Physiological Role and Substrates of Rnase D and Rnase BN in Escherichia Coli

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Physiological Role and Substrates of Rnase D and RNase BN in Escherichia Coli

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UNIVERSITY OF MIAMI

PHYSIOLOGICAL ROLE AND SUBSTRATES OF RNASE D AND RNASE BN IN ESCHERICHIA COLI

By Christie H. Taylor

A DISSERTATION

Submitted to the Faculty of the University of Miami in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Coral Gables, Florida December 2014

UNIVERSITY OF MIAMI A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy

PHYSIOLOGICAL ROLE AND SUBSTRATES OF RNASE D AND RNASE BN IN ESCHERICHIA COLI

Christie H. Taylor

Approved:

______Murray P. Deutscher, Ph.D. Arun Malhotra, Ph.D. Professor of Biochemistry Associate Professor of and Molecular Biology Biochemistry and Molecular Biology

______Feng Gong, Ph.D. Kurt R. Schesser, Ph.D. Associate Professor of Biochemistry Associate Professor of and Molecular Biology Microbiology and Immunology

______Zhongwei Li, Ph.D. M. Brian Blake, Ph.D. Professor of Biomedical Science Dean of the Graduate School Florida Atlantic University

TAYLOR, CHRISTIE H. (Ph.D., Biochemistry and Molecular Biology) Physiological Role and Substrates of RNase D (December 2014) and RNase BN in Escherichia coli

Abstract of a dissertation at the University of Miami.

Dissertation supervised by Professor Murray P. Deutscher.

No. of pages in text. (125)

There are many ribonucleases in Escherichia coli and, presumably, each one has an important role in RNA metabolism. After an individual RNA has been fully transcribed, the fate of that RNA is determined by its sequence, structure, and the availability of RNases which may be determined by conditions inside and outside of the cell. During periods of exponential growth, the majority of RNA degradation is carried out by RNase II, RNase R and polynucleotide phosphorylase, and the short RNA fragments generated by the degradation process are reduced to individual nucleotides by oligoribonuclease. Maturation of

RNAs is performed primarily by RNase III, RNase P, RNase E, RNase G, RNase

T and RNase PH. These processes of degradation and maturation represent most of RNA metabolism in cells, and can be accounted for by the aforementioned RNases. However, there are two additional RNases that have no known primary role in RNA metabolism – RNase D and RNase BN.

Here, I present in vivo and in vitro data showing that RNase D and, to a lesser extent, RNase BN are responsible for fine tuning regulatory elements in

cells by removing at least one RNA that is deleterious for growth and survival when present in excess. This is the mRNA for CsrA, an RNA binding protein that acts in concert with the sigma factor RpoS when cells are growing slowly or not at all. I will also show that cells lacking RNase D take more time to exit lag phase than do cells in which RNase D is present and that this defect is not due to aberrant maturation of ribosomal RNAs. In order to identify RNAs that are substrates for RNase D and RNase BN in vivo, Northern analyses of many stable and messenger RNAs, electrophoresis of radiolabeled RNA and tRNA nucleotidyltransferase assays were carried out, among others. These ruled out tRNA and rRNA as the primary substrates for RNase D and RNase BN during stationary phase and recovery from prolonged starvation. They also revealed that the mRNA for CsrA is upregulated 3- to 4- fold in cells lacking RNase D during early stationary phase and that RNase D and RNase BN are expressed most abundantly during rapid growth. Assays of growth, cell morphology and ability to synthesize critical cellular components like flagella and extracellular matrix were utilized concurrently with the previously noted molecular techniques. These showed that cells lacking RNase D have a reduced ability to synthesize extracellular matrix, show reduced motility, took longer to recover from prolonged starvation, and synthesize more of the messenger molecule cyclic-di-GMP than wild type cells. Cells lacking RNase D and RNase BN were at a competitive disadvantage when grown in low pH conditions. Together, these studies have revealed that these two RNases do indeed have a role in E. coli allowing cells to more efficiently enter and exit rapid growth by fine-tuning the availability of at

least one specific RNA that has a deleterious effect when overexpressed. These studies presented here indicate that the roles of RNase D and RNase BN are subtle and while many substrates have been ruled out, undoubtedly additional substrates remain to be discovered.

TABLE OF CONTENTS

Page

LIST OF FIGURES ix

LIST OF TABLES xii

Chapter

1. INTRODUCTION AND LITERATURE REVIEW 1

1.1. Overview 1

1.2. RNases in E. coli 3

1.3. RNase BN 6

1.4. RNase D 7

1.5. sRNA, tRNA, mRNA and rRNA 8

1.6. RNA metabolism 10

1.7. Summary 12

2. ElaD 14

2.1. Background 14

2.2. Experimental procedures 14

2.2.1. Bacterial Strains 14

2.2.2. Media 16

2.2.3. Individual growth analysis 16

iii

2.2.3.1. Growth curve analysis 16

2.2.3.2. Exponential to stationary phase transition analysis 17

2.2.3.3. Survival during stationary phase analysis 17

2.2.3.4. Lag phase analysis 17

2.2.4. Co-culture assay 17

2.2.4.1. Full cycle competition assay 18

2.2.4.2. Calculation of doubling times 18

2.2.5. Gene mapping 19

2.2.6. Phage P1 transductions 19

2.2.6.1. Phage P1 lysates 19

2.2.6.2. Phage titers 19

2.2.6.3. Multiplicity of infection calculations 19

2.2.6.4. Transduction procedure 19

2.2.7. KEIO collection and pCA24N plasmid 20

2.2.8. DIC microscopy 20

2.2.9. Statistical methods 20

2.2.9.1. Bivariate analysis 20

2.2.9.2. Multivariate analysis 20

2.3. Results 21

2.4. Discussion 25

3. Searching for Substrates via Molecular Techniques 28

3.1. Background 28

3.2. Experimental Procedures 29

3.2.1. RNA purification and fractionation 29

3.2.2. tRNA nucleotidyltransferase assay 29

3.2.3. PAGE for small and 38 cm gels 29

3.2.3.1. Gels 29

3.2.3.2. 32P labeled RNA 30

3.2.4. Ribosome gels 30

3.2.5. Northern Analysis 30

3.2.5.1. Primer design 30

3.2.5.2. Densitometry 30

3.3. Results 31

3.4. Discussion 48

4. Searching for Phenotypes 50

4.1. Background 50

4.2. Experimental Procedures 50

4.2.1. MTT assay 50

4.2.2. Antibiotic sensitivity and resistance 51

4.2.2.1. SensiDisc Assays 51

4.2.2.2. Gradient Plates 51

4.2.3. Spectinomycin Assay 51

4.2.4. Phage Burst Assay 51

4.3. Results 52

4.4. Discussion 68

5. CsrA Substrate and Clumping Phenotype 71

5.1. Background 71

5.2. Experimental Procedures 72

5.2.1. Alternative Carbon sources 72

5.2.2. Cloning of csrA into pHC79 plasmid 72

5.2.3. Cell adherence assays 72

5.2.4. Paper chromatography assay 72

5.2.5. Extracellular matrix formation assay 73

5.2.6. Detergent and buffers for plating procedures 73

5.2.6.1. Buffers for pH maintenance 73

5.2.6.2. Detergents 73

5.2.7. Microscopy for clumping 73

5.2.8. Western analysis 73

5.2.9. Transcription of csrA mRNA in vitro 73

5.2.10. in vivo and in vitro molecular analysis 74

5.2.10.1. Standards 74

5.2.10.2. in vivo analysis 74

5.2.10.3. in vitro analysis 74

5.3. Results 75

5.4. Discussion 86

6. Recovery from lag phase phenotype 88

6.1. Background 88

6.2. Experimental Procedures 89

6.2.1. Alternating nutrient conditions assay 89

6.2.2. Starvation shock assay 89

6.2.3. Starvation competition assay 89

6.2.4. Sucrose gradients 89

6.2.4.1. Subunit analysis 89

6.2.4.2. Ribosome analysis 90

6.2.5. 5' Primer Extension procedure 90

6.2.6. 3' RACE procedure 90

6.3. Results 90

6.4. Discussion 104

7. Possible influence of Crl 105

7.1. Background 105

vii

7.2. Experimental Procedures 105

7.2.1. Strains 105

7.2.2. Media 105

7.3. Results 106

7.4. Discussion 108

8. DISCUSSION AND FUTURE WORK 109

REFERENCES 117

viii

LIST OF FIGURES

Page

CHAPTER 2

Figure 2. 1 Growth of BEZ33 and MG1655 in individual culture. 21

Figure 2. 2 Optical density of original strain in individual culture. 22

Figure 2. 3. Graphic of gene map showing cut sites for BEZ33. 23

Figure 2. 4. Microscopic analysis of cell phenotypes. 24

Figure 2. 5. Northern analysis of 6S RNA. 25

CHAPTER 3

Figure 3. 1 Representative Northern blots of tRNA. 33

Figure 3. 2 Representatie Northern blots of sRNA. 38

Figure 3. 3 Representative Northern blots of mRNA & other RNA 43

Figure 3. 4 Northern blot of csrA mRNA. 44

Figure 3. 5 32cm PAGE of total RNA. 45

Figure 3. 6 A. Northern of RNase D mRNA in MG1655. 47

B. Northern of RNase D mRNA in CA265. 47

Figure 3. 7 A. Northern of RNase BN mRNA in MG1655. 48

B. Northern of RNase BN mRNA in CA265. 48

CHAPTER 4

Figure 4. 1 Spectinomycin resistance assay. 64

Figure 4. 2 Stab cultures for motility using TTC dye. 67

CHAPTER 5

Figure 5. 1 Survival during stationary phase. 75

Figure 5. 2 Quantification of cell aggregation. 76

Figure 5. 3 Northern blot of csrA mRNA. 77

Figure 5. 4 Western blot of CsrA protein. 78

Figure 5. 5 in vitro digestion of CsrA mRNA. 79

Figure 5. 6 in vivo half-life of CsrA mRNA. 80

Figure 5. 7 Phenocopy of csrA mRNA levels in vivo with pHC79::csrA.

Figure 5. 8 Quantification of cell aggregation with pHC79::CsrA. 82

Figure 5. 9 Quantification of extracellular matrix formation with crystal violet stain. 84

Figure 5. 10 Paper chromatography for c-di-GMP. 85

CHAPTER 6

Figure 6. 1 Effect of abrupt depletion of nutrients 91

Figure 6. 2 Long term growth and survival. 92

Figure 6. 3 Growth from lag phase. 93

Figure 6. 4 Northern of 16S rRNA. 94

Figure 6. 5 Northern of 23S rRNA. 95

Figure 6. 6 High magnesium ribosomal sucrose gradient. 96

Figure 6. 7 Low magnesium ribosomal subunit sucrose gradient. 97

Figure 6. 8 A. Primer extension of 5' ends of 16S rRNA. 98

B. Primer extension of 5' ends of 23S rRNA. 98

Figure 6. 9 A. RACE of 3' ends of 16S rRNA. 99

B. RACE of 3' ends of 23S rRNA. 99

Figure 6.10 A. Northern of CsrA mRNA. 100

B. Western of CsrA protein. 100

Figure 6. 11 Northern of 6S RNA . 101

Figure 6. 12 Low molecular weight radiolabeled RNA from starved and lag phase cells. 102

Figure 6. 13 38 cm PAGE of high molecular weight RNA. 103

CHAPTER 7

Figure 7. 1 Gel of Is1 element in MG1655*. 106

Figure 7. 2 Co-cultured growth and survival of Pro::Tn10 crl+ strains.

107

Figure 7. 3 Growth of Pro::Tn10 crl+, rpoS- strains. 108

LIST OF TABLES

Page

CHAPTER 2

Table 2. 1 Bacterial Strains 15-16

Table 2. 2 Phage Strains 16

CHAPTER 3

Table 3. 1 tRNA probes 31-32

Table 3. 2 Densitometry for tRNA 34-35

Table 3. 3 Results of tRNA Nucleotidyltransferase Assay 35

Table 3. 4 sRNA probes 36-38

Table 3. 5 Densitometry for sRNA 39-40

Table 3. 6 mRNA and other probes 41

Table 3. 7 Densitometry for mRNA and pseudogenes 42

CHAPTER 4

Table 4. 1 A. MTT assay quantification for PM1 & PM2A 52-55

B. MTT assay quantification for PM3B & PM4A 55-58

C. MTT assay quantification for PM9 & PM10 59-62

Table 4. 2 Antibiotic sensitivity and resistance 63

Table 4. 3 Phage burst 66

xii

Chapter 1. INTRODUCTION AND LITERATURE REVIEW

1.1 Overview

The central dogma of biology is that deoxyribonucleic acids (DNA) is the template for ribonucleic acids (RNA), which is then used to make protein. Broad swaths of science revolve around elements of this central process and the phenomena that exist to contradict it. Much is known about DNA, RNA and protein and the processes in which they participate in living organisms

(Korostelev, 2011).

RNA metabolism is the link between protein and DNA. It involves many components to bridge the gap because not only is RNA the molecule that contains the information from which the proteins are translated, but it also comprises the bulk of the very molecular machinery that assembles the protein. It is certainly interesting that many of the components involved in the journey from

DNA to protein contain ribozymes at their catalytic core or are made almost completely of RNA, as in the case of tRNA and ribosomes. These and similar phenomena are often the basis for the theory that RNA was the original molecule of life and replication in living systems (Neveu et al. 2013).

Needless to say, the fate of RNA is central to any biological system, particularly for RNAs that have outlasted their usefulness or are somehow defective. Many reviews on the topic of RNA decay are available (Bandyra &

1 2 Luisi, 2013) and yet many possible avenues for scientific exploration still exist

(Dogini et al. 2014, Morris et al. 2014).

The culmination of decades of research into the topic from molecular biology, biochemistry and genetics has created a detailed understanding of the relationships between various RNases and RNA in Escherichia coli. This dissertation focuses on two of the remaining RNases whose roles in the model organism E. coli remain unclear - RNase D and RNase BN.

RNA degradation typically occurs when an RNA substrate is defective or is no longer needed within the cell. Which RNase breaks down a particular RNA depends largely on the amount of secondary structure in the RNA and to a lesser extent on sequence (Bandyra, et al. 2013). Highly structured RNAs like tRNA and ribosomal RNA are degraded by PNPase and RNase R. PNPase associates with an RNA helicase in vivo to allow it to overcome secondary structure. RNase R requires a single stranded 3' overhang of 4 or more nucleotides to digest structured RNA and has the ability to resolve secondary structure without the aid of a helicase. RNAs with less secondary structure are degraded by RNase II, a close relative of RNase R. RNase II is sensitive to secondary structure in potential substrates and has no helicase activity of its own associated with the enzyme (Vincent & Deutscher 2009). These three exoribonucleases are responsible for the bulk of RNA degradation in E coli. The roles of RNase D and

RNase BN in E. coli have or had remained mysterious despite a number of papers that suggested possible roles for the enzymes in vivo (Perwez & Kushner

2006, Takaku & Nashimoto 2008, Dutta & Deutscher 2010). These studies have

3 shown a number of substrates for these enzymes in vivo and in vitro and also shown that the RNases that are synthesized are functional in vivo.

Because it is unlikely that an organism whose very survival can depend on minutes difference in doubling time, it would be evolutionarily advantageous for a cell to eliminate excess useless DNA from the chromosome and cease dedicating resources toward RNA and functional proteins that are apparently unnecessary. Several deletion mutants have been characterized and found that

RNase D and RNase BN play no obvious role in RNA metabolism within E. coli during rapid growth (Bandyra, et al. 2013). Still, these genes remain in the E. coli chromosome and the RNA and functional proteins are synthesized. It stands to reason that these proteins are therefore necessary for some as yet unknown function in E. coli.

1.2 RNases in E. coli

E. coli contains eleven endoribonucleases, including three toxins – RelE,

MazF and ChpBK, E coli also has eight exoribonucleases and three exoribonucleolytic toxins – YoeB, YafQ and RnlA (Kamada & Hanaoka, 2005;

Montiejunaite et al., 2007; Otsuka & Yonesaki, 2005). RNase BN can behave both as an endoribonuclease and as an exoribonuclease, but it is understood to behave primarily as an exoribonuclease in vivo (Dutta & Deutscher, 2009). These enzymes along with RNA polymerase comprise the bulk of enzymes associated with the processes involved in RNA metabolism.

4 The endoribonuclease toxins participate in mRNA degradation under stress (Li & Deutscher 2004). They were inherited by horizontal gene transfer based on bioinformatic analysis of bacterial genomes suggesting that their presence is due to genetic parasitism rather than an evolved function of E. coli.

This of course does not bar the possibility that E. coli can harness these enzymes as they evolve. (Escobar-Páramo, et al. 2004)

Other endoribonucleases in E. coli include RNase I, RNase III, RNase P,

RNase E, RNase G, RNase BN, RNase HI, RNase HII. RNase I is isolated to the periplasm of the cell as a nonspecific T2 family ribonuclease responsible for degrading exogenous RNAs – it serves as a defense from foreign and potentially infectious RNAs. RNase HI and HII are involved in DNA replication and DNA repair. RNase HI degrades the RNA strand of DNA:RNA hybrids and RNase HII cleaves RNA residues that have accidentally been incorporated into DNA during replication and repair. (Karginov, et al. 2004; Kitahara, et al. 2011)

The remaining endoribonucleases in E. coli – RNase III, RNase P, RNase

E, RNase G and under special circumstances, RNase BN – all participate in RNA metabolism via mRNA degradation, stable RNA maturation and quality control of defective RNAs.

The exoribonuclease toxins, YoeB and YafQ both participate in mRNA degradation under stress conditions like the endoribonuclease toxins. RnlA (also called RNase LS) participates in T4 mRNA degradation and in E. coli RNA metabolism at large instead of being active primarily under stress conditions like the other toxins (Otsuka and Yonesaki, 2005). Interestingly, RNase HI's activity

5 stimulates the RnlA toxin's activity. This synergistic relationship suggests that other synergistic interactions between RNases in E. coli are possible (Naka, et al.

2014).

The remaining RNases participate in the dominant elements of RNA metabolism. RNase II, a processive, hydrolytic RNase is proficient in degrading single stranded RNA but is inhibited by the presence of secondary structure as it participates in mRNA degradation and stable RNA maturation. Its paralogue,

RNase R is very much like RNase II in its protein sequence and structure, but its function is divergent in that it can invade secondary structure given a 3' overhang longer than 4 nucleotides (Vincent & Deutscher, 2009). It too participates in mRNA degradation, stable RNA degradation and quality control. RNase D,

RNase T and oligoribonuclease (ORN) are all members of the DEDD family of

RNases. RNase D is distributive and hydrolytic and it is able to participate in stable RNA maturation of single stranded substrates and possibly tRNA

(Zaniewski, et al. 1984). RNase T is much more active on these substrates as a distributive, hydrolytic enzyme. RNase T participates in tRNA end-turnover and stable RNA maturation. It is single strand specific and is unable to invade secondary structures or junctions in RNAs (Hsiao, et al. 2014). RNase T also discriminates against pyrimidines. ORN is hydrolytic like RNase D and RNase T, but it is a distributive enzyme. It shows an affinity for short oligoribonucleotides and is responsible for the completion of mRNA degradation in E. coli.

RNase BN is a distributive, hydrolytic enzyme that shows preference for tRNA precursors and tRNA molecules in which the 3'-CCA sequence has been

6 altered. PNPase and RNase PH are also paralogues with phosphorolytic activity.

PNPase participates in mRNA degradation and stable RNA degradation and quality control, but is only able to overcome secondary structure in RNA with the help of an RNA helicase in vivo, with which it associates. RNase PH participates in stable RNA maturation as a single strand specific RNase. (Dominguez-

Malfavon, et al. 2013)

1.3 RNase BN

RNase BN was first discovered when its absence made E. coli cells resistant to infection by the mutant T4 phage, Bu33. The head coat protein of the Bu33 phage contains an amber mutation that can be overcome by a viral encoded amber suppressor tRNA. RNase BN is the only enzyme in E. coli capable of efficiently maturing this amber suppressor. Maturation of this amber suppressor allows the phage variant Bu33 to kill the host cell. Without RNase BN, however,

T4 variant Bu33 is unable to synthesize its head coat protein and E. coli becomes resistant to the phage. This is an unusual case in that the absence rather than the presence of the RNase confers a selective advantage

(Deutscher, et al. 1983).

Subsequently, RNase BN has been characterized extensively in E. coli in vitro and also in vivo. It is best known for its ability to mature tRNA precursors.

RNase BN can discriminate based on the available sequence at the 3' end of tRNA to more efficiently yield the characteristic 3' CCA sequence. It does this by trimming the 3' tail based on the sequence of nucleotides. If there is a C, CC, or

7 CCA in front of the discriminator nucleotide, it will leave them intact but will remove any other residues. This interesting property of the enzyme prevents the inefficient removal and re-addition of the essential residues (Dutta, et al. 2013;

Deutscher 1990). RNase BN homologues by the same name and also called

RNase Z are often responsible for maturing tRNA by cleaving back to the discriminator residue in other organisms. E. coli RNase BN does this because of a very narrow catalytic channel. This narrow channel can only accommodate single stranded RNA. If altered, this channel can accommodate processive exoribonucleic activity on the ends of these transcripts (Dutta et al. 2013).

These properties can sometimes suggest a function. For example, processive enzymes are more often involved in degradation where endoribonucleases are more likely to participate in maturation of transcripts. This unusual suite of properties make RNase BN more elusive, rather than elucidating the role of the enzyme.

1.4 RNase D

RNase D is a 3'-5' exoribonuclease that is capable of maturing tRNA both in vitro and in vivo (Zhang & Deutscher, 1988). It is also known and was originally named for its ability to degrade denatured RNA in vitro, hence the designation "D". Overexpressing RNase D in strains mutated for tRNA nucleotidyltransferase is known to slow growth in E. coli (Zhang & Deutscher,

1988). Consistent with this deleterious effect, the expression of this RNase is very tightly regulated by an unconventional UUG start codon (Zhang &

8 Deutscher, 1989). Converting this codon to an AUG results in an eleven-fold increase in expression from single-copy plasmids. Additionally, an upstream hairpin structure seems to negatively influence RNase D expression, but to a lesser extent than the UUG codon (Zhang & Deutscher, 1992). Microarray data suggests that RNase D is expressed almost exclusively during exponential phase (Zhou & Rudd, 2013) RNase D protein has also been shown to be produced most abundantly during exponential growth (Dutta, unpublished). Tight regulation for a potentially toxic RNase, its low in vivo activity on tRNA and its expression pattern makes RNase D a curiosity among the RNases of E. coli.

1.5 sRNA, tRNA, mRNA and rRNA

There are several groups of RNA in E. coli, including but not limited to ribosomal RNA (rRNA), transfer RNA (tRNA), small non-coding RNA (sRNA) and messenger RNA (mRNA). Ribosomal RNA is the most abundant species of RNA in E. coli making up about 80-90% of the RNA during rapid growth. This makes it quite visible on agarose and polyacrylamide gels, forming the upper limit for large stable RNAs. Ribosomes consist of 2 subunits that include three RNA transcripts. The large subunit (50s) consists of 2 RNAs, a larger one (23S) and a smaller one (5S). The small subunits include only one RNA (16S). The rRNA in the 23S subunit is 2904 nucleotides long. The 5S transcript is 120 nucleotides long and the other RNA that comprises the16S subunit is 1541 nucleotides long.

These RNAs form most of the interface between the 2 subunits and act as scaffolding for the ribosomal proteins. The pathways for maturation and

9 degradation of ribosomes have been studied extensively and are largely known.

(Muller, et al. 2000; Halic, et al. 2006; Basturea, et al. 2013)

Transfer RNA is the next most abundant stable RNA in the cell. There are

86 known genes for tRNA in E. coli. Each corresponds to one of the 20 amino acids in the cells. Only 47 are required to recognize all of the codons due to wobble, but all are synthesized at stoichiometric ratio relative to the number of ribosomes in a cell. (Crick, 1966 Dinçbaş, et al. 1995; Smailov & Garilova

1985)They are among the smaller RNA transcripts in the cell. All are less than 90 nucleotides long but more than 50 nucleotides in length. These RNAs are also very consistent in their appearance on acrylamide and agarose gels forming the visible boundary at the bottom of gels for small stable RNAs. (Blattner et al.

1997)

Messenger RNAs have shorter half-lives compared to rRNA or tRNA, but are equally important for their role in translation. The lengths of these transcripts range greatly from just over a hundred nucleotides to several thousand. Their presence on stained agarose and acrylamide gels is rarely more than a faint smear, if they are visible at all. (Noiguera, et al. 2001)

Small non-coding RNAs include stable and non-stable transcripts. Their stability varies depending on environmental conditions. Most are fairly stable and serve as regulatory molecules unlike their functional tRNA and rRNA non-coding counterparts. Often small non-coding RNAs bind to other RNAs which are then targeted for destruction, but others have more novel mechanisms for controlling metabolism in E. coli. (Lalaouna, et al. 2013; Gottesman & Storz, 2011)

10 All of these RNAs are possible substrates for RNase D and RNase BN, though some are more likely than others. Because both enzymes seem to have an affinity for smaller substrates (they both can mature tRNA both in vitro and in vivo, e.g.), single stranded and denatured substrates, we can make some inferences about the roles of these molecules and potential substrates.

1.6 RNA metabolism

RNA metabolism in E. coli consists of transcription, maturation, quality control and degradation. Other processes like the formation of secondary structure and aminoacylation, addition of proteins, modification, translation and other processes contribute to the functionality of these RNAs. (Charette, 2000)

The oligomeric molecules of RNA consists of 2 kinds of purine – adenine and guanine – and two pyrimidines – uracil and cytosine. This is different from deoxyribonucleic acids (DNA) in terms of its composition and importance because the ribonucleic phosphate backbone twists at a slightly different pitch.

This disparity adversely affects the annealing efficiency of heterodimer and reduces the ability of heterodimers to form extended helical structures as efficiently as DNA homodimers. Uracil is also replaced by thymine in DNA.

Adenine complements uracil and cytosine complements guanine. These molecules comprise the single stranded RNA linked by their phosphate backbone. The single stranded RNA readily forms secondary and even tertiary structures. Secondary structures include dimers and hairpins that can frequently be observed in stable RNAs. Ribosomal RNAs form many complex tertiary

11 structures that contribute to their functionality including pseudoknots which include two stem-loop structures that intercalate with one another on part of the stem of each stem-loop (El Yacoubi et al. 2012).

Transcription is the process by which RNA polymerase transcribes a single strand of RNA from its cognate DNA strand and is the first step in gene expression. This process is regulated by the expression of various sigma factors which bind to the RNA polymerase and direct its specificity for certain genes designated by that particular sigma factor. The holoenzyme is able to bind the transcription initiation site and split the DNA double helix by breaking the hydrogen bonds that hold the two strands together. It moves down the DNA strand adding cognate RNA nucleotides according to the complementary DNA strand. As the difference in structure of the nascent RNA twists the RNA transcript off the DNA molecule, and the DNA reunites with its sister strand

(Fulcrand et al. 2013).

The newly transcribed RNA may have any number of destinations depending on its evolved structural, functional and sequence properties. Many

RNAs in E. coli are co-transcribed as an entire operon and must be cleaved to form individual transcripts by RNases before they can either form appropriate secondary structures or be translated. The purines and pyrimidines themselves may be altered, parts of the transcript may be methylated or the transcript may be defective and degraded in order for these transcripts to be fully functional (Li

& Deutscher, 2008).

12 1.7 Summary

Each cell regulates the processes of transcription on many levels. Precise regulation is at the center of efficient management of resources within a cell.

RNA metabolism is one of many finely tuned elements that link these processes.

Evolutionary pressures would result in deterioration of unnecessary genetic material, but instead we observed maintenance of RNase D and RNase BN genes and proteins. Thus suggesting that there is an evolutionary advantage for the persistence of RNase D and RNase BN in E. coli.

It is plausible that RNase D and RNase BN exist to support the more active RNases in cells. In living systems redundancy confers robustness. In E. coli RNA metabolism, RNase BN is known to be the least active on tRNA – tRNA is the substrate which RNase BN is known to mature and control for quality in other organisms. This limited activity in vivo may be indicative of a supportive role.

RNase D has a variety of substrates in vitro and can act on them in vivo if a number of other RNases are absent from the cell. However, it's activity during normal physiological activities is also very low and eliminating the gene from the cell has few obvious consequences during rapid growth.

We postulated that a small non-coding RNA was the most likely candidate substrate based on the properties of RNase D and RNase BN and their expression patterns. Because these RNAs are most important during stress conditions, it would follow that an RNase which was not apparently necessary during rapid growth might also be important under stressful conditions. It also

13 follows that two RNases which are known to act on small RNAs, one of which favors structured substrates and the other which was discovered for its activity on denatured substrates, might have specificity for this class of RNAs in vivo.

Chapter 2. ElaD

2.1 Background

The gene encoding RNase BN “rbn” was successfully mapped by our lab to an operon containing two genes – elaC and elaD. The gene elaC was subsequently renamed "rbn” (Ezraty et al. 2005). Subsequent work in our lab (R.

Fink, unpublished observations) observing the physiological properties of a strain mutated for the gene rbn using recombineering and PCR products suggested a

“6S RNA mutant-like” phenotype during stationary phase as reported by

Trotochaud and Wassarman (2004). Trotochaud & Wassarman showed that the absence of 6S RNA led to a decrease in viability during stationary phase growth.

The small non-coding 6S RNA is a molecule responsible for regulating various elements of stationary phase growth. This regulatory molecule is present in most prokaryotes. Its hairpin structure resembles an open promoter which binds to RNA polymerase to prevent RNA transcription. This part of the project sought to compare observed phenotypes of 6S RNA mutants with those of

RNase BN mutant cells. The fate of 6S RNA transcripts from stationary phase cells would be observed via Northern analysis. (Trotochaud & Wassarman, 2004;

Wassarman, 2007)

2.2 Experimental procedures

2.2.1. Bacterial Strains

Background strain is K-12 MG1655* (referred to in this work as MG1655 unless otherwise noted) and is known to be mutated for RNase I and Crl and possibly

14 15 GlpR. (Tavazoie et al. 2011) Strains from Ben Ezraty* and Kenn Rudd** are marked (Table 2.1.).

Phenotype Donor Recipient MG1655 “Wild type” n/a n/a BN* CamR, Bu33R, BN- PCR MG1655 BEZ33 CamR, BN-, Bu33R PCR MG1655 MV201** KanR, BU33R, elaC::kan-frt KEIO n/a MV274** KanR, BU33S, elaD::kan-frt KEIO n/a KEIO XT000 KanR, BU33R, elaC::kan-frt MG1655 elaC KEIO KK274 KanR, BU33S, elaD::kan-frt MG1655 elaD XT001 BU33R, ∆elaC pCP20 KK201 KO274 BU33S, ∆elaD pCP20 KK274 CK elaC KanR, BU33R, elaC::kan-frt MV201 CA265 CK elaD KanR, BU33S, elaD::kan-frt MV274 CA265 CO elaC BU33R, ∆elaC pCP20 CK elaC XT002 KanR, rnd::kan-frt KEIO** MG1655 KanR, BU33R, ∆elaC, XT003 KEIO** XT001 rnd::kan-frt XT004 ∆rnd pCP20 XT002 XT005 BU33R, ∆elaC, ∆rnd pCP20 XT003 XT006 relA::kan-frt KEIO MG1655 XT007 rbn::kan-frt KEIO CA265 XT008 rnd::kan-frt YK1829 CA265 MG1655*, RNase I-, PAP- MZ140** KEIO MG1655 (pcnB::kanFRT) MG1655, RNase I-, XT008 MZ140 MG1655 pcnB::kanFRT BU33R, ∆elaC, XT009 MZ140 XT001 pcnB::kanFRT XT010 Δrnd, pcnB::kanFRT MZ140 XT004 XT011 Δrbn, Δrnd, pcnB::kanFRT MZ140 XT005 XT012 relaxed, pcnB::kanFRT MZ140 CA265 XT013 relaxed, BN-, pcnB::kanFRT MZ140 CAN Table 2.1. Bacterial Strains. Symbol denotes strains from Ben Ezraty* and strains from KE Rudd** With exception of the

16 lab wild type MG1655, all other strains were constructed for this project. "Donor" indicates the source for genetic material used in P1 transduction, plasmid or if PCR was used to alter the strain. "Recipient" indicates the parent strain.

Strains were created with P1 transductions (as described) using linked markers or KEIO strains and 'cured' of antibiotic resistance using the pCP20 plasmid. See table 2. 2 for complete list of phage strains.

Strain Phenotype Application Plaques for wild type, Transducing phage P1vir* gene transduction at without lysogenic increased frequency genes T4 Plaques on wild type Ensure T4 sensitivity T4-Bu33 BN mutants are resistant Assess BN function Q-beta* Plaques Phage burst assay Table 2. 2. Phage Strains. Symbol denotes strains from KE Rudd*.

2.2.2. Media - Cells were grown at 37°C, unless otherwise noted, in M9 or LB

medium buffered with 10 mM TRIS, pH 7.8. Antibiotics, when present, were at

the following concentrations: ampicillin, 100 µg/mL; chloramphenicol, 35 µg/mL;

kanamycin, 50µg/mL.

2.2.3. Individual growth analysis

2.2.3.1. Growth curve analysis - To assess growth rate for both strains

independent of each other, cells were diluted from an overnight culture into fresh

LB and monitored by A600 and serial dilution at selected intervals. Colony forming

unit (CFU) per milliliter was calculated based on the dilution used and the

number of colonies was counted by hand.

2.2.3.2. Exponential to stationary phase transition analysis - To assess entry to stationary phase for both strains independent of each other. Cells were grown to an OD600 of 3.0 in LB and monitored by OD600 and serial dilution from mid log phase into stationary at 30 minute intervals. CFU per milliliter was calculated based on the dilution used and the number of colonies counted by hand. Cells were diluted to between 0.1 and 1 OD to maintain accuracy based on the liner range of the machine.

2.2.3.3. Survival during stationary phase analysis - To assess independent survival. Cells were observed by absorbance at 600 nm (A600) and serial dilution from early stationary phase at daily intervals. CFU per mL was calculated based on the dilution used and the number of colonies. Cells were diluted to between

0.1 and 1 absorbance to appreciate the liner range of the machine.

2.2.3.4. Lag phase analysis - To assess time for exit from lag phase for both strains independent of each other. Cells were grown eight hours after being diluted from an overnight culture into fresh LB. After the cells had recently entered stationary phase (about 8 hours to maintain roughly uniform numbers of

CFU/ml) they were diluted and monitored by absorbance and serial dilution until early to mid log phase. CFU per milliliter was calculated as in 2.2.3.3.. Cells were diluted to between 0.1 and 1 absorbance to maintain accuracy based on the linear range of the machine.

2.2.4. Co-culture assay - To assess significant differences in growth rate between wild type and mutant cells, overnight cultures were diluted and grown 4-

18 6 hours to ensure exponential phase cells. These cultures were grown at 37ºC after being diluted into 5 ml LB at 1:100. These cultures were diluted 1:100 and plated on LB and LB with antibiotic every 2 hours for differential screening by serial plating. CFU per milliliter was calculated based on the dilution used and the number of colonies counted by hand.

2.2.4.1. Full cycle competition assay - 25 ul from overnight cultures were inoculated to 2.5 ml of LB. These cultures were grown with aeration at 37 ºC. The culture was diluted, 50 ul into 2.5 ml of fresh media every 12 hours, and plated on LB and LB with antibiotic for differential screening by serial plating. CFU per milliliter was calculated based on the dilution used and the number of colonies was counted by hand.

2.2.4.2. Calculation of doubling times - Definitions: D = doubling time (time for the cells to divide); t = time interval in hours or minutes; n = # of times E. coli doubles during the time interval; No = # of E. coli at the beginning of the time interval; N = # of E. coli at the end of the time interval.

19 2.2.5. Gene mapping – gene maps came from EcoGene.org (Rudd et al. 2013)

2.2.6. Phage P1 transductions

2.2.6.1. Phage P1 lysates – Overnight cultures of donor strain (50mL) were combined with 1uL of phage (~109 PFU/mL) and grown at 37°C with aeration until debris became apparent indicating that cellular lysis has occurred. Typically, this takes only 5 hours. Plaque forming units (PFU) per milliliter was calculated based on the dilution used and the number of plaques counted by hand - PFU multiplied by the dilution factor and divided by the plated volume in mL..

2.2.6.2. Phage titers – phage lysates were diluted in TMC and inoculated to

42°C 0.6% agar with a phage sensitive strain of E. coli. These were plated on room temperature agar and observed for plaque forming units (PFU) after 16 hours. PFU/mL was calculated as PFU multiplied by the dilution factor and divided by the plated volume in mL.

2.2.6.3. Multiplicity of infection calculations – Multiplicity of infection (MOI) was 1.5 phage particles (based on PFU/mL) per cell (based on A600).

2.2.6.4. Transduction procedure – Transductions were performed at an MOI of 1.5:1 in a volume of 100 mL exponential culture of recipient cells, 100mL phage P1 dilution and 10 uL of 1 M CaCl2 [5 mM CaCl2]. Cells were grown without aeration at 37°C for 20 minutes. For antibiotic resistance genes requiring outgrowth, cells were spun down and grown in minimal media citrate (to chelate remaining calcium) for 20 minutes. Cells were diluted and plated on LB with antibiotic. Transductants were scored and colony PCR confirmed mutations.

2.2.7. KEIO collection and pCA24N plasmid – The KEIO collection of mutants

20 includes all non-lethal mutations of E. coli in a MG655 K-12 background. These mutations were created using a cassette for kanamycin resistance flanked by

FLP-FRT sequences. This means that these kanamycin resistance cassettes can be removed using a FLP-FRT recombinase (Baba et al. 2006).

2.2.8. DIC microscopy - Wide-field microscopy was performed to observe cell morphology. We used an Olympus fluorescence BX61 microscope equipped with

Nomarski differential interference contrast (DIC) optics (Olympus, Center Valley,

PA, USA), a U-Plan-Apo 100× objective (NA 1.35), a Roper CoolSNAP HQ camera (Photometrics, Roper Scientific, Pleasanton, CA, USA), and a 175 W

Xenon remote source with liquid light guide. Images were acquired using

SlideBook 4.01 (Intelligent Imaging Innovations, Denver, CO, USA).

2.2.9. Statistical methods

2.2.9.1. Bivariate analysis - Type 2 2-tailed t-tests were performed for experiments including only two variables to establish statistical significance.

2.2.9.2. Multivariate analysis – ANOVA followed by log-squared ratio,

Tukey's test or Pearson was used for all relevant data in experiments including more than one variable to assess statistical significance of results. Typically no y- intercept was used to calculate best fit models.

21 2.3 Results

In order to assess cell survival in rich media as previously reported by

Trotochaud & Wassarman (2004) and R. Fink (unpublished observations),

BEZ33 and MG1655 were grown individually in LB at 37ºC with aeration. They showed no obvious difference in growth until cells were starved for two or more days (Figures 2.1). This result is consistent both with those observed

Trotochaud & Wassarman under similar conditions and those results reported by our lab.

Figure 2.1. Growth of BEZ33 in and MG1655 in individual culture. Growth in buffered LB at 37°C with aeration as measured by viable count. Wild type MG1655 (WT) remains abundant where the RNase BN mutant BEZ33 drops off after a few hours in stationary phase. These are representative data from an experiment done twice.

Biomass from strain BEZ33 and MG1655 from the same cultures as in

Figure 2. 1. (grown in LB at 37ºC with aeration) were also estimated by optical density yielding puzzling results (Figures 2.2). The optical densities remained indistinguishable when comparing wild type to BEZ33 cells, while viable counts revealed significant differences (Figure 2.1). These conflicting data were resolved by bioinformatic analysis.

Figure 2.2. Growth of BEZ33 and MG1655 in individual culture. Growth in buffered LB at 37°C with aeration as measured by A600. Wild type MG165 (WT) and RNase BN mutant BEZ33 remain at similar optical density after a few hours of growth. These are representative data from an experiment done twice.

Strain BEZ33 was created with site directed mutagenesis and primers 5′-

GGGTTGCGATAACAGGGCAAGTTTCGCCCTGTTTTTAATAATAAGCAGCATA

TGAATATCCTCCTTA-3′ and

5′ATACTAGCCGGAATATTTTTTGAAACTGTATGAACTCATGGAATTAATTTGT

23 GTAGGCTGGAGCTGCTTC-3′. The underlined portion of the primer corresponds to E. coli gene addresses 2379593 - 2379642 and 2382575 –

2382528. The region encoding rbn lies between 2381608 and 2382525 and the coding sequence corresponding to elaC occupies the nucleotides between

2382713 and 2383924. The promoter for elaD corresponds to a sigma 70 and starts at position 2380676. These coordinates show that the ribosomal entry site, terminators and gene were deleted in the process (Figure 2.3).

Figure 2. 3. Gene map showing cut sites for BEZ33. The upper part of this graphic (not to scale) represents a gene map for the region containing rbn and elaD including gene addresses for the 3’- and 5’ends of both genes and the transcription start site (uppermost arrow) for elaD. The lower arrows represent the approximate location and corresponding gene addresses for the primers used to create the BEZ33 deletion mutant.

To test the hypothesis that elaD may be the source of the observed phenotype, we examined two mutants from the KEIO collection and assessed their phenotypes relative to those previously observed.

Two strains lacking rbn and elaD, respectively, were used to provide genetic material for P1 transductions into a common background (MG1655).

Both were compared in growth experiments and via DIC microscopy. The results

24 obtained by microscopy demonstrated that BEZ33 and strains lacking the elaD gene form filaments, whereas wild type MG1655 and strains lacking the elaC gene (rbn) form normal stubby rods that are typical of cells in stationary phase

(Figure 2.4).

Figure 2. 4. Microscopic analysis of cell phenotypes. MG1655 cells (WT) mutated for RNase BN (RBN), ElaC (ELA) compared to BEZ33 as visualized with DIC microscopy. These image are representative of all cells visualized. White bars represent 5 micrometers. These are representative data.

As expected, the growth profiles of the strains lacking elaC mirrored that of

BEZ33. The strain lacking RNase BN grew like the wild type (data not shown).

Because the results in Figure 2.1 so closely resembled those from

Trotochaud and Wassarman (2004), we performed molecular analysis of K-12 derivative elaC- BEZ33, MG1655 Δ elaD and MG1655 Δ elaC to ensure that 6S

RNA was not affected on a molecular level during stationary phase. Northern

Analysis of 6S RNA was performed in MG1655, K-12 Derivative ElaC- BEZ33,

25 MG1655 Δ elaD and MG1655 ΔelaC. No obvious differences were observed in migration or concentration of RNA. Three major species of RNA were observed with other minor species of RNA (Figure 2.5). This result is consistent with what is known about 6S RNA; three different isoforms of one nucleotide difference in length should be observed.

Figure 2. 5. Northern analysis of 6S RNA. RNA collected from stationary phase MG1655 cells (WT), cells mutated for RNase BN (RBN), ElaC (ELA) and BEZ33 (BEZ) as visualized by Northern analysis using a probe specific to 6S RNA. Notice there are three distinct species of RNA, 119, 120 and 121 nt long. These are representative data from an experiment performed twice.

2.4 Discussion

Based on the similarity between data from Trotochaud and Wassarman and an observation in our lab we tested the hypothesis that RNase BN mutants might have a similar phenotype to mutants for 6S RNA. We observed growth both by viable count and also via absorbence at 600 nanometers. The previous in vivo competition and individual growth and survival assays revealed encouraging results, but upon examining the phenotypic data more closely, we found the results confusing; similar absorbencies apparently conflicted with fold differences in viable count when observed in stationary phase BEZ33 cells.

Differential interference microscopy revealed that this was due to filamenting in

26 BEZ33 cells. The absorbence was the same, because the biomass was also similar. The disparity in the number of colony forming units was much lower in the filamenting cells.

Subsequent analysis of the primers used and gene mapping data available through EcoGene.org revealed that the gene for RNase BN was not the only one mutated in the BEZ33 strain. The region containing elaD had also been removed in the process. Subsequently, we followed up with comparison of the

BEZ33 strain with two in-frame mutants from the KEIO collection whose orientation preserved the promoters for all genes in the operon and the next operon – one mutated for RNase BN and one mutated for elaD. Growth assays revealed similarity between MG1655 and the cells lacking RNase BN. The optical density data reflected the viable counts, as expected. BEZ33 and cells lacking elaD also had similar profiles to one another, but that were different from those seen in MG1655 and cells lacking RNase BN. BEZ33 and elaD mutants showed a decline in CFU versus the cells lacking rbn and wild type when examined by viable count, but estimates of population by absorbance were indistinguishable.

Examining these cells via microscopy showed that BEZ33 and cells lacking elaD formed filaments, a possible septation defect, during stationary phase and that MG1655 and cells mutated for RNase BN did not and instead displayed typical stubby rod shaped cells. This cellular morphology accounts for the disparity in viable count versus optical density observed in the strains that form filaments.

27 Phenotypes observed during stationary phase, particularly filamentation, have been attributed to the ablation of expression of the gene elaD. The E. coli gene elaD has been shown to have structural similarity to deubiquitinases and also displays these properties in vivo (Catic, et al. 2007), but it is unlikely that this property observed on ubiquitin in other organisms would cause the filamenting phenotype in E. coli. This gene may play a role in another element of E. coli growth and survival, but it is not germane to the subject of this research.

The Northern analysis of 6S RNA using RNA from stationary phase cells reveals that the absence of RNase BN has no effect on the RNA transcript. All three species of 6S RNA are known to appear in E. coli and were visible and none was significantly increased or decreased relative to wild type strain

MG1655 versus background. This effectively rules out any involvement of RNase

BN on 6S RNA under these conditions.

Chapter 3. Searching for Substrates via Molecular Techniques

3.1 Background

It is known that RNase D and RNase BN can degrade RNA both in vivo and in vitro and yet other RNases are known to be more active on all of the previously examined substrates (Zhang, Deutscher 1989; Redko et al. 2007).

Because RNase D and RNase BN are capable of acting on RNA both in vivo and in vitro, we hypothesized that the substrate must be an RNA.

This section focuses on the search for a substrate at a molecular level primarily by using Northern Analysis. Based on the knowledge that RNase BN acts on small highly structured substrates. Accordingly, RNase D acts on small unstructured substrates in other organisms as well as in E. coli. Therefore, small non-coding RNAs were selected as likely substrates for RNase D. (Zhang &

Deutscher. 1988; Asha et al. 1983).

I will show that the tRNAs examined are not substrates for either of these enzymes under the conditions examined. Neither the Northern blots of the tRNA nor the tRNA nucleotidyltransferase assay to check for terminal A residue removal show any influence by these two enzymes. I will also show that many mRNAs and sRNAs are not substrates for these two enzymes during stationary phase. A long sequencing gel of total RNA from wild-type MG1655 and mutants lacking RNase BN and/or RNase D also shows no difference in migration of stable RNAs, suggesting that the absence of these ribonucleases does not cause gross changes in stable RNA.

28 29 The data will show that a small RNA, csrA mRNA, an RNA with structured and unstructured regions is a substrate for RNase D and RNase BN in vivo and in vitro. The absence of RNase D, in particular, elevates the concentration of the mRNA for CsrA protein 3- to 4-fold and doubles the half-life of the mRNA.

This section also examines possible regulatory elements of RNase D and

RNase BN by Northern blotting the mRNAs for these enzymes in a stringent and relaxed strain. I will show that RNase D is regulated in part by the stringent response and that RNase BN is expressed independently of ppGpp. I will also show that RNase D and RNase BN are expressed most abundantly during rapid growth and drop off dramatically during stationary phase.

3.2 Experimental Procedures

3.2.1. RNA purification and fractionation – RNA was purified by hot phenol extraction for total RNA and by direct phenol extraction as previously described by Hilderman & Deutscher (1974).

3.2.2. tRNA Nucleotidyltransferase assay - In 500 uL of buffer (50 mM glycine, pH 9.0; 10 mM MgCl2) 1 mM (14C) ATP and 400 mg tRNA and 0.5 ug of enzyme were combined and incubated for 30 minutes at 37°C. The contents of the tube were precipitated in ice cold 10%TCA with 5% yeast tRNA. The supernatant and pellet were chilled for 30 minutes and each assayed for activity.

3.2.3. PAGE for 12 cm and 38 cm gels

3.2.3.1. Gels - 6-15% acrylamide and 8M urea buffered with TBE were combined and solidified with TEMED and 10% APS. Gels were loaded with 2:1

30 dye to sample and run at 35 mv until the xylene cyanol reached the bottom of the gel for 12 cm and 38 cm gels. For 38 cm gels, the bottom third of the gel was retained for analysis. RNA was transferred via capillary action to filter paper for drying in a heat vacuum gel dryer or to nitrocellulose for Northern blotting.

3.2.3.2. 32P labeled RNA - E. coli cells were grown in low-phosphate medium supplemented with 32P (orthophosphoric acid) in order to radiolabel nucleic acids with 32P for analysis via densitometry in a 38 cm urea-PAGE gel.

3.2.4. Ribosome gels - 1.5% agarose in TAE or TBE buffer was solidified at room temperature and run at 60 v for 1.5 hours. Gels were stained with ethidium bromide and photographed under UV light.

3.2.5. Northern Analysis

3.2.5.1. Primers - Probes for Northern analysis were generated using oligos complementary to RNA, tested for self complementarity, self dimerization and hairpin formation and were purchased from Sigma. These oligos were labeled using γ -[32P] ATP and T4 polynucleotide kinase purchased from New England

BioLabs and used according to the manufacturer's instructions.

3.2.5.2. Densitometry – Radiolabeled materials were resolved in PAGE and visualized using a STORM 840 phosphoimaging device (GE Healthcare).

Quantification was carried out using Image J (National Institutes of Health).

31 3.3 Results

Because the most likely substrates for RNase D and RNase BN are RNA and most likely small RNA, systematic Northern analysis of RNA from mutant and wild type strains was performed using probes complementary to the rRNA transcripts, tRNA genes, mRNA genes and sRNA genes of interest. These were done on RNA collected from stationary phase cells because exponential phase growth is seemingly unaffected by mutating RNase D and RNase BN and that these RNases are more likely to be important during stationary phase growth or stress conditions.

A list of the tRNAs examined by Northern blot includes most of those in

MG1655 (Table 3.1.). Of the 73 tRNA transcripts probed, several were duplications and the probes corresponded to more than one tRNA resulting in multiple bands visualized in Northern analysis.

tRNA Primer Sequence (5'-3') AlaVUTXW ATCCCGCATAGCTCCACCA ArgZYVQ GCATCCGTAGCTCAGCTGG ArgX TAGCTCAGCTGGATAGAGCG AsnWVUT TCCTCTGTAGTTCAGTCGGT AspTUV GAGTCCCGTCCGTTCCGCCA CysT GACTCCGGAACGCGCCTCCA GlnYXVW TCCGGAACGCGCCTCCA GlyU GCGGGCGTAGTTCAATGGT GlyYXWV GCGGGAATAGCTCAGTTGGT HisR GGTGGTATAGCTCAGTTGG IleVUT AGGCTTGTAGCTCAGGTGGT IleXY GCCCCTTAGCTCAGTGGTTA LeuVTQP AAGTCCCCCCCCTCGCACCA LeuU AAGTCCCGTCCTCGGTACCA

32 LeuW AAGTCTCGCCTCCCGCACCA LeuX GAGTCCGGCCTTCGGCACCA LeuZ AAGTCCCGCTCCGGGTACCA LysZYQWVT AATCCTGCACGACCCACCA MetUT CTACGTAGCTCAGTTGGTTA MetWZYV GTAGCTCGTCGGGCTCATAA PheUV ATTCCGAGTCCGGGCACCA ProK GTGATTGGCGCAGCCTGGTA ProL GCACGTAGCGCAGCCTGGTA ProM GCGAGTAGCGCAGCTTGGTA SelC ACTCCTGTGATCTTCCGCCA SerT GAATCTCTGCGCTTCCGCCA SerU AA ATCCCCCTCTCTCCGCCA SerVXW GA ATCCCCGCCTCACCGCCA ThrTW GCTGATATAGCTCAGTTGGTA ThrU GACTTAGCTCAGTAGGTAGA ThrW ATAGCTCAGTTGGTAGAGCA ThrV CTGATATGGCTCAGTTGGTA TrpT AGTCTCTCCGCCCCTGCCA TyrVUT GAATCCTTCCCCCACCACCA ValZYXUT ATCCCGTCATCACCCACCA

Table 3.1 tRNA probes. Probes designed using sequence data from EcoGene were checked for complementarity to other regions of the genome using BLAST sequence alignment and then tested for self-complementarity, hairpin formation and self dimerization. Several bind to more than one duplicated tRNA gene. These will have more than one letter after the three letter amino acid abbreviation.

33 RNA samples were isolated by direct phenol extraction and fractionated using isopropanol to enrich for transfer RNA . Examples of blots are presented in

Figure 3. 1.

Figure 3. 1. Representative Northern blots of tRNA RNA was collected from stationary phase cells via direct phenol extraction and fractionated using isopropanol. Depending on the intensity of the signal achieved with preliminary trials using 12 ug of RNA per lane, up to 100 ug may have been used. Representative data are from experiment performed at least twice. All blots are summarized in table 3. 2.

When the tRNAs were probed using Northern blotting and phosphoimages were analyzed by densitometry, no additional bands appeared and no bands disappeared. Densitometry recorded in Table 3. 2. suggests that the absence of

RNase D and RNase BN does not affect any of these tRNA.

34 tRNA RBN/WT RND/WT MUT/WT MUTpcnB/WT

AlaVUTXW 1.05 1.09 0.80 1.06 ArgZYVQ 1.20 0.55 0.82 0.50 ArgX 0.74 0.68 0.90 1.05 AsnWVUT 1.00 0.88 1.02 0.85 AspTUV 0.93 0.95 0.76 0.91 CysT 0.95 0.94 0.68 1.00 GlnYXVW 1.09 1.36 0.68 0.62 GlyU 0.79 0.88 0.95 0.88 GlyYXWV 0.88 0.95 1.06 1.20 HisR 0.81 0.78 0.98 1.11 IleVUT 0.83 0.92 0.90 0.94 IleXY 1.12 0.72 1.24 1.00 LeuVTQP 0.74 0.66 1.49 1.04 LeuU 0.83 0.98 1.21 1.15 LeuW 1.10 1.01 1.53 0.76 LeuX 1.15 0.65 1.58 1.15 LeuZ 1.34 0.72 1.15 0.90 LysZYQWVT 1.34 0.95 1.15 0.88 MetUT 1.47 0.85 1.46 0.67 MetWZYV 0.70 0.92 1.15 1.13 PheUV 0.79 1.00 1.24 1.16 ProK 1.22 0.83 1.12 1.01 ProL 0.70 1.06 1.06 1.09 ProM 0.70 0.86 1.15 0.70 SelC 0.70 0.79 1.20 0.80 ThrTW 2.07 0.85 0.63 4.99 ThrU 0.68 0.76 0.76 1.15 ThrW 1.12 0.93 0.77 1.19 ThrV 1.06 1.19 1.12 0.82 TrpT 0.94 1.15 0.76 1.14 TyrVUT 1.03 1.07 3.05 0.84 ValZYXUT 1.15 0.79 1.51 0.87

35 Table 3. 2. Densitometry for tRNA. Northern blots using probes for tRNA were visualized using a phosphoimage analyzer and densitometry was calculated using background subtraction and compared to wild type. Densitometry listed includes ratios of MG1655 (WT) versus cells lacking RNase BN (RBN), RNase D (RND), both RNases (MUT) and both RNases and pcnB (MUTpcnB).

To ensure that the most sensitive assay was being used to assess tRNA

(particularly in light of the fact that many of the probes recognized more than one duplication of the same tRNA gene) and in order to examine whether the tRNAs were fully functional, we also determined whether terminal AMP residues might have been lost from the tRNA, a portion of the same tRNA preparation used for the Northern blots was assayed for AMP incorporation in the presence of tRNA nucleotidyltransferase. This assay would detect any differences in the amount of terminal A residues between the mutant and wild type cells. The data in Table

3.3 suggest that terminal A residues are not affected in a strain mutated for

RNase D and RNase BN under the conditions described.

Mutant for MG1655 RNase D and RNase BN % incorporation (based 18±2.1 15±2.4 on CPM/mg)

Table 3. 3. Results of tRNA nucleotidyltransferase assay. Incorporation of 14C ATP by tRNA nucleotidyltransferase was measured for tRNA from MG1655 and a strain lacking RNase BN and D. Specific activity of 4x103 CPM/nMol was added and incorporation was monitored by scintillation based on acid soluble and acid precipitable counts. Values are averaged from three separate experiments. Error was less than 16% for all species of RNA tested.

After RNase D and RNase BN were effectively eliminated as the primary enzymes responsible for maintaining, maturing and degrading tRNA during stationary phase, we moved on to small non-coding RNAs (sRNA). Probes used for Northern analysis of sRNA are summarized in Table 3.4. These studies revealed that none of the RNAs tested, except for csrA mRNA, are affected by deletion of RNase BN or RNase D.

Gene Primer Sequence (5'-3') ssrS 5' GCGAACATCTCAGAGAAATTTTGTCTTCA ssrS m GCTTCTCGGACGGACCGAGCATGCTCACCAAC ssrS 3' GATAAGAAGGGAATCTCCGAGATGCCGCCGC rprA AAGCATGGAAATCCCCTGAGTGAAACAACGAA micF GCTATCATCATTAACTTTATTTATTACC micC GTTATATGCCTTTATTGTCACAGATTTTATTTTC rydC CTTCCGATGTAGACCCGTATTCTTCGCCTGTACCA gadY GGTCCCCTATGCCGGGTTTTTTT rseX TGATGCTTCCGTTATTAGCCTTTTATCGTC istR-1 CGACTGACGAAACCTCGCTCCGGCGGGG dsrA CAUCAGAUUUCCUGGUGUAACGAAUUUUUUAAGUG omrA CCCAGAGGTATTGATTGGTGAGATTATTCG omrB CCCAGAGGTATTGATAGGTGAAGTCAACTTC ryhB AGACCCUCGCGGAGAACCUGAAAGC sgrS CCCCATGCGTCAGTTTTATCAGCACTATTTT micA GAAAGACGCGCATTTGTTATCATCATCCCT gcvB CCTGAGCCGGAACGAAAAGTTTT oxyS GAAACGGAGCGGCACCTCTTTTAACCCTT csrB GAGTCAGACAACGAAGTGAACATCAGGATGAT glmY GCUCAUUCACCGACUUAUGUCAGCCCCUUC glmZ GCTCATTCCATCTCTTATGTTCGCCTTA carB GAGTCAGACAACGAAGTGAACATCAGGATGAT csrC CGCTAACAGGAACAATGACTCAGGATGAG gcvA CCT GAG CCG GAA CGAAAAGTTTTATCGGAA

37 cyaR GCTGAAAAACATAACCCATAAAATGCTA dicF TTTCTGGTGACGTTTGGCGGTATCAGT eyeA GCAAGCAAGGATAAAGAGTGCGACG sokA AACTTTGATTTATAGTCAGGTGGG sokB GCTAGGTTCATTCGTTGGCCTCGGTTGATA sokC TTCAGCATATAGGAGGCCTCGGGTT spf GTAGGGTACAGAGGTAAGATGTTCTATC symR AGTCATAACTGCTATTCTCCAGGA rttR TCCCTGAACTTCCCAACGAATCCGC rybB ACTGCTTTTCTTTGATGTCCCCATTTTGTGGA ryeC GTGAGGGTTAGGGAGAGGTTTCCCCC ryeD GTGAGGGTAGAGCGGGGTTTCCCCCG rygC AGCGCTGAAACTTATGAGTAACAGTACA rygD ACAAGGGTGAGGGAGGATTTCTCCC rygE ACAAGGGTAAGGGAGGATTTCTCCC psrO CTAATGACAACCCTAACCCAGCTCTATG ryeA TAACAGATAAAAAGAGACCGAACACGATT ryeB GCTGATGACCACCACGCTTTTTATTGACCA sroB TCGCTTAAAGTGACGGCATAATAATAA sroH UGCAGCAGACUGAAGAAAUUC tff GACTTCCGATCCATTTCGTATACACA isrA TGAAATCTGTCACTGAAGAAAATTGGCAAC ryfD CAAGACGATCCGGTACGCGTGATTTTCT ryhA CGGCCTGAAAAACAGTGCTGTGCCCTT ryjA ATCAACACCAACCGGAACCTCCACCAC ryjB TCATCCGTCGTTGACTCCATGCCGATT psrN AAGGGGAGTAACTTCATTGCCGGTCGAT psrD TTTTTTTCCATCAGATATAGCGTATTG rdlA GTTCTGGTTCAAGATTAGCCCCCGTTCT rdlB GTCTGGTTTCAAGATTAGCCCCCGTTCTG rdlC GTCTGGTTTCAAGATTAGCCCCCGTTTTGTTGT rdlD GTCTAGAGTCAAGATTAGCCCCCGTGG rybA CTCAAGGATCGACGGGATTAGCAA rydB TTATCGCCCCTTCAAGAGCTAAGCCACTG ryfA GCCCTTTCCGCCGTCTCGCAAACG ryfB CGTTATTGAAGATTTTGCTGTGCTTTACAC ryfC TGCTGCACAAAATTAAAGTTAAAAAGTAAAA

38 Table 3. 4. sRNA probes. Probes designed using sequence data from EcoGene were checked for complementarity to other regions of the genome using BLAST sequence alignment and then tested for self-complementarity, hairpin formation and self dimerization.

For Northern blots of sRNA, total RNA was isolated by direct phenol extraction and fractionated using isopropanol to enrich for low molecular weight

RNAs (since rRNA makes up the majority of RNA in a cell). These RNAs were then resolved on acrylamide gels, blotted to nitrocellulose membranes and examined by Northern analysis as noted in the methods section. Example blots are shown in Figure 3. 2.

Figure 3. 2. Representative Northern blots of sRNA RNA was collected from stationary phase wild type (WT), mutant for RNase BN (RBN), mutant for RNase D (RND) and RNase D and RNase BN cells (MUT) via direct phenol extraction and fractionated using isopropanol.

39 Representative data from experiment performed twice. All blots are summarized in Table 3. 6.

When the sRNAs were probed, no additional bands appeared, no bands disappeared under the conditions tested and densitometry values recorded in

Table 3. 5. These showed no differences between the wild type and mutant strains.

Gene MUT/WT MUT/WTpcnB ssrS 5' 1.043 1.143 ssrS mid 1.038 0.884 ssrS 3' 1.515 0.906 rprA 0.665 0.948 micF 0.896 0.871 micC 1.345 0.922 rydC 0.932 0.948 gadY 1.143 0.805 rseX 1.027 1.109 istR-1 0.852 0.837 dsrA 0.894 1.128 omrA 1.353 1.235 omrB 1.119 1.123 ryhB 0.864 0.896 sgrS 1.022 0.313 micA 1.022 0.493 gcvB 0.499 0.311 oxyS 0.111 1.230 csrB 1.059 0.990 glmY 1.111 1.282 glmZ 1.296 1.217 carB 0.138 1.002 csrC 0.846 0.846 gcvA 1.088 0.922 cyaR 0.951 1.002 dicF 1.385 1.038 eyeA 0.217 1.048 sokA 0.948 1.381 sokB 1.027 0.894 sokC 0.102 1.349

40 spf 0.923 0.975 symR 0.493 0.846 rttR 1.017 1.166 rybB 0.929 1.321 ryeC 1.498 1.088 ryeD 0.922 0.922 rygC 1.587 1.166 rygD 1.031 0.894 rygE 1.088 0.511 psrO 0.871 1.217 ryeA 0.948 1.104 ryeB 1.094 1.435 sroB 0.975 1.084 sroH 1.337 0.990 tff 1.300 1.450 isrA 0.173 1.185 ryfD 0.995 0.846 ryhA 1.554 0.948 ryjA 0.951 1.119 ryjB 0.922 0.922 psrN 0.896 0.896 psrD 0.846 0.896 rdlA 1.247 1.059 rdlB 0.922 1.619 rdlC 1.053 1.143 rdlD 0.974 0.846 rybA 0.935 1.002 rydB 0.859 0.896 ryfA 1.163 1.181 ryfB 1.074 0.871 ryfC 0.940 1.033 Table 3. 5. Densitometry for sRNA. RNA was collected from stationary phase cells via direct phenol extraction and fractionated using isopropanol. Northern blots using probes for sRNA were visualized using a phosphoimage analyzer and densitometry was calculated using background subtraction and compared to MG1655. Depending on the signal strength of the radiolabeled probe in preliminary experiments using 12ug of RNA, up to

41 100ug was used. Values are averages of at least two experiments, but some were done three or more times. Densitometry listed includes ratios of MG1655 (WT) versus cells lacking RNase BN and RNase D (MUT) and both RNases and pcnB (MUTpcnB)Error was less than 50% for all species of RNA tested. These data suggest that none of the sRNAs tested are affected by deleting

RNase D and RNase BN based on a statistical analysis. Since these data effectively eliminated many other possible RNAs as substrates, we tested several mRNAs, pseudogenes and genes of unknown function (y-genes) using Northern analysis. Probes for the RNAs are listed in Table 3. 6.

Gene Primer Sequence (5'-3') luxS ATGCCGTTGTTAGATAGCTT cyaR GCTGAAAAACATAACCCATA fkpR ATGAAATCACTGTTTAAAGT lptD ATGAAAAAACGTATCCCCAC ssrA GGGGCUGAUUCUGGAUUCGACGGGAUUUGCG yjeT CATGTTAGGAAAACGATT ytfK AAAAGTGAAGGATAAGCT yfjL AACAAATGAAAGACTTAAA yeeP' TACCGTGACATTCTGCCCTGAA yjhG ATGTCTGTTCGCAATATTTTT yihP ATGAGTCACATCACAACGGAA rtcB GAATTACTGACCACTGAAAA csrA TCAGACAACGAAGTGAACATCAGGAT Table 3.6. mRNA and other probes. Probes designed using sequence data from EcoGene were checked for complementarity to other regions of the genome using BLAST sequence alignment and then tested for self- complementarity, hairpin formation and self dimerization.

42 Total RNA was isolated by hot phenol extraction and spun down to remove ribosomes. This was fractionated using isopropanol. The high molecular weight fraction was used in blots where RNAs larger than 120 nucleotides were being probed. The low molecular weight fraction was retained for use in blots where RNAs smaller than 120 nucleotides were being probed. RNA was resolved using 6-9% acrylamide gel electrophoresis. These mRNAs, pseudogenes and y- genes were blotted to nitrocellulose and subjected to Northern analysis. When the mRNA, pseudogenes and y-genes were probed, no additional bands appeared, no bands disappeared and densitometry values are recorded in Table

3. 7.

Gene MUT/WT MUT/WTpcnB luxS 0.888 0.955 cyaR 1.183 1.257 fkpR 1.238 0.948 lptD 1.138 1.188 ssrA 1.204 1.213 yjeT 1.238 1.446 ytfK 0.930 1.373 yfjL 1.048 1.309 yeeP' 1.093 1.105 yjhG 1.303 1.339 yihP 1.313 1.304 rtcB 1.467 1.183 csrA 1.170 1.388 Table 3. 7. Densitometry for mRNA and other RNA. RNA was collected from stationary phase cells via direct phenol extraction and fractionated using isopropanol. Northern blots using probes for sRNA were visualized using a phosphoimage analyzer and densitometry was calculated using background subtraction and compared to MG1655. Values are averages of two experiments, except

43 for csrA, ssrA and luxS. Error was less than 50% for all species of RNA tested. Representative Northern blots from Table 3. 7 are shown in Figure 3. 3.

Figure 3. 3. Representative Northern blots of mRNA and other RNA. RNA was collected from stationary phase cells via hot phenol extraction and fractionated using isopropanol after ribosomes were removed from lysates via high speed centrifugation. Each lane was loaded with up to 100 ug depending on the intensity of the signal from preliminary blots using only 12 ug. These are representative data from an experiment performed at least twice. All blots are summarized in Table 3. 7.

44 The mRNA for CsrA was the only RNA that was probed and found to be significantly different in mutants for RNase BN and RNase D when compared with wild type. A representative Northern blot shows the increase for the RNase

D mutant and to a lesser extent (if at all) RNase BN in Figure 3. 4. Also, a double

RNase D, RNase BN mutant reproducibly shows a greater increase than an

RNase D mutation by itself.

Figure 3. 4. Northern blot of csrA mRNA. RNA collected from stationary phase MG1655 cells (WT), cells mutated for RNase BN (RBN), RNase D (RND) and both (MUT) as visualized by Northern analysis using a probe specific to CsrA mRNA. 30 ug of RNA was added to each lane. Quantification ([RNA]) was done using ImageQuant. These are representative data from an experiment performed four times.

To observe whether any other stable RNA species are affected by the absence of RNase D and RNase BN, polyacrylamide gel electrophoresis of total

RNA labeled with 32P was run in 38 cm sequencing gels from cells collected after they had entered stationary phase. These gels revealed no species of RNA that was affected enough to be detected by this assay (Figure 3. 5.).

Figure 3. 5. 38 cm PAGE of total RNA. 32P labeled RNA fractionated with isopropanol and resolved in 12% PAGE with TAE. MG1655 (WT), mutants for RNase BN (RBN), cells mutated for RNase D (RND) and double mutant cells (MUT) during stationary phase. These are representative data from an experiment performed at least twice.

46 After surveying many possible in vivo substrates for RNase D and RNase

BN, we reasoned that expression patterns of RNase BN and RNase D mRNA might reveal some indications as to the when these two enzymes are important in E. coli. We also wanted to see which regulatory elements are involved in their expression. Northern blots were done on RNA isolated at several time points during lag phase, exponential and stationary phase growth from the wild type strain (MG1655) and a relaxed strain which lacks the ability to synthesize ppGpp

(CA265) using probes for the mRNA of RNase D and RNase BN (Figures 3. 6. and 3.7). These showed that RNase BN is not affected by (p)ppGpp. They also showed that RNase D is regulated by the increase of (p)ppGpp that accompanies stress. Most importantly, these data show that both RNase D and

RNase BN are expressed most abundantly during rapid growth (between 1 and 4 hours) and that levels of their mRNA drop off as growth begins to slow between 8 and 16 hours.

Figure 3. 6. A. Northern of RNase D mRNA in MG1655. RNA (100ug per lane) from MG1655 isolated via hot phenol extraction after high speed centrifugation to remove ribosomes and isopropanol fractionation to remove tRNA, resolved via PAGE, blotted to nitrocellulose and probed with a radiolabeled probe for RNase D mRNA. RNA was collected during lag phase (1 hour), exponential phase (2 & 4 hours) and during stationary phase (8 and 16). Concentrations of RNA [RNA] relative to the first time point were calculated by densitometry versus background. These are representative data from an experiment performed twice. B. Northern of RNase D mRNA in CA265. RNA (100ug/lane) from CA265 isolated via hot phenol extraction after high speed centrifugation to remove ribosomes and isopropanol fractionation to remove tRNA, resolved via PAGE, blotted to nitrocellulose and probed with a radiolabeled probe for RNase D mRNA. RNA was collected during lag phase (1 hour), exponential phase (2 & 4 hours) and during stationary phase (8 and 16). Concentrations of RNA [RNA] relative to the first time point were calculated by densitometry versus background. These are representative data from an experiment performed twice.

Figure 3. 7. A. Northern of RNase BN mRNA in MG1655. RNA (100ug per lane) from MG1655 isolated via hot phenol extraction after high speed centrifugation to remove ribosomes and isopropanol fractionation to remove tRNA, resolved via PAGE, blotted to nitrocellulose and probed with a radiolabeled probe for RNase BN mRNA. RNA was collected during lag phase (1 hour), exponential phase (2 & 4 hours) and during stationary phase (8 and 16). Concentrations of RNA [RNA] relative to the first time point were calculated by densitometry versus background. These are representative data from an experiment performed twice. B. Northern of RNase BN mRNA in CA265. RNA (100ug per lane) from CA265 isolated via hot phenol extraction after a high speed centrifugation to remove ribosomes and isopropanol fractionation to remove tRNA, resolved via PAGE, blotted to nitrocellulose and probed with a radiolabeled probe for RNase BN mRNA. RNA was collected during lag phase (1 hour), exponential phase (2 & 4 hours) and during stationary phase (8 and 16). Concentrations of RNA [RNA] relative to the first time point were calculated by densitometry versus background. These are representative data from an experiment performed twice.

3.4 Discussion

Northern analysis of tRNA, rRNA, mRNA, sRNA, pseudogenes and y- genes revealed only one RNA that was affected by eliminating RNase D and

49 RNase BN. csrA mRNA was upregulated 3.5 fold in double mutants on average and was primarily affected by the absence of RNase D. This will be discussed further in chapter 5.

The results for total RNA, low molecular weight and high molecular weight fractions in 38 cm gels did not reveal any significant increase, decrease or shifting of bands in strains mutated for RNase BN or D relative to wild type strains.

Northern blots of RNase D and RNase BN mRNAs suggest that RNase D expression is influenced by the presence of (p)ppGpp and that RNase BN is not.

RNase D and RNase BN are likely more important during exponential phase as they are most abundant during rapid growth. This will likely be a subject for further study and has already been confirmed that some sRNAs are more abundant in mutants lacking RNase D and RNase BN during rapid growth conditions (unpublished observations, Tanmay Dutta via direct communication).

Chapter 4. Searching for Phenotypes

4.1 Background

It is well known that geneticists delete genes in order to find a phenotype associated with that mutation. We began this work hypothesizing that mutants lacking RNase D and RNase BN must have a phenotype in E. coli. Previous work examining the role of RNase BN in a cell did not reveal any differences between wild and cells lacking RNase BN except for cells infected with the mutant T4 phage Bu33. Those studies therefore focused primarily on molecular evidence of the absence of the RNase (Callahan, et al. 2000, Callahan & Deutscher 1996,

Ruven, Deutscher 1993).

Finding a phenotype for cells lacking RNase BN or RNase D may suggest a role for these two enzymes and a suite of RNAs that may be their substrates in vivo. A number of conditions were tested including but not limited to cold and heat shock, osmolar shock, starvation shock, and long-term starvation. This chapter will show that mutants lacking RNase D are less motile and form quantifiable aggregates during stationary phase and that mutants lacking RNase

BN have slightly reduced survival in low pH conditions.

4.2 Experimental Procedures

4.2.1. MTT assay - BioLog® Phenotype MicroArray™ plates PM1 (carbon sources), PM2A (carbon sources), PM3B (nitrogen sources), PM4A

(phosphorous and sulfur sources), PM9 (osmolytes) and PM10 (pH) were all inoculated with MG1655 or XT003 (Δrbn, Δrnd) according to the manufacturer's

50 51 instructions (http://www.biolog.com/pmTechDesOver.html) and then compared for absorbency at A495 in a spectrophotometer after samples had been diluted to between 0.1 and 1.0 absorbance to ensure that readings were within the linear range of the device.

4.2.2. Antibiotic sensitivity and resistance

4.2.2.1. SensiDisc Assays - Antibiotic sensitivity was measured by applying

SensiDisc™ (BD Diagnostics) containing antibiotics to an LB agar plate topped with 2.5ml 0.6% LB agar that had been inoculated with the strain of choice.

Plates were scored visually after 16 hours.

4.2.2.2. Gradient Plates – gradient plates were created by pouring 18ml of plain agar in LB or M9 with 0.2% glucose with one edge of the plate resting on a pencil. After this agar had congealed, another 18mL of agar media was plated with an appropriate concentration of antibiotic while the plate rested on a flat surface.

4.2.3. Spectinomycin Assay – Performed as previously described by Sat &

Engleberg-Kulka (2001).

4.2.4. Phage burst assay - Phage burst size was measured by first calculating the PFU/mL for each phage stock being used. The MOI was adjusted to 2:1 phage to host. After adsorption in a calcium rich media, sodium citrate was added to chelate remaining calcium ions and deter additional phage binding and then PFU/mL was assessed again for the resulting supernatant and calculated as the burst size as predicted by the MOI and number of resulting PFU/mL. For a list of phage strains, see Table 2. 2.

52 4.3 Results

A BioRad phenotypic microarray with plates PM1A, PM2A, PM3B, PM4A,

PM9, PM10 was used to examine carbon sources, nitrogen sources, phosphorous and sulfur sources, osmolytes and pH conditions anticipating that one or more may reveal a disadvantage for cells lacking RNase BN and RNase

D. No dramatic differences were observed in any of the 564 conditions tested using the manufacturer’s suggested cutoff of 5-fold or a more stringent cutoff of

3-fold (Table 4. 1. A-C.).

Well Plate PM1A MUT/WT Plate PM2A MUT/WT A2 L-Arabinose 0.53 Chondroitin Sulfate C 1.06 A3 N-Acetyl-D-Glucosamine 0.71 a-Cyclodextrin 0.96 A4 D-Saccharic Acid 0.45 b-Cyclodextrin 1.01 A5 Succinic Acid 1.07 g-Cyclodextrin 1.00 A6 D-Galactose 0.56 Dextrin 0.98 A7 L-Aspartic Acid 0.98 Gelatin 0.96 A8 L-Proline 0.44 Glycogen 1.02 A9 D-Alanine 0.53 Inulin 1.06 A10 D-Trehalose 0.96 Laminarin 1.04 A11 D-Mannose 0.57 Mannan 0.97 A12 Dulcitol 1.05 Pectin 1.01 B1 D-Serine 1.04 N-Acetyl-D-Galactosamine 1.01 B2 D-Sorbitol 1.00 N-Acetyl-Neuraminic Acid 0.99 B3 Glycerol 0.99 b-D-Allose 1.02 B4 L-Fucose 1.01 Amygdalin 1.04 B5 D-Glucuronic Acid 1.01 D-Arabinose 1.05 B6 D-Gluconic Acid 1.02 D-Arabitol 1.04 D,L-a-Glycerol- B7 Phosphate 1.03 L-Arabitol 1.03 B8 D-Xylose 0.99 Arbutin 1.13 B9 L-Lactic Acid 1.04 2-Deoxy-D-Ribose 1.00 B10 Formic Acid 1.04 i-Erythritol 1.05 B11 D-Mannitol 1.04 D-Fucose 1.02

53 3-0-b-D-Galacto-pyranosyl- B12 L-Glutamic Acid 1.05 D-Arabinose 1.03 C1 D-Glucose-6-Phosphate 1.03 Gentiobiose 1.03 D-Galactonic Acid-g- C2 Lactone 1.02 L-Glucose 0.89 C3 D,L-Malic Acid 0.92 Lactitol 1.05 C4 D-Ribose 1.05 D-Melezitose 1.02 C5 Tween 20 1.01 Maltitol 1.00 C6 L-Rhamnose 0.98 a-Methyl-D-Glucoside 1.04 C7 D-Fructose 0.87 b-Methyl-D-Galactoside 0.99 C8 Acetic Acid 0.97 3-Methyl Glucose 0.99 C9 a-D-Glucose 1.03 b-Methyl-D-Glucuronic Acid 1.05 C10 Maltose 1.01 a-Methyl-D-Mannoside 0.99 C11 D-Melibiose 0.98 b-Methyl-D-Xyloside 0.96 C12 Thymidine 1.04 Palatinose 16.05 D1 L-Asparagine 0.96 D-Raffinose 1.00 D2 D-Aspartic Acid 0.98 Salicin 1.02 D3 D-Glucosaminic Acid 0.97 Sedoheptulosan 1.02 D4 1,2-Propanediol 1.05 L-Sorbose 1.04 D5 Tween 40 0.71 Stachyose 0.94 D6 a-Keto-Glutaric Acid 0.76 D-Tagatose 18.83 D7 a-Keto-Butyric Acid 1.05 Turanose 1.03 D8 a-Methyl-D-Galactoside 1.03 Xylitol 1.03 D9 a-D-Lactose 0.77 N-Acetyl-D-Glucosaminitol 1.00 D10 Lactulose 1.05 g-Amino Butyric Acid 1.04 D11 Sucrose 0.98 d-Amino Valeric Acid 1.04 D12 Uridine 0.93 Butyric Acid 1.01 E1 L-Glutamine 1.05 Capric Acid 0.98 E2 m-Tartaric Acid 1.01 Caproic Acid 1.00 E3 D-Glucose-1-Phosphate 1.03 Citraconic Acid 1.02 E4 D-Fructose-6-Phosphate 1.05 Citramalic Acid 1.04 E5 Tween 80 1.03 D-Glucosamine 0.96 a-Hydroxy Glutaric Acid- E6 g-Lactone 0.96 2-Hydroxy Benzoic Acid 6.44 E7 a-Hydroxy Butyric Acid 1.03 4-Hydroxy Benzoic Acid 0.95 E8 b-Methyl-D-Glucoside 0.96 b-Hydroxy Butyric Acid 0.92 E9 Adonitol 1.04 g-Hydroxy Butyric Acid 0.91 E10 Maltotriose 0.91 a-Keto-Valeric Acid 0.89

54 E11 2-Deoxy Adenosine 1.00 Itaconic Acid 0.91 E12 Adenosine 1.01 5-Keto-D-Gluconic Acid 0.96 F1 Glycyl-L-Aspartic Acid 1.03 D-Lactic Acid Methyl Ester 1.01 F2 Citric Acid 1.02 Malonic Acid 1.04 F3 m-Inositol 0.64 Melibionic Acid 0.99 F4 D-Threonine 0.96 Oxalic Acid 1.07 F5 Fumaric Acid 1.02 Oxalomalic Acid 1.05 F6 Bromo Succinic Acid 0.97 Quinic Acid 1.00 F7 Propionic Acid 1.01 D-Ribono-1,4-Lactone 0.97 F8 Mucic Acid 1.02 Sebacic Acid 1.19 F9 Glycolic Acid 0.98 Sorbic Acid 0.88 F10 Glyoxylic Acid 0.98 Succinamic Acid 1.03 F11 D-Cellobiose 1.02 D-Tartaric Acid 1.05 F12 Inosine 0.98 L-Tartaric Acid 0.82 G1 Glycyl-L-Glutamic Acid 1.00 Acetamide 0.90 G2 Tricarballylic Acid 0.99 L-Alaninamide 0.93 G3 L-Serine 1.00 N-Acetyl-L-Glutamic Acid 0.88 G4 L-Threonine 1.05 L-Arginine 1.00 G5 L-Alanine 0.97 Glycine 1.04 G6 L-Alanyl-Glycine 1.02 L-Histidine 1.00 G7 Acetoacetic Acid 0.97 L-Homoserine 1.02 N-Acetyl-b-D- G8 Mannosamine 0.99 Hydroxy-L-Proline 1.05 G9 Mono Methyl Succinate 0.97 L-Isoleucine 1.01 G10 Methyl Pyruvate 0.97 L-Leucine 0.96 G11 D-Malic Acid 0.94 L-Lysine 0.98 G12 L-Malic Acid 0.98 L-Methionine 1.05 H1 Glycyl-L-Proline 0.94 L-Ornithine 1.04 p-Hydroxy Phenyl Acetic H2 Acid 0.92 L-Phenylalanine 1.04 m-Hydroxy Phenyl Acetic H3 Acid 1.02 L-Pyroglutamic Acid 1.05 H4 Tyramine 1.02 L-Valine 0.75 H5 D-Psicose 1.04 D,L-Carnitine 0.74 H6 L-Lyxose 0.98 Sec-Butylamine 0.77 H7 Glucuronamide 1.02 D.L-Octopamine 0.70 H8 Pyruvic Acid 1.02 Putrescine 0.71 H9 L-Galactonic Acid-g- 1.03 Dihydroxy Acetone 0.74

55 Lactone H10 D-Galacturonic Acid 1.00 2,3-Butanediol 0.74 H11 Phenylethylamine 1.01 2,3-Butanone 0.80 H12 2-Aminoethanol 1.03 3-Hydroxy 2-Butanone 0.82

Table 4. 1. A. MTT assay quantification for PM1 and PM2A. Wild type MG1544 and double mutant cells were inoculated to separate Phenotype MicroArrays™ plates and incubated overnight at 37°C without aeration. Aliquots from the control wells (MG1655) were compared to the plates where the strain was mutated for RNase D and RNase BN. The fold difference was recorded in this table. This experiment was done only once.

Well Plate PM3B MUT/WT Plate PM4A MUT/WT A2 Ammonia 0.95 Phosphate 1.05 A3 Nitrite 0.58 Pyrophosphate 0.96 A4 Nitrate 0.61 Trimetaphosphate 1.03 A5 Urea 0.43 Tripolyphosphate 1.03 A6 Biuret 0.65 Triethyl Phosphate 0.98 A7 L-Alanine 0.61 Hypophosphite 0.92 Adenosine- 2’- A8 L-Arginine 0.55 monophosphate 1.04 Adenosine- 3’- A9 L-Asparagine 0.64 monophosphate 0.98 Adenosine- 5’- A10 L-Aspartic Acid 0.60 monophosphate 0.99 Adenosine- 2’,3’-cyclic A11 L-Cysteine 0.62 monophosphate 1.00 Adenosine- 3’,5’-cyclic A12 L-Glutamic Acid 1.01 monophosphate 1.04 B1 L-Glutamine 0.77 Thiophosphate 1.03 B2 Glycine 0.79 Dithiophosphate 1.05 B3 L-Histidine 0.48 D,L-a-Glycerol Phosphate 0.93 B4 L-Isoleucine 0.62 b-Glycerol Phosphate 0.95 B5 L-Leucine 0.61 Carbamyl Phosphate 0.98 B6 L-Lysine 0.50 D-2-Phospho-Glyceric Acid 1.03 B7 L-Methionine 0.62 D-3-Phospho-Glyceric Acid 0.93 Guanosine- 2’- B8 L-Phenylalanine 0.54 monophosphate 0.89

56 Guanosine- 3’- B9 L-Proline 0.38 monophosphate 0.99 Guanosine- 5’- B10 L-Serine 0.42 monophosphate 1.03 Guanosine- 2’,3’-cyclic B11 L-Threonine 0.58 monophosphate 1.03 Guanosine- 3’,5’-cyclic B12 L-Tryptophan 0.66 monophosphate 1.01 C1 L-Tyrosine 0.57 Phosphoenol Pyruvate 1.01 C2 L-Valine 0.52 Phospho- Glycolic Acid 1.01 C3 D-Alanine 0.49 D-Glucose-1-Phosphate 1.01 C4 D-Asparagine 0.63 D-Glucose-6-Phosphate 0.99 2-Deoxy-D-Glucose 6- C5 D-Aspartic Acid 0.52 Phosphate 0.98 D-Glucosamine-6- C6 D-Glutamic Acid 0.63 Phosphate 1.03 C7 D-Lysine 0.98 6-Phospho-Gluconic Acid 0.95 C8 D-Serine 0.73 Cytidine- 2’- monophosphate 0.83 C9 D-Valine 0.50 Cytidine- 3’- monophosphate 1.03 C10 L-Citrulline 0.82 Cytidine- 5’- monophosphate 1.00 Cytidine- 2’,3’-cyclic C11 L-Homoserine 0.55 monophosphate 1.04 Cytidine- 3’,5’-cyclic C12 L-Ornithine 0.59 monophosphate 1.05 N-Acetyl-L-Glutamic D1 Acid 0.58 D-Mannose-1-Phosphate 1.00 N-Phthaloyl-L- D2 Glutamic Acid 0.52 D-Mannose-6-Phosphate 0.99 D3 Cysteamine-S- D3 L-Pyroglutamic Acid 0.62 Phosphate 1.05 D4 Hydroxylamine 0.46 Phospho-L-Arginine 0.93 D5 Methylamine 0.55 O-Phospho-D-Serine 0.91 D6 N-Amylamine 0.55 O-Phospho-L-Serine 1.03 D7 N-Butylamine 0.66 O-Phospho-L-Threonine 0.99 D8 Ethylamine 1.43 Uridine- 2’- monophosphate 1.00 D9 Ethanolamine 0.43 Uridine- 3’- monophosphate 0.98 D10 Ethylenediamine 0.61 Uridine- 5’- monophosphate 0.97

57 Uridine- 2’,3’- cyclic D11 Putrescine 0.55 monophosphate 1.05 Uridine- 3’,5’- cyclic D12 Agmatine 0.46 monophosphate 1.04 E1 Histamine 0.56 O-Phospho-D-Tyrosine 0.63 E2 b-Phenylethyl- amine 0.64 O-Phospho-L-Tyrosine 0.97 E3 Tyramine 0.63 Phosphocreatine 0.98 E4 Acetamide 0.65 Phosphoryl Choline 1.04 E5 Formamide 0.63 O-Phosphoryl-Ethanolamine 1.00 E6 Glucuronamide 0.59 Phosphono Acetic Acid 0.98 2-Aminoethyl Phosphonic E7 D,L-Lactamide 0.45 Acid 0.99 Methylene Diphosphonic E8 D-Glucosamine 0.52 Acid 0.98 Thymidine- 3’- E9 D-Galactosamine 0.52 monophosphate 1.04 Thymidine- 5’- E10 D-Mannosamine 0.55 monophosphate 0.95 N-Acetyl-D- E11 Glucosamine 0.63 Inositol Hexaphosphate 0.97 N-Acetyl-D- Thymidine 3’,5’- cyclic E12 Galactosamine 0.60 monophosphate 1.05 N-Acetyl-D- F1 Mannosamine 1.00 Negative Control 1.05 F2 Adenine 1.38 Sulfate 1.00 F3 Adenosine 3.16 Thiosulfate 1.03 F4 Cytidine 0.60 Tetrathionate 1.05 F5 Cytosine 0.55 Thiophosphate 0.98 F6 Guanine 0.55 Dithiophosphate 1.04 F7 Guanosine 0.37 L-Cysteine 0.95 F8 Thymine 0.44 D-Cysteine 1.01 F9 Thymidine 0.42 L-Cysteinyl-Glycine 1.03 F10 Uracil 0.47 L-Cysteic Acid 1.04 F11 Uridine 0.55 Cysteamine 0.81 F12 Inosine 0.58 L-Cysteine Sulfinic Acid 0.98 G1 Xanthine 0.62 N-Acetyl-L-Cysteine 1.02 G2 Xanthosine 0.46 S-Methyl-L-Cysteine 0.97 G3 Uric Acid 0.54 Cystathionine 1.04 G4 Alloxan 0.57 Lanthionine 1.01

58 G5 Allantoin 0.61 Glutathione 1.04 G6 Parabanic Acid 0.59 D,L-Ethionine 0.91 D,L-a-Amino-N- G7 Butyric Acid 0.65 L-Methionine 0.83 g-Amino-N-Butyric G8 Acid 0.57 D-Methionine 1.00 e-Amino-N-Caproic G9 Acid 0.62 Glycyl-L-Methionine 1.05 D,L-a-Amino- G10 Caprylic Acid 0.63 N-Acetyl-D,L-Methionine 1.05 d-Amino-N-Valeric G11 Acid 0.52 L- Methionine Sulfoxide 0.94 a-Amino-N-Valeric G12 Acid 0.63 L-Methionine Sulfone 1.04 H1 Ala-Asp 0.61 L-Djenkolic Acid 1.05 H2 Ala-Gln 0.59 Thiourea 1.02 H3 Ala-Glu 0.53 1-Thio-b-D-Glucose 1.00 H4 Ala-Gly 0.40 D,L-Lipoamide 0.96 H5 Ala-His 0.38 Taurocholic Acid 1.01 H6 Ala-Leu 0.50 Taurine 1.00 H7 Ala-Thr 0.66 Hypotaurine 1.04 p-Amino Benzene Sulfonic H8 Gly-Asn 0.56 Acid 1.04 H9 Gly-Gln 0.46 Butane Sulfonic Acid 1.03 2-Hydroxyethane Sulfonic H10 Gly-Glu 0.59 Acid 1.05 H11 Gly-Met 0.56 Methane Sulfonic Acid 1.04 H12 Met-Ala 0.58 Tetramethylene Sulfone 0.52

Table 4. 1. B. MTT assay quantification for PM3B and PM4A. Wild type MG1544 and double mutant cells were inoculated to separate Phenotype MicroArrays™ plates and incubated overnight at 37°C without aeration. Aliquots from the control wells (MG1655) were compared to the plates where the strain was mutated for RNase D and RNase BN. The fold difference was recorded in this table. This experiment was done only once.

59 Well Plate PM9 MUT/WT Plate PM10 MUT/WT A1 NaCl 1% 0.58 pH 3.5 0.45 A2 NaCl 2% 0.44 pH 4 0.49 A3 NaCl 3% 0.43 pH 4.5 0.47 A4 NaCl 4% 0.45 pH 5 0.76 A5 NaCl 5% 0.91 pH 5.5 0.61 A6 NaCl 5.5% 0.62 pH 6 0.64 A7 NaCl 6% 0.39 pH 7 0.60 A8 NaCl 6.5% 0.54 pH 8 0.64 A9 NaCl 7% 0.85 pH 8.5 0.66 A10 NaCl 8% 0.94 pH 9 0.72 A11 NaCl 9% 0.64 pH 9.5 0.80 A12 NaCl 10% 0.78 pH 10 1.50 B1 NaCl 6% 0.79 pH 4.5 0.40 B2 NaCl 6% + Betaine 0.81 pH 4.5 + L-Alanine 0.52 NaCl 6% + N-N Dimethyl B3 Glycine 0.74 pH 4.5 + L-Arginine 0.54 B4 NaCl 6% + Sarcosine 0.76 pH 4.5 + L-Asparagine 0.95 NaCl 6% + Dimethyl pH 4.5 + L-Aspartic B5 sulphonyl propionate 0.57 Acid 0.64 pH 4.5 + L-Glutamic B6 NaCl 6% + MOPS 0.59 Acid 0.69 B7 NaCl 6% + Ectoine 0.74 pH 4.5 + L-Glutamine 0.89 B8 NaCl 6% + Choline 0.65 pH 4.5 + Glycine 0.61 NaCl 6% + Phosphoryl B9 Choline 0.56 pH 4.5 + L-Histidine 0.80 B10 NaCl 6% + Creatine 1.08 pH 4.5 + L-Isoleucine 0.76 B11 NaCl 6% + Creatinine 0.69 pH 4.5 + L-Leucine 0.88 B12 NaCl 6% + L- Carnitine 0.60 pH 4.5 + L-Lysine 0.48 C1 NaCl 6% + KCl 0.66 pH 4.5 + L-Methionine 1.05 pH 4.5 + L- C2 NaCl 6% + L-Proline 0.64 Phenylalanine 0.42 NaCl 6% + N-Acetyl L- C3 Glutamine 0.64 pH 4.5 + L-Proline 0.40 NaC1 6% + β-Glutamic C4 Acid 0.51 pH 4.5 + L-Serine 0.58

60 NaC1 6% + γ–Amino -N- C5 Butyric Acid 0.64 pH 4.5 + L-Threonine 0.37 C6 NaC1 6% + Glutathione 0.71 pH 4.5 + L-Tryptophan 0.60 C7 NaCl 6% + Glycerol 0.63 pH 4.5 + L-Citrulline 0.51 C8 NaC1 6% + Trehalose 0.54 pH 4.5 + L-Valine 0.54 NaC1 6% + pH 4.5 + Hydroxy-L- C9 Trimethylamine-N-oxide 0.46 Proline 0.81 NaC1 6% + C10 Trimethylamine 0.81 pH 4.5 + L-Ornithine 0.87 pH 4.5 + L- C11 NaCl 6% + Octopine 0.63 Homoarginine 0.64 C12 NaC1 6% + Trigonelline 1.01 pH 4.5 + L-Homoserine 0.98 pH 4.5 + Anthranilic D1 Potassium chloride 3% 0.88 Acid 0.93 D2 Potassium chloride 4% 0.52 pH 4.5 + L-Norleucine 0.87 D3 Potassium chloride 5% 0.58 pH 4.5 + L-Norvaline 0.47 pH 4.5 + α- Amino-N- D4 Potassium chloride 6% 0.46 Butyric Acid 0.48 pH 4.5 + p-Amino- D5 Sodium sulfate 2% 0.91 Benzoic Acid 0.51 D6 Sodium sulfate 3% 0.55 pH 4.5 + L-Cysteic Acid 0.63 D7 Sodium sulfate 4% 0.97 pH 4.5 + D-Lysine 0.52 pH 4.5 + 5- D8 Sodium sulfate 5% 0.57 Hydroxylysine 0.75 pH 4.5 + 5- D9 Ethylene glycol 5% 0.56 Hydroxyyryptophan 0.77 pH 4.5 + D,L- D10 Ethylene glycol 10% 0.88 Diaminopimelic Acid 0.92 pH 4.5 + D11 Ethylene glycol 15% 0.53 Trimethylamine-N-oxide 0.71 D12 Ethylene glycol 20% 0.59 pH 4.5 + Urea 0.55 E1 Sodium formate 1% 0.56 pH 9.5 0.65 E2 Sodium formate 2% 0.46 pH 9.5 + L-Alanine 2.97 E3 Sodium formate 3% 0.68 pH 9.5 + L-Arginine 0.48 E4 Sodium formate 4% 0.49 pH 9.5 + L-Asparagine 0.35 pH 9.5 + L-Aspartic E5 Sodium formate 5% 0.91 Acid 0.66 E6 Sodium formate 6% 0.52 pH 9.5 + L-Glutamic 0.90

61 Acid E7 Urea 2% 0.65 pH 9.5 + L-Glutamine 0.57 E8 Urea 3% 0.46 pH 9.5 + Glycine 0.38 E9 Urea 4% 1.03 pH 9.5 + L-Histidine 0.60 E10 Urea 5% 2.06 pH 9.5 + L-Isoleucine 0.95 E11 Urea 6% 0.60 pH 9.5 + L-Leucine 0.79 E12 Urea 7% 0.93 pH 9.5 + L-Lysine 0.90 F1 Sodium Lactate 1% 0.45 pH 9.5 + L-Methionine 0.89 pH 9.5 + L- F2 Sodium Lactate 2% 0.63 Phenylalanine 0.50 F3 Sodium Lactate 3% 2.72 pH 9.5 + L-Proline 0.51 F4 Sodium Lactate 4% 33.00 pH 9.5 + L-Serine 0.81 F5 Sodium Lactate 5% 0.61 pH 9.5 + L-Threonine 0.45 F6 Sodium Lactate 6% 0.39 pH 9.5 + L-Tryptophan 0.66 F7 Sodium Lactate 7% 1.05 pH 9.5 + L-Tyrosine 0.85 F8 Sodium Lactate 8% 0.74 pH 9.5 + L-Valine 0.75 pH 9.5 + Hydroxy-L- F9 Sodium Lactate 9% 0.93 Proline 0.86 F10 Sodium Lactate 10% 0.69 pH 9.5 + L-Ornithine 0.29 pH 9.5 + L- F11 Sodium Lactate 11% 1.07 Homoarginine 0.34 F12 Sodium Lactate 12% 0.95 pH 9.5 + L-Homoserine 0.86 Sodium Phosphate pH 7 pH 9.5 + Anthranilic G1 20mM 2.70 acid 0.40 Sodium Phosphate pH 7 G2 50mM 1.33 pH 9.5 + L-Norleucine 1.48 Sodium Phosphate pH 7 G3 100mM 0.36 pH 9.5 + L-Norvaline 0.46 Sodium Phosphate pH 7 G4 200mM 0.50 pH 9.5 + Agmatine 0.66 Sodium Benzoate pH 5.2 G5 20mM 3.98 pH 9.5 + Cadaverine 1.05 Sodium Benzoate pH 5.2 G6 50mM 0.69 pH 9.5 + Putrescine 0.75 Sodium Benzoate pH5.2 G7 100mM 0.34 pH 9.5 + Histamine 0.41 Sodium Benzoate pH 5.2 pH 9.5 + G8 200mM 0.88 Phenylethylamine 0.33 G9 Ammonium sulfate pH8 0.64 pH 9.5 + Tyramine 0.65

62 10mM Ammonium sulfate pH 8 G10 20mM 0.34 pH 9.5 + Creatine 0.38 Ammonium sulfate pH 8 pH 9.5 + G11 50mM 0.64 Trimethylamine-N-oxide 0.87 Ammonium sulfate pH8 G12 100mM 0.58 pH 9.5 + Urea 0.45 H1 Sodium Nitrate 10mM 0.89 X-Caprylate 0.48 H2 Sodium Nitrate 20mM 0.72 X–α-D-Glucoside 0.68 H3 Sodium Nitrate 40mM 0.28 X-β-D-Glucoside 0.88 H4 Sodium Nitrate 60mM 0.54 X-α-D-Galactoside 0.45 H5 Sodium Nitrate 80mM 1.02 X-β-D-Galactoside 0.59 H6 Sodium Nitrate 100mM 0.50 X-α- D-Glucuronide 0.22 H7 Sodium Nitrite 10mM 0.78 X-β- D- Glucuronide 0.46 H8 Sodium Nitrite 20mM 0.78 X-β-D-Glucosaminide 0.72 H9 Sodium Nitrite 40mM 0.74 X-β-D-Galactosaminide 0.99 H10 Sodium Nitrite 60mM 0.79 X-α-D-Mannoside 0.54 H11 Sodium Nitrite 80mM 0.66 X-PO4 0.83 H12 Sodium Nitrite 100mM 0.81 X-SO4 0.91

Table 4. 1. C. MTT assay quantification for PM9 and PM10. Wild type MG1544 and double mutant cells were inoculated to separate Phenotype MicroArrays™ plates and incubated overnight at 37°C without aeration. Aliquots from the control wells (MG1655) were compared to the plates where the strain was mutated for RNase D and RNase BN. The fold difference was recorded in this table. This experiment was done only once.

63 In order to ascertain if there was a disadvantage conferred by the absence of RNase BN and RNase D in the presence of antibiotics, both MG1655 and the double mutant strain were grown in liquid media, inoculated to soft agar and plated on LB. Individual SensiDiscs each containing ampicillin, carbenecillin, chloramphenicol, clindamicin, kanamycin, penecillin, neomycin or novobiocin were placed on the plates when the soft agar had solidified. After 24 hours of growth at 37°C the zone of clearing around the discs was measured in millimeters on 4 edges of the cleared zone and averaged (Table 4. 3). These data are all consistent with sensitivity for these antibiotics according to the manufacturer's specifications. There was no statistically significant difference between wild type and mutants in this assay.

Wild type zone Mutant zone Antibiotic of clearing of clearing (mm) (mm) ampicillin 14±2 15±2 carbenecillin 12±2.5 12±2.5 chloramphenicol 17±2 18±2 clindamicin 16±1.5 16±21.5 kanamycin 18±2 18±2 penecillin 22±2 22±2 neomycin 15±1 15±1 novobiocin 16±2 16±2

Table 4. 2. Antibiotic sensitivity and resistance. SensiDisc™ assay on MG1655 (wild type) and strain mutated for RNase D and RNase BN was grown to mid-log phase, inoculated to soft LB agar and plated over LB agar. Disc were placed on the soft agar and after 24 hours incubation at 37°C the zone of clearing was measured. This experiment was done twice.

64 A 2001 research paper by Sat, Engleberg-Kulka, et al. revealed that the deletion of RNase BN conferred a selective advantage to E. coli when treated with spectinomycin under specific conditions. This result was reproduced in our lab (Figure 4. 2.). We added the wash step (WASH) thinking that the effect of spectinomycin might result from the antibiotic that had bound to the ribosomes and that a wash could allow the antibiotic to dissociate from 30s ribosomal subunits. These data suggest that this is the case. An identical assay (including the wash step) substituting chloramphenicol for spectinomycin suggested that this phenomenon is unique to spectinomycin (data not shown).

Figure 4. 1. Spectinomycin resistance assay. Cells treated with high concentrations of spectinomycin in MG1655 (WT), an RNase BN mutant (MUT) and a strain mutated for RNase D and RNase BN (DBL). The 2 left panels are duplicates after treatment with spectinomycin (SPEC) and the right 2 panels are duplicates after aliquots from the treatment panels were washed with PBS (WASH). These are representative data.

Because previous experimentation had revealed differences when cells shift from rapid to slow growth, we devised an assay to perturb homeostasis by rapidly depleting nutrients from cells, starving them and then reintroducing them to medium containing a carbon source. This alternating nutrient upshift and downshift experiment was done in co-culture of cells to increase the sensitivity of the assay. We found that cells mutated for RNase D were outperformed by wild type cells or cells lacking RNase BN alone when reintroduced to rich media. A co-culture assay in mineral salts media containing a carbon source revealed that mutants for RNase D had a longer lag phase relative to wild type cells or cells lacking RNase BN alone. This prolonged lag phase accounts the differences observed during the upshift/downshift experiment. Long term competition where cells are allowed to cycle through lag, exponential and stationary phase growth in co-culture revealed these differences are apparent in rich media and mineral salts media with glucose (data not shown) after significant number of cycles (see

Chapter 6.).

66 RNase BN is known to participate in the maturation of phage encoded

RNA as in the case of the amber suppressor tRNA in the mutant T4 phage Bu33.

Therefore, we tested phage burst size in RNase BN and RNase D mutant strains to determine whether or not RNase D and RNase BN might play a role in phage infection or participate in resistance to infection by phage. A predetermined number of phage was combined with wild type and mutant cells and incubated long enough for one cycle of infection. Then PFU per mL was determined by plating in soft agar that had been inoculated with the wild type strain. There was no difference between wild type and double mutant strains indicating that RNase

BN and D are not important for infection with these phage strains. (Table 4. 3.)

Mutant for RNase D phage MG1655 and RNase BN T4 120±18 120±21 P1vir 81±15 83±12 Q-beta 168±8 162±13 Table 4.3. Phage Burst Assays. Phage burst values for MG1655 and ΔrbnΔrnd in the same background. Cells were combined with phage at a multiplicity of infection of adjusted to 2: 1 phage to host. After adsorption, washing and plating, PFU/mL was assessed again for the resulting supernatant and calculated as the rate infection as predicted by the MOI and number of resulting PFU/mL. These data are the average of three different experiments.

Serial infection of surviving mutant and wild type E. coli with the same phage did not reveal any differences that indicate improved resistance to infection with phage (data not shown) suggesting that no 'memory' of the infection was created by the presence or absence of either of the two ribonucleases.

67 We examined motility as an important aspect of E. coli biology whose role can be separate from rapid growth and biofilm formation. We measured motility by stab culture using 2,3,5-triphenyltetrazolium chloride (TTC) dye. This colorless dye is absorbed by a living cell and metabolized to create a red color. Cells were grown overnight in liquid culture with aeration at 37°C and then spent an additional 24 hours without aeration to encourage motility (Figure 4. 3.). The strain lacking RNase D was less motile than the wild type strain based on the qualitative migration of cells out from the original stab (darkest red) in the middle of the tube.

Figure 4. 2. Stab cultures for motility using TTC dye. Cells from wild type (WT) and RNase D mutant (RND) were grown overnight in LB with aeration and then left for an additional 24 hours without aeration all at 37°C. Cells were stabbed with a sterile needle into soft LB agar with TTC dye. Compare with the un-inoculated tube (B). Representative data from experiment performed at least twice.

68 A number of additional stress conditions and metabolic substrates were also examined in order to determine if RNase D and RNase BN were important for growth or survital under these conditions. The growth and survival of wild type and double mutant were compared during exponential, stationary, recovery from stationary phase and lag phase in rich media (LB) and minimal media (M9 with 0.2% glucose, succinate, acetate) individually. The double mutant and wild type were also compared during rapid growth in rich media (LB) at the following temperatures; 4°C, 21°C, 37°C, 41°C and with the antibiotics naladixic acid, rifampicin, amikacin and doxycycline (data not shown). They were also compared in concentrations of NaCl ranging from 0 to 2M, concentrations of NH4Ac ranging from 0 to 2M, and at pH values ranging from 4 to 10 in rich media buffered with

CAPS, MOPS, HEPES and TRIS (data not shown). Low pH showed a subtle, but reproducible disadvantage in cells mutated for RNase BN. Rapid growth in minimal media (M9); glucose, succinate, acetate, pyruvate; starvation in rich and minimal media, recovery from starvation in rich and minimal media also revealed no advantage for the wild type strain (data not shown). None of these showed any significant differences; however, some additional phenotypes will be presented in chapters 5 and 6.

4.4 Discussion

Because RNase BN and D do not confer any apparent advantage during exponential phase growth, a number of stress conditions and metabolic substrates were examined in order to determine if the enzymes were important

69 for growth or survival with respect to these metabolites or stress conditions. Co- cultures were used to improve the sensitivity of the assays for most of these growth conditions.

The phenotypic microarrays included a very broad range of metabolic and osmotic stress conditions. The only clear pattern was a subtle but consistent decrease in growth of the double mutant in acidic conditions. This result has been confirmed by another member of the lab (Tanmay Dutta, unpublished via direct communication). This defect is subtle in the microarray data. This may be consistent with the high frequency of false positive results associated with this kind of assay. These data were confirmed by growing cells in co-culture in rich media in pH ranging from 4 to 10 (HEPES and TRIS buffered); wild type MG1655 did appear to have a subtle advantage over mutants for RNase BN, but not

RNase D (data not shown).

The growth and survival of wild type and cells lacking RNase D and

RNase BN were compared during exponential, stationary, recovery from stationary phase and lag phase in rich media (LB) and minimal media (M9 with

0.2% glucose, succinate, acetate) individually and at the following temperatures;

4°C, 21°C, 37°C and 41°C. These experiments revealed no differences between mutant and wild type strains.

The double mutant and wild type were also compared during rapid growth in rich media (LB) with several antibiotics that rely on a range of mechanisms to inhibit bacterial growth and division. These assays revealed no differences between wild type and strains lacking RNase BN and RNase D.

70 They were also compared in concentrations of NaCl ranging from 0 to 2M and concentrations of NH4Ac ranging from 0 to 2M, which conformed to the corresponding results in the microarray table where no significant differences were observed. Rapid growth in minimal media (M9); glucose, succinate, acetate, pyruvate; starvation in rich and minimal media, recovery from starvation in rich and minimal media also revealed no advantage for the wild type strain, consistent with the phenotypic microarray data.

The observed phenotypes included the aggregation of cells in cells lacking

RNase D during stationary phase in rich media, which will be explored further in chapter 5, and the slowed recovery from lag phase observed in cells mutated for

RNase D and RNase BN, which will be explored in chapter 6. The reduction in motility and subtle defects in growth under high pH conditions also merit further research and discussion. Reproducing the results of Sat, Engleberg-Kulka et al. showed that mutants for RNase BN but not RNase D have some resistance to spectinomycin and that the phenotype seems to be unique to that antibiotic. This requires further study to determine the mechanism by which this is achieved.

This and other studies in this chapter certainly show that the effects of removing RNase BN and RNase D are subtle.

Chapter 5. csrA Substrate and Clumping Phenotype

5.1 Background

In chapter 3 we showed that CsrA mRNA concentrations were increased in cells lacking RNase D and to a lesser extent in cells lacking RNase BN. We will show that CsrA mRNA is a substrate for RNase D and RNase BN in vitro and in vivo.

We will also show that cells lacking RNase D aggregate during stationary phase based on DIC microscopy when compared with wild type MG1655 cells as previously discussed in chapter 4. In this section we show that the cell aggregates can be resolved with the addition of 0.04% TWEEN-20 prior to plating. The increase in csrA mRNA resulting from the ablation of RNase D expression was subsequently linked to the phenotype of aggregation. Increasing the concentration of csrA mRNA exogenously confirmed that the clumping phenomenon co-occurred with the increase in csrA mRNA which we'd previously linked to the absence of RNase D. We also show that other phenotypes associated with cellular processes not associated with increases in CsrA protein are also affected by deleting RNase D including synthesis of extracellular matrix, motility and concentrations of cyclic-di-GMP.

We will show that the synthesis of cyclic-di-GMP appears to be elevated in strains mutated for RNase D and motility is reduced in strains lacking RNase D.

This means that other aspects of metabolism controlled by unidentified regulatory elements are affected by mutating RNase D and to a lesser extent, by

RNase BN.

71 72 5.2 Experimental Procedures

5.2.1. Alternative Carbon sources - M9 mineral salts media was supplemented with 0.2% glucose, 50 mM pyruvate, or 50 mM succinate.

5.2.2. Cloning of csrA into pHC79 plasmid - The plasmid pHC79 was grown in

MG1655 and isolated by alkaline lysis. PCR was used to amplify the 186 base csrA gene and the 235 base region upstream. This fragment was gel purified. pHC79 was cleaved with ScaI. Blunt end ligation was used to integrate the PCR generated fragment into the plasmid.

5.2.3. Cell adherence assays - In a 96 well microtiter plate, MG1655 and the mutant for RNase BN and D were inoculated to M9 with 2% glucose and LB. The plate was incubated and wells were assayed in triplicate at 36 and 72 hours. The supernatant fluid was aspirated gently, treated with 0.2% TWEEN-20 and plated for enumeration. The remaining contents of the well were disrupted by pipetting with PBS, treated with 0.2% TWEEN-20 and plated for enumeration. The number of free cells (supernatant) and adherent cells (remaining contents) were used to calculate the rate of cell adherence.

5.2.4. Paper Chromatography assay - Strains were grown in LB with aeration at

37ºC or 32ºC and spotted onto 3 mM filter paper (Whatman). The paper was clipped to a rack and placed into a glass chamber with t-amyl alcohol: formic acid: ddi water at 6:4:1. Papers were air dried in a fume hood after the solvent had migrated within a few centimeters of the top of the paper. These were examined and photographed under UV light. The cyclic-di-GMP used as a standard was a generous gift from Delia Gutman.

73 5.2.5. Extracellular matrix formation assay - Crystal violet staining to quantify extracellular matrix formation was done according to O'Toole, 2011. Cells were grown in a microtiter dish in 40 µL of LB without aeration. Wells were inoculated with wild type or mutant strains that had been grown overnight in liquid culture

(~16 hours) at 37ºC with aeration and then allowed to sit for an additional 24 hours without aeration. After inoculation, microtiter plates were incubated for 24 hours before the liquid part of the culture was aspirated; wells were washed with

PBS, then stained with crystal violet, washed with PBS and then destained with

30% acetic acid. Aliquots of the destaining solution were measured at 550 nm.

5.2.6. Detergent and buffers for plating procedures

5.2.6.1. Buffers for pH maintenance – CAPS, TRIS or HEPES were used at

10 mM.

5.2.6.2. Detergents – TWEEN-20 was used at 0.04% to disrupt aggregates before plating. No cells were grown with detergents unless otherwise specified.

5.2.7. Microscopy for clumping - Cells were viewed under a Leica DMIL inverted contrasting microscope. The optimized phase contrast was very efficient for visualizing cells. Fields were isolated at random and every unit within the field was scored whether it contained only one cell or was an aggregate of 2 or more cells. At least 80 units were scored for each experiment.

5.2.8. Western analysis - performed according to Romeo, et. al. (2007) with help troubleshooting from W. Liang.

5.2.9. Transcription of CsrA mRNA in vitro - Synthesis of csrA transcript was done using the Ambion® MEGAshortscript™ Kit (AM1354) according to the

74 manufacturer's protocol. Product was cleaned with Sigma Aldrich® GenElute™ kit per the manufacturer's instructions.

5.2.10. in vivo and in vitro molecular analysis

5.2.10.1. Standards - 3H tRNA was used as a standard for methylene blue staining and Northern analysis. RiboRuler [Fermentas SM1831] Low Range RNA

Ladder was used to visualize RNA between 100 and 1000 bases. A probe for 5S

RNA (rrfA-H) is used to visualize 120 nt and used as a binding control.

Radiolabeled, probes prepared by Sigma were used to show 61, 35 and 25 nt.

5.2.10.2. in vivo analysis- Cells were grown to mid-log phase and then treated with 200 ug/mL rifampicin to stop further transcription. Aliquots were collected at selected time points. RNA was isolated by direct phenol extraction and fractionated using isopropanol. 6% SDS-PAGE gels were run at 35 mV with

5 ug of total RNA per lane.

5.2.10.3. in vitro analysis - 1 ug of enzyme and 2 ng substrate were combined in 10 mM Tris HCl (pH 7.5), 5 mM MgCl2, 200 mM KAcetate. Enzymes were provided courtesy of Arun Malhotra and Tanmay Dutta. Substrates were labeled with 32P or 3H.

75 5.3 Results

Long term growth and survival were shown to be affected by deleting

RNase D in chapter 4. To elucidate the nature of this defect we tracked growth over several days after cells had been previously starved. Cells were grown in rich media that had been buffered with TRIS via serial plating. Cells were plated both without (closed symbols) and with (open symbols) treatment with the non- ionic detergent TWEEN-20 before plating (Figure 5.1.). These data show that the differences in growth disappear upon first treating cells with TWEEN-20 prior to plating.

Figure 5. 1. Survival during stationary phase. MG1655 (WT), cells lacking RNase BN (RBN), cells lacking RNase D (RND) or both (MUT) were grown overnight. Cells were enumerated using viable count on rich agar once a day. Open symbols represent cells washed with detergent before plating to resolve aggregated cells. These are representative data from an experiment performed twice.

76 The unusual growth pattern observed in Figure 5. 1. was resolved by microscopic analysis. Wild type and mutant strains lacking RNase BN and/or

RNase D were starved for several days and cells were then observed via microscopy. Microscopic analysis revealed that the cells formed quantifiable aggregates when RNase D was absent (Figure 5. 2.). Also visible in Figure 5. 1. are cells washed with a gentle detergent and then placed in a vortex for thirty seconds which resolved cell aggregates, allowing them to be plated for more accurate enumeration. Note, however, that some cells did not survive the detergent treatment.

Figure 5. 2. Quantification of cell aggregation during stationary phase. MG1655 (WT), cells lacking RNase BN (RBN), cells lacking RNase D (RND) or both (MUT) were grown overnight. Cell aggregates were enumerated using light microscopy where fields were selected at random and the units in each field were scored whether the unit consisted of an individual cell or 2 or more cells in an aggregate. These data are representative from an experiment performed twice.

77 Chapter 3 (Figure 3. 4.) revealed that the mRNA for the protein CsrA was elevated in strains lacking RNase D and to a lesser extent RNase BN. To confirm this result, Northern blots of csrA mRNA were performed on the low molecular weight fraction of isopropanol fractionated RNA collected from stationary phase cells grown on rich media (Figure 5. 3.). These data show that on average the mRNA for csrA increases 3- to 4- fold in strains mutated for RNase D. The slight increase in csrA mRNA is also reproducible in the strain lacking RNase BN. The strain lacking RNase BN and RNase D mirrors this subtle increase due to the absence of RNase BN in addition to the significant increase attributed to the absence of RNase D.

Figure 5. 3. Northern blot of csrA mRNA. RNA was fractionated with isopropanol and resolved in 12% PAGE with TAE. The gel was blotted to nitrocellulose and then was incubated with a radiolabeled probe complementary to csrA mRNA. MG1655 (WT), mutants for RNase BN (RBN), cells mutated for RNase D (RND) and double mutant cells (MUT) were quantified using densitometry [RNA]. These are representative data from an experiment performed four times.

78 Existing data from Romeo, et al. (2013) suggest that increasing CsrA protein could perturb physiological phenomena including, but not limited to, biofilm formation (as alluded to by the observations in Figure 5. 2.). This led us to examine the increase in CsrA protein in the double mutant strain relative to the wild type using Western analysis with an antibody for CsrA protein (a generous gift from T. Romeo). Protein was collected from an aliquot of the same cells used to make the Northern blot shown in Figure 5. 3. This was used in Western blots of CsrA protein to search for any effect on the protein in mutant strains (Figure 5.

4.). The blots showed that CsrA increased 3.3- to 3.5-fold on average in an rnd mutant strain and a rbn, rnd double mutant. Only a small increase was observed in a rbn mutant strain.

Figure 5. 4. Western blot of CsrA protein. Western blot of CsrA protein using 12 ug protein fractionated by centrifugation to remove large proteins and run in SDS-PAGE. MG1655 (WT), mutants for RNase BN (RBN), cells mutated for RNase D (RND) and double mutant cells (MUT) were quantified using densitometry [protein]. These are representative data from an experiment performed three times.

79 To eliminate the possibility that the observed increase in RNA is not a direct effect of deleting RNase D, we determined whether purified RNase BN or

RNase D could act on csrA mRNA (Figure 5. 5.). These data show that RNase D can digest csrA mRNA in vitro. RNase BN is also capable of digesting csrA mRNA in vitro, but it is less active on the substrate.

Figure 5. 5. In vitro digestion of CsrA mRNA. Acid soluble counts released during an in vitro digestion of tritiated in vitro transcribed CsrA mRNA digested with purified RNase D (RND) or RNase BN (RBN) shows that in vitro transcribed CsrA mRNA is a substrate for RNase D and RNase BN. About 85% of the incorporated activity was released for RNase D. A no-enzyme control (CONTROL) was included without added enzyme. These data are representative from an experiment carried out three times.

80 The fact that csrA mRNA is a substrate for RNase BN and to a greater extent RNase D in vitro led us to examine the half-life for csrA mRNA in vivo.

Northern blots of csrA mRNA at various times after the addition of rifampicin (200 mg/mL) to halt further transcription allowed us to determine the half-life of the

RNA in living cells (Figure 5. 5.) The half-life in mutants (10 minutes) is roughly double that observed in wild type cells (5.5 minutes).

Figure 5. 6. In vivo digestion of csrA mRNA. Rifampicin was added to MG1655 (WT), cells lacking RNase BN (RBN), cells lacking RNase D (RND) and cells lacking RNase D and RNase BN (MUT). Aliquots were collected at selected time points (min), RNA was extracted via direct phenol extraction, fractionated with isopropanol and resolved by PAGE and probed in a Northern blot with a primer complementary to CsrA mRNA. Densitometry [RNA] was performed on the phosphoimage versus background. This is a representative result from an experiment carried out three times.

These two experiments showed that csrA mRNA is a substrate for RNase

D and to a lesser extent for RNase BN in vitro and in vivo. Subsequently we confirmed that the increase of the csrA mRNA and the increase in CsrA protein were directly responsible for the previously observed phenotype (chapter 4).

81 To ascertain whether or not this was the case, the gene for csrA was cloned into the pHC79 cosmid with a generous section of DNA predicted to contain promoter and terminators for the gene. The insertion of this gene disrupted the cosmid’s tetracycline resistance gene and left the second cassette conferring resistance to ampicillin intact, allowing for screening using selective media. The pHC79 cosmid is temperature sensitive and it's copy number correlates directly to the temperature and therefore the expression of the gene can be controlled by adjusting the temperature. The cosmid containing the construct (pHC79::csrA) was electroporated into competent cells and grown in rich media with ampicillin and compared for levels of csrA mRNA via Northern analysis (Figure 5. 6.).

Figure 5. 7. Phenocopy of csrA mRNA levels in vivo with pHC79::csrA. Wild type cells (WT) and cells containing the cosmid including the csrA mRNA sequence and native promoter (pHC), and RNase D mutant (RND) were grown overnight at 32°C or 37°C. RNA was collected, resolved by urea PAGE and probed with a oligo complementary for csrA mRNA. These results were quantified using ImageQuant. These data are averaged from multiple experiments.

82 These data indicate that the level of expression in the pHC79::csrA cosmid at 32°C is comparable to that seen at either 32°C or 37°C in cells mutated for

RNase D and RNase BN as well as RNase D alone. Because these values were similar, we proceeded to examine the previously detected phenotypes by comparing wild-type cells (WT) with wild type containing an empty vector

(pHC79) or wild type containing the pHC79::csrA cosmid (pHC79:: csrA) at 32°C.

We used the most direct measure of cell aggregation by visualizing the cells using microscopy (Figure 5. 8.). These data indicate that the elevation of csrA mRNA from a plasmid alone can cause an increase in aggregation relative to wild type cells.

Figure 5. 8. Quantification of cell aggregation with pHC79::csrA. Cells containing an empty vector (MG1655 (pHC79)) or the vector containing the csrA mRNA and its native promoter and compared with the control (MG1655) and grown at 32°C to express levels of csrA mRNA similar to those expressed by an RNase D mutant. Representative data from experiment performed at least twice.

83 Other elements of cellular metabolism and morphology known to be affected by

CsrA were also examined. Cultures were grown in LB and M9 0.2% glucose in 96 well microtiter plates at 37°C for 24 hours without aeration and assayed by aspirating the liquid supernatant. The supernatant was then serially diluted and plated for enumeration of viable cells. The wells were washed and settled and adherent cells were enumerated in the same fashion as the supernatant. The same amount of cells in strains lacking RNase BN and RNase D (6%±1 at 36 hours, 3%±1 at 72 hours) adhered to the walls of the wells relative to the wild type (4%±1 at 36 hours, 5%±1. at 72 hours). The amount of cells in the supernatant for cells lacking RNase BN, cells lacking RNase D and cells lacking both RNases were indistinguishable from the wild type (data not shown). Though these cells did not adhere to surfaces or remain in suspension in numbers significantly different from wild type as determined from ANOVA followed by

Tukey’s test, they did adhere to one another. Subsequently, we assayed the ability ot cells to form extracellular matrix.

84 The formation of extracellular matrix was measured by growing cells in microtiter dish wells overnight at 32°C (the optimal temperature to mimic csrA mRNA expression via pHC79::csrA copy number) without aeration. Biofilm formation was measured by aspirating the liquid part of the culture, washing the well with phosphate buffered saline (PBS) and then by staining the well with crystal violet. After washing the wells again with PBS, the wells were destained with 30% acetic acid and absorbance of the destaining fluid was measured at

A550 (Figure 5. 9.). The data showed that strains that have elevated csrA mRNA due to either deletion of RNase D or ectopic overexpression from a plasmid form extracellular matrix at a lower rate than wild type cells, cells lacking RNase BN or wild type cells containing an empty vector.

Figure 5. 9. Quantification of extracellular matrix formation with crystal violet stain. Wild type cells (WT), cells lacking RNase BN (RBN), cells lacking RNase D (RND), double mutant cells (MUT), wild type cells containing the empty vector (E) and vector containing the CsrA gene and native promoter (pHC) were grown overnight at 32°C without aeration, wells were washed, stained with crystal violet, washed again and destained with an acetic acid solution. The absorbance of this solution was measured at Å550. This is averaged data from an experiment performed four times. Statistical significance was confirmed with ANOVA.

85 Cyclic-di-GMP, a messenger molecule, is also affected by changes in levels of CsrA. CsrA expression is known to correlate indirectly to levels of cyclic- di-GMP (c-di-GMP) in vivo. To see if this was also true of our mutants for RNase

D, cells were grown overnight and subjected to paper chromatography using

CMP and c-di-GMP as markers (Figure 5. 9.). These data did not correlate with those seen for mutants of csrA. In strains with CsrA expressed from the pHC79 plasmid or the vector alone, there was a slight increase in c-di-GMP and strains where RNase D was mutated there was a dramatic increase in c-di-GMP.

Figure 5. 10. Paper chromatography for c-di-GMP. Cells from wild type (W), cells containing the empty vector (V), cells mutated for RNase D (D) and cells containing the pHC79::csrA plasmid (P) were grown overnight in LB with aeration and then spotted to Whatman filter paper. Compare with the CTP control and c-di-GMP control. Results were photographed using an ultraviolet lamp. These are representative data from an experiment performed twice.

86 5.4 Discussion

The relationship between RNase D and CsrA is clear - RNase D controls the levels of csrA mRNA early in stationary phase. The aggregation observed during stationary phase was quantifiable during microscopic examination. Cells mutated for RNase D showed a significant increase in the number of cells that aggregated during stationary phase relative to wild type MG1655 cells.

This phenotype coincided with an increase in csrA mRNA as shown in

Northern blots using RNA collected from wild type MG1655 cells and MG1655 cells mutated for RNase D and RNase BN. These showed that csrA mRNA levels were elevated 3.5-fold in cells mutated for RNase D. RNase BN contributed a small but reproducible increase in csrA mRNA. There was a corresponding 3.3- fold increase in the protein for CsrA when examined with Western blotting.

To ensure that csrA mRNA was indeed a substrate for RNase D and

RNase BN and to rule out pleiotropic effects of removing RNase D from a cell, csrA mRNA and RNase BN or D were combined in vitro. This confirmed that both

RNase D and RNase BN are capable of digesting in vitro transcribed csrA mRNA. In vitro enzyme activity was confirmed using tRNA-C*A for RNase BN and denatured tRNA for RNase D (not shown). These studies revealed that in vitro RNase D was much more active on in vitro transcribed csrA mRNA than

RNase BN. This result is consistent with the in vivo data.

We wanted to be sure that the effect was direct even though we've shown that csrA mRNA is a substrate for RNase D and RNase BN in vitro, so we looked at the mRNA for csrA in vivo. The half-life of csrA mRNA was determined in vivo

87 using Northern blots to visualize concentrations of the csrA mRNA at set time points after the addition of rifampicin, to stop transcription of the csrA mRNA. In the wild type strain csrA mRNA had a half-life of 6.5 minutes and the double mutant the half-life of mRNA coding for csrA was approximately 12 minutes. This demonstrates that the effect is direct and that CsrA mRNA is a substrate for

RNase BN and primarily RNase D in vivo.

Even though the csrA mRNA is a substrate for RNase BN and primarily

RNase D in vivo, we wanted to explore the previously observed phenotype of cell aggregation during stationary phase. Wild type and double mutant cells were grown in a 96-well microtiter dish to observe for cell adherence. The ratios of planktonic and adherent cells remained consistent in this assay, but this didn't necessarily tell us anything about the formation of extracellular matrix.

Subsequent crystal violet staining of the extracellular matrix did reveal that cells mutated for RNase D in particular formed less stainable matrix than wild type.

We also demonstrated that the elevation of csrA mRNA alone is sufficient to cause the phenotypic abnormalities of aggregation and limited ability to form extracellular matrix seen in mutants for RNase D but not the associated decrease in motility seen in figure 4. 3. or the levels of cyclic-di-GMP suggesting an avenue for further research. This combined with the in vivo and in vitro data showing that csrA mRNA is a substrate for RNase D (and to a lesser extent RNase BN) suggests that RNase D is responsible for regulating csrA expression early in stationary phase, allowing a more efficient transition from exponential growth into stationary phase.

Chapter 6. Recovery from Lag Phase Phenotype

6.1 Background

During recovery from lag phase after a period of prolonged starvation, cells mutated for RNase D were shown to recover more slowly. This phenotype was discovered during an assay alternating nutrient conditions to see how cells behaved under the stress of switching from starvation to the use of specific carbon sources. In this case the carbon source was glucose. It is known that defects in ribosomes can affect recovery from stationary phase (Basturea, et al.

2012; Hirokawa, et al. 2008; Kramer, et al.1999).

Because the data suggested that cells mutated for RNase D and to a lesser extent RNase BN were defective in their recovery from starvation, we considered ribosome maturation to be among the most likely possibilities. We also considered cells lacking RNase BN and D were failing to thrive relative to wild type cells because either a factor made during stationary phase that needed to be degraded was not being broken down efficiently in cells lacking RNase BN or RNase D or that the maturation of a factor necessary for rapid growth was delayed in cells where the RNases BN and D were not available to participate in maturation. While we saw ribosomal maturation as a likely cause for the observed defect in lag phase, we also considered defective maturation or degradation of sRNA and tRNA.

I will show that maturation of 23S rRNA, 16S rRNA and 5S rRNA is not affected by the absence of RNase BN or RNase D and that assembly of subunits

88 89 and ribosomes is also unaffected. I will also show that maturation of the 3' and 5' ends of the 23S rRNA and 16S rRNA are not affected by deleting rnd or rbn from cells.

6.2 Experimental Procedures

6.2.1. Alternating nutrient conditions assay - Strains were first grown overnight in

LB at 37 ºC with aeration, spun down and resuspended in M9, allowed to grow at

37 ºC for another 24 hours with aeration and then resuspended in M9 with 0.2% glucose. This cycling in M9 ± glucose was repeated for the indicated number of days.

6.2.2. Starvation shock assay - Overnight cultures were grown in 2.5ml of LB.

Cells from these cultures were spun down and washed 3 times with phosphate buffered saline (PBS) before being resuspended in 2.5ml of M9. The cultures were then incubated at 37 ºC with aeration. The culture was sampled and serially diluted every 24 hours for measurement of cell number by plating on LB

6.2.3. Starvation competition assay - The ability of strains to persist in starvation was tested in overnight cultures, grown in 2.5ml of LB. The cultures were spun down and washed 3 times with PBS before being resuspended in 2.5ml of M9.

The cultures were then incubated at 37ºC. The culture was sampled, serially diluted every 24 hours and plated for measurement of cell number.

6.2.4. Sucrose gradients

6.2.4.1. Subunit analysis - Gradients of 10% - 30% sucrose in a buffer

(20mM HEPES (pH 7.4), 60 mM NH4Cl, 2 mM Mg(OAc)2, 0.5 mM EDTA, 10 mM

90 mercaptoethanol) were made using a rotary gradient maker. Cell extracts were prepared by sonicating cells and spun down at 6K rpm to remove cellular membrane debris. The supernatant fraction was layered on a sucrose gradient and spun at 21K rpm in the Beckman Ultracentrifuge in SW28 for 18 hours for ribosome profiles.

6.2.4.2. Ribosome analysis - Gradients of 10% - 40% sucrose in a buffer

(20mM HEPES (pH 7.4), 60 mM NH4Cl, 20 mM Mg(OAc)2, 0.5 mM EDTA, 10 mM mercaptoethanol) were made using a rotary gradient maker. Cell extracts were as in 6.2.4.1. The supernatant fraction was layered on the sucrose gradient and spun at 21Krpm in the Beckman Ultracentrifuge in SW28 for 10 hours for ribosome and polysome profiles.

6.2.5. Primer extension procedure- This assay was performed using the

Promega Primer Extension System Cat.# M5101 according to the manufacturer's instructions and with help troubleshooting from S. Mohammed.

6.2.6. 3'-RACE procedure - This assay was performed using ExactSTART™ mRNA 5´- & 3´-RACE Kit from Epicentre Cat.#ES80910 according to the manufacturer's instructions.

6.3 Results

The nutrient upshift/downshift assay showed that mutant cells were outcompeted by wild type cells when reintroduced to rich media following starvation. This result from the alternating nutrient conditions suggested that there was some defect in cell growth during recovery from stationary phase

91 (Figures 6.1.). Either cells were deficient in some factor that allowed them to commence rapid growth or they were unable to eliminate some factor that was hindering the transition from stasis to rapid growth. We did ask why we hadn’t seen these differences in previous experiments where competitive growth had been done in rich media and mineral salts media with a carbon source (data not shown). Typically we had done these assays over a period of 5 days and subsequently we opted to do a similar experiment for a longer period of time.

Figure 6. 1. Effect of abrupt nutrient depletion. Co-cultured MG1655 (WT) and cells mutated for RNase D and RNase BN (MUT) grown in M9 mineral salts alternating with or without glucose for 24 hour intervals before being spun down, washed and resuspended in M9 mineral salts media with or without glucose. Cells were enumerated using viable count on rich agar. This is representative data from an experiment performed at least twice.

92 This effect is subtle during full-cycle competition in LB. Many cycles of growth were necessary to observe a trend in survival of cells lacking RNase D and RNase BN compared with wild type cells in LB (Figure 6.2.). This experiment shows that the defect might not be detectable until more than seven days have passed when cells are grown in competition in rich media.

Figure 6. 2. Long term growth and survival. Co-cultured MG1655 (WT) and cells mutated for RNase D and RNase BN (MUT) grown in LB for 24 hours before being spun down, washed and reinoculated to LB. Cells were enumerated using viable count on rich agar. This is representative data from an experiment done twice.

93 We determined that the defect in growth resulted from a prolonged lag phase in mutant cells by observing and comparing the initiation of growth of wild type cells with cells lacking RNase D and RNase BN (Figure 6.3.). This shows that the lag phase for the mutant cells is longer than for the wild type cells and that by the time the mutant cells are growing rapidly, the wild type has depleted the bulk of the available nutrients.

Figure 6. 3. Growth from lag phase. Co-cultured MG1655 (WT) and cells mutated for RNase D and RNase BN (MUT) grown in LB for 24 hours before being spun down, washed and starved for 3 days in mineral salts media and then re-inoculated to LB and observed for growth during lag phase. Cells were enumerated using viable count on rich agar. These are representative data from an experiment performed twice.

94 Because aberrant ribosome assembly is known to prolong lag phase, we looked at gross elements of ribosome processing and assembly. A Northern blot using a probe complementary to 16S ribosomal RNA over a time course showed no obvious differences between wild type and mutant cells (Figures 6.4.). This suggests that there are no gross differences in the processing of this RNA, but the trailer and leader sequences for 16S rRNA after it is cleaved from the cistronic message by RNase III are too small to be efficiently probed in the manner. 16S rRNA has 113 nt at the 5’ end and 33 nt at the 3’ end that must be removed in order to yield a mature transcript. (Jemolio, DK et al. 1996)

Figure 6. 4. Ribosomal RNA time course Northern for 16S rRNA. MG1655 (WT) and ∆rbn∆rnd cells (MUT) were grown for 28 hours in LB with aeration and then spun down, washed, resuspended in mineral salts media. Aliquots were taken after 72 hours of starvation (T=0), and then two (T=2) and four hours after the addition of glucose. RNA was isolated via hot phenol extraction, resolved in 0.6% agarose, blotted to nitrocellulose and probed with a DNA oligo complementary to 16S rRNA. This is a representative blot from an experiment performed twice. Values for densitometry were within 15%.

95 A similar Northern blot for rRNA 23S subunits over a time course showed no obvious differences between wild type and mutant cells (Figures 6.5.). The

23S rRNA transcript also is transcribed as a cistronic message and is cleaved by

RNase III. In this case, this particular assay is not sensitive enough to pick up small differences in the processing of the 5’ end of the transcript at 3 to 7 nt and on the 3’ end with a length of only 7-9 nt that must be removed in order to yield a mature 23S rRNA transcript (Jemolio, DK et al. 1996).

Figure 6. 5. Ribosomal RNA time course Northern for 23S rRNA. MG1655 (WT) and ∆rbn∆rnd cells (MUT) were grown for 28 hours in LB with aeration and then spun down, washed, resuspended in mineral salts media Aliquots were taken after 72 hours of starvation (T=0), and then two (T=2) and four hours after the addition of glucose. RNA was isolated via hot phenol extraction, resolved in 0.6% agarose, blotted to nitrocellulose and probed with a DNA oligo complementary to 23S rRNA. This is a representative blot from an experiment performed twice. Values for densitometry were within 12%.

96 After these Northern blots showed no gross abnormalities, we used sucrose gradients with high magnesium concentration to observe 70S ribosomes. These showed no obvious differences between wild type and mutant cells. Radioactive ribosomes were used as a control for sedimentation (Figure

6.6.). This suggests that there is no aberrant assembly of ribosomes or ribosomal proteins in strains lacking RNase D and RNase BN as their migration through sucrose is effectively identical.

Figure 6. 6. High magnesium ribosomal sucrose gradient. This shows the migration profile of ribosomes in a 10%/40% sucrose gradient with 20mM magnesium from starved MG1655 (WT) and ∆rbn∆rnd cells (MUT) cells 2 hours after the addition of glucose. Representative peaks are indistinguishable. Circle and cross points represents trace for MG1655 (traceWT) and mutant cells (traceMUT). These are representative data from an experiment performed three times.

97 A low magnesium (2mM) sucrose gradient (30%/10%) was performed using ribosomes collected after starvation (data not shown) and recovery from starvation. A trace of radioactive ribosomes was used as a control. Sucrose gradients with low magnesium to elucidate subunits showed no differences between wild type and mutant strains (Figure 6.7.).

Figure 6. 7. Low magnesium ribosomal subunit sucrose gradient. This shows the migration profile of ribosomes in a 10%/30% sucrose gradient with 2mM magnesium from starved MG1655 (WT) and ∆rbn∆rnd cells (MUT) cells 2 hours after the addition of glucose. Representative peaks are indistinguishable. Circle and cross points represents trace for MG1655 (traceWT) and mutant cells (traceMUT). These are representative data from an experiment performed three times.

98 Primer extension was performed to examine the 5' ends of the ribosomal RNA and showed no obvious differences between the wild type and the mutants (Figure 6. 8. A. & B. ) It should be noted that RNase BN is not normally active at the 5' end of RNA and that RNase D is not known to act on 5' ends of RNA.

Figure 6. 8. A. 5’ Primer extension of 16S rRNA. Primer extension of 16S rRNA reveals no difference in length of the 3' ends of these transcripts in the various backgrounds. The 23nt primer and expected 51nt PE product are present in all samples. Lane 1. MG1655 (WT); Lane 2. Δrbn (BN); Lane 3. Δrnd (D); Lane 4. ΔrbnΔrnd (MUT). B. 5’ Primer extension of 23S rRNA. Primer extension of 23S rRNA reveals no difference in length of the 3' ends of these transcripts in the various backgrounds. The 23nt primer and expected 54nt PE product were visible in all 4 samples. Lane 1. MG1655 (WT); Lane 2. Δrbn (BN); Lane 3. Δrnd (D); Lane 4. ΔrbnΔrnd (MUT).

Because it is more likely that RNase D and RNase BN are active on the 3' ends of their respective substrates, we performed RACE on the 3' ends of the ribosomal RNAs collected from wild type, single and double and mutant strains

(Figure 6. 9. A. & B. ) These experiments show no differences in the maturation of the 3' ends of the 23S and 16S rRNA.

Figure 6. 9. A. 3’ RACE of 16S rRNA. 3' RACE to observe possible differences in the ends of 16S rRNA from wild type and double mutant for RNase D and RNase BN from RNA isolated during lag phase. Lane 1. 1kb ladder (L); 2. MG1655 (WT); 3. ΔrbnΔrnd (MUT). Bands in lanes 2 and 3 correspond to a 70nt PCR amplified product. Representative data from experiment performed at least twice. According to manufacturers instructions, this is not a quantitative assay. B. 3’ RACE of 23S rRNA. 3' RACE to observe possible differences between wild type and double mutant for RNase D and RNase BN the ends of 23S rRNA isolated during lag phase. Lane 1, 1kb ladder (L); 2. MG1655 (WT); 3. ΔrbnΔrnd (MUT). Bands in lanes 2 and 3 correspond to a 82nt PCR amplified product. These are representative data from an experiment performed twice. According to manufacturers instructions, this is not a quantitative assay.

100 CsrA also had to be eliminated as a possible source for the prolonged recovery from starvation observed in cells lacking RNase D. A time course of csrA mRNA and CsrA protein showed that there is no difference between a double mutant and single mutant strain during the lag phase (Figure 6.10. A. & B.).

Figure 6. 10. A. Northern of CsrA mRNA. Northern analysis of CsrA mRNA. 6% urea-PAGE transferred to nitrocellulose and probed with 32P-labeled oligo. RNA collected from cells starved for 2 days in M9. MG1655 (WT) and double mutant (MUT) before addition of glucose (0 hours), and also 2 hours after addition of glucose at 2% (2 hours) and 4 hours after addition of glucose (4 hours). MG1655 was set to one (1.0) for each time point for densitometry [RNA] or none detected versus background values (ND). Representative data from an experiment performed at least twice. B. Western of CsrA protein. Western blot of CsrA protein. MG1655 (WT) and double mutant (MUT) before addition of glucose (0 hours), and also 2 hours after addition of glucose at 2% (2 hours) and 4 hours after addition of glucose (4 hours). MG1655 was set to one (1.0) for each time point for densitometry [protein] or none detected versus background values (ND). These are representative data from an experiment performed three times.

101 The non-coding RNA, 6S RNA, was also probed to eliminate the possibility of it's involvement in the lag phase defect. Wassarman (2007) demonstrated that

6S RNA could slow rapid growth when overexpressed. This is due to the ability of the RNA to form a hairpin that resembles an open promoter. The combined sequence and structure of this hairpin formed by 6S RNA binds efficiently to the

RNA polymerase holoenzyme containing the housekeeping factor sigma 70 .

(Figure 6. 11). These data show no obvious differences between mutant and wild-type cells at various time points during the lag phase. The transcript was not detectable at 4 hours of growth even when RNA fractionated using isopropanol to eliminate the high molecular weight RNA which comprises the bulk of RNA in the cell. The low molecular weight fraction is enriched for small RNA like 6S RNA, but this did not increase the sensitivity of the assay enough for these purposes.

Figure 6. 11. Northern of 6SRNA. Northern analysis of 6S RNA. 6% urea-PAGE transferred to nitrocellulose and probed with 32P-labeled oligo. 20Ug of RNA per lane was collected from cells starved for 2 days in M9. MG1655 (WT) and double mutant (MUT) after starvation and before addition of glucose (0 hours), and also 2 hours after addition of glucose at 2% (2 hours) and 4 hours after addition of glucose (4 hours). MG1655 was set to one (1.0) for each time point for densitometry or none detected versus background values (ND). These are representative data from an experiment performed twice.

102 Cells were starved for three days in mineral salts medium before and after the addition of 2% glucose to observe whether any aberrant stable RNA was present. RNA was collected from cells and fractionated into low and high molecular weight fractions using isopropanol. The low molecular weight fraction can be seen in Figure 6. 12. No additional bands were detected, no bands disappeared and concentrations of the most prominent bands selected showed no discernible differences when comparing wild type and strains lacking RNase

BN and/or RNase D.

Figure 6. 12. Low molecular weight radiolabeled RNA from starved and lag phase cells. 20ug of 32P labeled RNA fractionated with isopropanol and resolved in 12% PAGE with TAE. MG1655 (WT), mutants for RNase BN (RBN), cells mutated for RNase D (RND) and double mutant cells (MUT) during starvation (STARVED) and lag phase (LAG PHASE). Lag is four hours of growth in M9 with 2% glucose. These are representative data from an experiment performed twice.

103 The high molecular weight fraction can be seen in Figure 6. 13. No additional bands were detected, no bands disappeared and concentrations of the most prominent bands selected showed no discernible differences when comparing wild type and strains lacking RNase BN and/or RNase D.

Figure 6. 13. 38 cm PAGE of high molecular weight RNA. 20ug of 32P labeled high molecular weight RNA in 8% PAGE with TAE. MG1655 (WT) and MG1655 mutated for RNase D and RNase BN (MUT) during stationary (STAT) phase and lag phase growth (LAG). Representative data from experiment performed four times.

104 6.4 Discussion

Though defects in ribosome assembly are often cited as sources for an increase in the length of lag phase for E. coli, the experimental results show that ribosomal RNA maturation is not affected by eliminating RNase BN or RNase D.

Additionally, the effect seems to be entirely the result of the deletion of RNase D.

RNase BN does not have an obvious effect on the observed phenotype of a delay in exit from lag phase.

Both high magnesium (20 mM) sucrose gradient (40%/10%) and low magnesium (2 mM) sucrose gradients (30%/10%) of ribosomes collected from wild type and double mutant strains during recovery from starvation revealed indistinguishable 70s, 50s and 30s peaks with no apparent shifts suggesting that

RNase BN and D play no role in the maturation of these subunits and that these are likely not the source of the growth defect observed in the double mutant.

The molecular cause for this phenotype remains to be discovered.

The most likely cause tor this defect is likely to be an sRNA that either must be degraded for efficient exponential growth or one that must be synthesized to allow cells to transition efficiently from starvation to growth.

Chapter 7. Possible Influence of Crl

7.1 Background

A 2012 paper by Fredollino et al. details common mutations in laboratory strains of E. coli. The mutation of interest for this project is an insertion element in the gene crl. This mutation results in a phenocopy of an rpoS mutant according to their paper and other literature (Banta, et al. 2013). We demonstrate A. that the strain used for this study does contain the described mutation in crl; B. that this mutation in crl is necessary for previously observed molecular and physiological phenotypes that we tested; and C. that a mutation in rpoS can also create conditions that yield similar results to those observed in previous experiments.

7.2 Experimental Procedures

7.2.1. Strains

MG1655unseq, a K-12 derivative known to be free of the Is1 element in crl and

CAG18447; a strain with a proline synthetase ProA::Tn10 95% (tetracycline resistant phenotype, requires supplemental proline) linked to an intact crl and

DH79H1; and a rpoS mutant with a KEIO kanamycin cassette and flp-frt sites were all generous gifts from Kenn Rudd's collection and were originally constructed by Singer et al. in Carol Gross' lab (1989).

7.2.2. Media

Media was supplemented with 15 mM proline for strains deficient in proline synthetase.

105 106 7.3 Results

The existence of this mutation in the strain in use for this project was confirmed by PCR of the region of the gene in question using probes as detailed in Freddolino, et al. (2012) This PCR reaction revealed an insertion of ~750 base pairs, consistent with their findings of an 777bp IS1 element interrupting the crl gene in our 'wild type' strain. (Figure 7. 1)

Figure 7. 1. Gel of Is1 element in MG1655*. PCR products using probes for genomic crl. Ladder (L), MG1655unseq (WT) and MG1655* (MUT) run in 6% agarose. This experiment was performed twice.

107 The crl gene was restored with a P1 transduction from a donor strain with an intact crl gene 95% linked to proline synthetase gene proA, interrupted by a

Tn10 insertion, allowing for simple selection on media containing tetracycline and proline. The rescored crl was confirmed by PCR (data not shown). The recipient was MG1655* and the resulting progeny is the “wild type” strain used for all subsequent experiments in this chapter. It was clear that the mutation in crl is required for the clumping phenotype (observed via microscopy, data not shown) as well as the the defect in lag seen after starvation observed in strains mutated for RNase D (Figure 7. 2. A & B.).

Figure 7. 2. Co-cultured growth and survival of Pro::Tn10 Crl+ strains. MG1655* (pro-Tn10:: crl) (WT) and MG1655 * (pro-Tn10::Crl) mutated for RNase D and RNase BN (MUT) grown in alternating nutrient conditions mineral salts media and M9 media with 0.2% glucose supplemented with proline. This is a representative graph from an experiment performed twice. Because crl- is known to phenocopy an rpoS deletion, a P1 transduction was used to move a rpoS deletion from the KEIO collection into the MG1655* (pro-

108 Tn10:: crl) strains, These strains were tested to see if a mutation in rpoS was

sufficient to reproduce the observed phenotypes in the presence of exogenous

proline (Figure 7. 3.). This shows that crl is necessary for aggregation.

Figure 7. 3. Growth of Pro::Tn10 crl+, rpoS- strains. MG1655 (WT) and MG1655 mutated for RNase BN (BN), RNase D (RND) and both (BND) grown in alternating nutrient conditions mineral salts media and M9 media with 0.2% glucose. This experiment was done three times.

7.4 Discussion

The mutation of crl is necessary for all observed phenotypes in K-12 derivative MG1655* in chapters 3 through 6. Mutating the gene for rpoS produced a similar phenotype when RNase D and/or BN were mutated, confirming that the crl- mutation phenocopies the rpoS- mutation and that a mutation in crl or rpoS is requisite for the phenotypes in strains lacking either

RNase in this study. Some of the observed phenotypes have yet to be tested.

Chapter 8. Discussion and Future Work

The conclusions of chapter 2, while not directly relevant to the studies of

RNases in E. coli, that particular line of study did reveal a gene likely to be involved in cell division of E. coli. The apparent septation defect in elaD mutant strains resulting in filamentation of these cells under stationary phase conditions but not during rapid growth suggests a gene that is relevant to cell division during stress conditions but not rapid growth. Subsequent research from another lab suggests that this protein, ElaD, is a deubiquitinase (Catic et al. 2007). Since E. coli has no ubiquitin, the authors suggest that this protein is a virulence gene that allows pathogenic strains of E. coli to evade the immune system by effectively un-marking themselves for destruction. While this is interesting as it pertains to the study of virulence, it does not explain the septation defect observed in BEZ33 and the elaD- strain. This particular observation represents a potentially fruitful avenue of future study in the area of cell division under stress conditions.

Unfortunately, these data are not relevant to the study of RNase BN and D and their respective substrates. This led us to search for potential substrates and evidence for their primary roles in E. coli via molecular techniques.

The molecular techniques employed in the search for potential substrates primarily included Northern analysis. The plethora - 109 - of negative results in chapter 3 eliminate many possibilities for study under the conditions studied, particularly stationary phase. The expression profiles of RNase D and RNase BN suggest that testing these potential substrates during exponential phase may yield more and better data. That some sRNAs are affected during rapid

109 110 growth has been confirmed by recent results in our lab (Tanmay Dutta, unpublished results by communication). The fact that both RNases are expressed during exponential phase is certainly an indicator of their function.

The fact that their expression is downregulated during stationary phase and stress conditions by different mechanisms is another relevant finding. While

RNase D is downregulated by the stringent response in E. coli, the mechanism for RNase BN's regulation is still unknown. Because RpoS activity is also effectively ablated affected by the mutation in the gene encoding Crl, this mechanism is also unlikely to be responsible for reduced expression of RNase

BN during stress and stationary phase conditions. Additionally the sequence of

RNase BN mRNA lacks the target sequence “RUACARGGAUGU” for sequestration by binding to CsrA RNA binding protein. Another mechanism must be responsible for regulating the expression of RNase BN. It is likely, based on bioinformatic data (Salgado et al. 2012), that RNase BN's expression is controlled by the housekeeping RNA polymerase holoenzyme containing sigma

70. This suggests that the depletion of sigma 70 during entry to stationary phase and the natural half-life of the protein controls the increased expression during exponential phase and ablation of RNase BN during stationary phase.

Concurrent with the search for a phenotype using molecular techniques, we also subjected cells lacking RNase D and RNase BN to a variety of physiological conditions to search for a phenotype that might suggest possible substrates of RNase D and RNase BN. The data from Phenotype MicroArray plate PM4A and accompanying liquid culture growth experiments suggest a

111 subtle disadvantage in the ability of strains lacking RNase D and RNase BN to survive in acid media which has been confirmed by another member of our lab

(Tanmay Dutta, unpublished observation via direct communication). This presents one avenue for future study.

Other data from this avenue of study suggest that mutants for RNase D and to a lesser extent RNase BN aggregate during stationary phase growth.

Further analysis not only related this phenotype to the elevation of the mRNA for the gene csrA but also revealed that the phenotype was a direct result of elevated csrA mRNA and CsrA protein.

Messenger RNA for the protein CsrA was shown to be affected by deleting

RNase D in MG1655*. This gene is responsible for global regulation of carbon metabolism under starvation as well as a suite of other activities including biofilm formation and motility. The messenger RNA for the RNA binding protein CsrA is modulated by the enzyme RNase D during early stationary phase growth.

Analysis of the secondary structure of csrA mRNA showed that the 3' tail of csrA is relatvely unstructured and is rich in A and U residues. These findings may explain why csrA mRNA is a substrate for RNase D and RNase BN. The presence of consecutive C residues may explain why RNase BN is less active than RNase D.

The increase in csrA message and protein associated with the absence of

RNase D is sufficient to cause the aggregation of cells and reduced ability of cells to form extracellular matrix. A paper chromatography assay was done to examine if cyclic-di-GMP levels were depressed as one would expect in E. coli with an

112 abundance of csrA protein. This revealed a surprising increase in the levels of the signaling molecule that seemed independent of the presence of the increase in CsrA protein, but did coincide with cells lacking RNase D. Although it seems evident that this data regarding cyclic-di-GMP is a significant finding, it creates questions as opposed to answering them.

The phenotype of recovery from lag phase observed in chapter 4 inspired the studies in chapter 6. It was clear based on the agarose gels, Northern analysis, low and high magnesium sucrose gradients, 3' RACE and 5' primer extension of ribosomes and ribosomal subunits, that defects in ribosomal RNA or ribosome assembly was not the likely cause for the observed defect in recovery from prolonged starvation. It is evident that this mystery remains to be solved.

The final chapter explores a confounding variable for our study of the physiological role and functions of RNase D and RNase BN. The mutation in the gene encoding Crl is known to phenocopy a mutation in rpoS. Since much of this study centered on stress conditions and stationary phase growth, it was troubling that restoring the gene crl ablated all phenotypes that were tested. This begs the questions of whether or not RNase D and RNase BN exact a measure of control as backups for potentially deleterious effects of naturally occurring mutations in crl or rpoS or if they have their own function in E. coli. Future works done in a background where crl is intact may reveal more physiologically relevant phenotypes and substrates for RNase D and RNase BN.

Several questions remain unanswered. Namely, 1. what is responsible for

the defects in motility in RNase D mutant strains; 2. what is responsible for the

113 increase in cyclic-di-GMP in strains lacking RNase D; 3. what is responsible for the defect in exit from lag phase; 4. how does the lack of RNase BN reduce the ability of cells to survive acid conditions; and last but not least 5. what role do crl or rpoS play in these observations, if any. Firstly, it seems plausible that the defect in motility and the increase in cyclic-di-GMP observed in RNase D mutants are related (Weber, et al. 2006). An increase in cyclic-di-GMP is associated with a decrease in motility (Tagliabue et al. 2010), but it is also known that cyclic-di-

GMP expression is largely regulated by RpoS. Because the parent strain is known to be mutated for crl, that this mutation has a very similar phenotype to a mutation in rpoS and that several of the previously observed phenotypes in cells lacking RNase D and RNase BN were resolved by restoring the defective gene encoding Crl, it stands to reason that this phenotype (or phenotypes if they are indeed unrelated) might also be absent in a strain that contains an intact crl.

Even though the slow exit from lag phase is among the effects of deleting RNase

D from a cell that is also resolved by restoring crl, this finding still represents a meaningful mechanism to create redundancy and thereby robustness in strains where crl or rpoS have been affected by an insertion, deletion or frameshift event. Similarly, the activity of RNase BN during growth in low pH suggests that

RNase BN may be responsible for depleting the excess of an RNA that is deleterious for growth in low pH conditions or that it is involved in the maturation of an RNA that is necessary for optimal growth in acidic media, possibly gadE or arrS (Aiso et al. 2011; Méndez-Ortiz et al 2007).

114 Further research to understand the roles that RNase BN and D play in E. coli might include a tiled microarray of total RNA from wild type and double mutant strains obtained from starved cells and after the addition of glucose to observe changes that occur during lag phase, cells collected during exponential phase growth and also in cells grown in acidic media. Hits might include RNAs involved in the observed defect in lag phase of starved cells, RNAs whose potentially deleterious expression was suppressed by the RNases during exponential phase, RNAs whose increased expression might hinder the ability of cells to survive in acidic conditions, RNAs that affect motility and RNAs that affect cyclic-di-GMP degradation or synthesis.

While a handful of microarrays have been done on cells comparing stationary phase cells to exponentially growing cells, minimal media to rich, and no glucose versus 0.4% glucose, none of these microarrays were performed under the conditions of interest – prolonged starvation and lag phase – and none in strains mutated for RNase D. (Gutiérrez-Ríos et al. 2003; Shimada et. al.

2004; Tao et al. 1999) Moreover, no study looked directly at the entire genome in a mutant for RNase BN or RNase D. Most of the genes tested were known open reading frames and a few predicted ORFs. This is important because while any of the thousands of mRNA are possible substrates (and one indeed is), other stable and small RNA are also likely targets of these enzymes – those with either unstructured regions, highly structured regions or both. The number of 'hits' would likely be manageable due to the existence of a microarray of >1300 genes that revealed no differences between a mutant for RNase BN and a wild type

115 E.coli strain. (Schilling et al. 2004) Even so, it would only be necessary to examine a handful of hits to fulfill the premise of this study: to find the physiological roles and substrates of RNases BN and D. The mRNA and sRNA are too numerous to probe individually by Northern, primer extension and RACE.

Naturally this experiment would include a mutant for both RNases BN and RNase

D so the results would have to be dissected so that each hit could be attributed to the either RNase BN or RNase D during the verification process. A tiled microarray would be cost effective and time efficient method of finding the substrates or RNase BN and D. A pull down assay with catalytically inactive

RNase D and RNase BN might reveal possible substrates for RNase BN and D.

This is an alternative to a tiled microarray and would require sequencing of the purified RNA.

It is clear that RNase D and RNase BN have a role in the growth and survival of E. coli. My findings show that in the presence of a mutation in the genes encoding rpoS or crl, that RNase D can foster more robust growth and survival. I have also shown that in this background RNase D is necessary for efficient recovery from long term starvation, for efficient synthesis of extracellular matrix, for maintenance of the mRNA encoding the protein CsrA as well as motility and normal signaling with cyclic-di-GMP. My preliminary data along with

Tanmay Dutta's results show that RNase BN is necessary for efficient growth and survival in low pH environments. While these data are somewhat dispersed throughout various elements of E. coli biology, this only serves to accentuate that these two ribonucleases are relevant in E. coli growth and survival. In a

116 strain whose very existence can be dependent on minutes differences in doubling time, the regulatory fine-tuning offered by these two enzymes is meaningful.

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