An E. Coli Small Rna Inhibits Translation Initiation from a Distance

AN E. COLI SMALL RNA INHIBITS TRANSLATION INITIATION FROM A DISTANCE

MUHAMMAD SHAFIUL AZAM

DISSERTATION

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Microbiology in the Graduate College of the University of Illinois at Urbana-Champaign, 2019

Urbana, Illinois

Doctoral Committee:

Professor Carin K. Vanderpool, Chair Professor Rachel J. Whitaker Professor James M. Slauch Professor Gary J. Olsen ABSTRACT

In bacterial systems, small RNA (sRNA)-dependent translational repression is commonly carried out via sRNA-mRNA base pairing interactions near the Shine-

Dalgarno (SD) region. In this so-called “canonical” mechanism, the sRNA is the direct regulator; it competes with the initiating ribosomes while the chaperone protein Hfq plays a supporting role. Contrary to this widely accepted model, there are a few examples in the literature where the sRNA base pairs far from the SD region, yet translation of the target mRNA is still inhibited. Mechanistically, non-canonical translation regulation is one of the least understood aspects of sRNA biology. In the targetome of an E. coli sRNA

SgrS, manXYZ is a non-canonical target where SgrS base pairs at two distinct sites that

are far from the SD regions of manX and manY , yet translation of these two cistrons are

repressed by SgrS. We found that manX translation is controlled by a molecular role-

reversal mechanism where an Hfq binding site is directly adjacent to the manX ribosome

binding site. In this regulatory mechanism, SgrS plays the role of a guide to recruit Hfq to

the appropriate binding site to form the silencing complex. We also report that SgrS

forms a duplex with a uridine-rich translation-enhancing element in the manY 5'

untranslated region. Notably, we show that the enhancer is ribosome-dependent and

that the small ribosomal subunit protein S1 interacts with the enhancer to promote

translation of manY . In collaboration with the chaperone protein Hfq, SgrS interferes with

the interaction between the translation enhancer and the r-protein S1 to repress

translation of manY mRNA. Since bacterial translation is often initiated from a nonlinear

ribosome binding site, sRNA-mediated enhancer silencing could be a common mode of

gene regulation.

Dedicated to

Haseena Khan A Greek among Romans

iii

ACKNOWLEDGEMENTS

“I don't view myself as a practitioner of a particular skill or method. I'm constantly looking at what's the most interesting problem that I could possibly work on. I really try to figure out what sort of scientist I need to be in order to solve the problem I'm interested in solving.”

-Erez Lieberman Aiden

Aside from my mentors, most of my heroes are not some renowned professors. They are my friends, my partner, and my lab mates. Extraordinary humans disguised as ordinary people. They inspired me the most.

My PI and a group of professors influenced me in a meaningful way. Even though they were busy, they were always happy to have a science conversation with me. Some of them were not my committee members; they just cared. The next hundred pages or so is filled with some exciting biology, without the stories of the people that cared for me, inspired me. These few lines are my inadequate attempt to express their positive impact on my life.

TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION ...... 1 1.1 A Historical Perspective ...... 1

1.1.1 The roots of microbiology ...... 1

1.1.2 Escherichia coli and the rise of modern microbiology ...... 2

1.2 Gene Regulation in Bacteria ...... 3

1.2.1 Transcriptional and post-transcriptional regulation ...... 3

1.2.2 RNA-mediated regulation in bacteria ...... 4

1.2.3 The age of small RNA ...... 5

1.2.4 Regulation of translation by bacterial sRNA ...... 7

1.3 Protein Partners in sRNA Mediated Gene Regulation ...... 9

1.3.1 The chaperone proteins ...... 9

1.3.2 RNase E and the degradosome complex ...... 10

1.4 SgrS Rescues E. coli During Sugar-Phosphate Stress ...... 11

1.5 The Ribosomal Protein S1 and Its Role in Translation Initiation ...... 12

1.6 Aim of This Study ...... 14

CHAPTER 2: MATERIALS AND METHODS ...... 15

2.1 Strains and Plasmids ...... 15

2.2 Media and Reagents ...... 16

2.3 β-Galactosidase Assays ...... 16

2.4 In Vitro Transcription ...... 17

2.5 Protein Purification ...... 18

2.5.1 Purification of Hfq ...... 18

2.5.2 Purification of S1 ...... 18

2.6 Toeprinting Assays ...... 20

2.7 Footprinting Assays ...... 21

2.7.1 Hfq and SgrS footprints ...... 21

2.7.2 Ribosome footprints ...... 21

2.7.3 DEPC footprints ...... 22

2.8 Electrophoretic Mobility Shift Assays ...... 22

2.9 In Vitro Translation ...... 23

CHAPTER 3: TRANSLATIONAL REGULATION BY BACTERIAL SMALL

RNAS VIA AN UNUSUAL HFQ-DEPENDENT MECHANISM ...... 31

3.1 Introduction ...... 31

3.2 Results ...... 34

3.2.1 Hfq is essential for translational repression of manX ...... 34

3.2.2 Hfq inhibits formation of the translation initiation complex on

manX mRNA……………………………………………………………………. 43

3.2.3 manX is regulated by DicF sRNA ...... 44

3.2.4 Hfq binds next to the manX ribosome binding site ...... 46

3.2.5 DicF is a weaker regulator of manX ...... 50

3.3 Discussion ...... 52

CHAPTER 4: TRANSLATION INHIBITION FROM A DISTANCE: SgrS

SILENCES A TRANSLATION ENHANCER IN THE LEADER REGION…………..56

4.1 Introduction ...... 56

4.2 Results ...... 60

4.2.1 Hfq is essential for translation repression of manY ...... 60

4.2.2 The manY translation initiation region contains a ribosome-

dependent enhancer ...... 64

4.2.3 Genetic analysis of enhancer sequences ...... 66

4.2.4 Ribosome protection of the non-contiguous RBS and

translation enhancer ...... 70

4.2.5 Ribosomal protein S1 binds to the manY 5' UTR ...... 74

4.2.6 Evidence of SgrS competition with S1 for control of manY

translation ...... 77

4.3 Discussion ...... 80

CHAPTER 5: CONCLUSIONS ...... 84

REFERENCES ...... 87

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CHAPTER 1

INTRODUCTION

1.1 A Historical Perspective

1.1.1 The roots of microbiology

We know little about all the bacteria roaming in our planet. Because we have

sampled, sequenced, and studied a tiny fraction of them, most of the bacteria are

familiar to us from few bits of genomic information gathered from natural samples. To

this day, after a century of extensive research, basic research on the bacterium

Escherichia coli continues to generate unexpected biological phenomenon.

Nevertheless, the knowledge we have gathered can be compiled into volumes after volumes, and it is sufficient to initiate any scientific method to get a view into the inner workings of life.

As the twentieth century began, multiple disciplines of science: physics, biochemistry, genetics, and chemistry cross-pollinated to initiate a new discipline- molecular biology [56]. A significant highlight of this period was the research on bacterial growth physiology by Monod, the creation of the “Phage Group” by Delbrück and Luria, and a book “ What is Life?” by Erwin Schrödinger [90, 95, 120]. As the century rolled along, the field of molecular biology exploded with unprecedented breakthroughs on genetic analysis. With the explosion of biochemical approaches, it was possible to take full advantage of model organisms like Escherichia coli that, with relative ease, could be manipulated to prepare enzymatically active cellular extract and isolate macromolecules so that they could be tagged with radioisotopes. Quasi overnight, every molecular biology lab became engaged into exploration of major biosynthetic pathways, understanding of the concept of the gene and the genetic code, and most importantly, the molecular mechanisms of gene regulation.

Today, we have a remarkably detailed pictures of metabolic pathways; we know

how and where most of the macromolecules are made. The major players of gene

regulation are already identified, and the list is impressive. Nonetheless, there are

considerable limitations in our understanding of how these players communicate with

each other in response to signals to maintain cellular homeostasis and adapt to a

favorable or a harsh environment. Until we understand thoroughly the paradigm of gene

regulation, we will be handicapped in discovering the design principles of life.

1.1.2 Escherichia coli and the rise of modern microbiology

As a member of the gut microbiome, E. coli thrived within our ancestors for

almost five millions of years, predating Homo sapiens [73, 167]. In 1885, a German

pediatrician, Theodor Escherich, isolated a rod shaped bacteria from baby diapers that

thrived in almost all foods [34]. In his manuscript, he described this bacterium as

“slender, short rods” and named it Bacterium coli communis [34]. Few years later, when

Theodor Escherich passed away, scientists renamed it Escherichia coli [167] .

With a moderate sized genome (4.6 m bp) of ~4,300 genes, the Gram negative bacterium Escherichia coli K-12 is a member of the phylum Proteobacteria, a subgroup of the Enterobacteriaceae family [11]. A major portion of the genome (87%) encodes protein-coding genes, 0.8% consists of stable RNA genes, and 0.7% encodes repeat elements, leaving only a tenth of the genome for regulatory functions [11]. E. coli cells are rod shaped, about 2.5 µm long with a diameter of 0.8 µm [9]. It leads a luxurious life in the lower intestine of warm blooded animals and a member of the human microbiome

[9].

At a first glance, E. coli is an unglamorous gram-negative bacterium, it does not differentiate, sporulate, photosynthesize, grow in extreme environments, communicate through a QS (quorum sensing) system, or encode any CRISPR immunity (although the 2 first sequenced CRISPR array was from E. coli [55]); yet the study of this organism is intricately intertwined with the explosion of modern molecular biology. In 1922, a strain of E. coli (K-12) was isolated from a diphtheria patient in California; since then, it had been preserved by the Stanford University as a student laboratory strain [139]. This strain became the rock star model system when Joshua Lederberg, as a graduate student in Edward Tatum’s lab, started to study what he called “the long-shot gamble in looking for bacterial sex.” [69, 139] In the following decades, K-12 will become the organism of choice; scientists all over the world will run hundreds and thousands of experiments on it to learn some of the basic aspects of cell biology, including replication

[72], transcription [129], translation, and the genetic code [56, 96], resulting in its ubiquitous use in academia, pharmaceutical industry, genetic engineering, recombinant protein production, and experiments in microbial evolution [12].

1.2 Gene Regulation in Bacteria

1.2.1 Transcriptional and post-transcriptional regulation

In response to environmental variations, bacteria use a wide range of sophisticated mechanisms to tune expression of genes. Any step in the gene expression, starting at the initiation of transcription to post-translational regulation, can be modulated by cells. Often, changes in gene expression, which frequently propagate through the highly connected gene regulatory network, is essential to maintain cellular homeostasis, activate stress response pathways, and change flux through the metabolic pathways [4].

One way in which bacteria ensure a well-coordinated regulation of gene expression is transcriptional control. Due to a short half-life of RNA transcripts, a pause or inhibition in transcription leads to rapid cessation of protein synthesis [158]. Post- transcriptional gene regulation is employed when cells modulate the half-life and 3 translation of the mRNA transcript. Since the discovery of the regulatory RNAs, there has been an increasing appreciation for the significance of post-transcriptional gene regulation [130, 142].

1.2.2 RNA-mediated regulation in bacteria

In January of 1955, Francis Crick shared an unpublished paper among the RNA

Tie Club members titled “On Degenerate Templates and the Adaptor Hypothesis,” perhaps the most remarkable unpublished paper in the history of molecular biology [23,

156]. For years, Crick has been thinking about the relationship between DNA, RNA, and protein. In his adapter hypothesis paper, Crick argued that RNA was an intermediate between DNA and protein and developed a scheme to explain their relationship (known as the central dogma of molecular biology) [23]. This widely known hypothesis turned out to be true, and during the early days of molecular biology, scientists assumed that the sole purpose of mRNA is to serve as a template for translation. Research over the last few decades has discovered numerous RNA regulators (a heterogeneous group of

RNA molecules that include riboswitches, non-coding RNAs, and CRISPRs) and expanded our understanding of the function of RNA beyond this singular definition.

1.2.3 The age of small RNA

Fig. 1.1: Timeline of the major discoveries in small RNA research. In 1981, RNA I was first discovered, it inhibits the replication of ColE1 plasmid [142]. This discovery was quickly followed by first detection of a Tn10 encoded sRNA and a chromosomally encoded RNA MicF [86, 125], predating eukaryotic miRNA discovery [70, 108]. Based on experimental and computational predictions, current estimates suggests that the E. coli genome encodes more than a hundred sRNA [154].

Almost 37 years ago, the first regulatory non-coding RNA, RNA I, was discovered

in bacteria [142]. Since then, in all three domains of life, a surprisingly diverse yet poorly

characterized pool of regulatory RNAs has been discovered (Fig. 1.1). Bacterial sRNAs

are small transcripts, most in the range of 50-250 nucleotides; they act by imperfect,

non-contiguous base pairing with mRNA targets to control translation or mRNA stability.

Examples of positive regulation by sRNAs are less numerous compared to negative

regulation, but sRNAs have been shown to pair with the 5´ UTR of an mRNA to prevent

formation of a translation-inhibitory secondary structure [67, 91] or to pair with

ribonuclease recognition sequences, thus stabilizing the mRNA (Fig. 1.2) [40]. For

negative regulation, sRNAs often, but not always, operate as translational repressors by

directly pairing with sequences overlapping the SD, sequestering it from the incoming

ribosome (Fig. 1.2) [10, 80, 94].

Fig. 1.2: Positive and negative post-transcriptional regulation by bacterial sRNA. For positive regulation, sRNAs base-pair with the 5 ˊ-UTR making the SD-region easily accessible for the initiating ribosome. Negative regulation is achieved when the sRNA base pairs near the SD region and inhibits translation initiation by competing with the initiating 30S subunit. Often, sRNAs base pair with the target mRNA promoting subsequent degradation by RNase E and the degradosome complex.

To perform any of these regulatory tasks, sRNAs frequently depend on the chaperone protein Hfq. Hfq was initially discovered as the host factor for the replication of bacteriophage Q β, but over the last few decades, its pleiotropic role in cellular physiology has reignited the interest of the research community [39]. Hfq has emerged as a key factor in sRNA-mediated gene regulation, and in control of stability of mRNAs and sRNAs [79, 88]. Hfq is thought of as a matchmaking chaperone that promotes interaction between the sRNA and the target by binding to both RNAs. Another key role of Hfq is protection of sRNAs from RNase E-mediated degradation [57, 88, 126].

1.2.4 Regulation of translation by bacterial sRNA

Besides degradation, studies on sRNA mediated gene silencing have established translational repression as an important mode of gene expression regulation [17].

Studies over the last few years suggest that SgrS is capable of regulating multiple targets at the level of translation [76, 110]. The canonical model of translation initiation involves recognition of the SD by the incoming 30S ribosomal subunit, mediated by

RNA-RNA interaction between the purine rich Shine-Dalgarno (SD) sequence and the

16S rRNA of the 30S ribosomal subunit [124]. Generally, sRNAs target the translation initiation stage by occluding ribosome binding to a target mRNA. To inhibit initiation, however, the sRNA has to bind to a certain sequence window around the SD region (-20 to +19 with respect to initiation codon) where the incoming 30S ribosomal subunit binds

[10].

Fig. 1.3: The canonical mode of sRNA-mediated translation repression. In bacteria, translational repression by sRNAs involves sRNA-mRNA base pairing that occludes the SD region, directly preventing translation. In this mechanism, the sRNA is the direct regulator, while the RNA chaperone Hfq plays a supporting role by stabilizing the sRNA.

Despite substantial progress in understanding the mechanism of translational repression, some crucial questions still remain to be answered concerning how the position of the base pairing site results in regulatory outcomes of translational regulation and/or mRNA degradation. There are several cases where an sRNA can base pair with regions of the mRNA away from the RBS, including the coding region, and still inhibit translation independent of mRNA degradation [17]. Sometimes, these non-canonical repression mechanisms can be achieved by the host factor Q β (Hfq) by a ‘role-reversal’ mechanism. In such cases, Hfq assumes the primary role in translation inhibition in addition to its chaperone role. With the help of its partner sRNA, Hfq is recruited close to the SD region and prevents the 30S subunit loading [28].

In addition to this ‘role-reversal’ mode of action, sRNAs can also bind to an upstream “ribosome standby site” and repress translation. This type of mechanism was first described for an antisense RNA IstR-1, which base pairs approximately 100 nucleotides upstream of the ribosome binding site of tisB mRNA to represses translation

[24]. The ribosome standby model was proposed by de Smit and van Duin as a mode of translation initiation that can be used when the SD region is occluded in a secondary structure [25]. The key assumption of this model is that the 30S subunit binds to an accessible single-stranded site, most likely “non-specifically”, near the secondary structure occluding the SD in order to wait for a brief opening of the secondary structure.

A successful outcome of this interaction, the model proposes, is transient unfolding of the secondary structure, followed by relocation of the ribosome to the SD. If standby sites are occluded by an sRNA, translation will be inhibited.

Translation initiation can also be modulated by other elements known as translation enhancers. These elements can be targeted by sRNA to repress translation.

A recent study showed that Salmonella sRNA GcvB regulates various periplasmic

8 proteins using a conserved G/U-rich seed region that base pairs with target RNA sequences harboring C/A-rich repeat sequence [123]. When present, these C/A multimers stimulate translation in vivo and facilitate ribosome binding in vitro [123].

These translation-stimulatory sites can be located upstream or downstream of the

SD/start codon, a characteristic consistent with canonical translational enhancers. GcvB sequestration of the enhancer C/A-rich sequences therefore inhibits translation of target mRNAs.

1.3 Protein Partners in sRNA Mediated Gene Regulation

1.3.1 The chaperone proteins

A functional RNA molecule needs to maintain the correct conformation and avoid the tendencies to become kinetically trapped in inactive structures [115]. Over the last few decades, a diverse group of proteins has been discovered that help RNAs in reaching their correct conformation, facilitate interactions between two RNAs or, sometimes change the RNA secondary and tertiary structures by transient and non- specific binding [48, 88, 104, 143].

In bacteria, the ring-shaped homohexameric protein Hfq is perhaps the ‘poster child’ example of a RNA binding protein involved in sRNA-mediated gene regulation. Hfq was first identified as a host factor that unwinds phage Q β RNA for efficient replication [57,

58]. Since then, a volume of literature discovered its pleotropic role in endogenous gene control, ribosome biogenesis, and its role as a chaperone in sRNA mediated post- transcriptional gene regulation [165]. Hfq is a member of the Sm/LSm protein family that binds and protects sRNAs from degradation, and in the current standard model, if the sRNA possesses adequate complementarity with a target mRNA, promotes formation of an active silencing complex [159]. The ternary complex (composed of sRNA, mRNA, and Hfq) typically sequesters the SD region, inhibiting translation of the mRNA [46, 80]. 9

During the last decade, another chaperone protein ProQ has emerged as a new class of chaperone protein in Salmonella and E. coli [41, 45, 157]. Recent studies identified hundreds of transcripts that interact with ProQ and reinforced its role as a major regulator of bacterial motility, chemotaxis, and virulence genes [51, 157]. ProQ recognizes specific structural motifs of sRNAs and 3 ˊend of mRNAs--it has both strand exchange and RNA matchmaking activity [41, 51]. Although the mechanistic aspects of the chaperone protein ProQ and the cellular roles of the ProQ associated sRNAs remain mostly unresolved, current understanding highlights the importance of the chaperone proteins in bacterial physiology, growth, and virulence.

1.3.2 RNase E and the degradosome complex

A key player in sRNA mediated gene regulation is a single-strand specific endoribonuclease that is also involved in control and coordination of RNA metabolism including processing, maturation, and degradation of rRNA, tRNA, sRNA, and mRNA [1].

RNase E is an essential enzyme for E.coli that was first identified as a factor involved in processing of 9S rRNA precursor into mature 5S rRNA transcript [43]. The primary sequence of RNase E can be divided into two distinct regions: the amino terminal catalytic end, and the carboxy terminal RNA-binding region that harbors binding sites for

RNA and other components of the degradosome [19]. RNase E is frequently recruited by the sRNA (and/or the chaperone protein Hfq), which subsequently forms into a degradosome, an sRNA-induced mRNA degradation complex [93]. The initial cleavage in the AU-rich RNA motif is catalyzed by RNase E followed by a rapid exonucleolytic degradation by PNPase, another member of the degradosome complex [18]. A typical degradosome also contains an ATP-dependent RNA helicase RhlB, and a prominent glycolytic enzyme enolase [21, 106].

1.4 SgrS Rescues E. Coli During Sugar-Phosphate Stress

SgrS is a 227 nt long Hfq-dependent small RNA that promotes both positive and

negative effects on its targets [148]. It is involved in rescuing cells during a specific

condition called glucose-phosphate stress. High amounts of sugar-phosphate

accumulation inside bacterial cells cause toxicity through an unknown mechanism [153].

When cells are grown in presence of non-metabolizable sugar analogs, such as α-

methyl glucoside ( αMG) or 2-deoxy-D-glucose (2DG), it creates a similar situation where

phosphosugars are accumulated inside the cell, glycolysis is inhibited and the cells

display a bacteriostatic phenotype [92, 146]. Like their metabolizable counterparts, 2DG

and αMG enter the cell via the phosphoenolpyruvate phosphotransferase system (PTS).

2DG and αMG enter the cell through the mannose transporter (ManXYZ, EIIABCD Man ) and glucose transporter (PtsG, EIICB Glc ), and get phosphorylated. Accumulation of

sugars triggers an unknown stress signal that activates SgrR, a transcriptional activator

of SgrS [148]. Work published by our group showed that these newly synthesized SgrS

transcripts eventually base pair with the ptsG , and manXYZ transcripts and promotes degradation of the RNA duplex by RNase E [152].

Fig. 1.4: SgrS mediated regulation of manXYZ transcript. The PTS (PEP phosphotransferase) system uses a phosphorylation relay that transfers the phosphate from PEP to mannose transporter. When sugars are transported, they are phosphorylated. The accumulation of sugars inside the cell somehow activates SgrR, which activates the expression of SgrS. SgrS associates with the chaperone protein Hfq and base pairs with manXYZ . Base pairing stimulates RNase E-mediated degradation and translational repression.

1.5 The Ribosomal Protein S1 and Its Role in Translation Initiation

The 5 ˊ UTR region harbors two major determinants of translation initiation: i)

Shine-Dalgarno (SD) sequence, a purine rich 3-10 nt long sequence located

approximately 10 nt upstream of the start codon [124], and ii) A/U-rich sequence

upstream of the SD sequence that are frequently exploited by the ribosome to enhance

translation [14, 98, 137]. Several studies suggest that these translation enhancers

stimulate translation via a direct interaction with the small ribosomal subunit protein S1

[14, 31, 137].

S1 is not a typical ribosomal protein. It is unusually acidic, has high affinity for mRNA, and its association with the 30S ribosomal subunit is weak and reversible [30,

60, 133]. Given its weak affinity for the ribosome (a binding constant of 2×10 8 M-1), existence of free S1 in ribosome free cell fractions, and the molar stoichiometry (<1 for every 30S subunit) in purified ribosomes, it is often argued that S1 should be considered as a translation factor rather than a component of the small subunit [27, 133]. S1 protein consists of six OB-fold repeats and it interacts with various heterogeneous RNAs, as well as poly U, poly A, and poly C homopolymers [30, 133]. The structure of this protein is yet to be resolved, but a recent cryo-EM study proposed an elongated shape shown in figure 1.5 [122].

Fig. 1.5: The 3D structure of the r-protein S1. It is composed of six structural motifs (OB- folds, each approximately 70 amino acids in length) [122]. The domains are represented in different colors. Domains 1-3 at the N-terminal end interacts with the 30S ribosomal subunit (through r-protein S2) [31, 132].

A recent study on ‘minimal’ ribosomes found that ribosomes lacking the S1 protein can efficiently translate leaderless mRNAs [87]. Apart from the leaderless mRNAs, the role of S1 can be crucial where the interaction between ribosome- associated S1 and the 5 ˊ UTR ensure rapid initiation complex formation [47, 140]. Often, 13

S1 interacts with polypyrimidine-rich sequence near the SD region and melts secondary

structures and subsequently facilitates translation initiation complex formation [31].

1.6 Aim of This Study

In this study, we investigated the mechanism by which SgrS regulates the first

cistron of the manXYZ operon, manX . We observed previously that regulation of manX mRNA by SgrS involves base pairing 20 nucleotide downstream of the start codon, which lies outside the “five codon window” (a 15 nucleotide region, with respect to the start codon) [110]. We also characterized regulation of manX translation by another

sRNA regulator, DicF [5], a 53-nt long Hfq-dependent sRNA [35]. The DicF binding site

on manX mRNA is even further downstream than the SgrS binding site. We hypothesized that each of these sRNAs regulates manX translation by a “non-canonical” mechanism, since their binding sites are positioned too far downstream for sRNA-mRNA base pairing to directly occlude ribosome binding. To test this hypothesis, we addressed several questions. Does sRNA-mRNA duplex formation directly inhibit translation by preventing formation of the translation initiation complex? If not, then is Hfq required for translational repression? Does Hfq bind to the manX mRNA near the ribosome binding site?

SgrS base pairs with the manY 5ˊ UTR and the pairing region is 30 nucleotide upstream of the manY SD [109]. This is highly unusual since the base pairing site is positioned too far upstream for sRNA-mRNA base pairing to directly occlude ribosome binding and leads to the following obvious questions. Does sRNA-mRNA duplex formation directly inhibit translation by preventing the base pairing interaction between the 16S rRNA and the SD site? If not, then is Hfq required for translational repression?

Does Hfq bind to the manX mRNA near the ribosome binding site? Also, does this base pairing site (SgrS anti-seed region) interact with a cellular factor to modulate translation?

CHAPTER 2

MATERIALS AND METHODS

2.1 Strains and Plasmids

The strains, plasmids, and oligonucleotides used in this study are listed in Tables

2.1 and 2.2. Derivatives of E. coli K-12 MG1655 were used for all experiments. Alleles were moved between strains using P1 transduction [83] or λ-red recombination [164].

Translational LacZ fusions, under the control of an arabinose-inducible P BAD promoter,

were constructed by PCR amplifying DNA fragments using primers with into PM1205

using λ-red homologous recombination (Table 2.2). These fragments were integrated

into the chromosome by λ-red recombination using counterselection against sacB as

described previously [77].

SA1328, a strain with tet-Cp19-115nt-manX`-`lacZ, ΔsgrS, lacI q, kan R, Δhfq genotype was constructed in two steps. First, Δhfq::FRT-kan-FRT was transduced into

JH111 and pCP20 was used to flip out the kan R cassette. The tet-Cp19-115nt-manX`-

`lacZ cassette was then transduced into the latter strain.

Strains containing truncated manX translational fusions under the control of a

PBAD promoter, were constructed in strain PM1205 [77]. The P BAD -22nt-manX'-'lacZ and

PBAD -25nt-manX'-'lacZ fusions were generated by PCR amplifying DNA fragments with primer pairs O-SA178/O-SA176 and O-SA177/O-SA176 primer pairs respectively, containing 5´ homologies to pBAD and lacZ (Table 2.2). The PCR products were

recombined into PM1205 using λ-red homologous recombination as described

previously. The same fusions with mutations in the Hfq binding site (in strains SA1522

and SA1620), were created using the method above, but using oligonucleotides O-

SA177/O-SA176 and O-SA177/O-SA433 to obtain the PCR products.

The manY 5ˊ-UTR ˊ-ˊlacZ and thrS 5ˊ-UTR ˊ-ˊlacZ translational fusions with wild-

type 5 ˊ-UTR and with mutations in the enhancer region was generated using single 15

stranded oligos (containing 5´ homologies to pBAD and 3 ˊ homoogies to lacZ) into

PM1205 using λ-red homologous recombination. The same protocol was used to generate a set of manY 5ˊ-UTR ˊ-ˊlacZ strains with a combination of i) strong and weak

SD regions, and ii) wild-type and mutated enhancer elements (Table 2.1 and 2.2).

Plasmids encoding wild-type and mutated rpsA under the control of a weak and a

strong SD sequences were constructed in a pWKS30 vector by PCR amplifying rpsA

gene from pET28-rpsA (for WT S1) and pET28-mrpsA (for a mutated version of the S1

protein) using OSA774/OSA776 (for strong SD) and OSA775/OSA776 (weak SD) primer

pairs. Assembly of these PCR fragments with the pWKS30 backbone (amplified with the

primer pair OSA783/OSA784) were performed using the NEBuilder® HiFi DNA

Assembly Master Mix (following manufacturer’s instructions).

2.2 Media and Reagents

Unless otherwise stated, bacteria were cultured in LB broth or on LB agar plates

at 37°C. TB medium was used for β-Galactosidase assays. To induce Lac promoters,

0.1 mM IPTG (isopropyl β-D-1-thiogalactopyranoside) was used. L-arabinose was used

at concentrations of 0.001%, for solid media, and 0.002%, for liquid media, to induce

PBAD promoters. Antibiotics were used at following concentrations: 100 µg/ml ampicillin,

25 µg/ml chloramphenicol, 50 µg/ml spectinomycin, and 25 µg/ml kanamycin.

2.3 β-Galactosidase Assays

Strains with manX ˊ-ˊlacZ fusions were grown overnight in TB medium and subcultured 1:100 to a fresh medium containing Amp and 0.002% L-arabinose (for PBAD

promoters). Cultures were grown at 37°C with shaking to OD 600 ~0.2, and 0.1 mM IPTG was added to induce expression of SgrS or DicF. Cells were grown for another hour to

OD 600 ~0.5. β-Galactosidase assays were performed on these cells according to the 16

previously published protocol [82]. Unless mentioned otherwise, strains with manY

UTR ˊ-ˊlacZ fusions were grown overnight in TB medium and subcultured 1:100 to a

fresh medium containing Amp and 0.002% L-arabinose. Cells were grown at 37°C with

shaking to OD 600 ~0.5 and β-Galactosidase assays were performed.

For assays conducted on strains in Fig. 4.8, culture conditions were slightly

different. Strains with manY'-'lacZ, thrS'-'lacZ , and sodB'-'lacZ fusions harboring WT S1 or

mS1 plasmids were grown overnight in TB medium and subcultured 1:100 to a fresh

medium containing Amp. Cultures were grown at 37°C with shaking to OD 600 ~0.4, and

0.002% L-arabinose (final concentration) was used to induce expression of reporter fusions. Cells were grown for another 20 minutes and 0.1 mM IPTG (final concentration) was added to induce S1 or mS1 production. Cells were grown for an additional 10 min before β-Galactosidase assays were performed. SgrS was induced by adding 0.5% αMG

(final concentration) to the media.

2.4 In Vitro Transcription

For in vitro transcription, template DNA was generated by PCR using gene specific oligonucleotides with a T7 promoter sequence at the 5´ end of the forward primer. The following oligonucleotides were used to generate templates for RNA footprinting and gel shift assays: O-JH219/O-JH119 and O-JH218/O-JH169 to generate manX and SgrS template DNA. DNA template for DicF transcription was generated by hybridizing two oligos, DicFW and DicFC, in TE buffer. Oligonucleotide pairs

OSA753/OSA754 and OSA734/OSA735 were used to PCR amplify manY and thrS templates from MG1655 genomic DNA. Transcription of these DNA templates was performed using the MEGAscript T7 kit (Ambion) following manufacturer’s instructions.

2.5 Protein Purification

2.5.1 Purification of Hfq

Hfq-His protein was purified following a previously published protocol [76].

BL21(DE3) cells harboring pET21b-Hfq-His 6 was cultured in 400 ml LB medium. At

OD 600 ~0.3, 1 mM IPTG was added to the culture and incubation was continued for 2 hrs.

The cells were washed with STE buffer (100 mM NaCl; 10 mM Tris·HCl, pH 8.0; 1 mM

EDTA) and resuspended in 10 ml Equilibration buffer (50 mM Na 2HPO 4–NaH 2PO 4, 300

mM NaCl, and 10 mM imidazole). The suspension was treated with 25 mg lysozyme,

incubated on ice for 10 min and sonicated. The supernatant was collected after

centrifugation at 16,000 × g for 10 min at 4°C followed by incubation at 80°C for 10 min.

The sample was centrifuged again, at 16,000 × g for 10 min at 4°C. The supernatant

was fractionated using a Ni 2+ NTA agarose column following manufacturer’s instructions

(Roche) and checked by SDS-PAGE electrophoresis. The fractions containing Hfq were

pooled, dialyzed, and stored in a storage buffer (20 mM Tris·HCl pH 8.0, 0.1 M KCl; 5

mM MgCl 2, 50% glycerol, 0.1% Tween 20, and 1 mM DTT) at −20°C.

2.5.2 Purification of S1

A DNA fragment containing the rpsA gene was PCR amplified from MG1655

genomic DNA using OSA632/OSA633 primer pair. The pET28 backbone was PCR

amplified with OSA636/OSA637 primer pair using pET28-Scy-C as a template (a gift

from the Nair lab, University of Illinois at Urbana). The fragments were assembled into

pET28-rpsA plasmid using the NEBuilder® HiFi DNA Assembly Master Mix (following

manufacturer’s instructions). Primer pair OSA779/OSA780 was used to PCR amplify

pET28-rpsA and introduce mutations into the rpsA coding region. The amplified PCR product was circularized using the NEBuilder® HiFi DNA Assembly Master Mix to generate pET28-mrpsA (Table 2.1, Fig. 2.1).

Fig. 2.1: The pET28-rpsA expression vector encoding the His-tagged S1 gene under the control of a T7 promoter. The poly-histidine tags are preceded by a thrombin cleavage site.

S1 was purified following a published protocol with modifications [134]. E. coli

BL21 (DE3) cells with the pET28-rpsA vector was grown to late exponential phase.

Around OD of 0.6-0.8, the expression of the protein was induced with IPTG (1 mM, final

concentration) and the incubation was continued for 4hr at 37 0C. At this point, cells were harvested by centrifugation and the cell pellet was resuspended in 30 mL of extraction buffer (1X PBS, 0.5 M NaCl, pH 7.2) and lysed in a French press. The supernatant was collected after centrifugation at 16,000 × g for 10 min at 4°C. The supernatant was fractionated using a Hi-Trap Ni 2+ column (GE Healthcare) following manufacturer’s

instructions (Roche) and checked by SDS-PAGE electrophoresis. The fractions

containing S1 were dialyzed overnight in TGED buffer (10 mM Tris-HCl pH 8, 5%

glycerol, 0.1 mM EDTA, and DTT 0.015 mg/mL) and loaded on a mono-Q column (GE

Healthcare). The column was washed with TGED buffer and protein was eluted with a

linear gradient of TGED buffer with NaCl (0.1 M-1 M). The fractions containing the S1

protein were pooled, dialyzed, and concentrated using Centricon 10 concentrator

(Millipore-Sigma). The poly-histidine tag was removed using the Thrombin cleavage kit

(Millipore-Sigma) following manufacturer’s instructions. The cleaved proteins were

further resolved in a Superdex 200 column with TGED buffer containing 0.1 M NaCl. The

column fractions containing S1 were concentrated using Centricon 10 concentrator

(Millipore-Sigma), mixed with an equal volume of 100% glycerol and stored at -20 0C.

2.6 Toeprinting Assays

32 Toeprinting assays were performed using unlabeled manX ATG and P -end- labeled primer in the presence and absence of Hfq and SgrS following the previously published protocol [36]. For each reaction, 2 pmol of manX RNA and 1.6 pmol of end-

labeled primer (O-JH119) were heated for one min at 95°C in toeprint buffer (20 mM

Tris-HCl pH 7.5, 50 mM KCl, 0.1 mM EDTA, and 1 mM DTT). The mixture was chilled in

ice for five minutes, and 10 mM MgCl 2 and 1 mM of dNTPs were added. Purified Hfq and

in vitro synthesized SgrS RNA were added to the appropriate reaction mixtures and

incubated at 37°C for 10 minutes. Next, ribosomes (1.3 pmol, NEB) were added to this

reaction mixture, and the incubation was continued at 37°C for five minutes. Thirteen

picomoles of fMet-tRNA (Sigma) was added to this reaction and cDNAs were

synthesized using SuperScript III reverse transcriptase (Invitrogen). The reaction was

stopped by adding 10 µl of loading buffer II (Ambion). The reaction products were

analyzed on an 8% polyacrylamide-urea gel. Sequencing ladders were generated using

Sequenase 2.0 DNA sequencing kit (Affymetrix) .

2.7 Footprinting Assays

2.7.1 Hfq and SgrS footprints

In vitro RNA-footprinting reactions were performed as described previously [29] with some modifications. 0.1 pmol of 5´-end labeled manX mRNA was incubated at 37°C for 30 minutes in structure buffer (Ambion) containing 1 ng of yeast RNA (Ambion), in the presence or absence of 100 pmoles of unlabeled SgrS and 3.7 pmoles of Hfq. Lead acetate (Sigma) was added to perform the cleavage reaction (2.5 µM) and incubated at

37°C for two min. Reactions were stopped by adding 12 µL of loading buffer (Ambion).

The alkaline ladder was generated by incubating 5´-end labeled manX mRNA at 90°C for 5 minutes in alkaline buffer (Ambion). RNase T1 was used for 5 min at 37°C to generate the G-ladder. The samples were resolved on an 8% polyacrylamide-urea gel.

2.7.2 Ribosome footprints

0.1 pmol of 5´-end labeled manY and thrS mRNAs were incubated at 37°C for 30

minutes in structure buffer (Ambion) containing 1 ng of yeast RNA (Ambion), in the

presence or absence of 10 pmol E. coli 70S ribosome (NEB). Lead acetate (Sigma) was

added to perform the cleavage reaction (2.5 µM) and incubated at 37°C for two minutes.

Reactions were stopped by adding 12 µL of loading buffer (Ambion). The alkaline ladder

and the G-ladders were generated as described before. The footprinting samples were

resolved in a 8% polyacrylamide-urea gel.

2.7.3 DEPC footprints

The DEPC footprint experiment was performed following a published protocol with

some modifications [14]. In brief, 5 pmol of in vitro transcribed manY transcript was incubated with WT or mS1 protein (3 µM, final concentration) at 37 ºC for 10 minutes in binding buffer (20 mM Tris-HCl pH 7.6, 10 mM MgCl 2, 100 mM NH 4Cl, and 0.5 mM DTT).

1 µL DEPC (Sigma, undiluted) was added to the reaction mixture and incubated at room temperature with gentle shaking. Reactions were stopped with 40 µL stop solution (0.4 M

Na-acetate pH 5.2, and 20 mM EDTA). Modified manY transcripts were phenol extracted and precipitated with ethanol. Using a reverse primer (OSA754), primer extension was performed with α32 P dATP (Perkin-Elmer) in the reaction mixture. Sanger ladders were

generated using the Sequenase Version 2.0 DNA Sequencing Kit (Applied Biosystems).

2.8 Electrophoretic Mobility Shift Assays

RNA-RNA and RNA-protein gel electrophoretic mobility shift assays were

performed using 0.01 pmol of P 32 -labeled denatured manX RNA and the indicated amounts of SgrS (denatured at 95°C for 1 min) or Hfq in binding buffer (10 mM Tris-HCl pH 8.0, 0.5 mM DTT, 0.5 mM MgCl 2, 10 mM KCl, 5mM Na 2HPO 4–NaH 2PO 4 pH 8.0). The mixture was incubated at 37°C for 30 min, and non-denaturing loading buffer (50% glycerol and 0.1% bromophenol blue) was added. The samples were resolved on a 4.6% native polyacrylamide gel for 1.5 h at 10 mA. The fraction of manX RNA bound was determined using Fluorescent Image Analyzer FLA-3000 (FUJIFILM) to quantitate the intensities of the bands. The data were fit into Sigmaplot software to obtain the KD value.

Hfq-sRNA-target mRNA gel mobility shift assays were performed using 0.01

pmol of P 32 -labeled manY RNA and the indicated amounts of SgrS and Hfq in binding

buffer. The mixture was incubated at 37°C for 30 min. Hfq was removed from the

reaction with phenol extraction. A non-denaturing loading buffer (50% glycerol and 0.1% 22 bromophenol blue) was added to the aqueous phase and the samples were resolved on a 4.6% native polyacrylamide gel for 1.5 h at 10 mA. The fraction of manY RNA bound was determined using Fluorescent Image Analyzer FLA-3000 (FUJIFILM) to quantitate the intensities of the bands. The data were fit into GraphPad Prism7 software to obtain the KD value.

2.9 In Vitro Translation

In vitro transcribed myc tagged manY transcripts (1 pmol, with and without translation enhancer) were translated in vitro using the PURExpress translation kit

(NEB). Translation reactions were stopped by adding Laemmli sample buffer (Bio-Rad).

The samples were resolved in a NuPAGE gradient gel (Life Technologies) and transferred to Immobilon-PSQ membrane (Millipore-Sigma). The membranes were incubated in a blocking buffer with anti-myc antibody (Sigma). The signals were visualized with ECL western blotting substrate (thermo Scientific).

Table 2.1 Strains and plasmids used in this study

Strain Description Source DJ480 ∆lac X74 D. Jin, NCI NM2000 DJ480, mini λ::Cm (carries λ red recombination N. Majdalani, functions) NCI JH111 λattB:: lacI q-PN25 tetR -spec R ΔsgrS [110] JH184 tet-Cp19-ptsG`-`lacZ, ΔsgrS, lacIq, kanR [110]

SA1522 PBAD -25nt(mutated Hfq binding site)-manX’-‘lacZ, This study mal::lacIq JH175 tet-Cp19-115nt-manX`-`lacZ, ΔsgrS, lacIq, kanR [110] SA1328 tet-Cp19-115nt-manX`-`lacZ, ΔsgrS, lacIq, kanR, This study Δhfq JH178 tet-Cp19-65nt-manX`-`lacZ, ΔsgrS, lacIq, kanR [110] JH181 tet-Cp19-30nt-manX`-`lacZ, ΔsgrS, lacIq, kanR [110]

SA1403 PBAD -22nt-manX'-'lacZ, mal::lacIq This study

SA1404 PBAD -25nt-manX'-'lacZ, mal::lacIq This study

Table 2.1 (cont.)

Strain Description Source DB151 ΔsgrS, λattB::lacIq, tetR(ep), spR, hfq::cat, ptsG'- [109] 'lacZ

SA1620 PBAD 25nt(mutated Hfq binding site)-manX 19 ˊ- ˊlacZ, mal::lacIq R PM1205 lacI ˊ:: P BAD -cat-sacB-lacZ, mini λtet , ΔaraBAD [77] araC+, mal::lacI q SA1747 PBAD -manY UTR-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q q SA1735 PBAD -mut1-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1736 PBAD -mut2-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1737 PBAD -mut3-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1738 PBAD -mut4-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1739 PBAD -mut5-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1748 PBAD -mut6-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1749 PBAD -mut7-ATG '-'lacZ, ΔaraBAD araC+, mal::lacI This study

SA1801 PBAD -WT thrS UTR '-'lacZ, ΔaraBAD araC+, This study mal::lacI q q SA1811 PBAD -G-thrS'-'lacZ, ΔaraBAD araC+, mal::lacI This study q SA1815 PBAD -A-thrS'-'lacZ, ΔaraBAD araC+, mal::lacI This study

SA1516 PBAD -manY UTR( Δenh )'-'ATG-lacZ, ΔaraBAD This study araC+, mal::lacI q SA1517 PBAD -manY UTR( Δenh )'-'ATG-enh (in frame)-lacZ, This study ΔaraBAD araC+, mal::lacI q SA1806 PBAD -flip-enh manY UTR '-'lacZ, ΔaraBAD araC+, This study mal::lacI q SA1807 PBAD -manY UTR with a strong SD and mut3 This study enhancer '-'lacZ, ΔaraBAD araC+, mal::lacI q SA1808 PBAD -manY UTR with a lacZ SD and mut5'-'lacZ, This study ΔaraBAD araC+, mal::lacI q SA1809 PBAD -manY UTR with a weak SD '-'lacZ, ΔaraBAD This study araC+, mal::lacI q SA1810 PBAD -manY UTR with a strong SD '-'lacZ, This study ΔaraBAD araC+, mal::lacI q q SA1916 PBAD -mut9 ATG ˊ-'lacZ, ΔaraBAD araC+, mal::lacI This study PBAD -mut10 ATG '-'lacZ, ΔaraBAD araC+, This study SA1917 mal::lacI q PBAD -mut14 ATG '-'lacZ, ΔaraBAD araC+, This study SA1918 mal::lacI q PBAD -mut15 ATG '-'lacZ, ΔaraBAD araC+, This study SA1919 mal::lacI q SA1907 PBAD -manY UTR-ATG '-'lacZ, hfq::kan, ΔaraBAD This study araC+, mal::lacI q q SA1902 PBAD -sodB'-'lacZ, ΔaraBAD araC+, mal::lacI This study

Table 2.1 (cont.)

Strain Description So urce PBAD -manY UTR-ATG '-'lacZ, ΔaraBAD araC+, This study SA1755 mal::lacI q, rne131::kan SA1750 PBAD -mut1-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan

SA1751 PBAD -mut2-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan

SA1752 PBAD -mut3-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan

SA1753 PBAD -mut4-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan

SA1754 PBAD -mut5-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan

SA1756 PBAD -mut6-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan

SA1757 PBAD -mut7-ATG '-'lacZ, ΔaraBAD araC+, This study mal::lacI q, rne131::kan Plasmid pBRCS12 pHDB3, P LlacO , vector control [151] pLCV1 pHDB3 P LlacO ‐sgrS K12 [147] pBRCS22 pHDB3 P LlacO -sgrS St [151] pSUB11 3xFLAG::FRT-kan R-FRT [145] pET21b-Hfq- T7promoter-Hfq-(his) 6 This study

His 6 pLac-DicF pLac-DicF [5] pLac-DicF20 pLac-DicF20 (U41A, G42C) [5] pWKS30 PLlacO , vector control [155] pWKS30-SmS1 pLac-strong SD-mutated rpsA This study pWKS30- pLac-weak SD-mutated rpsA This study WmS1 pWKS30-WS1 pLac-weak SD-WT rpsA This study pWKS30-SS1 pLac-strong SD-WT rpsA This study pET28-rpsA T7 promoter –WT rpsA -thrombin site- (His) 12 This study pET28-mrpsA T7 promoter –mutated rpsA -thrombin site- (His) 12 This study

Table 2.2 Oligos used

Oligo Description Sequence 5 ˊ-3ˊ OSA178 Single-stranded oligo for ACCTGACGCTTTTTATCGCAACTCTCTACTGTTTCTCC recombineering manX 5ˊ UTR ATACGATAATAAAGGAGGTAGCAAGTGACCATT (with 22nt) into PM1205 OSA176 Reverse primer for TAACGCCAGGGTTTTCCCAGTCACGACGTTGTAAAACG recombineering manX (+39 nt ACAGCCCAACCATGTGTGCCTATAAC into the coding region) OSA177 Single-stranded oligo for ACCTGACGCTTTTTATCGCAACTCTCTACTGTTTCTCC recombineering manX 5ˊ UTR ATTTAACGCGGCGGGAGGAGGTAGCAAGTGACCATTGC with mutated Hfq binding site OSA433 Reverse primer for TAACGCCAGGGTTTTCCCAGTCACGACGTTGTAAAACG recombineering (manX with 19 ACTTTAAGCAACTGCTCTGCAGCCCAAC codons, with DicF binding site) into PM1205 OJH119 Reverse primer, manX in vitro CGATCCAGCCGACGTTTTTCCTGCTCGCCTAACAGCAT transcription T OJH219 Forward primer, manX in vitro TAATACGACTCACTATAGGGGCGAAACGCAGGGGTTTT transcription TGGTTGTAG DicFW Single-stranded oligo with T7 TAATACGACTCACTATAGGGTTTCTGGTGACGTTTGGC promoter for DicF in vitro GGTATCAGTTTTACTCCGTGACTGCTCTGCCGCCC transcription (Watson strand) DicFC Single-stranded oligo for DicF in GGGCGGCAGAGCAGTCACGGAGTAAAACTGATACCGCC vitro transcription (Crick strand) AAACGTCACCAGAAACCCTATAGTGAGTCGTATTA manYUTR Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGTTTTCACACTTAAGTCTTACGTAAACA into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut1 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut1 manY 5ˊ TTATCACTCAGGGTTCACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut2 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut2 manY 5ˊ TTATCACTCAGTTGGCACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut3 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut3 manY 5ˊ TTATCACTCAGGGGGCACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut4 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut4 manY 5ˊ TTATCACTCAGTTTTCACACGGAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut5 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut5 manY 5ˊ TTATCACTCAGAAAACACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut6 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut6 manY 5ˊ TTATCACTCATTTTTCACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

Table 2.2 (cont.)

Oligo Description Sequence 5 ˊ-3ˊ

mut7 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut7 manY 5ˊ TTATCACTCAATTTTCACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut9 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut9 manY 5ˊ TTATCACTCAGTTTTTACACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut10 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut10 manY 5ˊ TTATCACTCAGTTTTCGCACTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut14 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut14 manY 5ˊ TTATCACTCAGTTTTCACGCTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

mut15 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering mut14 manY 5ˊ TTATCACTCAGTTTTCACTCTTAAGTCTTACGTAAACA UTR into PM1205 GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

OSA628 T7 forward primer for in vitro TAATACGACTCACTATAGGGAGTCCGTAAGGTTTCCAC transcription of manY (-129 CGAT relative to the manY start codon)

OSA629 Reverse primer for in vitro TCGAGGATTGATCCCATACCTGCGATA transcription of manY (+74)

manY46UT Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCACTCAGTTTTCA R recombineering manY 5ˊ UTR CACTTAAGTCTTACGTAAACAGGAGAAGTACAATGGTC (46nt) into PM1205 GTTTTACAACGTCGTGACTGGG

Δenh Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATTTAAGTCTTACGT recombineering manY 5ˊ UTR AAACAGGAGAAGTACAATGGTCGTTTTACAACGTCGTG (Δenh) into PM1205 ACTGGG

mvENH Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATTTAAGTCTTACGT recombineering manY 5ˊ UTR AAACAGGAGAAGTACAATGCACTCAGTTTTCACACTTG (ATGenh) into PM1205 TCGTTTTACAACGTCGTGACTGGG

OSA003 T7 forward primer for in vitro TAATACGACTCACTATAGGGCGTATTGTGTTGATTATC translation of manY (-62 relative ACTCAGTTTTC to the manY start codon) OSA823 Reverse primer for in vitro TATTTGATGCCTCTAGAGTCTTACTTATCGTCATCGTC translation of manY -3XFLAG TTTGTAATCAATATCATGATCCTTGTAGTCTCCGTCGT (+351) GGTCCTTATAGTCGGTAATAGTACGAACGATGATGGTC AGTACCT

OSA321 T7 forward primer for in vitro TAATACGACTCACTATAGGGCGTATTGTGTTGATTATT translation of manY (-62 relative TAAGTCTTACGTAAACAGGAGAAGTAC to the manY start codon, Δenh)

Table 2.2 (cont.)

Oligo Description Sequence 5 ˊ-3ˊ

Y UTR lacZ Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA SD recombineering manY 5ˊ UTR TTATCACTCAGTTTTCACACTTAAGTCTTACGTAAACA with lacZ SD into PM1205 GGAAACAGCTATGGTCGTTTTACAACGTCGTGACTGGG

lacZ mut3 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGGGGGCACACTTAAGTCTTACGTAAACA (with lacZ SD and mut3 GGAAACAGCTATGGTCGTTTTACAACGTCGTGACTGGG enhancer) into PM1205 lacZ mut5 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGAAAACACACTTAAGTCTTACGTAAACA (with lacZ SD and mut5 GGAAACAGCTATGGTCGTTTTACAACGTCGTGACTGGG enhancer) into PM1205 flip-enh Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCTATTAGTTGTGT recombineering manY 5ˊ UTR TATGCACTCAGTTTTCACACTTCAAATGCATTCTGAAA (with the flipped enhancer) into GGAGAAGTACAATGGTCGTTTTACAACGTCGTGACTGG PM1205 G

sSD Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGTTTTCACACTTAAGTCTTACGTAAACA (with a strong SD) into PM1205 GGAGGAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

wSD Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGTTTTCACACTTAAGTCTTACGTAAATT (with a weak SD) into PM1205 GGAGTAGTACAATGGTCGTTTTACAACGTCGTGACTGG G

sSD+mut3 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGGGGGCACACTTAAGTCTTACGTAAACA (with a strong SD and the mut3 GGAGGAGTACAATGGTCGTTTTACAACGTCGTGACTGG enhancer) into PM1205 G

wSD+mut3 Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATCGTATTGTGTTGA recombineering manY 5ˊ UTR TTATCACTCAGGGGGCACACTTAAGTCTTACGTAAATT (with a weak SD and the mut3 GGAGTAGTACAATGGTCGTTTTACAACGTCGTGACTGG enhancer) into PM1205 G

WT thrS Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATTAATAAACAAATT recombineering thrS 5ˊ UTR (-66 TTTCTTTGTATGTGATCTTTCGTGTGGGTCACCACTGC to -1, relative to the start codon) AAATAAGGATATAAAATGGTCGTTTTACAACGTCGTGA into PM1205 CTGGG

A-thrS Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATTAATAAACAAAAA recombineering thrS 5ˊ UTR with AAACTTTGTATGTGATCTTTCGTGTGGGTCACCACTGC A-thrS (-66 to -1, relative to the AAATAAGGATATAAAatgGTCGTTTTACAACGTCGTGA start codon) into PM1205 CTGGG

G-thrS Single-stranded oligo for TCGCAACTCTCTACTGTTTCTCCATTAATAAACAAAGG recombineering thrS 5ˊ UTR with GGGCTTTGTATGTGATCTTTCGTGTGGGTCACCACTGC G-thrS (-66 to -1, relative to the AAATAAGGATATAAAATGGTCGTTTTACAACGTCGTGA start codon) into PM1205 CTGGG

OSA734 T7 forward primer for in vitro TAATACGACTCACTATAGGGGATTGCGAACCAATTTAG translation of thrS (-66 relative to CATTTGTTGG the thrS start codon)

Table 2.2 (cont.)

Oligo Description Sequence 5 ˊ-3ˊ

OSA753 T7 forward primer for in vitro TAATACGACTCACTATAGGGAAAATGATGGATCTGATC translation of manY (-98 relative AGCAAAATCGATAAGT to the manY start codon, for ribosome footprint, and reverse transcription of DEPC-modified transcripts) OSA754 Reverse primer for in vitro AGCGGACGGTGAAACTGAAATTC transcription of manY (+98 relative to the manY start codon, for ribosome footprint, and reverse transcription of DEPC- modified transcripts) OSA786 Forward primer for ACCTGACGCTTTTTATCGCAACTCTCTACTGTTTCTCC recombineering sodB (+1, ATATACGCACAATAAGGCTATTGTACGTATGC transcription start site) into PM1205 OSA787 Reverse primer for TAACGCCAGGGTTTTCCCAGTCACGACGTTGTAAAACG recombineering sodB (+120, ACGTTCAGGTTAGTGACATAAGTCTGATGGT relative to the start codon) into PM1205 OSA070 T7 forward primer for in vitro TAATACGACTCACTATAGGGACGTAAAGGAGGAATTCA transcription and translation of TGAGCAAAGGAGAAGAACTTTTCACTG gfp OSA071 Reverse primer for in vitro TACGTTTATTTGTAGAGCTCATCCATGCCATGTG transcription and translation of gfp OSA632 Forward primer to PCR amplify CTTTAAGAAGGAGATATACCATGACTGAATCTTTTGCT rpsA CAACTCTTTGAAGAG

OSA633 Reverse primer to PCR amplify CTGCTGCCGCGCGGCACCAGCTCGCCTTTAGCTGCTTT rpsA GAAAGCTT

OSA636 Forward primer for pET28 TTGATCTTTTCTACGGGGTCTGACGCT

OSA637 Reverse primer for pET28 CTTCTTGAGATCCTTTTTTTCTGCGCGT

OSA779 Primer to introduce Y205A, GTTGATCTGGGCGGCGTTGACGGCCTGCTGGCTATCAC F208A, and H219A mutations in TGACATGGCCTGGAAACG rpsA OSA780 Primer to introduce Y205A, TCAACGCCGCCCAGATCAACGGCTGCACCGGCGTCAGT F208A, and H219A mutations in GAGGTTCTTAACGATACCTTTAACTTCC rpsA OSA774 Forward primer with a strong SD TTGTGAGCGGATAACAATTTTCACACATTGGAGGAACA to PCR amplify rpsA and mrpsA ATATGACTGAATCTTTTGCTCAACTCTTTGAAG (to construct pWKS30-SS1) OSA775 Forward primer with a weak SD TTGTGAGCGGATAACAATTTTCACACATTCCAGGAACA to PCR amplify rpsA and mrpsA ATATGACTGAATCTTTTGCTCAACTCTTTGAAG (to construct pWKS30-WS1) OSA776 Reverse primer to PCR amplify GTTGTAAAACGACGGCCAGTATCCGGATATAGTTCCTC rpsA and mrpsA (to construct CTTTCAGCA pWKS30-WS1) OSA783 Forward primer for pWKS30 (for ACTGGCCGTCGTTTTACAACG Gibson assembly) OSA784 Reverse primer for pWKS30 (for AAATTGTTATCCGCTCACAATTCCACACA Gibson assembly)

Table 2.1 (cont.)

Oligo Description Sequence 5 ˊ-3ˊ

OJH218 T7 forward primer for in vitro TAATACGACTCACTATAGGGGATGAAGCAAGGGGGTGC transcription of SgrS CCCATG

OJH169 Reverse primer for in vitro AAAAAAAACCAGCAGGTATAATCTGCTGGCGGG transcription of SgrS

CHAPTER 3

TRANSLATIONAL REGULATION BY BACTERIAL SMALL RNAS VIA AN UNUSUAL

HFQ-DEPENDENT MECHANISM

3.1 Introduction

In bacteria, the canonical mechanism of translational repression by small RNAs

(sRNAs) involves sRNA-mRNA base pairing that occludes the ribosome binding site

(RBS), directly preventing translation. In this mechanism, the sRNA is the direct regulator, while the RNA chaperone Hfq plays a supporting role by stabilizing the sRNA.

There are a few examples where the sRNA does not directly interfere with ribosome binding, yet translation of the target mRNA is still inhibited. Mechanistically, this non- canonical regulation by sRNAs is very poorly understood. Our previous work demonstrated repression of the mannose transporter manX mRNA by the sRNA SgrS, but the regulatory mechanism was unknown [110].

In this study, we explored the regulatory mechanism of two Hfq-dependent sRNAs, SgrS and DicF, that negatively regulate manX translation by an unconventional

mechanism. The physiological condition that triggers expression of DicF is unknown, but

SgrS is expressed during a metabolic state called glucose-phosphate stress [147].

Sugars are critical nutrients that fuel central metabolic pathways: glycolysis, the pentose

phosphate pathway, and the TCA cycle, to generate precursor metabolites needed to

synthesize nucleotides, amino acids, and fatty acids. Nonetheless, accumulation of

excess phosphorylated sugar intermediates, and their non-metabolizable derivatives can

be growth inhibitory [33, 71]. For instance, when cells are grown in the presence of non-

metabolizable sugar analogs, such as α-methyl glucoside ( αMG) or 2-deoxy-D-glucose

(2DG), it creates glucose-phosphate stress; glycolysis is inhibited, and cells stop

growing [63, 147]. Under these conditions, E. coli induces expression of SgrS, an Hfq- 31

dependent small RNA with regulatory activities that restore cell growth. When produced

under glucose-phosphate stress conditions, SgrS base pairs with mRNA targets to

regulate their translation and stability [13, 101, 110, 147]. One of the key activities of

SgrS during glucose-phosphate stress is repression of mRNAs encoding

phosphotransferase system (PTS) sugar transporters, ptsG [147] and manXYZ [109,

110]. This repression inhibits new synthesis of PTS transporters and reduces uptake of sugars that are not being efficiently metabolized during stress. We have shown that base pairing-dependent repression of transporter synthesis by SgrS is required for continued growth under stress conditions [135].

Many sRNAs that repress gene expression do so by inhibiting translation initiation by preventing ribosome binding to target mRNAs. Since bacterial translation initiation requires RNA-RNA base pairing between the 16S rRNA and the SD region, sRNAs typically base pair with a site close to the SD region and compete with the 30S ribosomal subunit [127]. Initial research on the interaction between mRNA and the ribosome suggested that initiating ribosomes occupy ~40 nt, from -20 nt in the 5´ UTR to

+19 nt into the coding region, where numbering is with respect to the start codon [10,

54]. A study using synthetic antisense oligonucleotides tested the boundaries to define the region where a base pairing interaction could prevent formation of the translation initiation complex (TIC). This study indicated that oligos base pairing within 15 nucleotides downstream of the start codon can inhibit TIC formation [17]. This led to the

“five codon window” hypothesis that proposed that if an sRNA base pairs with nucleotides comprising the first five codons of the mRNA, it can directly inhibit binding of the 30S ribosomal subunit and repress translation initiation. Interestingly, some studies have uncovered apparent exceptions to this hypothesis where sRNAs repress translation, either directly or indirectly, by base pairing outside of the five-codon window

[13, 28]. For example, the Massé group found that binding of the sRNA Spot 42 at a site 32

~50 nt upstream of the sdhC start codon. Their evidence suggested that the Spot 42 itself does not directly compete with the initiating ribosome, but instead may recruit the

RNA chaperone Hfq to bind near the sdhC ribosome binding site (RBS) and act as the primary repressor [28].

In this study, we investigated the mechanism by which SgrS regulates the first cistron of the manXYZ operon, manX . We observed previously that regulation of manX mRNA by SgrS involves base pairing 20 nt downstream of the start codon, which lies outside the 5 codon window [110]. We also characterized regulation of manX translation by another sRNA regulator, DicF [5], a 53-nt long Hfq-dependent sRNA [35]. The DicF binding site on manX mRNA is even further downstream than the SgrS binding site. We hypothesized that each of these sRNAs regulates manX translation by a “non-canonical” mechanism, since their binding sites are positioned too far downstream for sRNA-mRNA base pairing to directly occlude ribosome binding. To test this hypothesis, we addressed several questions. Does sRNA-mRNA duplex formation directly inhibit translation by preventing formation of the translation initiation complex? If not, then is Hfq required for translational repression? Does Hfq bind to the manX mRNA near the ribosome binding site?

Our results demonstrate that Hfq is absolutely required for translational repression mediated by SgrS and DicF in vivo. In vitro, Hfq, but not the sRNAs, can specifically inhibit formation of the TIC on manX mRNA. RNA footprints confirmed that

SgrS and DicF have distinct binding sites in the manX coding region, and both of these sRNAs facilitate Hfq binding at a site close to the RBS. Taken together, our data demonstrate sRNAs mediate regulation of manX translation by a non-canonical mechanism involving recruitment or stabilization of Hfq binding at a site where it can directly interfere with translation.

3.2 Results

3.2.1 Hfq is essential for translational repression of manX

Previously, we identified manXYZ mRNA, which encodes a mannose and

(secondary) glucose transporter (EII Man ), as a target of SgrS [109, 110]. The polycistronic

manXYZ mRNA is negatively regulated post-transcriptionally via two independent SgrS-

manXYZ mRNA base pairing interactions [109]. The physiological outcome of this

regulation is repression of EII Man transporter synthesis, which helps rescue cell growth during glucose-phosphate stress [110]. One of the SgrS binding sites was mapped to the coding region of manX , and this binding site was shown to be necessary for translational repression of manX [109, 110]. Although we have identified the base pairing sites for

SgrS on the manXYZ transcript and established that translational regulation of this target

is important for cell growth during glucose-phosphate stress, the exact mechanism of

SgrS-mediated manX translational repression is unknown. The SgrS binding sites on the manXYZ mRNA are too far from the RBS of the manX and manY cistrons [109] to directly occlude ribosome binding (Fig. 3.1). We hypothesized that manX translation might be repressed by a non-canonical mechanism where Hfq serves as the direct repressor of translation while SgrS plays an accessory role, perhaps recruiting Hfq to bind stably near the SD.

Fig. 3.1: Base pairing between A) SgrS and a canonical target ptsG, and B) SgrS and a non-canonical target manX. SgrS base pairs on the ribosome binding site shielding it from the initiating ribosome. SgrS, on the other hand, base pairs downstream into the manX coding region.

To test this hypothesis, we first investigated whether translational repression of

two different SgrS targets, ptsG and manX, was dependent on Hfq in vivo. Aiba and

coworkers have already shown that SgrS can inhibit translation of ptsG in the absence of

Hfq [76], consistent with the canonical model for repression where the sRNA pairs near

the TIR and directly occludes ribosomes. Here, we utilized manX ′-′lacZ and ptsG ′-′lacZ translational fusions that we have already demonstrated are good reporters for SgrS- dependent regulation of these targets [110]. We monitored fusion activities after SgrS production was induced in wild-type and Δhfq backgrounds. Since stability of E. coli

SgrS (SgrS Eco ) is greatly reduced in the absence of Hfq [99], we utilized the Salmonella

SgrS allele (SgrS Sal ) which is more stable than SgrS Eco in the Δhfq background and has very similar seed region that allows it to efficiently regulate E. coli SgrS target mRNAs

([6] and Fig. 3.2). In agreement with our previous study, we found that SgrS Sal can

complement a ΔsgrS mutant for regulation of manX (Fig. 3.3A) [6]. Consistent with the

direct ribosome occlusion mechanism for translational regulation of ptsG, SgrS Sal 35

efficiently repressed the ptsG ′-′lacZ fusion in both the wild-type and Δhfq backgrounds

(Fig. 3.3B). In contrast, while SgrS efficiently repressed manX in a wild-type background, it failed to repress the fusion in an hfq mutant background, even at high levels of inducer

(Fig. 3.3C). These data indicate that SgrS cannot regulate manX in vivo in the absence of Hfq.

Fig. 3.2: A) SgrS Eco and B) SgrS Sal do not share detectable sequence similarity. However, for manX , the seed regions and the anti-seed region these two homologs base pair with are almost identical.

Fig. 3.3: A) Both Salmonella SgrS and E. coli SgrS can repress manX translation. A strain (JH175) with in locus manX’-‘lacZ translational fusion, under the control of

Fig. 3.3 (cont.)

heterologous Cp-19 promoter. Both SgrS Sal and SgrS Eco were induced with a final concentration of 0.1 mM IPTG. The samples were grown for another 60 min and assayed for β-galactosidase activity. B) Translation repression of a canonical target in SgrS targetome, ptsG, can be achieved by SgrS Sal in the absence of the pleotropic chaperone protein Hfq. β-Galactosidase activities in the samples were normalized to the levels the vector control strains to yield percentage relative activity. C) In the absence of Hfq, plasmid borne SgrS Sal , even with 0.5 mM IPTG (final concentration), SgrS cannot repress manX translation in the absence of Hfq.

If the role of SgrS in manX regulation is to recruit or enhance binding by the

putative primary or direct regulator Hfq, it follows that Hfq should bind close to the manX

TIR. To determine if there were elements in the 5 ′ UTR, such as a putative Hfq binding

site, that were required for translational regulation, we constructed a series of manX

translational fusions with truncations in the 5 ′ UTR (Fig. 3.4A). Activity of these fusions

was measured in the presence and absence of ectopically expressed sgrS . The

constructs contained 65, 30, 25, and 22 nt of the manX 5′ UTR region, and in the hfq +

background, all four fusions showed a similar pattern of regulation compared to the

construct containing the full-length 115-nt manX 5′ UTR (Fig. 3.3A). Further truncation of the manX 5′ UTR was not possible without interfering with the RBS. Nevertheless, the

fact that all truncated fusions were regulated similarly to wild-type suggested that the

putative Hfq binding site resided downstream of the 5 ′ boundary defined by the 22-nt

fusion (Fig. 3.4A). An A/U-rich motif just upstream of the manX RBS is similar to the motif that was shown in other studies to be preferentially bound by Hfq [38, 121]. To test the role of this motif in the regulation of manX translation, we constructed a mutant manX fusion where the A/U-rich motif was converted to a C/G-rich motif (Fig. 3.4B). In contrast with the wild-type manX fusion, which was efficiently repressed when SgrS was

ectopically expressed, SgrS failed to alter translation of the mutated fusion (Fig. 3.4B). It

is important to note that the A/U-rich putative Hfq binding motif is far upstream from the

38 known SgrS base pairing site (Fig. 3.4B). Thus, the loss of regulation in the mutant suggests that regulation of manX translation is not directly mediated by SgrS base pairing, but may instead involve Hfq binding to a site near the manX RBS.

Fig. 3.4

Fig. 3.4 (cont.)

Fig. 3.4: Genetic localization of hfq binding site A) Wild type manX UTR in E. coli is 115 nt. Four translation fusions with decreasing lengths of UTR were constructed where the heterologous promoter was moved closer and closer to the TIR. B) An A/U rich motif, right upstream of the manX RBS was mutated (mut-1, CGGCGGGA) and fused with lacZ (see text). In a hfq+ background, two lacZ translational fusion fusions- a 25 nt WT UTR and a mutated UTR, were assayed for β-galactosidase activity after SgrS Eco expression from a plasmid. The specific activities, quantified in Miller units, were normalized to the vector control to yield relative activity values. The RBS is in blue letters, and the manX start codon is in green. C) Beta galactosidase assay of fusions of 25 nt UTR-manX´-´lacZ with two nucleotides altered in the Hfq binding motif. The mutated Hfq motifs are shown below the X-axis.

If this A/U rich motif is the true Hfq binding site, we envisaged that abrogation of translation repression should also be observed with fewer nucleotide changes. We tested this prediction by constructing four strains, each harboring two nucleotide mutations (Fig. 3.4C). We noticed that the Hfq binding motif is too close to the RBS region and mutations in this motif is likely to change the expression and/or stability of the reporter gene. As expected, these two nucleotide mutations were sufficient to eliminate

Hfq binding, and we observed loss of SgrS mediated regulation in these mutated constructs (Fig. 3.4C). Interestingly, under two circumstances we noticed a drop in the basal activities of the lacZ reporter fusion- i) when hfq was deleted (Fig. 3.3C), and, ii) when the Hfq binding site was mutated (Fig. 3.4B, 3.5A).The Hfq binding site is next to the RBS, and it is not surprising that mutations in this motif could cause altered translation rate. If that is the case, then it is crucial to demonstrate that the loss of regulation results from the lack of Hfq binding and not due to low translation rate. To investigate this, we assayed beta-gal activities of the lacZ fusions containing two nucleotide mutations in the Hfq binding motif (Fig. 3.5B). Two constructs, containing

ATAATAGG and ATGGTAA motifs, showed similar basal reporter activity as the wild type (Fig. 3.5B) and these two mutations also abolished SgrS mediated repression (Fig.

3.4C). These data suggest that the lack of Hfq binding, not the altered translation rate of the mutant, abrogates SgrS mediated regulation of the fusions. Next, we asked whether, in the absence of hfq, the drop in the fusion activity is the outcome of reduced stability of the transcript. To test this possibility, we measured the lacZ fusion activities of the constructs mentioned above in the rne131 background. Interestingly, the beta gal activity does not drop when the strains harbored hfq::cat allele suggesting that in the absence of

Hfq, manX transcript becomes unstable in a RNase E-dependent manner (Fig. 3.5B).

This is an exciting finding, and we plan to investigate the effect of the RNase E-Hfq relationship on the stability of manX transcript in the future.

Fig. 3.5: Hfq impact translation and stability of manX mRNA. A) The basal β-gal activity of the manX´-´lacZ fusions with WT (ATAATAA) and mutated binding motifs, in presence or absence of Hfq. B ) The beta galactosidase activities of the WT and the mutated motif containing manX´-´lacZ fusions in rne131 background.

3.2.2 Hfq inhibits formation of the translation initiation complex on manX mRNA

If Hfq is the direct repressor of manX translation it should compete with the ribosome for binding to the mRNA in vitro. We used toeprinting assays [36] to test whether Hfq or SgrS could directly inhibit translation initiation. In a toeprinting assay, stable binding of the 30S ribosomal subunit and tRNA fMet to the RBS blocks a primer

extension reaction and produces a product with a characteristic size. Since native manX

has a weak GTG start codon that does not stably associate with commercially available

preparations of 30S ribosomes, we changed the start codon to the canonical ATG to

ensure strong initiation complex formation in vitro . We showed previously that this

construct, manX ATG , was efficiently repressed by SgrS [110]. The toeprint assay was

32 fMet performed by mixing manX ATG mRNA, P end-labeled primer, ribosomes, and tRNA in the presence and absence of Hfq. Reverse transcriptase was then added to begin the primer extension reaction. In the positive control reaction, we saw the characteristic toeprint signal caused by termination of reverse transcription at position +15/+16 relative to the start codon (Fig. 3.6A). With the addition of increasing concentrations of Hfq, the formation of the TIC was completely inhibited (Fig. 3.6A and 3.6B). However, when increasing concentrations of SgrS were added in the absence of Hfq, TIC formation was unperturbed (Fig. 3.6B). When the same concentrations of SgrS was added to a toeprint reaction using ptsG as a template, we observed inhibition of the toeprint signal (Fig.

3.6C). These results are consistent with the in vivo studies, and add further evidence

supporting our hypothesis that Hfq itself directly inhibits manX translation at the initiation

stage.

Fig. 3.6: Repression of the manX translation initiation complex formation by Hfq, lane 4, 5, and 6 (0.15 µM, 0.5 µM, and 1 µM). A) Hfq alone, in the absence of SgrS, can prevent manX translation initiation complex formation. Sequencing ladders, indicated by TGCA, were generated with the same oligo OJH119. The toeprint signal is indicated +15/16 relative to the start codons. B) Increasing amount of hfq was able to inhibit the initiation complex formation, whereas SgrS was not (100 nM, 250 nM, and 500 nM). Addition of ribosome and initiator tRNA is indicated by ‘+’ symbol. C) Repression of the ptsG translation initiation complex formation by SgrS, lane 4, 5, and 6 (100 nM, 250 nM, and 500 nM respectively). SgrS alone is able to prevent ptsG translation initiation complex formation. Sequencing ladders, indicated by TGCA. The toeprint signal is indicated +16 relative to the start codons.

3.2.3 manX is regulated by DicF sRNA

In a previous study, manX was identified as a putative target of another sRNA,

DicF [5]. To further investigate the regulation of manX by DicF and determine the regulatory mechanism, we monitored the activity of a manX'-'lacZ translational fusion

(under the control of a constitutive promoter to rule out indirect effects on manX transcription) in control cells and cells where DicF was ectopically expressed. Cells expressing dicF showed ~40% reduced β-galactosidase activity compared to control 44

cells (Fig. 3.7A). Compared to SgrS, which reduces manX translation by about 70% (Fig.

3.4B), DicF is a rather weak regulator.

Previous studies have demonstrated that sequences at either the 5 ′ or the 3 ′ end

of DicF can base pair with mRNA targets [5, 141]. We identified a potential base pairing

interaction between the 3 ′ end of DicF and the coding region of manX just downstream

of the known SgrS binding site (Fig. 3.7B) . To test this base pairing prediction, we made

a mutation in nucleotides of DicF that should disrupt the base pairing interaction (Fig.

3.7B). This mutant allele, dicF20 , lost the ability to regulate the manX'-'lacZ translational

fusion (Fig. 3.7C), consistent with the base pairing prediction. If DicF also base pairs

within the manX coding sequence, well outside the window that would allow direct

interference with ribosome binding, then like SgrS, DicF may also repress manX translation by influencing Hfq binding in the manX TIR. To test whether DicF-mediated regulation requires the putative Hfq binding site near the RBS, we constructed a mutant version of the manX'-'lacZ fusion (containing the putative DicF base pairing site) where the A/U-rich region next to the RBS is changed to G/C-rich ( mut6 ). In contrast with the wild-type manX fusion, which was repressed upon dicF expression, activity of the mut6 fusion with the mutation in the putative Hfq binding site was not altered by DicF (Fig.

3.7A). This observation is consistent with the model that DicF-mediated regulation of manX is similar to SgrS in that it requires Hfq binding proximal to the RBS where it acts as the direct translational repressor.

Fig. 3.7: DicF, a prophage encoded sRNA, also regulates manX translation. A) β- Galactosidase assays using manX’-‘lacZ fusions with WT (JH175) and mutated Hfq binding site containing UTRs (SA1620) under the control pf pBAD promoter. The specific activities were normalized to the corresponding empty vector control values to yield percent relative activity. B) The pairing sites of SgrS and DicF on the manX mRNA. Both sRNAs pair into an overlapping region downstream of the manX start codon. C) Mutations in DicF ( dicF 20 ) disrupt complementarity and regulation.

3.2.4 Hfq binds next to the manX ribosome binding site

We predicted that DicF base pairs at a site just downstream of the SgrS binding site, from residues G145 to C162 (Fig. 5B). Our genetic analyses suggest that Hfq binds in the 5’ UTR just upstream of the RBS to act as the direct repressor of manX translation for both SgrS- and DicF-mediated regulation (Figs 3.4B and 3.7A). To further test this prediction, we performed in vitro footprinting experiments with labeled manX RNA to identify the Hfq binding site(s) occupied in the presence of each individual sRNA. As we showed previously, SgrS protects its binding site from C139 to G152 on manX mRNA even in the absence of Hfq, and the addition of Hfq does not change the footprint (Fig.

3.8A) [110]. Notably, SgrS alone does not affect the structure around the RBS or start

codon. Consistent with our prediction (Fig. 3.8B), DicF protects manX mRNA from G150

to C167 in the absence and presence of Hfq. Again, DicF only impacted the reactivity of

nucleotides comprising its binding site in the manX coding region, and the structure

upstream in the TIR was unaffected. In the presence of either sRNA, Hfq clearly

protected manX mRNA nucleotides A97-A103 (Fig. 3.8A). Note that this region is the

same A/U-rich region that we predicted as the Hfq binding site (Fig. 3.8B) and that when

mutated, prevented SgrS- and DicF-dependent regulation (Figs. 3.4B and 3.7A,

respectively). Additionally, we observe some weak protections for residues U91-C95.

These findings demonstrate that Hfq binds at precisely the same location on manX

mRNA, adjacent to the RBS, regardless of which sRNA is present in the sRNA-mRNA-

Hfq ternary complex.

Now, this leads to a broader question regarding the nature of the secondary role

played by the sRNA when the regulation happens in a role-reversal fashion. Ectopically

expressed Hfq does not inhibit manX translation in the absence of the sRNA partner

(data not shown). We envisaged that the presence of the sRNA partner could be

essential if it functions as a guide to change the affinity of Hfq for the manX transcript. To test this possibility, we performed the footprint with a low concentration of Hfq in presence and absence of SgrS, and in agreement with our conjecture, we observed the protection by Hfq only when SgrS was present in the reaction (Fig. 3.8C). We used a mutated manX transcript where the binding motif was changed to CGGCGGG, and we did not observe any Hfq mediated protection, even in the presence of SgrS confirming our genetic data that indicated this RBS-adjacent motif as the Hfq binding site (Fig.

3.8D).

Fig. 3.8

Fig. 3.8 (cont.)

Fig. 3.8: Footprinting to map SgrS, DicF and Hfq binding sites on manX RNA. A) In vitro transcribed manX mRNA containing 115 nt UTR and the coding region containing 51 bases past the predicted DicF base pairing region was end labeled with P 32 and incubated with unlabeled SgrS and Hfq to perform footprinting reactions. T1 indicates RNase T1 ladder; OH, alkaline ladder; PbAc, lead acetate. Interaction between DicF: manX and Hfq-manX were mapped as described above. B) Putative structure of manX following interaction with Hfq and SgrS or DicF. C-D) Footprinting to investigate Hfq recruitment in presence or absence of SgrS with WT(ATAATAAA) Hfq binding motif and the mutated binding motif (CGGCGGGA).

3.2.5 DicF is a weaker regulator of manX

Compared to DicF, SgrS is a stronger repressor of manX translation (compare

repression in Fig. 3.4B to Fig. 3.7A). Our data suggest that each of these sRNAs

mediates translational regulation indirectly, via facilitating or stabilizing Hfq binding to a

site in the 5 ′ UTR adjacent to the RBS (Fig. 3.8A). To explore the basis for the different

efficiencies of regulation, we conducted experiments to measure the affinity of sRNA-

mRNA interactions and sRNA-Hfq interactions. We reasoned that differences in the

binding affinity of the sRNAs for manX mRNA and/or Hfq, could be important

determinants of regulatory efficiency for each sRNA-mRNA-Hfq interaction.

Electrophoretic mobility shift assays (EMSAs) were used to measure the specific binding

of SgrS and DicF individually to manX mRNA in vitro. We found that SgrS base paired

with manX mRNA, with a dissociation constant KD of 4.53 µM (Fig. 3.9A). DicF

interacted less strongly with manX mRNA, with a KD of 21.8 µM (Fig. 3.9A). The higher

KD for DicF-manX mRNA is consistent with our observation that compared to SgrS, DicF

is a weak repressor of manX translation. In a previous study, we found that in vivo, the

KD for SgrS binding to full-length manXYZ mRNA (with both manX and manY binding sites) was 2.3 µM [37]. Thus, our in vitro measurement is in good agreement with the in vivo data for SgrS.

EMSAs to monitor interactions of each sRNA with Hfq also revealed differences between SgrS and DicF. The Hfq-SgrS interaction was relatively strong, with a calculated KD of 3.37 nM (Fig. 3.9B). Hfq bound DicF less tightly with a calculated KD of

22.0 nM (Fig. 3.9B). The dissociation constant values we calculated for Hfq and SgrS or

DicF are in a similar range as those reported previously for the binding of Hfq to OxyS,

RyhB, DsrA, and Spot 42 sRNAs [42, 89, 126, 166]. Taken together, our data suggest

that SgrS is a more efficient regulator of manX translation than DicF and that differences

in sRNA-mRNA binding interactions and sRNA-Hfq interactions could influence the

efficiency of regulation.

Fig. 3.9: A) Native gel electrophoresis assay to examine the difference in binding of the manX mRNA with SgrS and DicF sRNAs.In vitro transcribed labeled manX mRNA (0.01 pmol) was mixed with indicated amount of cold SgrS and incubated at 37 0C for 30 min. The reaction mixture was resolved on a chilled native acrylamide gel. B) Gel mobility shift assay for SgrS and DicF. SgrS has higher affinity for Hfq. Measured band densities (n replicates, top left) were plotted to determine the dissociation constants.

3.3 Discussion

Our study shows that two sRNAs, SgrS and DicF, base pair with manX mRNA at distinct sites in the coding region, outside the window that would allow translational repression via a direct ribosome occlusion mechanism. Instead, we propose that Hfq acts as the direct regulator of manX translation and that the sRNAs play an accessory or secondary role. This model is supported by multiple lines of evidence presented in this study. In vivo, SgrS cannot repress manX translation in the absence of Hfq (Fig. 3.3C).

This is in contrast with regulation of another SgrS target, ptsG , which is known to occur

via the canonical (direct) mechanism of translational repression [59, 147]. SgrS

efficiently represses ptsG translation in an hfq mutant background (Fig. 3.3B). Loss of

regulation by both sRNAs is seen in an hfq+ background when the Hfq binding site

upstream of the manX RBS is mutated (Figs 3.4B, 3.7A). Structural analyses clearly demonstrate that SgrS and DicF bind to sites in the coding region of manX mRNA and have no impact on the structure near the TIR. In contrast, in the presence of either SgrS or DicF, Hfq binds to the same site on manX mRNA, directly adjacent to the RBS (Fig.

3.8A). The mapped Hfq binding site is also in agreement with previously published high- throughput studies [81, 143]. Differences in the relative strength of manX translational regulation promoted by SgrS and DicF were correlated with the strength of sRNA-mRNA and sRNA-Hfq interactions (Fig. 3.9A, B). Collectively, our data are consistent with a non-canonical mechanism of regulation where the sRNAs play a guide-like role in regulation by promoting Hfq binding to a site near the manX RBS so that Hfq itself

directly interferes with ribosome binding (Fig. 3.10). This model is in stark contrast to the

canonical model of bacterial sRNA-mediated translational repression where sRNAs are

the direct competitors of ribosome binding, while the chaperone Hfq assumes the

secondary role. Here, the chaperone Hfq swaps its role with the RNA partner.

Fig. 3.10: A model for the non-canonical roles played by two distinct sRNAs, SgrS and DicF, to repress manX translation in a Hfq dependent mechanism.

A recent study reported a non-canonical mode of translation regulation where the sRNA Spot 42 binds its target sdhC mRNA far upstream of the RBS. Spot 42 itself

cannot compete with the incoming ribosome for binding to sdhC mRNA, but instead

seems to facilitate binding of Hfq to a site near the TIR in order to block translation

initiation [28]. Another study by Blasi and colleagues also reported that in the absence

of any sRNA, Hfq causes translation repression of ompA mRNA where Hfq competes with the 30S ribosomal subunit [150]. Here, we show that sRNAs can employ similar mechanism and recruit Hfq to bind near the RBS even if they base pair downstream in the mRNA coding region. Footprinting of manX mRNA with SgrS, DicF, and Hfq

53 revealed that SgrS base pairs with nucleotide +24 to 36 on the manX transcript, while

DicF base pairs little downstream, with nucleotides +34 to +51, but both sRNAs recruit

Hfq at the same location near RBS (Fig. 3.8). In other words, the identity and the location of base pairing sites do not seem to be the determining factor for non-canonical sRNA-based regulation effected by Hfq binding near the TIR. We propose that the sRNAs carry out the task of substrate recognition that subsequently allows the protein partner to be recruited at a particular binding site (Fig. 3.8C). Similar mechanisms are widely utilized by prokaryotic CRISPR guide RNAs, and eukaryotic non-coding RNAs, including small interfering RNAs, microRNAs, and small nucleolar RNAs. All of these types of RNAs act as a part of a ribonucleoprotein complex where the RNA component recognizes the substrate nucleic acid while the protein component performs catalysis.

Regulation of manX translation by DicF, an sRNA encoded on the cryptic prophage Qin on the E. coli chromosome, was confirmed in this study. Like other small

RNAs encoded on horizontally acquired genetic elements like prophages and pathogenicity islands, DicF is poorly characterized. However, research over the last decade, suggests that horizontally-acquired sRNAs are crucial regulators of bacterial physiology, growth, and stress responses [49, 100, 105]. For instance, the sRNA InvR, encoded in Salmonella pathogenicity island 1, is a major regulator of outer membrane porin OmpD [103]. IpeX, an sRNA encoded on the cryptic prophage DLP12 in E. coli , is a regulator of outer membrane porins, OmpC and OmpF [20]. DicF was identified in the

1980s when it was observed to cause a filamentation phenotype when expressed from a multi-copy plasmid [16]. We recently demonstrated that DicF directly regulates translation of mRNA targets encoding diverse products involved in cell division and metabolism [5]. These include mRNAs encoding the tubulin homolog FtsZ, xylose uptake regulator XylR, and pyruvate kinase PykA [5, 141]. Our current study extends the

DicF targetome to include manX . Though SgrS and DicF share a common target in 54

manX , these sRNAs are not expressed under the same conditions. We did not see DicF expression when cells were challenged with αMG or 2DG (data not shown). Under standard laboratory growth conditions, the dicBF operon is not expressed, and we do not

yet know the signal that triggers the expression of this operon. Further research aimed at

uncovering the physiological conditions stimulating DicF production may provide insight

into the biological role of DicF-mediated manX regulation.

CHAPTER 4

TRANSLATION INHIBITION FROM A DISTANCE: SgrS SILENCES A TRANSLATION ENHANCER IN THE LEADER REGION

4.1 Introduction

Microbes respond to changes in environmental conditions using a variety of

mechanisms to modify gene expression resulting in changes in cell structure and

function. Small RNAs (sRNAs) are found in microbes across the tree of life and are

major regulators responsible for post-transcriptional control of gene regulation. sRNAs

regulate target mRNA translation and stability by a variety of molecular mechanisms that

depend on sRNA-mRNA duplex formation. Early studies characterizing sRNA-mRNA

interactions reported sRNA base pairing with sequences around the Shine-Dalgarno

(SD) sequence [86, 119]. In vitro structure probing of sRNA-mRNA duplexes and toeprinting assays probing translation initiation complex formation [2, 17, 42, 89], have revealed that a common mechanism of sRNA-mediated translational repression is steric occlusion of ribosome binding. However, a growing number of examples of translational repression by sRNAs involve base-pairing interactions far from the SD sequence [3, 28,

52, 123]. Several mechanisms have been reported for regulation of translation via sRNA-mRNA duplex formation distant from ribosome binding sites. Th90e first involves sRNA binding at a site upstream or downstream of the translation start resulting in recruitment of the RNA chaperone Hfq to bind at a site near the SD so that Hfq occludes ribosome binding [3, 28]. Another mechanism involves sRNA sequestration of CA-rich sequences that act as translation enhancer elements [123, 162]. The sRNA GcvB represses translation of several mRNAs using a GU-rich seed region that base pairs with

CA-rich target sites located at variable distances upstream of the SD. In the absence of

GcvB, the CA-rich sequences enhance mRNA translation by an unknown mechanism.

With GcvB present, these enhancer sequences are sequestered and translation of 56

mRNA targets is reduced. A third mechanism of sRNA-mediated translational repression

from a distance involves ribosome standby sites. Certain mRNAs with stable secondary

structures around the ribosome binding site use upstream ribosome standby sites to

promote translation. Ribosome standby sites are located in single-stranded regions

where a 30S ribosomal subunit can bind and kinetically compete with RNA folding to

access the downstream translation initiation region [25, 128, 131]. The toxin-encoding

tisB mRNA has a ribosome standby site far upstream of the SD, and this site can

promote translation of tisB only in the absence of an antisense sRNA, IstR, which

sequesters the standby site [24, 114]. These examples illustrate the many mechanisms

of translational repression mediated by sRNAs binding to mRNAs within and outside the

translation initiation region.

The rate-limiting step in protein synthesis is generally considered to be binding of

the small ribosomal subunit to the mRNA upstream of the start codon to form the

translation initiation complex [64, 84]. In most cases, the mRNA leader region contains a

purine-rich SD sequence [124] just upstream of the start codon that forms a short duplex

with the 3'-end of the 16S rRNA (anti-SD sequence). Some leader regions may contain

additional sequence features that can also determine the rate of translation initiation

[85]. Pyrimidine-rich sequences that are sometimes present immediately upstream of the

SD can interact with the 30S subunit ribosomal protein S1 and act as translational

enhancers [14, 15, 31]. For these mRNAs, formation of the preinitiation complex involves

RNA-RNA interactions and RNA-protein interactions that both contribute to the efficiency

of translation initiation [31, 78, 137]. sRNAs could interfere with translation initiation by

disrupting any of the interactions – RNA-RNA or RNA-protein – that are important for

formation of the translation initiation complex.

In this study, we aimed to define the molecular mechanism by which an E. coli sRNA, SgrS, regulates translation from a distance. SgrS represses translation of all the 57 cistrons encoded by the manXYZ mRNA, which encodes the mannose PTS transporter

[109, 110]. SgrS binds to two distinct sites on manXYZ mRNA [3, 109, 110]. The first site is located within the manX coding sequence, and we showed recently that SgrS represses translation of manX via a non-canonical mechanism involving SgrS- dependent recruitment of Hfq to a binding site that overlaps the manX SD sequence – making SgrS the chaperone-like partner and Hfq the direct repressor of translation [3].

SgrS base pairs at a second site in the manX-manY intergenic region, 30 nucleotides upstream of the manY start codon (Fig. 4.1A). SgrS binding at this site represses manY and manZ translation ( manZ translation is coupled to that of manY ) by an unknown mechanism [109]. The manY binding site is outside the -20 to +20 (relative to the start codon) region protected by the translation initiation complex [10, 17, 54] suggesting that the mechanism of regulation by SgrS is not the canonical mechanism of occlusion of the

SD sequence.

Here, we probed the roles of Hfq, SgrS and the ribosome, particularly ribosomal protein (r-protein) S1, in modulating the translation of manY . We found that Hfq is required for SgrS-dependent regulation of manY , not only because it stabilizes SgrS [6], but also because it promotes SgrS-manY mRNA duplex formation. We discovered that the AU-rich sequences that comprise the SgrS binding site upstream of manY act as a translational enhancer. The enhancer promotes higher levels of translation independent of whether the SD is strong or weak. We performed genetic and biochemical experiments that provide evidence for ribosomal protein S1 binding to the enhancer sequence to make the manY SD more accessible. Our data are consistent with the model that manY translation is controlled by an r-protein S1-dependent enhancer, and

SgrS represses translation by interfering with S1 binding to the enhancer sequence, reducing translation efficiency.

Fig. 4.1: SgrS-dependent regulation of manY translation requires Hfq . A) A model showing the two SgrS binding sites on manXYZ mRNA. B) SgrS base pairing at the site in the manX-manY intergenic region, 30 nt upstream of the manY start codon. C) Wild- type (SA1747) and hfq::kan (SA1907) strains harboring a manY'-'lacZ translational fusion (under the control of a P BAD promoter) were transformed with a vector or plasmid containing one of two SgrS orthologs – SgrS Eco from E. coli , or SgrS Sal from Salmonella . Cells were grown and β-Galactosidase assays were performed as described in the Materials and Methods. Error bars represent standard deviation from three biological replicates. P values (unpaired t test) are shown as: *, P < 0.05; **, P < 0.005; ***, P < 0.0005;****, P < 0.0001; NS, not significant. 59

4.2 Results

4.2.1 Hfq is essential for translation repression of manY

During glucose-phosphate stress, translation of the manXYZ operon is repressed

by SgrS via base pairing interactions at two distinct sites on manXYZ mRNA (Fig. 4.1A)

[3, 109, 110]. The base pairing interaction between SgrS and the manX-manY intergenic

region (Fig. 4.1A, 4.1B) is 30 nucleotides upstream of the manY start codon [109].

Previous studies [10, 17, 54] have suggested that base pairing interactions outside the -

20 to +20 window (relative to the start codon) cannot regulate translation by steric

occlusion of ribosome binding, so the precise mechanism of SgrS-mediated translational

repression of manY has remained unknown. To investigate the molecular details of the

SgrS-manY interaction, we first looked at the role of the RNA chaperone Hfq [68, 89,

166]. For these experiments, we used a manY ʹ-ʹlacZ translational reporter fusion

construct described previously [110] that contains only the SgrS binding site in the

manX-manY intergenic region (Fig. 4.1C). We monitored activity of the fusion in wild-

type and Δhfq host strains carrying SgrS expression plasmids. Previously, we showed

that the stability of E. coli SgrS (SgrS Eco ) is reduced in the absence of hfq , but

Salmonella SgrS (SgrS Sal ), which has a similar seed region and demonstrates

comparable effectiveness in regulating targets in E. coli , is more stable in a Δhfq

background [6]. Consistent with our previous studies [3, 6], in a wild-type host, both

SgrS Eco and SgrS Sal repressed manY ʹ-ʹlacZ fusion activity to less than 50 % of the activity in vector control cells (Fig. 4.1C). In the Δhfq background, the basal level of manY ʹ-ʹlacZ

activity was reduced, and neither SgrS Eco nor SgrS Sal repressed translation of manY ʹ-

ʹlacZ further (Fig. 4.1C).

To test whether Hfq is required to promote annealing between SgrS and manY mRNA, we performed footprinting reactions in the presence and absence of Hfq. End- labeled manY transcripts incubated with and without SgrS and Hfq were subjected to 60 enzymatic (with RNase T1) and chemical (lead acetate) digestion. We performed an

SgrS-manY footprinting experiment by mixing denatured end-labeled manY mRNA and unlabeled SgrS (with or without Hfq), followed by incubation under conditions that allow for annealing and then digestion. Using this procedure, we observed SgrS-dependent protection in the absence of Hfq (Fig. 4.2A, compare lanes 3 and 4) and in the presence of Hfq (Fig. 4.2A, lanes 5 and 6). The protection covered residues -30C to -44A (Fig.

4.2A, numbering relative to start codon), with hypersensitivity of the three U residues (-

36U to -38U) in the center of the base pairing interaction that are not involved in duplex formation (Fig. 4.1B). These results are perfectly consistent with the SgrS binding site on manY mRNA that we previously mapped using genetic methods [109].

Fig. 4.2: Hfq promotes duplex formation between SgrS and manY mRNA. A) Footprinting assays were performed using in vitro transcribed labeled manY mRNA mixed with SgrS and then denatured prior to addition of binding buffer and varying concentrations of Hfq. Lane 1 – “T1” – contains the G-ladder generated by

Fig. 4.2 (cont.)

RNase T1 digestion, Lane 2 – “OH” – contains the ladder generated by alkaline hydrolysis, and lanes 3-9 – “PbAc” – are lead acetate digested samples. Nucleotides protected by SgrS are labeled on the right. B) Footprinting assays were performed using in vitro transcribed – non-denatured – labeled manY mRNA mixed with SgrS and varying concentrations of Hfq. Lanes are labeled as described for part A. C) EMSAs using native gel electrophoresis were performed with P 32 labeled manY and unlabeled SgrS that were hybridized in a binding buffer without prior denaturation. The samples were resolved in a chilled native acrylamide gel. Error bars represent standard deviation from (n=2) replicate experiments. D) EMSAs for SgrS and manY in the presence of Hfq. Hfq was removed by phenol extraction prior to running samples on a chilled native acrylamide gel. Error bars represent standard deviation from (n=3) replicate experiments. For both C and D, the measured band densities were plotted using the GraphPad Prism software to calculate the dissociation constants.

We next conducted footprinting experiments where SgrS and manY RNAs were not denatured prior to annealing and chemical probing (Fig. 4.2B). Under these conditions, we did not see SgrS-mediated protection of manY mRNA in the absence of

Hfq (Fig. 4.2B, compare lanes 3 and 4) or when Hfq was present in the annealing

mixture at lower concentrations (Fig. 4.2B lanes 5 and 6). In contrast, when Hfq was

present at a higher concentration, we observed SgrS-dependent protection of manY

mRNA over the known binding site – from residues -30C to -44A (Fig. 4.2B, lane 7).

When reactions were performed using a limiting amount of Hfq and increasing amounts

of SgrS (Fig. 4.2B, lanes 9, 10, and 11), we did not see protection. Thus, the

requirement for Hfq to promote SgrS-manY mRNA interactions under these conditions

could not be overcome by increasing concentrations of SgrS.

The data presented so far suggest that Hfq facilitates intermolecular SgrS-manY mRNA base pairing. To corroborate this finding, we conducted a set of electrophoretic mobility shift assays (EMSAs) to quantify SgrS-manY RNA interactions in the presence and absence of Hfq. For these experiments, in vitro transcribed manY and SgrS RNAs were mixed in a binding buffer without a prior denaturation step. We found that in the absence of Hfq, SgrS formed a complex with manY mRNA with a KD of 7.1 µM (Fig.

4.2C). When end-labeled manY RNA was incubated with increasing amounts of SgrS

and equimolar concentrations of Hfq (which was subsequently removed by phenol

extraction before the samples were resolved in native polyacrylamide gels), we obtained

a KD of 0.7 µM, 10-fold lower than in the absence of Hfq (Fig. 4.2D). Altogether, the data

suggest that Hfq promotes SgrS-mediated regulation of manY translation by facilitating

SgrS-manY duplex formation.

4.2.2 The manY translation initiation region contains a ribosome-dependent enhancer

SgrS acts as a translational repressor by binding at a site upstream of the two

major determinants of ribosome binding – the Shine Dalgarno (SD) and initiation codon.

We hypothesized that the region bound by SgrS might be a translation enhancer, similar

to those described in mRNA targets of other sRNAs [123, 162]. To test the role of the

putative enhancer located within the SgrS binding site, we constructed a set of lacZ

translational fusions. These fusions were driven by a heterologous promoter (P BAD ), and contained the manX-manY intergenic region, including the SgrS binding site (Fig. 4.3A, green region). The wild-type fusion had a high basal level of activity at ~4000 Miller Units

(Fig. 4.3A, “WT”). The fusion lacking the SgrS binding site and putative enhancer had a greatly reduced basal level of activity (Fig. 4.3A, “ Δenh”), consistent with the idea that

this sequence has a stimulatory effect on manY translation. A third fusion where the

putative enhancer was moved downstream of the ATG (maintaining the reading frame)

had similarly low activity (Fig. 4.3A, “ATGenh”), suggesting that the translation

stimulatory effect of this sequence is position-dependent.

To determine whether the enhancer activity of the region upstream of manY requires cellular factors other than the translation machinery itself, we performed in vitro

translation assays with 3XFLAG-tagged manY transcripts. The amount of ManY-

3XFLAG protein produced from wild-type transcripts containing the enhancer sequence

in the 5' UTR was substantially more than the protein produced from an equivalent

amount of Δenh transcripts (Fig. 4.3B). These results suggest that the enhancer activity of the sequences upstream of manY is not dependent on Hfq, RNase E, or other cellular

factors. Instead, we propose that the enhancer can stimulate translation of manY directly

through interaction with the ribosome.

Fig. 4.3: The translation enhancer is ribosome-dependent. A) Activities of three manY'- 'lacZ fusions were compared to assess the regulatory role of the putative enhancer sequence. All fusions are controlled by the heterologous P BAD promoter. “WT” contains the wild-type manX-manY intergenic region with the enhancer sequence (indicated by the green box) positioned ~30 nt upstream of the manY start codon. “ Δenh” contains a deletion of the enhancer sequence, “ATGenh” has the enhancer sequence deleted from the upstream region and replaced just downstream of (and in frame with) the start codon. Strains containing fusions were grown for 2 hr until reaching the mid-log phase and β-galactosidase assays were performed. Error bars represent standard deviation from three biological replicates. P values (unpaired t test) are shown as: *, P < 0.05; **, P < 0.005; ***, P < 0.0005;****, P < 0.0001; NS, not significant. B) Translation reactions

Fig. 4.3 (cont.)

were performed for 30 minutes using manY-3XFLAG transcripts. “WT” is wild-type enhancer sequence and “ Δenh” contains a deletion of the enhancer sequence (as in part A). Products of translation were detected by Western blot using anti-FLAG and anti-GFP antibodies. As a control, reaction mixtures from each tube were analyzed on an SDS- polyacrylamide gel and stained with Coomassie Blue (the band for EF-Tu is shown).

4.2.3 Genetic analysis of enhancer sequences

The putative enhancer region is U-rich (Fig. 4A, -28 to -29 and -35 to -38), similar to the motifs found in previous studies that interact with the small ribosomal subunit [15,

31, 78, 137]. We also noticed a “CACA” motif, similar to those reported to act as translation enhancers in other sRNA target mRNAs [123, 162]. To further characterize the sequence determinants of the putative enhancer, we constructed reporter strains derived from the WT manY'-'lacZ fusion, each with different nucleotide substitution

mutations in the putative enhancer region (Fig. 4.4A). Mutations in the U residues of the

putative enhancer had the strongest effect on the basal levels of translation. Changing -

37U and -38U to G residues (Fig. 4.4A, B, mut1 ) reduced activity by ~30%. Mutation of -

35U and -34U to G residues (Fig. 4.4A, B, mut2 ) had no effect on activity, but mutation of all four U residues to G residues (Fig. 4.4A, B, mut3(G) ) reduced activity of the fusion

by ~85% compared to the wild-type fusion. Importantly, mutation of the U residues from -

35 to -38 to A residues (Fig. 4.4A, B, mut5(A) ) yielded a fusion with activity equivalent to

the wild-type fusion. Mutation of U residues at positions -28 and -29 to G residues also

had a strong impact on basal levels of activity (Fig. 4.4A, B, mut4 ). Other mutations in

this putative enhancer region, including residues in the “CACA” motif, had no effect on

basal levels of translation (Fig. 4.4A, B , mut6, mut7, mut9, mut10, mut14, mut15 ). These

results suggest that U and A residues in this region are particularly important for the

translation enhancing effect.

Fig. 4.4

Fig. 4.4 (cont.)

Fig. 4.4: Mutational analyses of the AU-rich translation enhancer. A) A series of manY translational fusions containing wild-type or mutant enhancer sequences were assayed for β-Galactosidase activity. The top sequence (WT) is the wild-type sequence for the

Fig. 4.4 (cont.) manX-manY intergenic region. The SD is indicated in blue and start codon in bold black letters. The sequences involved in base pairing with SgrS are indicated by the line under the sequences. Red letters in mutant sequences indicate positions of substitutions. B) β- Galactosidase assays of cultures containing indicated carrying the indicated fusions. Cells were grown for 2 hr until reaching the mid-log phase and β-galactosidase assays were performed. Error bars represent standard deviation from three biological replicates. P values (unpaired t test) are shown as: *, P < 0.05; **, P < 0.005; ***, P < 0.0005;****, P < 0.0001; NS, not significant. C) Indicated reporter fusions harbored in the degradosome mutant strain ( rne131::kan ) and assayed for β-Galactosidase activity as described in part B. D) manY'-'lacZ reporter fusions containing different combinations of SD and enhancer sequences were constructed and assayed for β-Galactosidase activity as described in part B. Symbols and corresponding sequences are defined in the insets below the graph. Fusions are labeled 1 through 10 and discussed in text accordingly. Error bars represent standard deviation from three biological replicates.

A number of studies have described AU-rich enhancer sequences located upstream of the start codon and SD, where r-protein S1 binds and enhances efficiency of translation initiation [15, 31, 62, 78, 137]. We hypothesized that the putative enhancer might similarly interact with S1 to promote translation of manY . If this were true, then the enhancer activity of this upstream sequence should be independent of SD-anti-SD interactions, and the enhancer should promote translation regardless of the strength of the SD. To test this possibility, we tested reporter constructs with various combinations of enhancer and SD sequences (Fig. 4.4C). As observed in the previous experiment, mutation of U residues to G residues ( mut3(G) ) strongly reduced the activity of the reporter fusion with the native manY SD (Fig. 4.4C, compare activity of reporters 1 and

2). A construct containing the native manY SD and the reverse complement of the enhancer region – now A-rich rather than U-rich – had high levels of translation (Fig.

4.4C, reporter 3), comparable to the wild-type reporter. This result suggests that the translation-enhancing effect of this region is not strictly sequence-dependent, but that

AU-rich sequences in general are stimulatory. This is consistent with the known sequence preferences of r-protein S1.

We next tested whether the enhancer could stimulate translation from a weaker

SD. The native manY SD (5'-AGGAG-3') was swapped for the slightly weaker lacZ SD

(5'-AGGA-3'), leaving the rest of the sequence in the reporter the same. As expected, the activity of this reporter was reduced compared to the reporter with the stronger manY

SD (Fig. 4.4C, compare activity of reporter 1 and reporter 4). The mut3(G) enhancer sequence strongly reduced activity (Fig. 4.4C, reporter 5), while the mut5(A) enhancer sequence restored the activity (Fig. 4.4C, reporter 6) of the constructs with the lacZ SD.

The same patterns were seen for two additional SD variants (Fig. 4.4C, reporters 7, 8, 9 and 10). Regardless of the SD sequence, the wild-type U-rich enhancer promoted higher levels of translation, and mutation of U residues to Gs dramatically reduced the translation activity. These results suggest that the enhancer functions independent of the

SD to promote translation.

4.2.4 Ribosome protection of the non-contiguous RBS and translation enhancer

The analyses presented so far suggest that an upstream enhancer sequence plays a positive role in manY translation. We hypothesized the enhancer interacts with the initiating ribosome, specifically r-protein S1, and that this interaction increases translation initiation. Studies of thrS mRNA, encoding threonyl-tRNA synthetase, showed that the 5' UTR of thrS contains a bipartite ribosome binding site where an upstream AU- rich element interacts with r-protein S1 ([31, 113], Fig. 5A). The AU-rich S1 binding site acts as a translational enhancer for thrS mRNA [15, 31, 78, 137]. We used this mRNA as a comparison and positive control in the next experiments to further examine the role of S1 in regulation of manY translation. We made thrS translational fusions with the wild- type sequence and mutants where U residues were changed to either Gs or As (Fig.

5A). β-galactosidase assays showed the same pattern we observed for the manY fusions. The wild-type thrS fusion (Fig. 5B, “WT-thrS ”) had a high level of activity, which 70

was strongly reduced in the mutant fusion with G residues (Fig. 5B, “G-thrS ”) and restored in the mutant fusion with A residues (Fig. 5B, “A-thrS ”). Footprinting reactions performed using RNase T1 (enzymatic) and lead acetate (chemical) probing with end- labeled mRNA and purified E. coli ribosomes showed the expected bipartite RBS for thrS mRNA, with protection around the SD and at the upstream AU-rich enhancer (Fig.

5C). We then performed footprinting reactions with end-labeled manY mRNA and purified ribosomes. In the absence of ribosomes, the pattern of cleavage by lead acetate suggested that the sequences around the manY SD and start codon are not very accessible and may be sequestered in a secondary structure. This is consistent with other examples of S1-dependent translation of mRNAs with highly structured translation initiation regions [14, 114]. In the presence of ribosomes, we observed a bipartite footprint, similar to the thrS footprint, with protection of the SD and start codon as well as protection of the upstream region containing the U-rich enhancer (Fig. 5D). These results are consistent with our hypothesis that the manY leader contains an enhancer upstream of the SD, which interacts with the ribosome, possibly with r-protein S1.

Fig. 4.5

Fig. 4.5 (cont.)

Fig. 4.5: Like known r-protein S1 target thrS, the manY leader contains a bipartite RBS. A) Graphical representation of thrS mRNA, including the bipartite RBS. The AU-rich region upstream of the SD is known to encompass a binding site for r-protein S1. Substitution mutations in the S1 binding site are indicated above in red nucleotides – “G- thrS ” and “ A-thrS .” B) thrS'-'lacZ translational fusions were constructed. “WT-thrS ” indicates the wild-type fusion, “ G-thrS ” and “ A-thrS ” are the mutant fusions with sequences as indicated in part A. Error bars represent standard deviation from three biological replicates. P values (unpaired t test) are shown as: *, P < 0.05; **, P < 0.005; ***, P < 0.0005;****, P < 0.0001; NS, not significant. C) Ribosome footprint analysis of thrS mRNA. In vitro transcribed mRNA was end-labeled with 32 P and incubated with and without E. coli ribosomes. Lanes are labeled as described in Fig. 2A legend. Positions of the start codon and SD are indicated on the left. Ribosome protected nucleotides are indicated on the right. D) The manY UTR and early coding region sequences are shown. The sequences comprising the SgrS binding site and enhancer are indicated with a line above the sequence. The SD and start codon are indicated in blue and red print, respectively. The regions protected by the ribosome are indicated with a green line below the sequences. The ribosome footprint is labeled as in part C.

4.2.5 Ribosomal protein S1 binds to the manY 5' UTR

A large body of literature has characterized S1 RNA-binding activity [66, 133,

136, 163]. In gram-negative bacteria, S1 orthologs are composed of six S1 motifs that

are structurally similar (Fig. 4.6A), but diverge in amino acid sequence [117, 133]. A

cryo-EM based approach found that the first two motifs of S1 at the N-terminus interact

with the r-protein S2 and function as a platform for ribosome binding [74]. Using proteins

with mutations in the S1 motifs, the Marzi group determined that the first three N-

terminal domains are crucial for S1 binding to rpsO mRNA [31]. Several residues of the

third motif (Y205, F208, H219, and R254) were found to be crucial for interaction with Q β

RNA [138]. We aligned S1 proteins from 11 organisms belonging to the Proteobacteria

and found that three out of these four residues important for RNA binding (Y205, F208,

H219) were highly conserved (Fig. 4.6B). Alignment of S1 motifs from seven other

diverse E. coli RNA binding proteins revealed that residues F208 and H219 were still

fairly well conserved (Fig. 4.6C). These results led us to test whether mutation of

conserved residues would impact S1 RNA binding activity. To address this question, we

performed EMSAs with purified wild-type S1 and mutant S1 (mS1, Y205A, F208A,

H219A) proteins and in vitro transcribed thrS and manY RNAs. Wild-type S1 bound to

thrS mRNA with a KD of 3.1 nM, while mS1 had a higher KD of 16 nM (Fig. 4.6D). This

result is consistent with the idea that mS1 is slightly defective for binding a known target.

The interaction between wild-type S1 and manY mRNA was weaker than for thrS, with a

KD of 81 nM. As for thrS, the interaction between mS1 and manY RNA was diminished

(KD = 590 nM, Fig. 4.6E). Though higher than for thrS mRNA, these dissociation constants for manY mRNA are in a similar range to those reported previously for other

S1 targets [107]. Altogether, the data suggest that the third S1 motif is important for

binding of thrS and manY mRNAs.

Fig. 4.6: Characterization of wild-type and mutant S1 (mS1) protein binding to thrS and manY mRNAs . A) A structure of S1 (showing six S1 motifs, the N-terminal motif is on the right) predicted using I-TASSER homology modeling algorithm based on the deposited cryo-EM structure [8]. S1 motif 3 is indicated with a dotted circle. B) A global alignment of S1 motif 3 from diverse bacterial species. The alignment was calculated using the Clustal Omega algorithm [75] and represented using the ESPript 3 web tool [112]. α- helices are displayed as squiggles. β-strands are marked with arrows, strict β-turns with the letters TT. The ղ symbol was used for 3 10 -helices. C) A global alignment of S1 motif 75

Fig. 4.6 (cont.)

3 with amino acid sequences of S1 motifs from other E. coli RNA binding proteins. D-E) In vitro transcribed thrS and manY mRNAs were labeled at the 5'-end with P 32 and mixed with WT S1 or mS1 in a binding buffer for EMSAs. Band intensities were measured for three replicates to determine the dissociation constants.

Fig. 4.7: S1 unfolds the secondary structures in the manY UTR. The manY transcript was preincubated with S1 or mS1 before chemical modification. DEPC modified in vitro transcribed manY transcripts were used as a template for primer extension with a reverse transcriptase in the absence or presence of the protein (S1 and mS1), and resolved in a polyacrylamide-urea gel. Sanger ladders (T, G, C, and A) were generated with dideoxy-sequencing. The enhancer region, the SD, and the start codon are indicated on the left.

To further study the interaction of r-protein S1 and manY mRNA, we performed additional footprinting reactions using diethyl pyrocarbonate (DEPC), which primarily modifies exposed adenine residues [32]. After treatment with DEPC, a reverse transcription reaction using a labeled primer is performed. Bands represent positions that were DEPC-modified, causing reverse transcriptase to stop. In the absence of S1 protein, there was very little modification of A residues around the SD and start codon

(Fig. 4.7, lane 2), again suggesting that the manY mRNA translation initiation region might be sequestered in a secondary structure. In the presence of wild-type S1 protein, there was substantially more modification, particularly in the region between the enhancer and the SD. This suggests that S1 binding to this region might remodel the secondary structure to make it more accessible for formation of the translation initiation complex. In the presence of mS1, we also saw new modifications signifying structural rearrangements, but these were distinct from the changes seen in the presence of wild- type S1 (Fig. 4.7). This result is consistent with the EMSA (Fig. 4.6E) and suggests that mS1 binds differently (and more weakly) to the manY mRNA.

4.2.6 Evidence of SgrS competition with S1 for control of manY translation

We hypothesized that SgrS represses manY translation by competing with r- protein S1 for binding at the enhancer sequence. To test this hypothesis in vivo, we constructed plasmids for ectopic expression of S1 and mS1, reasoning that a brief pulse of expression could alter the competition between endogenous SgrS and S1 protein.

Previous work demonstrated that S1 stimulates translation of target genes in a narrow range of concentrations – at higher concentrations, S1 represses target translation [14,

27]. Since manY and thrS have different affinities for S1, we reasoned that S1- dependent stimulation of translation would require different levels of induction in vivo. 77

Thus, we made two different types of S1 expression constructs – one with a strong SD

sequence, and one with a weak SD sequence. We transformed plasmids with S1 or mS1

into strains harboring thrS'-'lacZ (positive control), sodB'-'lacZ (negative control), and manY'-'lacZ, and performed β-galactosidase assays after a brief induction. Since thrS

has a higher affinity for S1, we used expression constructs with the weak SD. In the

thrS'-'lacZ strain, ectopic production of wild-type S1 resulted in a very slight, but not statistically significant increase in β-galactosidase activity. Production of mS1 had a more pronounced effect – decreasing the activity by 16% (Fig. 4.8A). This result is consistent with the idea that the ectopically produced mS1 protein interferes with endogenous S1 activity and inhibits S1-mediated translational activation of a known target. In the strain with negative control sodB'-'lacZ, we used S1 and mS1 constructs with the strong SD sequence. There was no difference in β-galactosidase activity between control and S1- or mS1-producing strains (Fig. 4.8B).

In the manY'-'lacZ strain, we used the S1 and mS1 constructs with the strong SD sequence. We also performed the experiments in the absence and presence of α-methyl glucoside, to induce production of SgrS [147]. In the absence of SgrS, we saw the same

S1-dependent patterns of activity as for thrS . Production of wild-type S1 slightly increased the activity of manY'-'lacZ and production of mS1 had an inhibitory effect (Fig.

4.8C, compare the black bars). As predicted if SgrS and r-protein S1 compete for binding to manY mRNA, the efficiency of SgrS-dependent repression varied depending on S1 production. In the strain with the vector control, SgrS induction caused a 26% reduction in β-galactosidase activity (Fig. 4.8C). (Note, this is a more modest repression ratio than seen in other experiments because we used a much shorter time for SgrS induction.) In the strain producing wild-type S1, the SgrS-dependent reduction in activity was only 16%. This suggests that wild-type S1 overproduction can modestly protect manY from SgrS-dependent translational repression. Most strikingly, production of mS1 78 left manY more susceptible to SgrS-dependent repression, with a repression value of

49% (Fig. 4.8C). These data are all consistent with the model that r-protein S1 stimulates manY translation, and that SgrS binding inhibits translation by interfering with

S1-dependent activation.

Fig. 4.8: Short-term ectopic production of S1 or mS1 modulates the efficiency of SgrS- dependent translational regulation of manY. Low copy plasmids carrying rpsA encoding wild-type or mutant r-protein S1 under the control of an IPTG-inducible promoter were transformed into strains with thrS'-'lacZ (A), sodB'-'lacZ (B) or manY'-'lacZ (C) fusions (reporter fusions were under the control of arabinose-inducible P BAD promoter). Cultures

Fig. 4.8 (cont.)

were grown at 37°C to OD 600 ~0.4 and the reporter fusion was induced with L-arabinose. Cells were grown for another 20 minutes before IPTG (to induce S1 or mS1) was added. Cultures were grown for an additional 10 min before β-galactosidase assays were performed. For manY'-'lacZ (C) fusions, SgrS and S1 (or mS1) was induced instantaneously. Error bars represent standard deviation from three biological replicates. P values (unpaired t test) are shown as: *, P < 0.05; **, P < 0.005; ***, P < 0.0005;****, P < 0.0001; NS, not significant.

4.3 Discussion

Fig. 4.9: Model of SgrS-mediated enhancer silencing. Our results suggest that the manY translation initiation region contains structural elements that are remodeled in part by r- protein S1 binding to a translational enhancer upstream of the manY SD to promote high levels of translation in the absence of the sRNA SgrS. Under glucose-phosphate stress conditions, when SgrS is produced, it base pairs with sequences in the enhancer, preventing formation of the translation initiation complex. SgrS could inhibit translation by sequestering the enhancer and preventing binding of r-protein S1 or by altering a crucial the secondary structure that interacts with ribosome-bound S1.

In this chapter, we show that the E. coli sRNA SgrS, in collaboration with the

RNA chaperone Hfq, represses manY translation by base-pairing with sequences comprising a translational enhancer. The enhancer stimulates manY translation in vitro,

suggesting that the effect is mediated by ribosomes and not by other cellular factors.

The AU-rich enhancer increases translation regardless of the strength of the SD

sequence. The elements within the manY leader region resemble other so-called

bipartite RBSs where ribosomes interact with both the SD and start codon region as well

as a non-contiguous upstream region. Our data implicate r-protein S1 as an important

mediator of manY translation initiation via interactions with the AU-rich enhancer

sequence. Our data are consistent with a model where SgrS competes with S1 for

binding to the enhancer sequences, thereby reducing S1-dependent translation and

resulting in the observed SgrS-dependent repression of manY (Fig. 9).

The RNA chaperone Hfq is essential for manY translation repression by SgrS in vivo (Fig. 1C). Hfq is a pleiotropic global regulator of gene expression, owing to its dynamic and complex interaction patterns with many cellular RNAs }[81, 149, 160]. The involvement of Hfq in sRNA-mediated translational repression has been attributed to its ability to remodel RNA secondary structures and overcome energetic barriers presented by one or both RNA partners, and to enhance local concentrations of potential binding partners to increase the rates of association [118]. A recent report illustrated an example of Hfq-dependent structural remodeling of dgcM mRNA to allow duplex formation with

OmrA or OmrB sRNAs, which results in translational repression of dcgM [50]. Similar mechanisms have been found for other sRNA-mRNA pairs that require Hfq-dependent structure remodeling [22, 161]. In this study, Hfq was required in vivo for SgrS- dependent regulation of manY (Fig. 1C). In vitro, the presence of Hfq promoted SgrS- dependent protection of manY mRNA (Fig. 2A, B), and adding Hfq to in vitro annealing reactions resulted in a 10-fold reduction in SgrS-manY KD (Fig. 2C, D). However, we did

not observe Hfq-mediated structural changes in our footprint experiments (Fig. 2A, B).

We hypothesize that SgrS sequesters the enhancer from ribosome-associated S1

protein (Fig. 9). This model could explain why Hfq is essential for SgrS-mediated 81

silencing of the enhancer in vivo. S1 forms a complex with manY with a KD of 80.5 nM

(Fig. 6E) while SgrS forms a complex at a higher stoichiometry ( KD of 7.1 µM, Fig. 2C).

In the presence of Hfq, the SgrS-manY complex has a submicromolar dissociation constant ( KD of 0.77 µM, Fig. 2E), which could allow SgrS to effectively compete with S1

for binding to the same site on manY mRNA.

The manY mRNA leader possesses a bipartite RBS. The RBS of a gene often extends beyond the start codon and the SD [64, 124, 127] with a non-contiguous architecture where additional sequences in the 5' leader establish interactions with ribosomal proteins and modulate translation initiation complex formation [26, 44, 65, 97,

144]. The threonyl-tRNA synthetase ( thrS ) mRNA contains a bipartite RBS, with an upstream single-stranded U-rich enhancer that is separated from the SD region by a stem-loop structure. For thrS mRNA, the enhancer is recognized by r-protein S1 [116].

This is just one example of an S1 binding site, and there are many other varieties [14,

31, 114, 116], making it difficult to identify S1 binding sites based on sequence or structural characteristics. The manY enhancer is centered at ~25 nt upstream of the SD

and we see a bipartite pattern of ribosome-dependent protection (Fig. 5D) similar to the

pattern for known S1 target thrS mRNA (Fig. 5C). In the absence of ribosomes, the manY SD and start codon region is not accessible to cleavage by lead acetate (Fig.

5D), suggesting it is structured. This is common for mRNAs whose translation is activated by r-protein S1, as S1 binding upstream can promote unfolding of secondary structure to allow formation of ribosome preinitiation complexes [7, 61, 107]. Our DEPC modification and footprinting experiments also showed that the SD and start codon region of manY mRNA is rather inaccessible to modification, and that addition of S1 slightly enhances accessibility of these sequences (Fig. 7). Consistent with our results, other studies have demonstrated S1-mediated unfolding of secondary structures with free S1 proteins [7, 107]. In an elegant study, the Marzi group demonstrated that 82

ribosome-bound S1 could unfold rpsO mRNA and expose the SD region more efficiently

than free S1 protein [31]. Another recent study showed how ribosome-associated S1

prevents secondary structure formation and exposes an otherwise inaccessible SD of

the tisB mRNA [114]. With assistance from neighboring ribosomal components, S1 might

play a more prominent role in remodeling the secondary structure of manY mRNA in its

natural ribosome context. Nevertheless, our data are consistent with a model where

manY translation is enhanced by r-protein S1 binding to a region upstream of the SD in the absence of SgrS (Fig. 9).

Both technical challenges and opportunities continue to arise as we discover more and more examples of sRNA-mediated translational regulation from a distance, i.e., outside the boundaries where canonical steric occlusion of the SD and start codon are possible. There are many mRNAs with very long and highly structured 5' leaders, and doubtless an array of new regulatory mechanisms controlling mRNA translation and stability are yet to be discovered. Our study and the numerous recent global analyses of

RNA-RNA interactomes [3, 28, 123] suggest that there are dozens of targets for most sRNAs, and we must expand the search window beyond the SD and start codon region when looking for sRNA target binding sites. Mechanistic characterization of 5' UTR- targeting sRNAs could reveal new functional elements involved in translation initiation and help us uncover the complex and dynamic interactions between mRNAs and ribosomes.

CHAPTER 5

CONCLUSIONS

In bacteria, we have yet to uncover the mechanistic details of regulation carried out by the vast majority of sRNAs, but of those for which mechanisms have been established, sRNAs are typically the primary effectors of regulation. This raises some intriguing questions. Is the “canonical” mechanism of sRNA-mediated regulation with sRNA as primary regulator truly the most common, or have computational and experimental approaches used to study sRNAs been biased toward discovery of these mechanisms because they were the first type described? Regardless of the prevalence of each of these two different mechanisms, what features distinguish them and make one or the other more favorable for regulation of a given mRNA target?

One advantage of sRNA-mediated regulation that involves base pairing interactions outside the TIR could be that it provides a larger and more diverse sequence space to evolve new regulatory interactions. We have found that regulation of ptsG , the primary glucose transporter, by SgrS is conserved among E. coli relatives

where SgrS orthologs were found [53, 151]. The SgrS-ptsG mRNA interaction involves

pairing between the most highly conserved seed region of SgrS and the ptsG RBS, a region where the sequence is highly conserved for ribosome binding. In contrast, SgrS- dependent regulation of manX involves a less well-conserved portion of SgrS (adjacent to the conserved seed) and the coding sequence of manX, and this interaction is not entirely conserved among enteric species. For instance, E. carotovora SgrS and Y. pestis SgrS do not regulate their cognate manX orthologs [110] because the SgrS sequences that pair with manX mRNA are not conserved in these organisms. Analyses by Peer and Margalit [102] indicate that the SgrS-ptsG mRNA interaction evolved first,

84 with the binding sites on both mRNA and sRNA co-appearing in evolutionary time [102].

Their data suggest that SgrS-manX mRNA interaction evolved much later. So, SgrS first established a regulatory interaction with ptsG in an ancestral organism of the order

Enterobacteriales, which established this sRNA regulator in the genome and allowed evolution of interactions with additional targets. Other recent work on sRNA evolution suggests similar target acquisition mechanisms where sRNAs establish one target and gradually establish other interactions with the concurrent evolution of Hfq [102, 111].

Perhaps flexibility in regulatory mechanisms, e.g., where the sRNA can act as either a primary or accessory regulator along with Hfq, facilitates rapid evolution of additional sRNA-target interactions.

A long-held notion about sRNA-mediated gene regulation in eukaryotes is that the primary role of sRNAs is target recognition, while the associated protein partners perform the primary regulatory function of gene silencing or translational repression. In bacteria, the prevailing model has been the opposite that the sRNA is the primary regulator and associated proteins play secondary roles in promoting RNA stability or making the regulation irreversible (in the case of mechanisms involving mRNA degradation). Our findings, along with one other recent report on a similar non-canonical mechanism of regulation in bacteria [28] suggest that bacteria can utilize a broader range of sRNA-mediated regulatory strategies than previously suspected. So, while there are considerable differences among the domains in terms of the mechanisms of translation initiation, sites of sRNA binding, and the nature of the ribonucleoprotein complexes carrying out regulation, sRNA-mediated regulation of mRNA targets appears to be a common mode of regulation in all three domains of life.

The data presented in Chapter 4 proposes an intriguing mechanism where the sRNA sequesters a protein binding site. Similar to the transcriptional UP element,

translation enhancers could also be exploited by cells to modulate translation.

Translation enhancers can potentially influence either ribosome binding or ribosome

liberation from the SD element. The position requirement of enhancer sequence for the

functioning of the enhancer suggests that it ensures an anchor point and subsequently

facilitates TIC formation. A recent study by Takahashi and colleagues reported that

translation has no effect on the kon , rather it increases the koff between the ribosome and the SD sequence increasing translation efficiency [137]. Here, the enhancer/S1 interaction works against the SD/anti-SD interaction and shift the equilibrium towards the elongation state, which is a striking example of negative allosteric modulation. Some of the data presented in this report tentatively suggests that similar mechanism could be in play for manY enhancer as well.

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