of integration was the snrpD3 gene encoding the

small nuclear ribonucleoprotein SmD3. SmD3 is a component of the spliceosome that,

together with other Sm family proteins, forms a heteroheptameric ring around snRNAs

U1, U2, U4, and U5 to generate small nuclear ribonucleoproteins (snRNPs) essential for

32

the removal of introns from pre-mRNA (125). PCR was performed to confirm integration

of the ROSAβgeo sequence into snrpD3 (Figure 3.2B). Reactions with forward and

reverse primers designed to snrpD3 resulted in PCR products for both WT and 6H2

cDNA, indicating that each cell type maintains at least one intact allele. As expected, no

product was detectable in WT cells when snrpD3 forward and ROSAβgeo reverse

primers were used, but this set of primers produced the expected PCR product from the

fusion transcript in 6H2 cells. These PCR results confirm our 5’ RACE identification of

the disrupted gene and suggest that 6H2 cells are haploinsufficient for snrpD3.

Consistent with this model, quantitative real-time PCR (qRT-PCR) revealed a ~50%

reduction in relative snrpD3 mRNA (Figure 3.2C) and western blotting revealed a

corresponding ~50% reduction of SmD3 protein (doublet at 15 kDa and 18 kDa) in 6H2

relative to WT cells (Figure 3.2D). Thus, expression of snrpD3 in 6H2 cells is consistent

with a model in which integration of the ROSAβgeo provirus disrupted one of two alleles

for snrpD3.

Targeted knockdown of SmD3 recapitulates 6H2 phenotype. To confirm that the

palmitate-resistant phenotype in 6H2 cells is due to diminished SmD3 protein

expression, we used shRNA to knockdown SmD3 in WT CHO cells and tested for

associated changes in palmitate sensitivity. SmD3 protein levels were measured by

western blot following isolation of individual stable clonal knockdown lines. Two

independently isolated clonal lines showed 47% (sh1) and 64% (sh2) knockdown

relative to scrambled (contr) shRNA transfected cells (Figure 3.3A and 3.3B).

Knockdown clones were protected from palmitate-induced death as measured by PI

33

staining, and the degree of protection was proportional to the degree of knockdown

(Figure 3.3C). These data provide independent genetic evidence that loss-of-function of

SmD3 protects against lipotoxicity.

SmD3 disruption protects cells from generalized oxidative stress induction.

Lipotoxicity is known to involve fatty acid (FA) import and the generation of oxidative

stress (8, 60). To test whether 6H2 cells acquired resistance through diminished

capacity to take up palmitate, initial rates of FA uptake were quantified in WT and 6H2

cells. There was no significant difference between WT and 6H2 cells (Figure 3.4A),

indicating that resistance to lipotoxicity in 6H2 cells did not result from failure to take up

exogenous FA. To probe downstream aspects of the lipotoxic response, we quantified

palmitate-induced ROS in WT and 6H2 cells by CM-H2DCFDA (DCF) staining and flow

cytometric analysis. At 5 and 16 h following palmitate supplementation, ROS induction

was significantly blunted in 6H2 cells (Figure 3.4B). SmD3 knockdown clones were also

protected from palmitate-induced ROS (Figure 3.4C). More direct induction of oxidative

stress following exposure to H2O2 or menadione also resulted in blunted ROS levels in

6H2 cells compared to WT (Figure 3.4D), indicating 6H2 cells are protected not only

from palmitate-induced ROS but also from generalized oxidative stress induction or

amplification.

SmD3 regulates intronic non-coding RNA expression. The ROS resistance

phenotype observed in 6H2 cells is similar to the previously described 6F2 mutant.

Given the related phenotypes of these two mutants, and the well-appreciated

34

interactions of SmD3 with RNA, we assayed for the expression of the rpL13a snoRNAs

in 6H2 cells by RNase protection assay. Following palmitate treatment, WT cells show

the expected increase in snoRNA expression by RNase protection (Figure 3.5A). Under

the same conditions, snoRNA induction is blunted in 6H2 cells. Similarly, SmD3

knockdown clones were impaired in rpL13a snoRNA induction relative to control (Figure

3.5B). While snoRNAs are thought to be produced in the nucleus and canonical box

C/D snoRNAs function in that location, our data demonstrates that rpL13a snoRNAs

accumulate in the cytosol during metabolic stress (Chapter 2). Fractionation of WT and

6H2 cells by sequential detergent solubilization revealed that under basal conditions

snoRNAs are detectable in the nucleus and the cytosol but are substantially more

abundant in the nucleus (Figures 3.5C and 3.5D). qRT-PCR analysis of these fractions

reveals reduced rpL13a snoRNAs in the cytosol in 6H2 cells under palmitate-treated

conditions, and reduced levels of these snoRNAs in the nucleus under both basal and

palmitate-treated conditions (Figures 3.5E and 3.5F). The observation that

haploinsufficiency of SmD3 caused impairment of basal expression and lipotoxic

cytosolic accumulation of the intronic rpL13a snoRNAs, a deficit known to cause

resistance to lipotoxicity, is consistent with the ROS resistant phenotype observed in

6H2 cells. Furthermore, the observation that nuclear levels of the rpL13a snoRNAs are

decreased in 6H2 cells under basal as well as lipotoxic conditions implicates a defect in

the nuclear production of the snoRNAs.

To test whether 6H2 cells have a general defect in expression of intronic

snoRNAs, we measured basal nuclear expression of intronic box C/D snoRNAs U50,

U57, U60, and U21 and intronic box H/ACA snoRNAs U17b, U64, and ACA28 (Figure

35

3.5G). Expression of each of these snoRNAs was reduced in the nuclei of 6H2 versus

WT cells. Furthermore, we measured the expression of intronic splicing-

dependent/Drosha-independent pre-miRNAs to probe an unrelated class of intronic

ncRNAs (6, 89). Mirtrons miR-1224 and miR-1225 showed reduced expression in 6H2

cells. In contrast, independently transcribed, splicing-independent, non-intronic

snoRNAs U8 and U13 and pre-miR-23a showed no difference in expression between

WT and 6H2. Taken together, these data suggest that WT levels of SmD3 are generally

required for effective expression of splicing-dependent intronic ncRNAs.

SmD3 knockdown perturbs snRNP biogenesis. Disruption of multiple components of

the snRNP assembly pathway has previously been shown to alter snRNP expression

(98, 130). To test whether reduced levels of SmD3 affect snRNA expression, we used a

chimeric locked nucleic acid/DNA (LNA/DNA) oligonucleotide to specifically knockdown

SmD3 in murine fibroblasts. Following knockdown of SmD3 to ~50% of control SmD3

levels, we observed reductions in rpL13a snoRNAs similar to our mutant, (Figures 3.6A

and 3.6B). In SmD3 knockdown cells, qRT-PCR revealed that U4 and U5 snRNA

expression was decreased, but levels of other snRNAs were indistinguishable from

control (Figure 3.6C). Since unbound snRNAs are unstable compared to snRNAs in

snRNP complexes, quantification of snRNA levels provides insight into snRNP integrity

(91, 131). Consistent with this, U4 and U5 snRNAs were reduced following

immunoprecipitation with the α-Sm protein Y12 antibody, indicating that U4 and U5

snRNPs are also diminished (Figure 3.6D). By contrast, 50% knockdown of SmB,

another Sm protein family member, produced more modest decreases in snoRNA

36

production (Figure 3.6F). SmB knockdown cells remained sensitive to palmitate-induced

death (Figure 3.6G) and displayed a different pattern of alteration of snRNA expression,

characterized by decreases in U2 and U4 and increase in U4atac (Figure 3.6H). Our

data suggest that a reduction in SmD3 expression disrupts a specific set of snRNPs.

While there is some overlap with the effects of SmB knockdown on snRNA levels, the

pattern with SmD3 is distinct.

6H2 cells display normal splicing efficiency. Given that SmD3 levels affect snRNP

expression, we hypothesized that differences in intronic ncRNA expression caused by

SmD3 haploinsufficiency in the 6H2 cells could be explained by a defect in splicing. To

assess splicing related to the rpL13a snoRNAs, we measured the expression of

endogenous rpL13a splicing precursors and spliced products. There was no difference

in rpL13a pre-mRNA levels between WT and 6H2 cells under basal or palmitate-treated

conditions, as assessed by amplifying random hexamer-primed cDNA with primer pairs

designed across the junction between exon 3 and intron 3 (Ex3/Int3, Figures 3.7A and

3.7B). Quantification of the endogenous rpL13a mRNA, using oligo-dT-primed cDNA

and primers designed across the splice junction between exons 7 and 8, showed similar

expression between WT and 6H2 cells (Ex7/8, Figures 3.7A and 3.7B). Furthermore,

priming across four splice junctions formed by the removal of snoRNA-encoding introns

(Ex 2/3, 4/5, 5/6, and 6/7) displayed similar expression between WT and 6H2 cells

(Figures 3.7A and 3.7B).

To assess splicing efficiency more broadly, we transfected WT and 6H2 cells

with a previously validated reporter construct for expression of firefly luciferase from two

37

exons separated by a β-globin intron (128). Functional splicing results in the formation

of a luciferase mRNA encoding a protein with measurable luminescence and a

processed β-globin intron lariat. In the absence of splicing, a truncated luciferase protein

without enzymatic activity is formed due to multiple in-frame stop codons. Following

transfection of this construct, no difference in luciferase production was detected

between WT and 6H2 under untreated or palmitate treated conditions or in the presence

of clotrimazole, a known splicing inhibitor (128) (Figure 3.7C). To test whether the

presence of an intronic snoRNA affected splicing of flanking exons in 6H2 cells, we

replaced the intron in the split luciferase vector with the murine rpL13a intron 2

containing U32a. Following transfection with this construct, there was no difference in

luciferase levels between WT and 6H2 cells, although expression of the exogenous

murine U32a was reduced in 6H2 cells (Figures 3.7D and 3.7E). Similarly, we detected

no difference in luminescence using a Δsno split luciferase splicing reporter containing

mutated rpL13a intron 2 sequences lacking the entire 83 nucleotide U32a, and pre-

mRNA from the reporters with the intact U32a intron and the Δsno intron were

comparable (not shown). Furthermore, we quantified mRNA expression of the host

genes containing the intronic non-coding elements quantified in Figure 3.5G. We

detected no differences in mRNA levels from any of these host genes between WT and

6H2 cells (Figure 3.7F). Together, these data indicate that differences in intronic non-

coding RNA levels are not attributable to defective splicing of exons in 6H2 cells.

SmD3 knockdown does not alter alternative splicing. Previously, it was reported

that 85% or more knockdown of the survival of motor neuron (SMN) protein disrupts the

38

snRNP assembly pathway and leads to widespread differences in snRNA levels and

alternative splicing in a number of tissues (130). To test whether reduced levels of

SmD3 affected snoRNA processing through broad alterations in alternative splicing,

exon utilization in control (GFP) and SmD3 knockdown cells was analyzed using the

Affymetrix GeneChip mouse exon 1.0 ST microarray. Due to the lack of publically

available exon microarrays containing hamster sequences, we used our LNA/DNA

oligonucleotides to generate murine fibroblasts with haploinsufficiency of SmD3 to

phenocopy the 6H2 cells that have 50% wild type CHO levels for this protein (Figures

3.6A and 3.6B). Following knockdown, RNA was harvested from three independent

samples of GFP and SmD3 knockdown cells. Among the 266,200 probesets supported

by putative full-length mRNA, ∼190,000 probesets with significant signals above

background and representing exons of ~16,000 out of ~30,000 genes in the mouse

genome were included in the analysis. With a false discovery rate set at less than 0.1,

no genes were identified as having potential splicing pattern changes (fold change ≥

1.5). Plotting probeset intensity values from control (GFP LNA) samples against SmD3

knockdown samples revealed a highly linear relationship (R2 = 0.98475), consistent with

little variance between the two groups (Figure 3.8). These data show that a 50%

reduction in SmD3 expression does not have global effects on alternative splicing.

SmD3 controls intron lariat abundance. Box C/D snoRNAs, box H/ACA snoRNAs,

and mirtrons are each defined by unique consensus sequences, protein assembly

factors, and position within the intron (6, 38, 69, 88, 89). Nonetheless, these intronic

non-coding RNAs all require pre-mRNA splicing, intron lariat formation, debranching,

39

and exonucleolytic trimming prior to formation of a functional ribonucleoprotein. We

used qRT-PCR to probe snoRNA precursors to determine the level of processing at

which 6H2 cells are defective. Intron lariats from each rpL13a snoRNA-containing intron

were quantified by qPCR primers reading across the branch point with a sense primer

designed to a 3’ region of the intron and an antisense primer to a 5’ region (116). This

approach revealed a 38-63% decrease in lariats from each of the snoRNA-containing

rpL13a introns under untreated and palmitate treated conditions in 6H2 cells (Figure

3.9A). Validity of this approach was confirmed by the observation of a single PCR

product for each intron lariat (Figure 3.9B) and sequence analysis of the PCR products

(Figure 3.9C). Each primer pair read across the branch point and allowed for mapping

branch sites in rpL13a introns, each defined by an adenosine and 6 of 7 nucleotides

correlating with the consensus major spliceosome branch site. Interestingly, we were

unable to detect PCR products for endogenous rpL13a intron lariats lacking snoRNAs, a

finding consistent with recent work from others showing that intronic sequences lacking

snoRNAs are degraded more quickly than introns containing snoRNAs (126). On the

other hand, we were able to quantify both snoRNA-containing and non-snoRNA-

containing introns when they were overexpressed in cells transfected with the split

luciferase vector construct containing the U32a intron and the snoRNA-deleted U32a

sequence, respectively. In WT cells the presence of the intronic snoRNA was

associated with increased lariat abundance, whereas this apparent increase was not

observed in 6H2 cells (Figure 3.9D). These findings suggest that SmD3 contributes to

expression of intronic snoRNAs by enhancing intron lariat formation or stability.

40

CONCLUSIONS

Through the use of a genetic screen in CHO cells, our laboratory has identified

loci involved in lipid-induced cell death, a process in which oxidative stress is a central

feature. Lipotoxic and oxidative stress critically involve cytosolic expression of intronic

snoRNAs from the rpL13a genomic locus (72). We provide additional insight into the

underlying mechanisms of metabolic stress responses through the characterization of a

mutant cell line with disruption of one allele of the gene encoding SmD3. Our data show

that reduced cellular levels of SmD3 decrease the propagation of oxidative stress and

protect cells from palmitate-induced death. Although mutant 6H2 cells are distinct from

previously described palmitate-resistant mutants from our screen, mutation at the SmD3

locus confers a related molecular phenotype in that 6H2 cells fail to induce rpL13a

snoRNAs under lipotoxic stress. We show that SmD3, beyond its known role in splicing,

regulates expression of the intronic snoRNAs. Compared to WT levels of SmD3,

reduced levels of SmD3 lead to decreased intron lariat abundance and decreases in U4

and U5 snRNPs necessary for intron lariat formation. These perturbations decrease the

basal levels of the rpL13a intronic snoRNAs and subsequently blunt their induction

during lipotoxicity.

SmD3 together with six other Sm proteins form a heptameric ring around the

uridine-rich Sm-binding site found in snRNAs. These snRNPs form the major building

blocks of the spliceosome. Sequential assembly of the hetero-oligomers of SmD1-D2,

SmE-F-G, and SmB/B’-D3 is a highly regulated process (27, 84). Although these

proteins are critical for proper snRNP assembly, our data indicate that

haploinsufficiency of SmD3 is sufficient to maintain splicing of a specific endogenous

41

pre-mRNA and an exogenously provided splicing reporter under normal growth

conditions. Consistent with an ability to support overall wild type capacity for mRNA

splicing, the growth of mutant 6H2 cells is indistinguishable from parental wild type cells.

These findings suggest that excision of introns during the catalytic stage of splicing

goes to completion and haploinsufficient levels of SmD3 in these cells is not limiting for

production of mRNAs. Furthermore, ~50% knockdown of SmD3 does not cause global

changes in exon utilization. These findings indicate that the phenotype of SmD3

haploinsufficiency does not relate to global defects in formation of mRNAs.

To our knowledge, this is the first report of the loss-of-function phenotype for

SmD3. The finding of a second mutation that reduces rpL13a snoRNA expression and

leads to resistance to lipotoxicity further highlights the important role of these ncRNAs in

metabolic stress responses. More importantly, our study provides novel insights into the

broader molecular cell biology of intronic non-coding RNA elements. Future studies of

the precise molecular interactions of SmD3 within individual snRNPs are likely to

elucidate mechanisms through which this protein regulates non-coding RNA biogenesis.

42

Figure 3.1

Figure 3.1 6H2 cells are resistant to palmitate-induced cell death. (A) WT and palm-

resistant 6H2 mutant cells were incubated with 500 µM palm for 48 h; or 10 µM

camptothecin (camp), 80 nM staurosporine (staur), or 2 µM actinomycin D (actD) for 24

h. Cell death was quantified by propidium iodide (PI) staining and flow cytometry. Data

are expressed as mean fluorescence ± standard error (SE) for 3 independent

experiments with 104 cells/sample. *, p<0.005 for 6H2 versus WT.

UT palm camp staur actD0

20

40

60

80 WT6H2

PI p

ositi

ve c

ells

(%)

*BAA

43

Figure 3.2

Figure 3.2 6H2 cells are haploinsufficient for SmD3. (A) Autoradiogram shows

Southern blot analysis of WT (lanes 1-3) and 6H2 (lanes 4-6) genomic DNA digested

with restriction enzymes Bgl II, NcoI, or XbaI. Blot was probed with a 32P-labeled

fragment corresponding to the ROSAβgeo sequence. (B) PCR was performed on cDNA

from WT (lanes 1 and 2) and 6H2 (lanes 3 and 4) cells with reactions containing no

cDNA as controls (lanes 5 and 6). Forward (F) and reverse (R) primers for snrpD3 were

designed to detect endogenous snrpD3 (lanes 1, 3, and 5). Forward snrpD3 primer and

reverse primer for the proviral sequence were used to detect fusion transcript (lanes 2,

4, and 6). (C) RNA was isolated from WT and 6H2 cells and reverse transcribed using

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either random hexamers (ran hex) to prime total RNA or oligo dT to prime mRNA.

snrpD3 expression was determined by quantitative real-time PCR (qPCR) and

normalized to β-actin expression. (D) Protein expression in WT and 6H2 cells was

determined by western blotting and quantified by densitometry. Bands at 15 kDa and 18

kDa likely reflect known post-translational modification of SmD3. Representative blot

shown for SmD3 and β-actin control. On bar graphs, data are expressed as mean ± SE

for three independent experiments. *, p < 0.05 for 6H2 versus WT.

45

Figure 3.3

Figure 3.3 Targeted knockdown of SmD3 confers palmitate-resistance. Stable

clonal cell lines were generated following transfection with a scrambled (contr) or

snrpD3-targeting shRNA (sh1 and sh2). (A) SmD3 and β-actin protein expression was

determined by western blot. Blot shows three independent protein samples from each

respective cell line. (B) SmD3 expression relative to β-actin was quantified by

densitometry of blots as shown in A. Graph shows mean ± SE for three independent

samples. *, p < 0.05 for knockdown versus scrambled. (C) Scrambled and knockdown

cells were treated with palm for 48 h and cell death was assessed by PI staining and

flow cytometry. All data are expressed as mean fluorescence ± SE for three

independent experiments with 104 cells/sample. *, p<0.005 for knockdown versus

scrambled.

contr sh1 sh20

20

40

60 UTpalm

PI p

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contr sh1 sh2

SmD3

*-actin

A B

C

18 kDa15 kDa

42 kDa

46

Figure 3.4

Figure 3.4 SmD3 disruption protects cells from palmitate-induced and generalized

oxidative stress induction. (A) WT and 6H2 cells were incubated with 14C-palmitate

under lipotoxic conditions (500 µM palm). Mean initial rates of palmitate uptake are

expressed per µg protein (± SE) for 3 independent experiments. (B,C,D) WT and 6H2

cells were incubated with palmitate (B, 5 hr and 16 h), scrambled (contr) and

knockdown (sh1 and sh2) cells were treated with palmitate (C, 16 h), or WT and 6H2

cells were untreated (UT) or treated with menadione (mena) or H2O2 (D, 2h). ROS

induction was assessed by CM-H2DCFDA (DCF) labeling and flow cytometry. Graphs

show mean fluorescence ± SE for 3 independent experiments with 104 cells/sample. *,

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p<0.05 for 6H2 versus WT, or for knockdown versus scrambled.

48

Figure 3.5

Figure 3.5 Intronic non-coding RNA expression is disrupted in 6H2 cells. (A,B) WT

and 6H2 cells (A) and scrambled control and knockdown cells (B) were untreated or

supplemented with palm for 48 h. Small RNA was harvested and used in RNase

protection assay with 32P-labeled rpL13a snoRNA probes or miR-16 probe as control.

Protected probe was analyzed by autoradiography. (C,D,E,F)  WT and 6H2 cells were

untreated or treated with palm for 9 h. Cells were separated into cytosolic (CYT) and

nuclear (NUC) fractions by sequential detergent solubilization. (C) Fractions were

analyzed by western blotting or PCR visualization for cytosolic markers, hsp90 and

tRNAGlu, and for nuclear markers, fibrillarin and U6 RNA. (D) Molar quantities of

CYTOSOL

WT 6H2 WT 6H2 WT 6H20

1

2

3

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U32a U33 U35a

snoR

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: 5-a

ctin

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NUCLEUS

WT 6H2 WT 6H2 WT 6H20.0

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UTpalm

U32a U33 U35a

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snoR

NA

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ctin

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A

A B C

U32a

U33

U35a

miR-16

palm - + - +WT 6H2

F

U32a

U33

U35a

miR-16

palm - + - + - +contr sh1 sh2

hsp90

fibrillarin

CYT NUC

E

U50 U57 U60 U21 U8 U13U17b U64

ACA28

miR-1224

miR-1225miR-23a

0.0

0.5

1.0

1.5WT6H2

snoRNA pre-miRNA

C/Dintronic

C/Dnon-intronic

H/ACAintronic

mirtron miRNAnon-intronic

* ** * * * * *

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100

200

300

400

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49

rpL13a snoRNAs from cytosolic and nuclear fractions were quantified relative to a

standard curve. Plot represents relative molar ratio. (E,F) Total RNA was prepared from

the cytosolic (E) and nuclear (F) fractions and analyzed for rpL13a snoRNA abundance

relative to β-actin by qRT-PCR. (G) Total RNA was prepared from nuclear fractions of

WT and 6H2 cells and analyzed for intronic and non-intronic box C/D snoRNAs; intronic

box H/ACA snoRNAs; and splicing-dependent/Drosha-independent intronic and non-

intronic miRNAs. All data are expressed as mean ± SE for three independent

experiments. *, p < 0.05 for 6H2 versus WT.

50

Figure 3.6

Figure 3.6 SmD3 knockdown disrupts U4 and U5 snRNPs. (A,B,C,D) NIH 3T3 cells

were transfected with LNA/DNA oligonucleotides specifically targeting GFP or SmD3.

(A) Protein expression in GFP and SmD3 transfected cells was determined by western

blotting and quantified by densitometry. Representative blot shown for SmD3 and β-

actin control. (B) Nuclear RNA was isolated and analyzed for SmD3 mRNA and rpL13a

snoRNA expression by qRT-PCR relative to 36B4. (C) Total RNA was isolated and

A B

C D

E F

RN

A e

xp

res

sio

n

(re

l u

nit

s)

SmD3 U32a U33 U35a0.0

0.5

1.0

1.5

GFP LNA

SmD3 LNA

* * * *

U1 U2 U4 U5 U6 U11 U12 U4atacU6atac0.0

0.5

1.0

1.5

sn

RN

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xp

res

sio

n

(re

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nit

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SmD3 LNA

* *

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RN

A e

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n

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/Ig

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U1 U2 U4 U5 U60.0

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1.0

1.5

GFP LNA

SmD3 LNA

* *

SmB U32a U33 U35a0.0

0.5

1.0

1.5

RN

A e

xp

res

sio

n

(re

l u

nit

s)

contr siRNA

SmB siRNA

* * * *

U1 U2 U4 U5 U6 U11 U12 U4atacU6atac0.0

0.5

1.0

1.5

2.0

sn

RN

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xp

res

sio

n

(re

l u

nit

s)

contr siRNA

SmB siRNA

* *

*

GFP LNASmD3 LNA0.0

0.2

0.4

0.6

0.8

1.0S

mD

3 p

rote

in

(re

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nit

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*

GFP

LNA

SmD3

LNA

GFP

LNA

SmD3

LNA

SmD3

4-actin42 kDa

15 kDa

G

H

GFP LNASmD3 LNA0.0

0.2

0.4

0.6

0.8

1.0

Sm

B p

rote

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(re

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nit

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contr

siRNA

SmB

siRNA

contr

siRNA

SmB

siRNA

SmB

4-actin42 kDa

25 kDa

UT 24 h 48 h

0

20

40

60

80

PI p

ositiv

e c

ells (%

)

contr siRNA

SmB siRNA

palm

ns

51

analyzed for snRNA expression relative to 36B4 by qRT-PCR. (D) Cell lysates were

immunoprecipitated using α-Sm Y12 antibody or IgG control. RNA was isolated

following immunoprecipitation and analyzed by qRT-PCR for α-Sm immunoprecipitated

snRNA relative to control (IgG) precipitated. (E,F,G,H) NIH 3T3 cells were transfected

with control (contr) siRNA or siRNA targeting SmB. (E) Protein expression in control and

SmB siRNA cells was determined by western blotting and quantified by densitometry.

Representative blot shown for SmB and β-actin control. (F) Nuclear RNA was isolated

and analyzed for SmB and rpL13a snoRNA expression by qRT-PCR relative to 36B4.

(G) Control and SmB siRNA cells were incubated with 500 µM palm for 24 h or 48 h.

Cell death was quantified by PI staining and flow cytometry. (H) Total RNA was isolated

and analyzed for snRNA expression by qRT-PCR relative to 36B4. Data are expressed

as mean ± SE for three independent experiments. *, p < 0.05 SmD3 versus GFP or

SmB versus control; ns, not significant.

52

Figure 3.7

Figure 3.7 Host gene expression is normal in 6H2 cells. (A) rpL13a locus is shown

with non-coding region in black lines, exons in black boxes and snoRNAs in gray ovals.

Location of primers for qPCR analysis of pre-mRNA and mRNA are noted. (B) WT and

6H2 cells were untreated (UT) or treated with palm for 9 h. For analysis of rpL13a pre-

mRNA expression, total RNA was reverse transcribed using random hexamers, and

amplified using primers that span the junction between exon 3 and intron 3. For analysis

of rpL13a mRNA expression, total RNA was transcribed using oligo-dT and amplified

using primers that span exon-exon junctions. Primer pairs are as indicated in (A). (C)

WT and 6H2 cells were transfected with a split luciferase reporter containing a β-globin

A B

D E

F

Snhg5 Nop56 U60hg RPL5 Snhg3 RPS2 EIF5 VWA5B2 PKD10.0

0.5

1.0

1.5

ncR

NA

hos

t gen

e:

6-ac

tin R

NA

WT6H2

ns

(U50) (U57) (U60) (U21) (U17b) (U64) (ACA28) (miR1224)(miR1225)

intronic box C/D snoRNA host genes

intronic box H/ACA snoRNA host genes

mirtron host genes

ns nsnsnsnsns ns ns

WT 6H2 WT 6H20.0

0.5

1.0

1.5

sno 5 sno

splicing reportersnoRNA

*

snoR

NA

: luc

expr

essi

on (r

el u

nits

)

WT 6H2 WT 6H20.0

0.5

1.0

1.5

2.0

5 snosnosplicing reporter

ns ns

lucluc

lum

ines

cenc

e / l

ucex

pres

sion

(rel

uni

ts)

lucluc

WT 6H20.0

0.5

1.0

1.5UTpalm

*

clor

*

nsns

ns

lum

ines

cenc

e / l

ucex

pres

sion

(rel

uni

ts)

C

Ex3/Int3 Ex2/3 Ex4/5 Ex5/6 Ex6/7 Ex7/8 Ex3/Int3 Ex7/80.0

0.5

1.0

1.5

RN

A:6

-act

in R

NA

(rel u

nits

)

ns

WT6H2

ns ns ns ns ns ns ns

UT palm

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53

intron. 24 h post-transfection, cells were untreated or treated with palm or clortrimazole

(clor) for 4 h. Luminescence was measured and normalized to luciferase pre-mRNA

expression by qPCR. (D,E) WT and 6H2 cells were transfected with a split luciferase

construct containing the intact U32a intron (sno) or the U32a intron lacking the 83

nucleotide U32a snoRNA (Δsno). Total RNA was analyzed for luciferase pre-mRNA and

U32a snoRNA expression. Luminescence (D) and U32a snoRNA (E) were normalized

to luciferase pre-mRNA expression. Note that sequences differences between the

murine intronic sequences in the reporter construct and endogenous hamster

sequences enable discrimination between exogenous murine and endogenous hamster

snoRNAs using species-specific PCR primers. (F) Total RNA was prepared from WT

and 6H2 cells and analyzed for host genes of endogenous intron-encoded snoRNAs

and mirtrons by qRT-PCR relative to β-actin mRNA. All data are expressed as mean ±

SE for three independent experiments. *, p < 0.05; ns, not significant.

54

Figure 3.8

Figure 3.8 Alternative splicing is normal in 6H2 cells. Exon array analysis was used

to predict differences in alternative splicing. Relative probeset intensity values from

exon array were plotted for SmD3 LNA versus GFP LNA transfected cells using data

from three independent samples/arrays for each condition.

SmD3

LNA

0 1.4 2.8 4.2 5.6 7 8.4 9.8 11.2 12.6 140

1.4

2.8

4.2

5.6

7

8.4

9.8

11.2

12.6

14

GFP LNA

R2 = 0.98475

55

Figure 3.9

Figure 3.9 SnoRNA-containing intron lariats are decreased in 6H2 cells. (A) WT

and 6H2 CHO cells were untreated or treated with palm for 9 h. Nuclear RNA was

isolated and reverse transcribed using random hexamers. rpL13a intron lariat

abundance was determined by qRT-PCR using primers reading across branch points

(red arrows in diagram). Quantification of lariat PCR product is normalized to β-actin

mRNA. (B,C) PCR products generated by intron lariat primers in CHO WT cells were

visualized on 2% agarose gel (B) or cloned and sequenced (C). (B) Lanes with cDNA

(+) contain RT product as template. Lanes without cDNA (-) were negative control PCR

reactions with no RT product provided as template. (C) Sequencing revealed the branch

A B

C

WT 6H2 WT 6H2 WT 6H2 WT 6H20.0

0.5

1.0

1.5UTpalm

intron 2 intron 4 intron 5 intron 6

rpL13a lariats

* * * *

laria

ts: 3

-act

in R

NA

snoRNA U32a U33 U34 U35a

WT 6H2 WT 6H20.0

0.5

1.0

1.5

2.0

2.5

sno 2 sno

luciferase lariats

*

ns

*

ns

laria

ts: l

uc e

xpre

ssio

n (r

el u

nits

)

D

U32a intron 2

intron 4

intron 5

intron 6

CU

GU

GU

GU

UCUUcAc polyY

CUCUGAa polyY

polyY

polyY

CUCUGAg

UCCUuAC

CU

AG

AG

AG

U33

U34

U35a

YNYURAYBranch site

5'SS 3'SS

intron 2 intron 4 intron 5 intron 6cDNA + - + - + - + -

200 bp -100 bp -

56

site adenosine (red) for each intron. Branch site nucleotides showing conservation

(uppercase) are indicated. (D) WT and 6H2 cells were transfected with a split luciferase

construct containing the intact murine U32a intron (sno) or the U32a intron lacking the

83 nucleotide U32a snoRNA (Δsno). Intron lariat abundance was determined by qRT-

PCR using primers specific for the murine lariat sequences and normalized to luciferase

pre-mRNA expression. All data are expressed as mean ± SE for three independent

experiments. *, p < 0.05; ns, not significant.

57

Figure 3.10

Figure 3.10 Role for SmD3 in snoRNA production. A model for SmD3 role in

snoRNA production is shown. Open boxes denote exons; black lines denote intronic

sequences; and snoRNA sequences are shown in yellow and blue. Colored balls

represent snRNPs. In the presence of wild type levels of SmD3, the complement of

snRNAs is sufficient to support splicing of pre-mRNAs into mature mRNAs and intron

lariats that are sufficiently long-lived to produce snoRNAs. While haploinsufficiency of

SmD3 is able to support wild type levels of mRNA production from splicing, associated

decreases in the abundance of the of U4 and U5 snRNPs results in decreased intron

lariat abundance and decreased levels of intronic snoRNAs (yellow, blue).  

SmD3+/+

+ +

SmD3+/-����

����

����

����

����

����

58

CHAPTER 4

Superoxide triggers cytosolic snoRNA accumulation

INTRODUCTION

Under physiological conditions, ROS serve a number of critical functions.

Superoxide (O2-) and hydrogen peroxide (H2O2) are known to regulate gene expression

by signaling through phosphatases, JAK/STATs, MAP kinases, and transcription factors

including NF-κB (34). ROS levels are kept in check by antioxidant enzymes and

scavengers including dismutases, peroxiredoxins, glutathione peroxidases, and

catalase.

Under pathophysiological conditions, when ROS generation exceeds cellular

antioxidant defenses, oxidative stress results. The rpL13a snoRNAs are critical for ROS

propagation during lipotoxic stress (72). NADPH oxidase activation is also important for

FFA-induced oxidative stress, since inhibition of NADPH oxidase blocks lipotoxic cell

death (11). Lipotoxic conditions can activate NADPH oxidase through ceramide

synthesis or, in the absense of de novo ceramide synthesis, through protein kinase C

(PKC) dependent pathways (45, 61).

Nox enzymes generate ROS by transferring electrons from NADPH to molecular

oxygen. O2- is the primary product of Nox enzymes, and Nox4 is also able to produce

H2O2. In many cell types including cardiomyocytes and cardiac fibroblasts, Nox2 and

Nox4 are the predominant NADPH oxidases (67). Nox2 is normally quiescent, and its

activation requires stimulus-induced membrane translocation of cytosolic regulatory

subunits, including p47phox, p67phox, p40phox, and Rac1, a small GTPase. This

59

translocation is triggered by phosphorylation of p47phox by PKC (2). Nox4 is

constitutively active when assembled as a heterodimer with p22phox. Nox4 ROS

generation does not require association of cytosolic factors but is regulated at the gene

expression level.

Given that NADPH oxidase activation is a critical contributor to ROS generation

during lipotoxicity, we hypothesized that specific ROS (e.g. O2-) may be linked to ROS

propagation driven by the rpL13a snoRNAs. Here, we demonstrate a link between O2-

induction and cytosolic rpL13a snoRNA accumulation. Our results extend the known

relationship between ROS and rpL13a snoRNAs, suggesting a role for these snoRNAs

in additional oxidative stress-related diseases.

RESULTS

Palmitate-induced superoxide precedes cytosolic snoRNA accumulation.

Following palmitate treatment, rpL13a snoRNAs accumulate in the cytosol in a time-

dependent fashion in H9c2 cells (Figure 4.1A). By 6 hr following palmitate treatment,

U32a, U33, and U34 are elevated ~2-fold in the cytosol, and U35a is elevated 1.5-fold.

Since rpL13a snoRNAs are critical for ROS propagation during lipotoxicity, we

hypothesized that rpL13a snoRNAs participate in a feed-forward ROS cascade initiated

by ROS production following palmitate treatment. O2- is produced during lipotoxicity

through NADPH oxidase (31). To test how quickly O2- is generated following palmitate

treatment, we quantified O2- production in cells treated with palmitate in the presence of

the O2- detector MCLA. Palmitate treatment rapidly raised O2

- levels, and levels

remained high compared to untreated cells over the course of 1 hr (Figure 4.1B) These

60

data indicate O2- induction is an early response to lipotoxicity and precedes cytosolic

snoRNA accumulation.

Direct superoxide induction causes rapid cytosolic snoRNA accumulation. To test

whether O2- generation can induce cytosolic snoRNA accumulation independent of

metabolic stress, we treated H9c2 cells with chemical inducers of O2-. Since O2

- can be

rapidly dismutated into H2O2, we initially examined the relative production of H2O2

following treatment with menadione and doxorubicin. We loaded cells with DCF to track

H2O2 generation and observed a dose dependent increase in DCF signal following

menadione treatment (Figure 4.2B). In comparison, doxorubicin generated less than

half the H2O2 of menadione even at high concentrations, suggesting doxorubicin is not a

potent inducer of H2O2.

We confirmed that doxorubicin generates O2- in H9c2 cells by treating cells with

doxorubicin for 1 hr followed by O2- detection by MCLA. We detected a dose dependent

increase in O2- production (Figure 4.3A). Since doxorubicin generates O2

- while only

producing modest levels of H2O2, we wanted to test whether doxorubicin could induce

cytosolic snoRNA accumulation. Following treatment with doxorubicin, we observed

rapid accumulation of rpL13a snoRNAs in the cytosol (Figure 4.3B). U32a, U33, and

U34 were up-regulated by at least 2.5 fold as early as 20 min following palmitate

treatment. Additionally, the magnitude of cytosolic snoRNA expression was higher than

observed following palmitate treatment in H9c2 cells or other cell types (72, 94). These

data suggest that direct O2- induction can rapidly and robustly induce cytosolic snoRNA

accumulation.

61

Superoxide levels correlate with cytosolic snoRNA accumulation. Conversion of

O2- to H2O2 is predominately regulated by superoxide dismutases (SOD). Isoforms of

these antioxidant enzymes are located throughout the cell and extracellular space.

SOD1 is located in the mitochondrial intermembrane space and cytosol, SOD2 is

located in the mitochondrial matrix, and SOD3 is tethered to the extracellular matrix

(95). We hypothesized that chemical manipulation of these enzymes would alter O2-

levels and subsequently alter levels of rpL13a snoRNAs in the cytosol. To increase

levels of O2-, we co-treated cells with doxorubicin and diethyldithiocarbamate (DETC),

an SOD inhibitor (Figure 4.4A) We also co-treated cells with doxorubicin and MnTBAP,

an SOD mimic, to increase O2- scavenging and subsequently decrease O2

- levels

(Figure 4.4A). RpL13a cytosolic snoRNA expression correlated with these changes in

O2- levels (Figure 4.4B). Relative to doxorubicin treatment alone, U32a, U33, and U34

all showed increased cytosolic accumulation following co-treatment with doxorubicin

and DETC and decreased cytosolic accumulation following co-treatment with

doxorubicin and MnTBAP. U35a was unaltered by doxorubicin treatment but was up-

regulated following co-treatment with doxorubicin and DETC. Although doxorubicin co-

treatment with MnTBAP lowered cytosolic snoRNA levels relative to doxorubicin

treatment alone, snoRNA levels remained higher in the presence of doxorubicin and

MnTBAP than in H2O-treated control cells, despite lower O2- levels in the MnTBAP

treated cells. These data suggest O2- contributes to cytosolic snoRNA localization, but

other factors likely contribute and may be important for localization of U35a in particular.

62

CONCLUSIONS

The rpL13a snoRNAs are critical for ROS propagation in response to metabolic

stress. Cytosolic localization of these snoRNAs is thought to be an important feature of

the function of these non-coding RNAs during lipid-induced oxidative stress. Here, we

demonstrate a link between ROS induction and cytosolic snoRNA accumulation.

Specifically, O2- induction precedes cytosolic snoRNA accumulation under lipotoxic

conditions. Doxorubicin induced O2- also causes rpL13a snoRNAs to localize to the

cytosol independent of metabolic stress. Manipulation of O2- demonstrates that O2

-

levels directly correlate with cytosolic snoRNA expression. Our data are consistent with

a model in which palmitate and direct O2- inducers stimulate higher levels of O2

- in the

cells. This O2- provides a signal to nuclear snoRNAs, through a pathway yet to be

elucidated, causing translocation of snoRNAs to the cytosol (Figure 4.5)

Additional experimentation will be required to determine the source of O2- under

lipotoxic conditions. We hypothesize that NADPH oxidase is that source of O2-.

Inhibition of NADPH oxidase or Nox enzyme knockdown will enable determination of the

role of NADPH oxidase in cytosolic snoRNA accumulation. Apocynin is an NADPH

oxidase inhibitor that prevents the assembly of NADPH oxidase subunits. Diphenylene

iodium (DPI) is a non-specific inhibitor of flavoenzymes including NADPH oxidase,

quinone oxidoreductase, cytochrome P450 reductase, and nitric oxide synthase (59).

Inhibition with either of apocynin or DPI could be used to test whether NADPH oxidase

is generally involved in cytosolic snoRNA localization. Additionally, directed knockdown

of Nox2, p22phox, or p47phox would help elucidate whether specific NADPH oxidase

components are related to snoRNA localization. Nox2 knockdown would directly target

63

a predominant source of O2-. Knockdown of p22phox would test whether any of the Nox

enzymes 1-4 are involved, and knockdown of p47phox would test whether a key cytosolic

component required for Nox1-3 activity is related to snoRNA expression.

Additionally, we propose that palmitate-induced oxidative damage is amplified by

cytosolic snoRNA accumulation (72). Following O2- reduction by MnTBAP or NADPH

oxidase inhibition/knockdown, we hypothesize that oxidative damage will be reduced

during lipotoxicity. To test this hypothesis, protein carbonylation and palmitate-induction

of cell death will be quantified in future studies in cells expressing various levels of O2-.

Finally, it will be of great interest to determine the specificity of relationship

between specific ROS species and the broader genomic program of snoRNAs. We

hypothesize that specific ROS regulate different snoRNAs. Preliminary data from our

lab suggests a number of additional snoRNAs are found in the cytosol during lipotoxicity

(not shown). Even though U32a, U33, U34, and U35a are produced from the same pre-

mRNA, our data suggest that the kinetics or the specificity of ROS-stimulated

relocalization to the cytosol differs among these snoRNAs. Since oxidative stress is the

result of the propagation of various ROS induced at different times, we propose that

individual snoRNAs respond specifically to different ROS. To test this hypothesis, we

will use a custom Agilent microarray developed in our lab that enables quantitative

detection of approximately 300 known and predicted snoRNAs to determine the

program of snoRNAs that accumulate in the cytosol following specific induction of O2-

vs. H2O2 vs. other ROS species. Changes identified in snoRNA expression will be

confirmed by qPCR and in situ hybridization.

64

Together, our data suggest an intricate connection between ROS and snoRNA

localization. Future studies will be needed to elucidate the mechanisms through which

O2- and other ROS signal to snoRNAs and will broaden our understanding of the

contributions of these non-coding RNAs to metabolic stress.

65

Figure 4.1

Figure 4.1 Palmitate induces superoxide. (A) H9c2 cardiomyoblast cells were treated

with BSA or treated with 500 µM palmitate for 3, 6, 12, or 24 h. Cytosolic fractions were

isolated from cells by sequential detergent solubilization. Total RNA was prepared from

the cytosolic fraction and analyzed for rpL13a snoRNA and mRNA abundance relative

to 36B4 by qRT-PCR. Graph shows mean ± SE (n = 3) from a representative

experiment. (B) H9c2 cells were untreated (UT) or treated with palmitate (palm) in 2-

methyl-6-(4-methoxyphenyl)-3,7-dihydroimidazol[1,2-A]pyrazin-3-one, hydrochloride

BSA 3 h 6 h 12 h 24 h0

2

4

68

18

RN

A:3

6B4

mR

NA

(rel u

nits

)

U32aU33U34U35arpL13a mRNA

palm

0 10 20 30 40 50 600

500

1000

1500

time (min)

MC

LA lu

min

esce

nce

(rel u

nits

)

UTpalm

A

B

66

(MCLA) buffer for 60 min. Luminescence was measured every 30 sec and normalized to

a superoxide dismutase-treated control.

67

Figure 4.2

Figure 4.2 Doxorubicin is not a potent H2O2 inducer. (A) Superoxide (O2-) is rapidly

dismutated by the antioxidant enzyme superoxide dismutase (SOD), leading to the

formation of hydrogen peroxide (H2O2) and molecular oxygen. (B,C) H9c2 cells pre-

loaded with CM-H2DCFDA (DCF) dye were untreated (UT) or treated with menadione

(mena) or doxorubicin (dox) for 60 min. H2O2 induction was assessed by DCF

fluorescence every 2 min using a Tecan microplate reader.

0 10 20 30 40 50 600

100020003000400050006000

time (min)

DC

F flu

ores

cenc

e UT5 uM mena100 uM mena

0 12 24 36 48 600

100020003000400050006000

time (min)

DC

F flu

ores

cenc

e UT5uM dox20uM dox

2 O2- + 2 H+ � H2O2 + O2

B

C

A SOD

68

Figure 4.3

Figure 4.3 Doxorubicin induces rapid cytosolic rpL13a snoRNA accumulation. (A)

H9c2 cells were untreated (UT) or treated with doxorubicin (dox) for 60 min. Superoxide

induction was assessed by MCLA. Luminescence was measured every 30 sec and

integrated luminescence over 3 min is reported normalized to an SOD treated control.

(B) H9c2 cells were untreated (UT) or treated with 20 µM doxorubicin for 20 min, 60

min, 12 h, or 24 h. Cytosolic fractions were isolated from cells by sequential detergent

solubilization. Total RNA was prepared from the cytosolic fraction and analyzed for

UT 5uM 20uM7000

8000

9000

10000M

CLA

lum

ines

cenc

e(A

UC

)

dox

UT 20' 60' 12h 24h0

10

20

30

RN

A:3

6B4

mR

NA

(rel

uni

ts)

dox

U32aU33U34U35arpL13a mRNA

A

B

69

rpL13a snoRNA and mRNA abundance relative to 36B4 by qRT-PCR. Graph shows

mean ± SE (n = 3) from a representative experiment.

70

Figure 4.4

Figure 4.4 Manipulation of superoxide alters cytosolic rpL13a snoRNA

accumulation. (A,B) H9c2 cells were untreated (UT) or treated with 5 µM doxorubicin

(dox) for 60 min. Cells were co-treated with H2O, 400 µM diethyldithiocarbamic acid

(DETC), or 400 µM manganese(III) tetrakis(4-benzoic acid)porphyrin chloride

(MnTBAP). (A) Superoxide induction was assessed by MCLA. Luminescence was

measured every 30 sec and integrated luminescence over 3 min is reported normalized

to an SOD treated control. (B) Cytosolic fractions were isolated from cells by sequential

detergent solubilization. Total RNA was prepared from the cytosolic fraction and

U32a U33 U34 U35a 0

3

6

9101520

RN

A:3

6B4

mR

NA

(rel

uni

ts)

H2Odoxdox + DETCdox + MnTBAP

A

B

16000

7000

8000

9000

10000

11000

MC

LA lu

min

esce

nce

(AU

C)

H2Odoxdox + DETCdox + MnTBAP

71

analyzed for rpL13a snoRNA and mRNA abundance relative to 36B4 by qRT-PCR.

Graph shows mean ± SE from a representative experiment.  

72

Figure 4.5

Figure 4.5 Superoxide is linked to cytoplasmic snoRNA accumulation. A model for

the role of superoxide in cytoplasmic snoRNA accumulation is shown. Treatment with

palmitate or doxorubicin causes increased levels of superoxide radicals. Through an

unknown mechanism, superoxide signals nuclear snoRNAs to accumulate in the

cytoplasm. SnoRNAs are shown in green, purple, yellow, and blue.

 

O2-

Palmitate

•

Direct O2-inducer

•

?

73

CHAPTER 5

Discussion

Genetic screens provide powerful approaches to dissect cell biological

phenomena. Inherent in this approach is the advantage that genes are identified on the

basis of their functional contributions to the pathway of study. Since multiple mutations

may affect different components in the pathway, genetic approaches also have the

ability to elucidate interacting elements. Our screen for mutations that render cells

resistant to lipotoxicity identified rpL13a snoRNAs as critical mediators of metabolic

stress. Identification of a second, independent mutation that disrupts the expression of

these snoRNAs highlights the importance of ncRNAs in lipotoxicity. With this second

mutant, we identified SmD3 as an upstream regulatory element necessary for the

expression of rpL13a snoRNAs. Thus, in an unbiased genetic screen, we identified

unexpected elements of the lipotoxic response.

Phenotypic characterization of the 6F2 and 6H2 mutants revealed that rpL13a

snoRNAs are also involved in the general response to oxidative stress. Previous

biochemical studies in diverse cell types have correlated lipotoxic stress with the

generation of ROS (7, 45). The rpL13a snoRNAs provide evidence of a functional link

between the progress of lipotoxic cell death and the deleterious cellular response to

oxidative stress. While transcriptional, posttranslational, and signaling mechanisms are

known to contribute to the cellular response to oxidative stress (33, 41), our studies are

the first to implicate snoRNAs in the response to environmental perturbations.

A key observation that facilitated our understanding of the contributions of the

rpL13a locus to lipotoxicity was the extent of sequence conservation of this gene

74

beyond the protein-coding exons. Four box C/D snoRNAs, U32a, U33, U34, and U35a

located in rpL13a introns 2, 4, 5, and 6, respectively, are highly conserved across

mammalian species both in terms of their primary sequence and their position within the

locus. While box C/D snoRNAs function as guides for 2’-O-methylation in yeast (Lowe

and Eddy, 1999), the sequences of the U32a, U33, U34, and U35a snoRNAs and their

genomic organization diverge substantially from yeast to mammals. In mammals, a role

in the modification of ribosomal RNAs has not been demonstrated for these snoRNAs.

We show here in a stable mutant cell line that decreased basal expression of U32a,

U33, and U35a snoRNAs and loss of palmitate induction of these snoRNAs is not

associated with changes in 2’-O-methylation of predicted rRNA targets, yet is

associated with resistance to lipotoxicity. While it is possible that these snoRNAs

contain more than one functional guide sequence to target multiple substrates including

rRNAs (13, 110), their accumulation in the cytosol during lipotoxicity suggests a non-

nucleolar function for these snoRNAs.

Movement of snoRNAs to the cytosol is not without precedent, since the

independently transcribed box C/D snoRNA, U8, has been shown to be exported from

the nucleus (121). The present study provides the first direct evidence for intronic

snoRNAs in the cytosol. Our data are most consistent with a model in which the rpL13a

snoRNAs function in lipotoxicity and oxidative stress response pathways as non-

canonical box C/D snoRNAs. Our results demonstrate that U32a, U33, and U35a

function coordinately in propagation of oxidative stress in vivo. Thus, our study provides

evidence for snoRNA regulation of metabolic stress response pathways.

75

Cytosolic functions of snoRNAs

The cytosolic function of snoRNAs has yet to be elucidated. We hypothesize that

snoRNAs serve to guide the modification of cytosolic target RNAs during metabolic

stress. Cytosolic ncRNAs, particularly tRNAs, are known targets for modification. tRNAs

are subject to over 75 different modifications throughout their sequence. Modifications

to the anticodon loop can influence the recognition of codons and maintenance of the

translational reading frame (32). Although pre-mRNA are known to undergo 5’ m7G

capping, 3’ polyadenylation, and RNA splicing, modification of individual nucleotides

within cytosolic mRNAs is not well described. The abundance of modifications in tRNA,

rRNA, and snRNA raises the possibility that mRNAs might also be modified.

SnoRNAs could function in the regulation of mRNA translation, possibly through

targeted modification or sequestration of specific mRNAs. Recently, it was

demonstrated that targeted pseudouridylation of termination codons suppresses

translation termination in vitro (48). Given the manifold of changes in gene expression

during the lipotoxic and oxidative stress responses (10), the coordinate function of the

rpL13a snoRNAs (72), lack of a well-defined long stretch of perfect antisense homology

to targets in snoRNAs (53), and the possibility that snoRNA-directed modifications of

RNA targets may not be reflected by changes in abundance of those RNAs,

identification of specific targets for each of these snoRNAs will be a complex task and

the subject of future investigations. To identify individual targets of rpL13a snoRNAs,

transfection of cells with synthetic biotin-labeled snoRNA molecules could facilitate pull-

down of RNA species that interact specifically with rpL13a snoRNAs. While initial

approaches may utilize biochemical isolation of specific snoRNA-targetRNA complexes,

76

it will also be of great interest to examine 2’-O-methylation and pseudouridylation on a

genome-wide level.

Identification of precise snoRNA localization within the cytosol will also be useful

for understanding the function of snoRNAs during metabolic stress. Moreover, our initial

studies suggest that snoRNAs are subject to cellular export. Treatment of cultured cells

with palmitate leads to detectable snoRNA expression in culture media. Additionally,

snoRNAs are stably expressed in human and mouse serum, and treatment of 12-week-

old mice with LPS results in increased serum snoRNA expression (not shown). Thus,

snoRNAs may be readily exported from cells. A number of recent studies have shown

that miRNAs are released from cells after cell death, in apoptotic bodies, through

microvesicles including exosomes, and as part of protein complexes giving precedence

for extracellular ncRNAs (106, 113, 114). If future studies confirm a role for snoRNA

secretion in response to lipotoxicity and oxidative stress, circulating snoRNAs could

ultimately serve as a biomarkers for diseases characterized by oxidative stress.

Regulation of snoRNA processing

Identification of SmD3 through our genetic screen unmasks a previously

unsuspected role for this protein in production of intronic snoRNAs. The precise

mechanisms through which SmD3 regulates intronic snoRNA expression remain to be

determined. One possibility is that SmD3 has a direct role in snoRNA biogenesis

unrelated to its function in snRNPs. Unlike SmD3 haploinsufficient cells, SmB

knockdown cells remain sensitive to palmitate-induced death suggesting that SmD3 has

a function distinct from other Sm protein family members. Additionally, SmD3 has been

77

observed to function independently from the Sm core in the cytoplasm (30). SmD3 has

also been previously shown to bind small Cajal body RNAs (scaRNAs), a class of

snoRNAs specifically localizing to Cajal bodies, and human telomerase (hTR) via a CAB

box site (28). However, H/ACA snoRNAs U17b and U64, which are affected by

haploinsufficiency of SmD3, both lack CAB box sequences. Additional studies will be

required to decipher whether SmD3 can function independently from the Sm core during

snoRNA biogenesis.

Our data are most consistent with a model in which SmD3 does not directly

interact with the maturing snoRNA during processing, but rather controls processing

indirectly through a role in snRNP assembly and/or stabilization (Figure 3.10).

Knockdown of SmD3 results in decreased expression of U4 and U5 snRNPs that are

required for precatalytic spliceosome formation and initiation of an active spliceosome

capable of intron lariat formation (68). Furthermore, reduced expression of snoRNA

lariat precursors, resulting from decreased lariat production or stability, leads to

decreased abundance of the mature intronic snoRNAs and failure to support their

induction during lipotoxicity in 6H2 cells. These differences in intron lariat abundance

and snRNP expression suggest that the phenotype observed in 6H2 cells is related to

spliceosomal machinery, even though haploinsufficiency of SmD3 is sufficient to

maintain production of mRNAs.

We also observed that wild type levels of SmD3 are critical for the ability to

express a number of intronic non-coding RNA elements including other box C/D

snoRNAs, H/ACA snoRNAs, and mirtrons. These ncRNAs do not share significant

sequence similarities and each assembles with a unique set of proteins during

78

processing, making it unlikely that SmD3 directly recognizes and associates with the

nascent RNP. It has been shown that the C-terminal tail of SmD3 interacts with the

central Tudor domain of splicing factor SPF30 (62), indicating that SmD3 may be

important for the assembly of splicing factors specific to individual snRNPs. Individual

Sm proteins may be critical for the assembly of specific factors within different snRNPs

consistent with the different snRNA expression profiles observed following SmD3

knockdown versus SmB knockdown. In follow up, it will be of interest to determine

whether Sm protein-splicing factor interactions are critical for SmD3’s role in snRNP

assembly and/or stabilization and to better define the distinct but overlapping roles of

SmD3 and SmB in assembly and stabilization of snRNPs.

Regulation of snoRNA localization

SnoRNAs are processed in the nucleus and canonically reside within the

nucleolus. Our studies indicate that nuclear snoRNA levels are multiple orders of

magnitude higher in the nucleus than the cytoplasm even during metabolic stress.

Unlike snRNAs, snoRNAs are not thought to undergo a cytosolic phase during

maturation (69), suggesting that accumulation of snoRNAs in the cytosol requires an

active mechanism. We demonstrate that O2- induction precedes cytosolic snoRNA

accumulation consistent with the hypothesis that O2- functions as an upstream trigger of

snoRNA translocation from the nucleus to the cytosol. O2- participates in multiple

signaling pathways (34). ROS driven signal transduction includes induction of gene

expression, protein phosphorylation, and alteration of redox status (73). Examination of

79

O2- signaling pathways, particularly in the nucleus, will be of great interest in future

studies of ROS regulated snoRNA localization.

O2- diffuses only short distances before reacting with a target, so it is likely that

the source of O2- related to snoRNAs is generated in close proximity to its intended

target. Although NADPH oxidases are implicated to be involved in lipotoxicity, the

source of O2- leading to cytosolic snoRNAs remains to be elucidated. Nox enzymes

each have distinct subcellular localizations. Nox2 is found predominantly on the plasma

membrane, whereas Nox4 is located in intracellular membranes. O2- can also be

generated from several additional sources, including xanthine oxidases, peroxidases,

lipoxygenases, cyclooxygenases, and mitochondrial respiratory chain complexes (129).

Determination of the source of O2- will help elucidate how O2

- is linked to cytosolic

snoRNA accumulation.

 Summary

Together, our data demonstrate that snoRNAs are highly regulated in response

to metabolic stress and contribute significantly to the pathogenesis of oxidative stress.

To carry out this role, snoRNAs must be efficiently processed and properly localized.

Elucidation of the regulatory steps in snoRNA production and localization and

identification of targets for non-canonical snoRNAs will advance our understanding of

the pathogenesis of common human diseases. Moreover, regulation of snoRNAs

provides potential therapeutic targets for oxidative stress-related diseases including

type 2 diabetes.

 

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CHAPTER 6

Materials and Methods

Materials. Palmitate was from Nu-Chek Prep. 14C-palmitate and α-32P-UTP were from

PerkinElmer Life Sciences. Camptothecin, actinomycin D, hygromycin, and MnTBAP

were from Calbiochem. Staurosporine, H2O2, menadione, phloretin, clotrimazole,

doxorubicin, and DETC were from Sigma-Aldrich. Fatty acid-free bovine serum albumin

(BSA) was from SeraCare. Propidium iodide, CM-H2DCFDA, DHE, and MCLA were

from Invitrogen. All synthetic oligonucleotides were from IDT (see Table 1 for primer

sequences). Restriction enzymes were from New England BioLabs.

Cell Culture. CHO-K1 cells (American Type Culture Collection) and CHO-derived cell

lines were maintained in high glucose (4.5 mg/ml Dulbecco’s modified Eagle’s medium

and Ham’s F-12 nutrient mixture (1:1)) media with 5% non-inactivated fetal bovine

serum, 2 mM L-glutamine, 50 units/ml penicillin G sodium, 50 units/ml streptomycin

sulfate, and 1 mM sodium pyruvate. C2C12 myoblasts (American Type Culture

Collection) were maintained in high glucose (4.5 mg/ml Dulbecco’s modified Eagle’s)

medium with 10% heat-inactivated fetal bovine serum, 50 units/ml penicillin G sodium,

and 50 units/ml streptomycin sulfate. NIH 3T3 cells (American Type Culture Collection)

were maintained in high glucose (4.5 mg/ml Dulbecco’s modified Eagle’s medium)

media with 10% bovine calf serum, 2 mM L-glutamine, 50 units/ml penicillin G sodium,

and 50 units/ml streptomycin sulfate. H9c2 cardiomyoblast cells (American Type Culture

Collection) were maintained in high glucose (4.5 mg/ml Dulbecco’s modified Eagle’s

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medium) media with 10% fetal bovine serum, 2 mM L-glutamine, 50 units/ml penicillin G

sodium, and 50 units/ml streptomycin sulfate. For lipotoxicity experiments, cell culture

media was supplemented with 500 µM (WT CHO and 6H2) or 250 µM (shRNA

transfected cells) palmitate complexed to BSA at a 2:1 M ratio, as described previously

(61). For ROS induction, media was supplemented with the indicated concentrations of

H2O2, menadione, or doxorubicin.

Generation of CHO Cell Mutants. Vesicular stomatitis virus G protein pseudotyped

murine retrovirus encoding the ROSAβgeo retroviral promoter trap was generated as

described previously (26, 81). CHO cells were transduced with retrovirus at a low

multiplicity of infection (1 integration per 10 genomes on average) and mutants were

isolated as described previously (7). Number of retroviral insertions within the mutant

cell genome was assessed by Southern blotting. Genomic DNA was digested with

restriction enzymes, separated by agarose gel electrophoresis, transferred to nylon

membranes, and probed with a 32P-labeled probe corresponding to the ROSAβgeo

proviral sequence.

Cell Death Assays. Cell death was assessed by membrane permeability to propidium

iodide (PI) staining and flow cytometry (61). Following treatments, cells were harvested

by trypsinization and stained with 1 µM PI. Analyses were performed on 104

cells/sample.

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14C Palmitate Uptake Assay. 2 × 106 cells were resuspended in 1 ml phosphate-

buffered saline (PBS) containing 500 µM 14C-palmitate complexed to 250 µM BSA and

incubated for one minute at 37°C. Cells were washed with 10 ml PBS containing 0.1%

BSA and 500 µM phloretin, filtered, and cell-associated 14C was quantified by

scintillation counting. A parallel aliquot of cells was used for quantification of protein by

bicinchoninic acid assay (Pierce).

Identification of Trapped Gene. The endogenous gene disrupted by retroviral insertion

was identified by 5’ rapid amplification of cDNA ends (RACE) using an oligonucleotide

tag and ROSAβgeo sequences (SMART RACE cDNA amplification kit; Clontech). The

5’ RACE product was TA cloned and sequenced, and tested for sequence similarity by

NCBI BLAST. PCR was used to verify retroviral integration within the snrpD3 gene.

Quantitative Real Time PCR (qPCR). RNA was isolated using TRIzol or TRIzol LS

reagent (Invitrogen) and reverse transcribed to cDNA using the SuperScript III First-

strand Synthesis System for RT-PCR (Invitrogen) following the manufacturer’s

instructions. Reverse transcription was performed with oligo(dT) to detect mRNA or

random hexamers to detect pre-mRNA and intron lariats. For quantification of snoRNAs

and pre-miRNAs, RNA was isolated using Trizol LS (Invitrogen). cDNA synthesis was

primed with hairpin stem-loop oligos as previously described (72), with overhang

complementarity to the 3’ end of the processed snoRNA or pre-miRNA. cDNA was

amplified for 40 PCR cycles using SYBR Green PCR master mixture (Applied

Biosystems) and 100 nM template specific primers in a ABI Prism 7500 Fast Real-Time

83

PCR System. Relative quantification of gene expression was performed using the

comparative threshold method as described by the manufacturer.

Flow Cytometry Detection of Reactive Oxygen Species. Cells (2 × 105) were plated

in 12-well plates 24 h prior to various treatments. Cells were rinsed with PBS and

incubated with PBS containing 0.5 mM MgCl2, 0.92 mM CaCl2, and 1µM (C2C12 cells)

or 3 µM (CHO cells) 5-(and-6)-chloromethyl-2’,7’-dichlorodihydrofluorescein diacetate,

acetyl ester (CM-H2DCFDA, Invitrogen) in the dark at 37°C for 1 h. Cells were then

rinsed with PBS, harvested by trypsinization, and quenched with culture media. Mean

fluorescence was determined by flow cytometry on 104 cells/sample.

Generation of SmD3 antibody. For immunoblot detection of SmD3, polyclonal rabbit

anti-peptide antibody was generated from ProSci Incorporated. Animals were

immunized with the unique peptide NH2-CTGEVYRGKLIEAED-OH (murine sequence)

conjugated to KLH. Animals received six rounds of immunization followed by affinity

purification of serum.

Immunoblot Analyses. Whole cell protein lysates were prepared using RIPA buffer (50

mM Tris-Cl, 150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS,

and 5 mM EDTA) containing 1 mM phenylmethylsulfonyl fluoride and 1 × Protease

Complete inhibitor mixture (Roche). Subcellular fractions were isolated by sequential

detergent solubilization as described previously (40). Proteins (40 µg) were resolved by

15% (for rpL13a, SmD3, and SmB) or 12% (for hsp90, fibrillarin, lamin B1, CHOP-10,

84

and β-actin) SDS-PAGE gel electrophoresis and transferred to nitrocellulose membrane

(Whatman). Membranes were probed with antibodies to rpL13a (1:500), SmD3 (1:500),

β-actin (A 2066, Sigma, 1:5000), hsp90 (SPA-846, Stressgen, 1:2000), fibrillarin (MMS-

581S, Covance, 1:500), lamin B1 (ab16048, Abcam, 1:1000), CHOP-10 (F-168, Santa

Cruz, 1:500), and SmB (S0698, Sigma, 1:1000). Proteins were visualized using

appropriate horseradish peroxidase-conjugated secondary antibodies (Jackson

ImmunoResearch Laboratories, 1:10,000) and chemiluminescence reagents

(PerkinElmer Life Sciences). Band intensities were quantified by densitometry (Bio-Rad

Image Lab Software).

snoRNA Probe Synthesis and RNase Digestion. Hamster- and mouse-specific

snoRNA probes were generated with Megashortscript kit (Ambion). dsRNA templates

were generated for probe for each rpL13a snoRNA by PCR amplification of cloned

hamster or mouse rpL13a genomic sequence templates using primers containing the T7

RNA polymerase promoter and used for in vitro RNA transcription of 32P-labeled

snoRNA probes. miR-16 probes were synthesized using templates from mirVana

miRNA Detection kit (Ambion). RNA was isolated from cells using mirVana miRNA

isolation kit (Ambion) and hybridized to 32P-labeled RNA probes (mirVana miRNA

Detection kit) overnight at 42-52°C, followed by RNase digestion and ethanol

precipitation. RNA was separated by 10% or 15% polyacrylamide electrophoresis and

visualized by autoradiography.

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snoRNA Knockdown in vitro. Anti-sense oligos (ASOs) were designed to specifically

target murine U32a, U33, U35a, U50, U57, and U60 snoRNA sequences according to

Ideue et al. (44). For snoRNA “knock-down” experiments, 106 C2C12 myoblasts were

nucleofected using Nucleofector Kit V (Amaxa) and a total of 600 pmol of ASO.

Mapping of 2’-O-methyl modification by primer extension. Protocol was based on

methods from Lowe and Eddy (64). Total RNA from Trizol extraction was annealed with

32P end-labeled primers at 55°C for 4 min. Primer extension reactions were carried out

in the presence of 50 mM Tris-Cl (pH 8.6), 60 mM NaCl, 9 mM MgCl2, 10 mM DTT, 1

mM dNTP, and using avian myeloblastosis virus reverse transcriptase for 30 min at

37°C. For rRNA sequencing, ddNTPs were used in individual reactions. For 2’-O’methyl

mapping, reactions were carried in “High” (4mM), or “Low” (0.004 mM) dNTP

concentrations, and 5 mM MgCl2. Reaction products were separated by 6%

polyacrylamide electrophoresis (PAGE) and visualized by autoradiography.

In situ hybridization of snoRNA probes. Synthesis and labeling of sense and

antisense RNA probes were adapted from Darzacq et al. (16). C2C12 myoblasts were

fixed in PBS containing 3% paraformaldehyde for 10 min, rinsed with PBS and

permeabilized with 0.1% Triton X-100 for 10 min at room temperature. Cells were

dehydrated serially in 70% ethanol, 95% ethanol, 100% ethanol and then air-dried.

Sequence-specific oligonucleotide probes containing aminoallyl UTP were generated

using the FISH Tag RNA Kit (Invitrogen) and labeled with an amine-reactive Alexa Fluor

594 dye. Fluorescent probe (0.5 ng/µL) was denatured in hybridization buffer (2x SSC,

86

50% formamide, 10% dextran sulfate, 20 mM vanadyl ribonucleoside complexes) for 10

min at 80°C. Cells were incubated with probe/hybridization buffer at 37°C for 10 hr,

followed by sequential washes with 2x SSC, 1x SSC, 0.1x SSC all containing 1% SDS

for 15 min each at 37°C. Nuclei were counter-stained using SYTOX Green (Invitrogen).

Slides were mounted with SlowFade antifade reagent (Invitrogen). Images were

captured on a ZEISS LSM 510 META confocal laser scanning microscope using

constant pinhole size, detector gain, and offset for each probe.

Mouse model of LPS-mediated oxidative stress and in vivo snoRNA knockdown.

Female FVB mice were obtained from Charles River Laboratories and housed in

Washington University Division of Comparative Medicine facilites. Diet was standard 6%

fat breeding chow supplied ad libitum, and food was withheld at the time of LPS

injection. Between 10 and 16 weeks of age, LPS was administered at 8 mg/kg

intraperitoneally (IP), and animals were euthanized 12-24 h later. For in vivo knockdown

experiments, LNA-modified ASOs were purchased from Exiqon and used to specifically

target snoRNAs U32a, U33, and U35a. An ASO targeting GFP was used as a control.

Mice were injected IP with a total of 2.5 mg/kg of LNA every other day for a total of three

injections, and then dosed with LPS as above 48 h after the last LNA injection.

Individual snoRNA ASO concentrations were 1.25, 0.5, and 0.75 mg/kg per dose,

targeting U32a, U33, and U35a respectively (empirically determined based on

knockdown). Liver tissue was divided and either snap-frozen in liquid nitrogen (for

RNA), fixed in 10% neutral buffered formalin, or frozen in O.C.T. Compound (Tissue-

Tek). Experimental procedures were approved by the Washington University Animal

87

Studies Committee and were conducted in accordance with USDA Animal Welfare Act

and the Public Health Service Policy for the Humane Care and Use of Laboratory

Animals.

In vivo detection of ROS. Formalin-fixed, paraffin-embedded samples or liver frozen in

O.C.T. were mounted on slides in serial sections by the Washington University

Anatomic and Molecular Pathology Core Lab. Detection of superoxide was performed

on frozen sections using dihydroethidium (DHE; Invitrogen, Cat# D11347). Sections

were incubated with 2 µM DHE for 30 min at 37°C or pre-treated with 200 Units/ml

PEG-SOD (Sigma), followed by co-incubation of 2 µM DHE and 200 Units/ml PEG-SOD

for 30 min at 37°C to verify the specificity of staining as indicated. For each animal (n =

4 for GFP ASO; n = 5 for SNO ASO), intensity of staining was quantified in three

independent fields from each of six sections using ImageJ software. Protein carbonyls

were detected by immunoblot using the OxyBlot Protein Oxidation Detection Kit

(Chemicon) according to the manufacturer’s instructions. Blots were quantified by

densitometry using actin as a loading control. Tissue oxysterols (7-ketocholesterol, 7-

keto; 3β,5α,6β-cholestantriol, triol) per mg liver protein were quantified using LC/MS/MS

as described (87).

Generation of SmD3 shRNA Clones. Hamster snrpD3 cDNA sequence was used to

design siRNA oligonucleotides using Ambion’s siRNA Target Finder Program

(ambion.com/techlib/misc/siRNA_finder.html). shRNA oligonucleotides were designed

from siRNA sequences that conferred effective knockdown in transient transfection

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assays, and each was cloned into a pSilencer 4.1-CMV hygro vector (Ambion)

containing a hygromycin resistance cassette. shRNA vectors were transfected into CHO

cells with Lipofectamine 2000 reagent (Invitrogen). Cells were plated at limiting dilution

and treated with 80 µg/ml hygromycin. Clonal lines were isolated, and SmD3

knockdown assessed by immunoblot.

Luciferase Plasmids and Transient Transfection. The split luciferase vector

containing a β-globin intron was as described previously (128). All constructs generated

by PCR or Quick-Change (Stratagene) mutagenesis were confirmed by sequencing.

The β-globin intron in the split luciferase reporter was replaced with rpL13a intron 2

containing snoRNA U32a or the rpL13a intron 2 lacking the 83 nucleotide U32a

snoRNA sequence. Cells were transfected with Lipofectamine 2000 (Invitrogen) as per

the manufacturer’s protocol and assayed 20 h post-transfection.

Luciferase Detection. Cells (3 × 104) were plated in triplicate in 96-well plates.

Following transfection of luciferase reporters as described above, cells were lysed with

Dual-Glo Luciferase Reagent (Promega) according to the manufacturer’s protocol.

Luciferase was detected using a Tecan Infinite M200 microplate reader and Magellan

software.

Microarray Sample Preparation and Data Analysis. NIH 3T3 cells (2 × 105) were

plated in 6-well plates 24 h pre-transfection. Cells were transfected with 40 pmol

LNA/DNA oligonucleotides specifically targeting GFP or SmD3 (Exiqon) using

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lipofectamine 2000 (Invitrogen) as per the manufacturer’s protocol, and total RNA was

harvested 23 h post-transfection using TRIzol reagent (Invitrogen). Resulting RNA was

quantified by A260 and A280 readings using a Nanodrop spectrophotometer (Nanodrop

Technologies) and qualitatively assessed using a BioAnalyzer 2100 (Agilent

Technolgies). cDNA was prepared using the NuGen Ovation System and microarray

data was generated with Affymetrix GeneChip Mouse Exon 1.0 ST arrays containing

266,200 probesets in the Siteman Cancer Center Molecular and Genomic Analysis

Core at Washington University. Partek Genomics Suite 6.5 was used to calculate

probeset intensities from .CEL files using the RMA algorithm with default settings at

both the gene level and probeset level. Probesets with RMA intensity below 3 across all

samples were excluded to eliminate probesets with low expression levels. Alternative

splicing multiway ANOVA was applied using Partek defaults to identify alternative

splicing events with a false discovery rate (FDR) < 0.1. Core exon level analysis was

also applied at the exon level to determine differential expression of exons not grouped

by transcript.

Transient Knockdown. For SmD3 knockdown, NIH 3T3 cells (2 × 105) were plated in

6-well plates 24 h pre-transfection. Cells were transfected with 40 pmol LNA/DNA

oligonucleotides specifically targeting GFP or SmD3 (Exiqon) using lipofectamine 2000

(Invitrogen) as per the manufacturer’s protocol, and RNA was harvested 23 h post-

transfection. For SmB knockdown, NIH 3T3 cells (1 × 105) were plated in 6-well plates

24 h pre-transfection. Cells were transfected with 50 pmol control (Ambion) or snrpb

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(s74100, Ambion) Silencer Select siRNA using lipofectamine RNAiMAX (Invitrogen) as

per the manufacturer’s protocol, and RNA was harvested 24 h post-transfection.

Immunoprecipitation. For snRNA immunoprecipitation, NIH 3T3 cells were plated and

transfected as performed for SmD3 transient knockdown. 23 h post-transfection, cells

were harvested in lysis buffer (50 mM Tris-Cl pH 8, 150 mM NaCl, 0.5% Nonidet P-40)

containing 1 mM phenylmethylsulfonyl fluoride and 1 × Protease Complete inhibitor

mixture (Roche) and incubated on ice for 30 min. Cells were sonicated with five 5-

second pulses using a Branson Sonifier 250, incubating on ice for 20 sec between each

pulse. Cell lysates were centrifuged at 15,000 × g for 30 min at 4°C to remove insoluble

material. Cell lysates were immunoprecipitated using α-Sm Y12 antibody (ab3138,

Abcam, 1:100) or IgG control. RNA was isolated from immunoprecipitated samples

using TRIzol (Invitrogen) and reverse transcribed to cDNA using the SuperScript III

First-strand Synthesis System for RT-PCR (Invitrogen) following the manufacturer’s

instructions. Quantification of α-Sm immunoprecipitated snRNA expression relative to

control IgG precipitated snRNA was performed by qRT-PCR.

Superoxide Detection in vitro. Cells (8 × 104) were plated in 96-well opaque white

microplates 24 h prior to treatment. Cells were incubated for 1 hr in reaction buffer

containing 125 mM KCl, 10 mM HEPES, 5 mM MgCl2, 2 mM K2HPO4, and 5 µM 2-

methyl-6-(4-methoxyphenyl)-3,7-dihydroimidazol[1,2-A]pyrazin-3-one, hydrochloride

(MCLA) during continuous treatment (palmitate) or rinsed with PBS and incubated in

reaction buffer immediately following treatment (doxorubicin). MCLA light emission was

91

quantified using a Tecan Infinite M200 microplate reader and iControl software. The

photomultiplier was set with an integration time of 1000 ms. The MCLA signal was

quantified in real-time or as an integral of the signal measured every 30 sec over 3 min.

Real-time Reactive Oxygen Species Detection in vitro. Cells (5 × 104) were plated in

12-well plates 24 h prior to CM-H2DCFDA loading. Cells were rinsed with PBS and

incubated with PBS containing 0.5 mM MgCl2, 0.92 mM CaCl2, and 10µM CM-

H2DCFDA in the dark at 37°C for 10 min. Cells were then rinsed with PBS prior to

various treatments in PBS containing 0.5 mM MgCl2 and 0.92 mM CaCl2. Mean

fluorescence was quantified every 3 min with excitation/emission wavelengths of

492/517 nm using a Tecan Infinite M200 microplate reader and iControl software.

 

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 Table 5.1: Primer/oligo sequences application primer/oligo primer/oligo sequence qRT-PCR snrpD3 forward 5'-CGGTGTGCCGATTAAAGTCT-3' snrpD3 reverse 5'-TAACATGGGTGCGTTTTTCA-3' snrpb forward 5'-TCTTCATCGGGACCTTCAAAGCCT-3' snrpb reverse 5'-ACTCGCTTCTCTTCCCTTTCTGCT-3' β-actin forward 5'-GGCTCCCAGCACCATGAA-3' β-actin reverse 5'-GCCACCGATCCACACAGAGT-3' luciferase exon

1/intron forward 5'-CAGGTAAGTATCAAGGTTCCCG-3'

luciferase intron reverse

5'-CTCTTGGAAGGCAGATCTCTTG-3'

Snhg5 forward 5'-CAGTGAGTGAAATGCCAGCT-3' Snhg5 reverse 5'-GACAGGTGCGTTTGAAGACA-3' Nop56 forward 5'-TGAGGAACGGCTGTCTTTCT-3' Nop56 reverse 5'-TCTTCAAGCGCTTCTTCTCC-3' U60hg forward 5'-CCCTTTTTGCAGGTAAGGTG-3' U60hg reverse 5'-AAAATCCTGTTGCCTGGATG-3' RPL5 forward 5'-CCCGAACTACAACTGGCAAT-3' RPL5 reverse 5'-CCGATGAACTTCTGCATTGA-3' Snhg3 forward 5'-AAGAAGTCGCATCGGTACTGC-3' Snhg3 reverse 5'-GGTGCAAGTCCCCAAAGTTA-3' RPS2 forward 5'-GAAAATCATGCCAGTGCAGA-3' RPS2 reverse 5'-CAAGACCAACGTGACCATTG-3' EIF5 forward 5'-AGGCATGCTTGACACACATC-3' EIF5 reverse 5'-CAGAGCCATTTTCCTTGTCC-3' VWA5B2 forward 5'-GGCAGCAACCATGACTACCT-3' VWA5B2 reverse 5'-TCTTGAGGAATGCACACAGC-3' PKD1 forward 5'-GCTTCAGCAAGGTCAAGGAG-3' PKD1 reverse 5'-TGGATCCATTCCTTCAAAGC-3' 36B4 forward (rat) 5'-CAGAGGTGCTGGACATCACAGA-3' 36B4 reverse (rat) 5'-AGTGAGGCACTGAGGCAACAG-3' rpL13a qRT-PCR

rpL13a exon 2 forward

5'-CATTGTGGCAAGCAAGTGCTACTG-3'

rpL13a exon 3 reverse

5'-AATGTTGATGCCTTCACAGCGCAC-3'

rpL13a exon 3 forward

5'-CGGAAGGTAGTCGTTGTGC-3'

rpL13a intron 3 reverse

5'-TGCTTGTCCAGAACTTAGGTC-3'

rpL13a exon 4 forward

5'-TTTCCTCCGAAAGCGGATGAACAC-3'

rpL13a exon 5 reverse

5'-GGATCCCATCCAACACCTTGAGG-3'

rpL13a exon 5 forward

5'-CCTCAAGGTGTTGGATGGGATCC-3'

rpL13a exon 6 reverse

5'-GCTTCAGCCGCACAACCTTG-3'

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rpL13a exon 6 forward

5'-CAAGGTTGTGCGGCTGAAGC-3'

rpL13a exon 7 reverse

5'-GGCCTTTTCCTTGCGTTTCTCC-3'

rpL13a exon 7 forward

5'-TGGGGTCGGGTGGAAATACC-3'

rpL13a exon 8 reverse

5'-CTTTTCTGCCTGTTTGCGTAG-3'

rpL13a forward (rat)

5'-GGCTGAAGCCTACCAGAAAG-3'

rpL13a reverse (rat)

5'-CTTTGCCTTTTCCTTCCGTT-3'

snoRNA/miRNA/tRNA reverse transcription

U32a SLRT (hamster)

5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCCAAGTCTC-3'

U32a SLRT (mouse) 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCCGAGTCTC-3'

U33 SLRT (hamster)

5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCCCAGCCTC-3'

U33 SLRT (mouse) 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCACAGCCTC-3'

U35a SLRT (hamster)

5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCTACTGGCA-3'

U35a SLRT (mouse) 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCTCCTGGCA-3'

U50 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCGTCTCA-3'

U57 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCTGGATCAG-3'

U60 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCCCAAGC-3'

U21 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCGCCGCC-3'

U17b SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCTTTTGT-3'

U64 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCACCTGTGG-3'

ACA28 SLRT 5’-GTGTGGTCCCGACCACCACAGCCGCCACGACCACGCGTTTGTTA-3'

pre-miR-1224 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCCTGAGG-3'

pre-miR-1225 SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCCTGGGG-3'

pre-miR-23a SLRT 5'-GCGTGGTCCCGACCACCACAGCCGCCACGACCACGCGGTCAG-3'

Glut RNA SLRT2 5'-CTCAGCGGCTGTCGTGGACTGGGTGCTGCCGCTGAGTGGTTCCCTGA-3'

snoRNA/miRNA/tRNA PCR

U32a forward (hamster)

5'-AAGTCAGTGATGAGCAACAATCACCATC-3'

U32a forward (mouse)

5'-GAGTCCATGATGAGCAACACTCACC-3'

U33 forward 5'-CAGCCTATGATGAAGCGATCTCCCAC-3'

94

(hamster) U33 forward

(mouse) 5'-AGCTTGTGATGAGACATCTCCCACT-3'

U35a forward (hamster)

5'-GGCACATGATGTTCTTATTCTCACGATGGT-3'

U35a forward (mouse)

5'-GGCACATGATGTTCTTATTCTCACGATGGT-3'

U50 forward 5'-TCTATGATGATCCTATCCCGAAC-3' U57 forward 5'-GATGAACGAACTTGGCCTGACCTTC-3' U60 forward 5'-CCAAGCCTGTGATGAATTGC-3' U21 forward 5'-GTAGTTGGTCCTTTGATTGCATGTGATGT-3' U8 forward 5'-CCTTACCTGTTCCTCCTTTCG-3' U8 reverse 5'-GAGCAACCAGGATGTTGTCA-3' U13 forward 5'-TTTCTGGTTCATAAAGCGTGA-3' U13 reverse 5'-TGTCAGACGGGTAATGTGC-3' U17b forward 5’-AACGGGAGCTTAGGGCATT-3’ U64 forward 5'-CTTTGGATTGGCCTCACAGT-3' ACA28 forward 5'-AAGCAACACTCTGTGGCAGATGAA-3' pre-miR-1224

forward 5'-GCGCTGTTTCAGCTCGCTTCTC-3'

pre-miR-1225 forward

5'-GTGCAGCCGGACTGACTGA-3'

pre-miR-23a forward

5'-GGTTCCTGGGGATGGGATTTGATC-3'

Glu tRNA forward 5'-TAGTGGTTAGGATTCGGCGCTCTCAC-3' Universal reverse

primer 5'-TCCCGACCACCACAGCC-3'

Universal reverse primer 2

5'-GGCTGTCGTGGACTGGGTG-3'

intron lariat PCR

CHO rpL13a intron 2 sense

5'-GCTCTGAGACTTGACGGTCC-3'

CHO rpL13a intron 2 antisense

5'-CTCATCACTGACTTGCAGGC-3'

CHO rpL13a intron 4 sense

5'-ATCTCCCACTGGTGTTCGAG-3'

CHO rpL13a intron 4 antisense

5'-CAGATTCCCCATTCTCCATG-3'

CHO rpL13a intron 5 sense

5'-TCCTTCCCAGGTGATGCT-3'

CHO rpL13a intron 5 antisense

5'-TGGAAAGCTAGGCTTGCG-3'

CHO rpL13a intron 6 sense

5'-ATAATGCCACAGGCTCAGCT-3'

CHO rpL13a intron 6 antisense

5'-TCTGCCAGTCTACAGGTCCC-3'

mouse rpL13a intron 2 sense

5'-TGTGAGATCAACCCATGCAC-3'

mouse rpL13a intron 2 antisense

5'-CGAAAGATGGTGAGTGTTGC-3'

snRNA PCR U1 forward 5'-GATACCATGATCACGAAGGTGGTT-3' U1 reverse 5'-CACAAATTATGCAGTCGAGTTTCC-3'

95

U2 forward 5'-TTTGGCTAAGATCAAGTGTAGTATCTGTTC-3'

U2 reverse 5'-AATCCATTTAATATATTGTCCTCGGATAGA-3'

U4 forward 5'-GCGCGATTATTGCTAATTGAAA-3' U4 reverse 5'-AAAAATTGCCAATGCCGACTA-3' U5 forward 5'-TACTCTGGTTTCTCTTCAGATCGTATAAAT-3' U5 reverse 5'-AATTGGTTTAAGACTCAGAGTTGTTCCT-3' U6 forward 5'-GCTTCGGCAGCACATATACTAAAAT-3' U6 reverse 5'-ACGAATTTGCGTGTCATCCTT-3' U11 forward 5'-GTGCGGAATCGACATCAAGAG-3' U11 reverse 5'-CGCCGGGACCAACGAT-3' U12 forward 5'-AACTTATGAGTAAGGAAAATAACGATTCG-3' U12 reverse 5'-CCGCTCAAAAATTCGTCTCACA-3' U4atac forward 5'-AGCGCATAGTGAGGGCAGTACTG-3' U4atac reverse 5'-GCACCAAGGTAAAGCAAAAGCTCTA-3' U6atac forward 5'-AGGTTAGCACTCCCCTTGACAA-3' U6atac reverse 5'-TGGCAATGCCTTAACCGTATG-3' 5’ RACE ROSAβgeo reverse 5’-CAAGGCGATTAAGTTGGGTAACGC-3’ PCR snrpD3 forward 5’-GCCGTGTAACGTTTGTGCC-3’ snrpD3 reverse 5’-GGACACAGTTCTCTTCCAC-3’ ROSAβgeo reverse 5’-CTCAGGTCAAATTCAGACGG-3’ siRNA snrpD3 sense 5’-GCUCAUUGAAGCAGAGGACUU-3’ snrpD3 antisense 5’-GUCCUCUGCUUCAAUGAGCUU-3’ scrambled control

sense 5’-AAGAUGAGCAUAGGAUGUU-3’

scrambled control antisense

5’-AACAUCCUAUGCUCA-3’

LNA/DNA oligos GFP 5'-+T*+C*+A*+C*+C*T*T*C*A*C*C*C*T*C*T*+C*+C*+A*+C*+T-3'

U32a 5 ' +G*+C*+G*+G*+T*G*C*A*T*G*G*G*T*T*G*+A*+T*+C*+T*+C 3'

U33 5' +T*+G*+G*+T*+A*G*T*G*C*A*T*G*T*A*G*+A*+G*+T*+C*+A 3'

U35a 5' +T*+T*+A*+G*+C*C*T*T*T*G*G*C*A*T*T*+A*+T*+C*+G*+G 3'

snrpD3 5'-+T*+C*+T*+G*+T*A*T*G*T*G*A*C*T*G*T*+G*+A*+T*+G*+T-3'

RNase protection & FISH

CHO U32 forward

5’-TAC TGG GTA AGT TTC ATT CAG-3’

CHO U32 reverse 5’-GTA ATA CGA CTC ACT ATA GGG AGG AAG GAG TCC AGG AGG G-3’

CHO U33 forward 5’-GGG TGC CAT GGA GAA TGG G-3’

CHO U33 reverse 5’-GTA ATA CGA CTC ACT ATA GGG AAG

96

CGC TCT TAG CCC AGA TC-3’

CHO U34 forward 5’-GCA AGC CTA GCT TTC CAC AG-3’

CHO U34 reverse

5’-GTA ATA CGA CTC ACT ATA GGG CTG GGA AGG AGG CTG GTG G-3’

CHO U35 forward 5’-TTG CAG AGT GGT CTA GGT GG-3’

CHO U35 reverse

5’-GTA ATA CGA CTC ACT ATA GGG ACA TAT CCC CCT ATA CAG GAG-3’

FISH mouse U3 forward 5’-TGT AGA GCA CCC GAA ACC AC-3’ mouse U3 reverse 5’-TCC ACT CAG ACT GCG TTC C-3’ mouse rpL13a intron

1 forward 5’-AAT TAA CCC TCA CTA AAAG GGA GCA ATA AAC AGG GTG GCT GT-3’

mouse rpL13a intron 1 reverse

5’-GTA ATA CGA CTC ACT ATA GGG TCC TCA GAT GCT CAA GCA GA-3’

Primer extension

U32a, U33: 18S target

5’- GTAACTAGTTAGCATGCCAGAGTCTCG -3’

U32a: 28S target 5’- GCTACGGACCTCCACCAGAG-3’ U35a: 28S target 5’-TCGTACTGAGCAGGATTACCATGGC-3’ Z17a, Z17b, U45a,

U45b: 18S target 5’- CCCGTCGGCATGTATTAGCTCTAG-3’

*, phosphorothioate linkage; +, LNA residue  

97

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