The Modification State of Elongation Factor-P in Bacillus Subtilis and Pseudomonas

The Modification State of Elongation Factor-P in Bacillus subtilis and Pseudomonas

aeruginosa

Thesis

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in the Graduate School of The Ohio State University

Sarah Beth Tyler

Graduate Program in Chemistry

The Ohio State University

2015

Thesis Committee:

Dr. Michael Ibba, Advisor

Dr. Karin Musier-Forsyth, Advisor

Dr. Ross Dalbey

Sarah Beth Tyler

2015

Abstract

In order for life to function properly, proteins must be correctly translated. One instance that poses as a threat to translation is stretches of poly prolines in the transcript. Because of the pyrrolidine ring in proline, proline is a poor imino acceptor and donor, and a string of three or more prolines in a row can cause the ribosome to pause. The translation factor

Elongation Factor-P (EF-P) binds to the ribosome and stimulates bond formation between the prolines, alleviating the pause and allowing translation to continue. EF-P is post translationally modified on a conserved residue, and while EF-P or an EF-P homolog is found in all domains of life, there is variety in the post-translational modifications from one bacterial organism to another. Thus far, it has been discovered that EF-P in E. coli is modified with (R)-ß-Lysine, with dTDP-Rhamnose in P. aeruginosa, and a hypusine in the eukaryotic homolog, eIF5A. These modifications currently known in bacteria only represent a small amount of bacteria, and the question of what other modifications are remains unanswered. It has been confirmed by mass spectrometry that Bacillus subtilis

EF-P is post-translationally modified. The identity of this modification however, still remains elusive. In this work, methods including mass spectrometry, isoelectric focusing, and in vitro reactions are used to probe what the modification on B. subtilis EF-P is.

Besides getting closer to understanding what the modification on EF-P is, these techniques are also used to reveal potential regulation of EF-P. Finally, this work

addresses some of the experiments performed in identifying the modification on P. aeruginosa EF-P, including the synthesis of the substrate for modification of EF-P, dTDP-rhamnose.

iii

This document is dedicated to my parents, Ann and Steven Tyler, who have instilled in

me a deep appreciation for education.

Acknowledgments

I would like to thank my advisor, Dr. Michael Ibba for mentoring me during the time I was in his lab. He is a good leader and a scientist whom I wish to emulate. I would also like to acknowledge and thank Andrei Rajkovic for all of his training and guidance,

Tammy Bullwinkle, Kyle Mohler, and Sara Elgamal for their insight in experiments, and the rest of Ibba lab for their input and support. Lastly, I would like to acknowledge The

Ohio State Chemistry and Biochemistry Program and the Chemistry and Biochemistry

Interface Program for acceptance into the program and for the funding of my research.

Vita

June 2009 ...... East High School

2013...... B.S. Chemistry, University of Utah

2013 to present ...... CBIP Trainee, Department of Chemistry and

Biochemistry, The Ohio State University

Fields of Study

Major Field: Chemistry

Table of Contents

Abstract ...... ii

Acknowledgments ...... v

Vita...... vi

Fields of Study ...... vi

Table of Contents ...... vii

List of Tables ...... x

List of Figures ...... xi

Chapter 1. Introduction ...... 1

1.1. Aminoacyl-tRNA Synthetases ...... 1

1.1.2. Alternative Synthetase Functions ...... 3

1.2. Translation Machinery ...... 4

1.2.1. Initiation, Elongation, and Termination ...... 5

1.2.2. Regulation by mRNA ...... 8

1.3. Elongation Factor-P ...... 9

1.3.1. eIF5A ...... 11

vii

1.3.2. Modification Pathways of EF-P and eIF5A ...... 12

1.3.3. Bacterial responses to ∆EF-P ...... 15

Chapter 2. Materials and Methods ...... 16

2.1. Bacterial Strains and Plasmids ...... 16

2.2. Growth Curves ...... 18

2.3. Purification of EF-P ...... 18

2.4. Isoelectric focusing and Western Blotting as a Tool ...... 20

2.5. In Vitro aminoacylation of EF-P ...... 21

2.6. Glycoprotein Stain ...... 22

2.7. Lectin Binding Assays ...... 22

2.8. [14C] dTDP-Rhamnose Synthesis ...... 23

Chapter 3. Results ...... 25

3.1. Modification Status of EF-P in Bacillus subtilis...... 25

3.2. Mass Spectrometry ...... 26

3.2. Characterization of B. subtilis EF-P by Isoelectric Focusing ...... 30

3.3. In-vitro assays probing the identity of the modification substrate ...... 32

3.4. Glucose as a Potential Regulator ...... 35

3.5. EF-P in P. aeruginosa...... 38

viii

3.5. Glycoprotein Staining ...... 38

3.6. Lectin Binding Assays ...... 39

3.7. [14C]Rhamnose Synthesis ...... 40

Chapter 4. Future Work ...... 43

4.1. Identifying potential modifying enzyme in B. subtilis ...... 43

References...... 46

List of Tables

Table 1. A list of the B. subtilis strains used in this work...... 16

Table 2. P. aeruginosa strains used in this work ...... 17

List of Figures

Figure 1. Translation cycle ...... 8

Figure 2. Comparison of EF-P to tRNA ...... 11

Figure 3. Structure of Human eIF5A compared to EF-P ...... 12

Figure 4. Different modification pathways of EF-P and homolog eIF5A ...... 14

Figure 5. Exemplary Mass spectrometry data...... 28

Figure 6. Growth curves of B. subtilis strains ...... 30

Figure 7. Western blot from IEF gel showing EF-P charge states ...... 31

Figure 8. Isoelectric focusing gel to identify modification ...... 34

Figure 9. Modification status of EF-P as a function of OD600 ...... 37

Figure 10.The glycoprotein staining reaction ...... 39

Figure 11. The synthesis pathway for dTDP- rhamnose ...... 42

Chapter 1: Introduction

How genetic information is transferred from DNA to RNA to protein has been a long standing question that, despite many advances in knowledge over the past ~100 years, still has not been fully answered. While there are many questions about the details of the process, there is now a general understanding of the overall flow of genetic data, known as the central dogma (1). The central dogma explains how genetic information flows between DNA, RNA, and proteins. Beginning as DNA, genetic information can either be replicated into DNA by DNA polymerase, or transcribed into RNA by RNA

Polymerase. Then from RNA it can either be reverse transcribed, back into DNA by reverse transcriptase, or translated into proteins via the ribosome and its accompanying translation factors. Translation is done using the ribosome to catalyze the reaction, with mRNA transcripts as a template for synthesis, amino acids as the building blocks for proteins, and tRNAs as the adaptors between RNA and protein.

1.1 Aminoacyl-tRNA Synthetases

After the discovery that DNA was a double stranded helix, Crick made several highly accurate predictions about what would become known as the central dogma (2).

One of these hypotheses is the adaptor hypothesis. Postulated in the mid 1950’s, Crick predicted that RNA would be translated by the use of a template, with the amino acid

being carried to the template by an adaptor that possibly contained nucleotides. He also predicted that certain proteins would be necessary to join the amino acid to an adaptor, and that the adaptors would be highly specific. It has since been discovered that tRNA is the molecule that serves as the adaptor between the amino acid and its three-letter mRNA code (3). Each amino acid has a corresponding tRNA, and before the amino acid can be incorporated into a growing polypeptide chain, it must first be aminoacylated (or charged) onto its cognate tRNA. This is done via an aminoacyl-tRNA synthetase (aaRS), which uses a two-step reaction to charge the amino acid. The first step is activation of the amino acid with ATP, followed by esterification of the amino acid onto the 3’ end of the tRNA.

푎푎 + 푎푎푅푆 + 퐴푇푃 ↔ 푎푎 ∙ 퐴푀푃 + 푎푎푅푆 + 푃푃푖 (a)

푎푎 ∙ 퐴푀푃 + 푎푎푅푆 + 푡푅푁퐴 ↔ 푎푎 − 푡푅푁퐴 + 푎푎푅푆 + 퐴푀푃 (b)

Equation. The two step reaction of aminoacylation. a.) An aminoacyl-tRNA synthetase (aaRS) catalyzes the reaction of an amino acid (aa) and an adenosine triphosphate (ATP) to form an activated amino acid (aaAMP) and an inorganic pyrophosphate (PPi). b.) The aaRS catalyzes the reaction of an activated amino acid and a tRNA to form an aminoacyl-tRNA (aa-tRNA) and an adenosine monophosphate (AMP).

Once the amino acid is charged onto the tRNA, the aminoacyl-tRNA is released by the synthetase and escorted to the ribosome by EF-Tu.

Because certain amino acids only have small differences in size and shape, it is possible for a synthetase to charge an amino acid onto a non-cognate tRNA. One example of this is with isoleucine and valine, which only differ in structure by one methyl group.

Mischarging of amino acids onto non-cognate tRNA introduced a need for the cell to evolve proofreading, or editing mechanisms. There are two different ways that editing can occur: pre-transfer and post-transfer. Pre-transfer editing occurs before the activated amino acid has been transferred onto its tRNA. Post transfer editing, on the other hand, occurs after the activated amino acid has been transferred to the tRNA. Post transfer editing can occur either in cis or in trans: cis editing by the synthetase and trans by a separate protein. Certain synthetases have evolved editing sites as well as the site for aminoacylation, and aa-tRNAs can feely associate with and dissociate with the synthetase to be edited (4). Other synthetases have evolved trans editing mechanisms, in which editing occurs on a different protein that has evolved specifically for editing. An example of this is when alanine gets mischarged onto tRNAPro. The mischarged Ala-tRNAPro dissociates from ProRS and the protein ProXp-Ala edits and hydrolyzes the proline from tRNAPro (5).

1.1.2 Alternative Synthetase Functions

While the main function of an aaRS is to deliver amino acids to the ribosome to be incorporated into the growing polypeptide chain, other functions have been discovered. Because EF-Tu’s ability to bind to aa-tRNAs is not as strong as once believed (4), it is possible for aa-tRNAs to be shuttled to other locations in the cell and be

used for other purposes. These functions include antibiotic biosynthesis, tagging for proteolysis, and using aa-tRNAs to modify the properties of membrane lipids. For example, in Staphylococcus aureus, lysylphophatidyl-glycerol synthetase (LysPGS) transfers lysine from Lys-tRNALys to phosphatidylglycerol (PG) to form lysyl- phosphatidylglycerol (Lys-PG) (6). In addition to alternate functions, aminoacyl-tRNA synthetases have homologs that perform similar, albeit different functions. One example of this is PoxA, a homolog of bacterial class II-type LysRS. PoxA adds an (R)-ß-Lysine moiety onto the translation factor Elongation Factor-P (EF-P), which is a tRNA mimic that alleviates translational stalling due to poly-proline motifs (27). While the amino acid- binding pockets of PoxA and LysRS are structurally similar, PoxA cannot aminoacylate tRNA-Lys, and LysRS cannot aminoacylate EF-P (25).

1.2 Translation Machinery

Once the amino acids are charged onto their cognate tRNAs, they are delivered to the ribosome by elongation factor-Tu (EF-Tu), where peptide bond formation between amino acids is catalyzed. The ribosome is a large ribonucleoprotein that is made up of two RNA subunits and several protein factors. In bacteria, the ribosome is made up of the small 30S subunit and the large 50S subunit. The 30S subunit is made up of a 16S rRNA and 21 proteins, and the 50S subunit is made up of the 23S rRNA, the 5S rRNA, and 34 proteins (8,9). During translation, the 30S subunit binds to mRNA transcript and the anticodon stem loops of the tRNA. It monitors the interaction of the anticodon with the codon of incoming tRNAs. The 50S subunit contains the peptidyl-transferase center

(PTC), which catalyzes the formation of peptide bonds in the growing polypeptide. When assembled, the ribosome has three aminoacyl-tRNA binding sites: the A site, where aminoacyl-tRNAs enter the ribosome, the P site, where the peptidyl transfer occurs, and the E site, where de-acylated tRNA stays until it is ejected from the ribosome (10,11).

Other necessary translation machinery include EF-G, three initiation factors, IF1, IF2, and IF3, as well as three release factor, RF1, RF2, and RF3, and the ribosome recycling factor (RRF).

1.2.1 Initiation, Elongation, and Termination

Translation occurs in three steps: initiation, elongation, and termination (Figure

1). The probable first step to occur in initiation is the binding of initiation factor 3 (IF-3) to the 30S subunit of the ribosome. Upon IF-3 binding to the 30S subunit, the leftover mRNA and deacylated tRNA from the previous translation cycle are released from the

30S subunit. IF3 also prevents early interaction with the 50S subunit. The 30S subunit then has to bind to the Shine Dalgarno sequence on the mRNA, a ribosome specific binding site located a few nucleotides upstream of the AUG site. This sequence base pairs with the 16S rRNA of the 30S subunit, positioning the ribosome on the AUG codon

(9). There are cases in which there is no Shine-Dalgarno sequence, as in the case of leaderless mRNA that completely lack a 5’ untranslated region (UTR). In these cases, the mechanism of ribosome positioning is still not well understood (12). Once the ribosome has been properly positioned, the initiation factors IF-1, and IF-2 bind the complex. IF-1 binds the A site of the 30S subunit, preventing tRNA from binding, and IF-2 is a GTPase

that binds to fMet-tRNAfMet and then binds to the P site in the 30S subunit. While tRNAs in the translation cycle besides fMet-tRNAfMet bind to the A site of the ribosome, fMet- tRNAfMet binds to the P site so that the initiation complex will have an open A site for the next aminoacylated tRNA to bind to. When the 50S subunit then binds the GTP bound to

IF-2 is hydrolyzed, releasing all of the initiation factors from the ribosome and forming the initiation complex.

Elongation is the step in which the polypeptide is lengthened, and is comprised of three repeated steps known as the elongation cycle until the polypeptide has reached the correct length. Once the initiation complex is completely assembled, EF-Tu shuttles the first aa-tRNA to the E-site of the ribosome. EF-Tu is a GTPase that has evolved an equal specificity for each of the aminoacyl-tRNAs. It has developed recognition for either the amino acid, critical bases in the tRNA, or both. In order to bind all of the aa-tRNAs with similar specificity, it binds each of them relatively weakly, as, if it binds too strongly then it will not release them at the ribosome and if it binds too weakly then it will not deliver the aa-tRNA to the ribosome (13). It has recently been shown that, contrary to what was previously thought, aminoacyl-tRNAs are reversibly bound to EF-Tu, and it is possible for the aminoacyl-tRNA to dissociate to be edited or to be taken to a different pathway

(4). The GTP-EF-Tu- aa-tRNA complex interacts with the A site of the ribosome where the anticodon of the tRNA can bind to the decoding site. If the anticodon and codon are correct and Watson-Crick base pair interactions are made, then the GTP bound to EF-Tu is hydrolyzed introducing a conformational change in EF-Tu (8). This conformational change causes EF-Tu to dissociate from the aa-tRNA and the anticodon of the tRNA

more strongly binds to the codon of the mRNA. At this point, the amino acid is put into contact with the peptidyl-transferase center of the 50S subunit. (14, 11). The ribosome catalyzes the peptide bond formation between the nucleophilic α-amino group of the incoming amino acid and the carboxyl group of the peptide chain in the P site, which also releases the amino acid in the P site from it’s conjugate tRNA. The de-acylated tRNA remains in the P site and the polypeptide chain in attached to the tRNA in the A-site. The final step during elongation is translocation, when the ribosome shifts the mRNA one codon, so that the tRNA in the A site is moved to the P site, and the deacylated tRNA moves from the P site to the E site where it is released from the ribosome into the cytosol.

In order for this to happen, GTP-bound EF-G binds the ribosome, moving the tRNA in the A site to an A/P hybrid state, and the tRNA in the P site to a P/E hybrid state. Upon binding to the ribosome, EF-G undergoes a conformational change resulting in GTP hydrolysis. This conformational change and GTP hydrolysis to GDP moves the tRNA that was in the A/P hybrid state fully into the P site and allows translation to continue (9).

The inactivated EF-G leaves the ribosome, leaving an open A-site for a tRNA to bind.

Translation is signaled to end when the ribosome comes upon a stop codon in the mRNA. This is the beginning of termination. There are three release factors that aid in termination of translation, RF-1, RF-2, and RF-3. RF-1 recognizes the stop codons UAG and UAA, and RF-2 recognizes the stop codons UAA and UGA. Whichever of these two release factors bind to the A site then induces the hydrolysis and release of the peptide from the tRNA and therefore from the ribosome. Once the peptide is released from the

ribosome, the ribosome disassembles into the 30S and the 50S subunits. RF-3’s function is still unconfirmed, however it is thought to dissociate the ribosomal subunits (8).

Figure 1. Translation cycle, adapted from (8)

1.2.2) Regulation by mRNA

mRNA’s main role is to serve as a transcript for translation. However there are several cases where mRNA can be used as a tool not just for translation, but also for regulation. While most translation regulation takes place during initiation, there are instances when the mRNA transcript aids in regulation. One example of this is known as the “ramp” in translation- translation is predicted to be slow for the first 30-50 codons in

order to prevent ribosome congestion. After the first 30-50 codons, translation speeds up to a faster rate for the rest of translation (15). Another example of the mRNA transcript participating in regulation is in the case of SecM, which regulates SecA expression. In E. coli, the Sec pathway is the pathway that is responsible for transportation or integration of most of the pre-secretory and membrane proteins across the inner membrane. The

SecYEG complex is a universally conserved transport channel. SecA is a bacterial- specific membrane protein that promotes translocation through the SecYEG complex.

SecYEG expression appears to be constitutively expressed, however SecA is regulated by secM. secM has a carboxy-terminal sequence that causes the ribosome to stall, allowing for a rearrangement of the secM-secA mRNA. The rearrangement frees up the ribosome- binding site for the downstream secA gene, increasing the amount of secA transcribed

(16,17).

1.3) Elongation Factor-P

Elongation Factor-P (EF-P) is a soluble translation factor that rescues paused ribosomes. Found in all domains of life, with the eukaryotic homolog eIF5a, EF-P was first identified in 1975 as a factor that catalyzed peptide-bond synthesis in vitro between ribosomal bound N-formyl-Met-tRNA and puromycin (15). It has since been discovered to aid in elongation rather than initiation (18), specifically in cases when the ribosome pauses due to stretches of polyprolines in the transcript. Proline, one of the 20 canonical amino acids, has been shown to be a poor imino acceptor and donor (19). Due to the limited ability to change conformation because of the ring in its structure, when stretches

of three or more prolines pass through the ribosome, they cause the ribosome to pause

(20). EF-P binds the ribosome when it is stalled due to polyprolines and stimulates bond formation to continue translation (19).

EF-P has been characterized as a small protein composed of three ß-barrel domains (I-III). The N-terminal domain is comprised of an SH3-like domain, which binds proline-rich motifs (21). The C-terminal ends of domain II and III have an OB fold, which is commonly found in nucleic acid- binding proteins (22,23). The crystal structure of EF-P from Thermus thermophilus exists as a monomeric, 20 kDa protein and is composed of 16 ß-strands that fold to form the three previously mentioned domains (24).

The structure of T. thermophilus EF-P revealed that the three domains of EF-P create an overall structure that is remarkably similar to tRNA. EF-P’s arms are 65Å long and 53Å long, compared to tRNAPhe, which has arms that are 65Å and 70Å long (Figure 2).

Domain I of EF-P mimics the acceptor arm of tRNA, and Domains II and III exhibit an oligonucleotide-biding fold.

Figure 2. Comparison of EF-P to tRNA. Adapted from (24)

1.3.1) eIF5A

As previously mentioned, there are eukaryotic and archaeal homologs of EF-P- eIF5a and aif5a. While eIF5a and EF-P are very similar, there are clear differences between them. eIF5a is composed of two ß-barrel domains, with the N-terminal domain made of an SH3-like domain and the C-terminal made of an OB fold, just as in EF-P.

When eIF5A is superimposed onto EF-P, these two domains are the same between both proteins, however eIF5a lacks EF-P’s third domain (Figure 3)(25).

A. B.

) )

Figure 3. A.) Structure of Human eIF5A. B.) eIF5A, aIF5a, and EF-P overlaid on each other to show similarities. Adapted from (23).

1.3.2) Modification Pathways of EF-P and eIF5A

EF-P is not only a structural mimic to tRNA; it is also post-translationally modified in a pathway that mimics tRNA aminoacylation. Thus far, three different modification pathways for EF-P or eIF5A have been identified. In E. coli, the modification pathway is as follows: (S)-α-Lys is converted into (R)-β-Lys by the protein

YjeK. PoxA then attaches (R)-β-Lys onto K34 on EF-P (27, 28, 29). Finally, YfcM adds a hydroxyl group onto the K34 residue (Figure 4).

The PoxA and YjeK pathway is encoded in about 26% of all known bacterial genomes. While there are still unknown modification pathways, a second vastly different modification machinery has been found in bacteria in the γ-Proteobacterial orders, such as Pseudomonas aeruginosa. After phylogenetic analysis, a subfamily of EF-P was discovered that had a conserved arginine (R32) residue replacing the conserved lysine found in E. coli and Salmonella enterica (K34). A similar phylogenetic analysis 12

identified EarP, the modification machinery for this particular subfamily of EF-P. EarP is a rhamnosyltransferase, and rather than an (R)-(ß)-Lys modification, organisms with this subfamily of EF-P such as S. oneidensis and P. aeruginosa are modified with a rhamnose molecule (30).

While EF-P displays diversity in the types of molecules that it is modified with among bacterial organisms, eIF5A is modified with a hypusine molecule among all eukaryotes (31). This modification occurs in two sequential enzymatic steps; first, deoxyhypusine synthase transfers a 4-aminobutyl moiety from the polyamine spermidine to the ε-amino group of the conserved lysine side chain to form deoxyhupusine. This reaction is dependent on NAD. In the second step, the second carbon of the deoxyhupusine residue is hydroxylated by deoxyhypusine hydroxylase (Figure 4) (32).

A.)

B.)

C.)

Figure 4. Different modification pathways of EF-P and homolog eIF5A. A.) Modification pathway of E. coli EF-P. B.) Modification pathway of P. aeruginosa EF-P. C.) Modification pathway of Eukaryotic eIF5A.

1.3.3) Bacterial responses to ∆EF-P

An early attempt to show whether or not EF-P was essential was performed by inserting a temperature-sensitive kanamycin resistance (KanR) gene near the N-terminal end of EF-P. This construct was used to show that at permissive temperatures, the cells exhibited a wild type phenotype, but at non-permissive temperatures the cells didn’t grow. This was further supported by monitoring the amount of [S35]Met-puromycin formed from cell extracts from the temperature-sensitive control groups (33). This assay confirmed to them that EF-P was essential as cell extracts grown at non-permissive temperatures formed very small amounts of [S35]Met-puromycin, whereas formation was observed from cell extracts grown at permissive temperatures. Other studies have since shown that EF-P is not essential in E. coli, including in 2006, when EF-P was identified as a non-essential gene during the creation of the Keio collection (34). While EF-P isn’t an essential protein in bacteria, loss of EF-P or the modification on EF-P has varying detrimental effects on organisms, depending on what proteins in the genome contain poly proline stretches (19). Among other things, EF-P has been found to play a role in controlling virulence and is necessary for membrane integrity in Salmonella. (35,29). It is also necessary for virulence in Shigella flexneri (36), and for swarming motility in

Bacillus subtilis (37).

Chapter 2: Materials and Methods

2.1) Bacterial Strains and Plasmids

B. subtilis bacterial strains used in this work are listed in table 1. All mutations were made chromosomally, supplied either by Daniel Kearns’ lab, or purchased from the

Bacillus Genetic Stock Center.

Strain Genotype 3610 Wild Type DS354 ∆efp::tet DK2182 ∆efp amyE::PyqhS-efp cat DK2248 ∆efp amyE::PyqhS-efpK32A cat DK2466 ∆efp amyE::Physpank-efp-Flag spec BKE31060 gbsA::erm trpC2 BKE31070 gbsR::erm trpC2 Table 1. A list of the B. subtilis strains used in this work.

While most B. subtilis strains used had chromosomal mutations in them, the P. aeruginosa strain mutations were made using a clean deletion of the either efp or earP followed by insertion of the mutant with a plasmid. These strains and their plasmids are listed in table 2.

Plasmid Strain Genotype inserted P. Aeruginosa

PAK# wild type None # AJDP721 ∆EarP None AJDP751# ∆efp None # AJDP731 wild type pHERD20T AJDP743# wild type pAJD2217* AJDP1044# ∆EarP pHERD20T AJDP746# ∆EarP pAJD2217* AJDP761# ∆efp pAJD2217* AJDP752 ∆efp pAJD2218 (R32A) AJDP753 ∆efp pAJD2218 (R32K)

E. coli ST01 wild type pST101

ST02 wild type pST102 Table 2. P. aeruginosa strains used in this work. *pAJD2217=pHERD20T encoding N- term His-tagged P. aeruginosa EF-P. pAJD2218 and pAJD2219= pHERD20T encoding N-term His-tagged mutants, R32A and R32K, respectively. pST101 and pST 102=pET33B(+) encoding P. aeruginosa rmlA and His-tagged P. aeruginosa rmlA. #Strains supplied from Andrew Darwin (NYU).

Mutant strains AJDP752 and AJDP753 were created purifying the pAJD2217 vector, amplifying and mutating efp, and ligating it back in. The following primers were created for mutagenesis: R32A forward primer: 5’-CGAGTTCAACAAGTCCGGC

GCTAACGCTGCCGTCGTCAAG-3’, and the reverse primer: 5’-CTTGACGACGGCA

GCGTTAGCGCCGGACTTGTTGAACTC-3’. The R32K forward primer: 5’-

GAGTTCAACAAGTCCGGCAAGAACGCTGCCGTCGTCAAG-3’, AND R32K reverse primer: 5’-CTTGACGACGGCAGCGTTCTTGCCGGACTTGTTGAACTC-3’.

Site-directed mutagenesis PCR reactions were performed using Pfu Turbo DNA

polymerase. Following PCR, products were digested with dPN1 and transformed into

XL1-Blue cells, as well as XJB cells. Each mutation was confirmed by sequencing.

2.2) Growth Curves

Growth curves were performed in a variety of different media conditions. Cells grown in Lysogeny Broth (LB) were grown as follows: 5 mL starter cultures were inoculated and grown overnight at 37˚C. These starters were used to inoculate 50 mL LB cultures. LB cultures were supplemented with either 0.3% glucose or 0.3% glycerol.

Cells grown in minimal media followed the same protocol (minimal media has the following components: 5x M9 salts, 3.68 µg/mL CaCl2, 0.2 mg/mL MgSO4, 10 µg/mL

FeCl3, 0.3% glucose or glycerol, 0.1 mg/mL sodium citrate, filled to volume with water).

Cell density was monitored by observing the OD600.

2.3) Purification of EF-P

EF-P from wild type P. aeruginosa, the ∆EarP strain of P. aeruginosa, and the mutant strains R32A and R32K of P. aeruginosa were all purified using a His-tag affinity column. Cultures were grown in 1.5 L of LB induced with 0.2% arabinose, and allowed to grow for ~16 hours at 37˚C. Cultures were pelleted by centrifugation at 7,500xg for 8 minutes, and re-suspended in Buffer 1 [50 mM Tris pH 7.3 at 4˚C, 1 M NaCl, 10 mM imidazole, 2 mM ß-Mercaptoethanol, supplemented with protease inhibitor (Roche

Diagnostics)]. Cells were then lysed via sonication and clarified by centrifugation at

74,500 xg for 30 min, transferred to a clean tube, and the centrifugation step was

repeated. Purification of clarified lysate was performed using TALON Metal affinity resin from Clontech in a column at 4˚C. Beads were equilibrated by washing them with

20 column volumes of water followed by 20 column volumes of Buffer 2 [50 mM Tris, pH 7.4, 1 M NaCl, 10 mM imidazole]. After clarification of the lysate by sonication, the lysate was poured over the beads in a column. Washes of 50 column volumes were performed with buffer 2, and protein was eluted in Buffer 3 [50 mM Tris, pH 7.4, 1 M

NaCl, 160 mM imidazole]. Purified EF-P was identified on an SDS-PAGE gel, and put into one 4-hour dialysis in Buffers 4 [10% glycerol, 25 mM Tris, 150 mM NaCl, 2 mM ß- mercaptoethanol], and one overnight dialysis in Buffer 5 [50% glycerol, 25 mM Tris, 150 mM NaCl, 2 mM ß-Mercaptoethanol]. Purified protein was stored in 20 µL aliquots at -

80˚C.

Recombinant B. subtilis EF-P was grown in 1.5 L of Auto-induction media. After inoculation, cells were grown at 20˚C for 16 hours. Cells were then pelleted by centrifugation at 7,500 xg for 8 minutes and re-suspended in Buffer 6 [25 mM Tris pH 8,

600 mM NaCl, 2 mM ß-Mercaptoethanol, supplemented with protease inhibitor (Roche

Diagnostics)] and lysed by sonication, and clarified by centrifugation at 74,500 xg for 30 minutes at 4˚C, lysate was transferred to a clean centrifuge tube, and the centrifugation at

74,500 xg for 30 minutes at 4˚C was repeated. Chitin affinity resin from NEB were equilibrated by washing with 20 column volumes water followed by 20 column volumes

Buffer 7 [25 mM Tris pH 8, 600 mM NaCl]. Following clarification, lysate was poured over chitin resin and allowed to flow through. This was followed by two 50-column volume washes in Buffer 7. Elution was performed by flowing 3-column volumes of

Buffer 8 [25 mM Tris, pH 8, 600 mM NaCl, 100 mM DTT] over the column, and then leaving the resin in 1 column volume of Buffer 8 overnight. Following complete elution, protein was concentrated by 3 kDa centricon and dialyzed for 4 hours in buffer 4 and overnight in buffer 5. Purified protein was stored in 20 µL aliquots in -80˚C.

B. subtilis Flag-tagged EF-P was purified using ANTI- FLAG® M2 Magnetic beads. Cells were grown in 1.5 L of LB or M9 minimal media and induced with IPTG at an OD600 of 0.3 and allowed to grow for 2 hours. Cultures were pelleted by centrifugation at 7,500 xg for 8 minutes, and re-suspended in buffer 9 [50 mM Tris pH 7.4, 150 mM

NaCl, supplemented with protease inhibitor (Roche Diagnostics)]. Cells were then lysed using 0.25 mg/mL lysozyme at 37˚C for 30 minutes, followed by addition of 16 mg/mL

DNase for 30 minutes at 37˚C. Cells were then clarified by centrifugation at 20,000 xg for 15 minutes. ANTI-FLAG M2 Magnetic Beads then used as directed. Following purification, FLAG-EF-P was concentrated by 3kD centricon and stored at -80˚C.

Because these samples were prepared for mass spectrometry, no glycerol was added to the storage buffer.

2.4) Isoelectric Focusing and Western Blotting as a Tool

Either whole cell lysates or purified protein was used as sample when performing isoelectric focusing (IEF) gel experiments. Preparation of lysates was as follows: cultures were grown in 50mL flasks in either LB or M9 Minimal Media. Once cells had reached the desired OD600, cultures were pelleted by centrifugation at 7,500 xg for 8 minutes and re-suspended in Buffer 10 [10% glycerol, 25 mM Tris-HCl pH 8, 150 mM NaCl, 2 mM

ß-mercaptoethanol, supplemented with protease inhibitor (Roche Diagnostics)]. Cells were lysed either by sonication or by treatment with lysozyme, DNase, and RNase. After lysis, lysate was centrifuged at 21,000 xg for 30 minutes.

Native IEF gels were made using GE Healthcare Pharmalyte carrier ampholytes pH range 4.5-5.4. Gels were allowed to solidify for two hours, and pre-run at 100 V for

15 minutes, after which the gel was loaded. Once the gel was loaded samples were run at

200 V for 1 hour, 300 V for 1 hour, and 500 V for 30 minutes using 25 mM NaOH in the upper chamber and 20 mM acetic acid in the lower chamber. Following the run, the protein was transferred to a nitrocellulose membrane by a semi-dry transfer method, applying 0.8 mA per 1 cm2 of gel for 1 hour. Transfer buffer was composed of 20% ethanol in 1 x SDS-PAGE Buffer. The membrane sat in blocking buffer at 4˚C overnight.

Detection or EF-P was performed using anti-EF-P antibodies supplied from Daniel

Kearns (Indiana University).

2.5) In Vitro Aminoacylation of EF-P

In vitro assays were performed at 37˚C in 100 mM Hepes (pH 6, 6.8, 7.2, 7.4, 8, and 9), 30 mM KCl, 10 mM MgCl2, 2 mM ATP, 2 mM DTT, 50 μM [C]-lysine, inorganic pyrophosphate (Roche Diagnostics), 0.2mM [14C] Threonine, 26µM recombinantly purified EF-P, and cell-free B. subtilis extract. Timepoints were taken after

3, 5, 10, 20, 30 minutes, 1 hour, and 6 hours. Each timepoint was taken and immediately put into 5x protein loading buffer to quench the reaction. After the final timepoint, samples were run on a 13% SDS PAGE gel and visualized with a phosphorimager.

2.6.) Glycoprotein Stain

The glycoprotein-staining assay was performed using the Invitrogen Pro-Q

Emerald 488 Glycoprotein Gel and Blot Stain Kit. Purified EF-P from the wild type P. aeruginosa and ∆EarP strains were loaded into an SDS-PAGE gel. The gel was fixed in

Fix Solution (50% methanol and 5% glacial acetic acid in distilled water) and allowed to incubate with gentle agitation for one hour at room temperature. This step was repeated once. This was followed by two 20-minute washes in Wash Solution (0.3% glacial acetic acid in distilled water), and then the gel was placed in Oxidizing Solution (250 mL of 3% acetic acid combined with the supplied bottle of periodic acid) and allowed to incubate with gentle agitation for 20 minutes. The oxidation step was followed by three, 20-minute wash steps with Wash Solution and gentle agitation. The gel was stained by incubating the gel in the dark (wrapped with tin foil) with gentle agitation in the Pro-Q Emerald 488

Staining Solution (0.5 mL of DMSO to one vial of provided component A, followed by a

50 fold dilution into Pro-Q Emerald 488 staining buffer (supplied Component B) for 2 hours. Finally, the gel was washed with Wash Solution at room temperature with gentle agitation once for 20 minutes, and twice for 35 minutes. The gel was then visualized in a

Typhoon FLA 9000 at a fluorescence excitation wavelength of 470 nm.

2.7) Lectin Binding Assays

Purified P. aeruginosa EF-P from wild type and ∆EarP strains (in amounts of 1

µg, 100 ng, 20 ng, 0.4 ng, and 0.08 ng of purified EF-P) were spotted onto an Amersham nitrocellulose membrane. The CandyCane glycoprotein molecular weight standards mix was also spotted as a control. Non-specific binding was blocked by incubating the 22

membrane with gentle agitation in the provided Carbo-Free Blocking Solution for 30 minutes at room temperature. The membrane was then washed twice with gentle agitation with PBS for 1 minute each wash. Next the membrane was incubated with gentle agitation in 2 µg/mL biotinylated lectin for 30 minutes at room temperature, and then washed with TPBS (PBS +0.05% TWEEN 20) for 5 minutes with gentle agitation. The

VECTASTAIN elite ABC reagents were prepared according to kit instructions and the membrane was incubated in them with gentle agitation for 30 minutes at room temperature. The membrane was washed with TPBS and the substrate was applied for visualization by chemiluminescence.

2.8) [14C] dTDP-Rhamnose Synthesis

Synthesis of dTDP-[14C]Rhamnose was modeled after (37) with minor changes. A his-tagged rmlA strain of PAK was created by PCR amplifying the P. aeruginosa rmlA gene using the following primers: forward: 5’-CCATGGATGCACCACCACCACCAC

CACAAACGCAAGGGCATCATC-3’ and reverse: 5’-GGTGGTCTCGAGTCAGTA

CACGGTCTCGG-3’. The insert and the pET-33b(+) plasmid were digested using the restriction enzymes NCO1 and Xho1. The insert was then ligated into the pET-33b(+) plasmid using DNA Ligase and transformed into E. coli XJB and XL1 Blue cells.

α-D-glucose-1-phosphate thymidylyltransferase (RmlA) was prepared from the above P. Aeruginosa rmlA expressed in E. coli. The reaction was performed by first vacuum drying 50 μCi (442/mCi/mmol, 113nmol) of [U-14C] sucrose in a tube. Then the rest of the reaction components (16 μl of 1 M KH2PO4, pH 7.0, 80 μl (0.5 units) of sucrose phosphorylase, 10 μl of 40 mM TTP, 10 μl (2 units) of inorganic 23

pyrophosphatase (Roche Diagnostics), 200 μl of crude lysate of E. coli BL21 with P.

Aeruginosa rmlA cloned in pET-33b(+), 7 μl of 50 mM NADPH, 20 μl of 1 M HEPES buffer at pH 7.0, 8 µL of 500 mM MgCl2, and 49µL ddH2O, total volume of 400 µL) were combined and the reaction was allowed to proceed for 1 hour at 37˚C. From there, the rest of the RmlA enzymes were added- supplied from E. coli BL21 XJB crude lysate

(200µL) and additional NADPH (35µL of 10mM), and the reaction was allowed to proceed for 30 minutes longer at 37˚C to fully convert the dTDP-[14C]Glc to dTDP-

[14C]Rha. Upon completion, the reaction was filtered using an Amicon Ultra-0.5 3KDa

Centrifugal Filter Device to remove large proteins and the filtrate was vacuum dried until a final volume of about 250µL.

Chapter 3: Results

3.1.) Modification Status of EF-P in Bacillus subtilis

Bacillus subtilis is a member of the Firmicutes, a phylum of gram-positive bacteria. Unlike E. coli and P. aeruginosa, that have a large number of poly-proline motifs in their genome, B. subtilis was chosen as a model organism to study EF-P because of the small number of polyproline stretches predicted in its proteome. While

EF-P is needed for vegetative growth in E. coli, B. subtilis does not require EF-P activity for vegetative growth, however EF-P is essential for swarming motility (37). B. subtilis is missing both the PoxA and the EarP pathways, and the modification status of EF-P in this organism is currently unknown. The small amount of poly-proline stretches in the B. subtilis genome and the fact that B. subtilis lacked the modification machinery found in

E. coli and P. aeruginosa led to the question of whether or not EF-P in B. subtilis has a modification pathway. The modification on EF-P has shown to be necessary for EF-P to function (28, 35). Several approaches have been taken to address this question, including mass spectrometry, site directed mutagenesis, and isoelectric focusing that will be discussed in this work.

3.2) Mass Spectrometry

Mass spectrometry is a powerful analytical technique used to study the composition of chemicals. A chemical, once ionized, can be manipulated by applying electric and magnetic fields. Using the electric and magnetic fields, ions can be sorted and selected for, and their mass/charge (m/z) ratio can be measured. Mass spectrometry has been a particularly valuable asset in the establishment of a modification on EF-P in B. subtilis. Identifying modifications on proteins via mass spectrometry is a process that begins with purification of the protein of interest. Preliminary purification and mass spectrometry of B. subtilis EF-P was performed by Andrei Rajkovic.

Two different approaches of mass spectrometry, bottom up and top down were performed by Andrei Rajkovic to identify whether or not B. subtilis EF-P is post- translationally modified. Top down mass spectrometry is an approach that gives an overview of the system; there is no digestion or fragmenting of the protein. It provides general information about a protein but no information on the fragments other than their overall mass. Combining the data collected from a top down approach allows one to reassemble the protein of interest and obtain its mass. Top down data revealed that there is an overall extra mass on native, purified EF-P compared to the theoretical mass (Figure

5). Because there was a difference, the theoretical mass was subtracted from the mass obtained from native EF-P, resulting in an extra mass of 101 Da, which is attributed to be the mass of the modification. Figure 5B shows the mass spectrometry data for the recombinantly expressed EF-P in E. coli, which is missing the 101 Da extra mass, supporting the idea that the 101 Da is a result of the post-translational modification. More

information was needed to identify where the unexpected mass was located. Bottom down mass spectrometry is used to further characterize proteins. After the protein is digested and detected by the mass spectrometer, the protein enters a collision chamber, where high-energy collisions with gasses cause the protein fragments to be further broken up. Specific fragments can be isolated and others rejected, so it is possible to isolate and further fragment the peptide containing the conserved K32 residue, the site that is believed to be modified. The fragments can then be returned to the detector, obtaining more information about specific fragments. Bottom up mass spectrometry data revealed that the modification was indeed on the conserved residue K32, and the predicted mass of the modification on B. subtilis EF-P obtained from mass spectometry data is 101 Da

(Figure 5).

Figure 5. Mass spectrometry performed by Andrei Rajkovic. A.) Relative abundance vs. the m/z ratio of recombinant EF-P compared to native B. subtilis EF-P. B.) Identification of the extra mass attributed to the modification on B. subtilis EF-P on conserved K32.

Purification of native B. subtilis EF-P lacked the desired purity for mass spectrometry, so a strain of B. subtilis was created that had a FLAG-tag added onto EF-P.

FLAG-tagged EF-P was purified as described above and sent for further mass spectrometer analysis. An additional mass spectrometry experiment that could prove to be useful in identification of the modification is to include one more fragmentation step.

Once the fragment with the modification on it has been detected, it is possible to send it 28

to another collision chamber and fragment the modification off the peptide and once again return it to the detector. From this experiment, it would be possible to obtain information on the structure of the modification, depending on how the modification fragmented.

While the mass spectrometry data showed that the modification was on the conserved K32 residue, site directed mutagenesis was performed as a second way to confirm the position of the modification. Daniel Kearns’ lab performed site directed mutagenesis to mutate the conserved Lysine 32 to an Alanine (K32A). Growth curves were performed for wild type, ∆EF-P, and the K32A mutant of B. subtilis (Figure 6), which show that K32A exhibits the same growth pattern as ∆EF-P, but not the same as wild type B. subtilis.

Figure 6. Growth curves of wild type B. subtilis (Blue), ∆EF-P B. subtilis (Green), and K32A B. subtilis (Red).

3.3) Characterization of B. subtilis EF-P by Isoelectric Focusing

Isoelectric focusing is a technique that uses a charge gradient to separate proteins by their isoelectric focusing point. Applying a charge to ampholytes, or zwiterionic molecules that contain both acidic and basic groups, causes the ampholytes to separate according to their isoelectric focusing point, creating a pH gradient that can be used to separate proteins. While modified and unmodified forms of E. coli and P. aeruginosa EF-

P both have the same charge state, B. subtilis EF-P displayes a different charge state from modifeid to unmodified EF-P (Figure 7). Therefore, isoelectric focusing gels are a valuable tool for monitoring the modification state of B. subtilis EF-P and has been employed in several assays in this work. 30

Isoelectric focusing also suggested that the K32A residue is the site of modification on B. subtilis EF-P. Cell lysates from native B. subtilis, purified recombinantly expressed EF-P from E. coli, and the mutated K32A EF-P cell lysate were the run on an isoelectric focusing gel and transferred to a nitrocellulose membrane. The

K32A mutant runs in a different spot than both recombinant EF-P and wild type EF-P as it is missing the charge from the modification as well as the charge from the lysine. This is what is seen from the isoelectric focusing gel (Figure 7). Further support that the modification is indeed on K32 is that the K32A mutant exhibited different colony morphology than wild type B. subtilis. While the wild type cells have rough colonies,

K32A displays smooth colonies, as does the ∆EF-P strain.

1 2 3

Wild Type EF-P

Purified EF-P recombinantly expressed in E. coli K32A

Figure 7. Western blot from IEF gel showing the bands of K32A EF-P (lane 1), B. subtilis wild type EF-P (lane 2), and recombinant EF-P produced in E. coli (lane 3)

3.4) In-vitro assays probing the identity of the modification substrate

In vitro assays were performed to attempt to identify possible modifications. The first substrate that was chosen to pursue was threonine, as there were several pieces of evidence supporting the idea that threonine was the modification. The first piece of evidence is that the mass of threonine is the correct mass corresponding to the data obtained from the mass spectrometer. The second piece of evidince is that B. subtilis has two genes encoding for threonyl-tRNA synthetase, thrS and thrZ. ThrS is predominantly used during vegetative growth when there is an abundance of threonine (38). When threonine concentrations become low, thrS is suppressed and thrZ is induced (38, 39). It is possible that one of these proteins aminoacylates EF-P with threonine. This idea is especially plausible when compared to the E. coli modification pathway- PoxA is a lysyl- tRNA synthetase and very closely resembles lysyl-tRNA synthetase (40). While PoxA cannot aminoacylate tRNALys or lysyl-tRNA synthetase aminoacylate EF-P (26), PoxA and lysyl-tRNA synthetase are structurally very similar (41).

In vitro aminoacylation assays were performed on recombinant EF-P, with purified ThrZ or ThrS as the modifying enzyme and [14C] Threonine as the substrate by a previous student. However these experiments were unsuccessful in showing that either

ThrS or ThrZ could modify recombinantly expressed EF-P. Similar reactions were performed using recombinant EF-P, [14C] Threonine and wild type B. subtilis lysate, with the hope that the enzyme catalyzing the modificaiton of EF-P was abundant enough in the lysate to perform the reaction as described in the methods. Reactions were performed using wild type B. subtilis lysate, ∆EF-P B. subtilis lysate, and wild type lysate that was

run over sephacel beads to remove as much tRNAThr as possible so as to avoid seing aminoacylation of tRNA rather than of EF-P. Six assays were performed over a pH range of 6-9, over a course of 6 hours. When these results again lacked any activity, more substrates were tried, many of which were possible precursors instead of substrates. Other potential compounds we attempted to aminoacylate EF-P with were homoserine and betaine as potential substrates. Thus far, no substrates have been shown to modify B. subtilis EF-P in-vitro.

Other than threonine, there are several biological molecules with the same molecular weight that have a biosynthetic pathway in B. subtilis. Because radioactive substrates were not readily available for in-vitro assays, genetic approaches were taken to discover the identity of the modification. The biosynthetic pathways of potential substrates were investigated using the corresponding deletion strains that were available for purchase from the Bacillus Genetic Stock Center. The strains were grown to a specific

OD600 , followed by lysis, isoelectric focusing, and western blotting. Although this did not directly remove the modification from EF-P, deletion of genes in the biosynthetic pathways of substrates provided a way to access the modification state of EF-P when the substrates should be missing from the cell. One of the molecules that this was attempted with was betaine, which, besides having the appropriate mass also has an appropriate charge state to account for the charge differences seen in the isoelectric focusing gels.

Betaine is synthesized in B. subtilis by the uptake of choline (42) by the OpuB and the

OpuC ABC transporters followed by a two-step oxidation process. The enzymes GbsA and GbsB are responsible for the oxidation process. The opuB and the opuC operons are

known to be osmoinducible, and choline suppresses opuC during normal cellular growth.

GbsR is a regulatory protein that controls the transcription of both gbsA and gbsB, as well as for opuB and opuC. It has been shown that choline suppresses opuC expression during normal growth, and that in the absence of GbsR; opuB is also suppressed by choline (43).

Two mutants were purchased, a GbsA deletion and a GbsR deletion. The strains were grown and their crude lysate run on an IEF gel followed by western blotting. Both of these deletion strains produced modified EF-P (Figure 8), leading us to believe that betaine is not the substrate for B. subtilis EF-P modification. This approach was attempted for several substrates, however none of the deletion strains supplied unmodified EF-P.

Figure 8. IEF gel followed by electroblotting to identify the modification status of EF-P in ∆gbsA and ∆gbsR strains of B. subtilis. Lane 1.) Recombinant EF-P 2.) Wild type B. subtilis EF-P, from OD600= 1.4 3.) Wild type B. subtilis EF-P, from OD600= 1.4. 4.) ∆gbsA, OD=0.2 5.) ∆gbsR, OD=0.2 6.) ∆gbsA, OD=1.5 7.) ∆gbsR, OD=1.5

3.5) Glucose as a Potential Regulator

While performing isoelectric focusing gels followed by western blotting, it was noted that the ratios of the modified EF-P to unmodified EF-P were variable. This variability was traced to the differences in when the cells were harvested. If the cells were lysed when they were at early or mid-log phase, then there was a large majority of modified EF-P. However when the cells were lysed after they had grown into stationary phase, there was about an equal amount of modified to unmodified EF-P. While there is no evidence to suggest that EF-P is regulated in other systems, it is possible that EF-P could be regulated in B. subtilis. Independently, in a study using pulse-chase labeling to study the degradation targets of the Clp proteases, it was shown that during B. subtilis growth in glucose-starved conditions, the state of EF-P changes. Whether EF-P is modified or degraded during this study is not clear, and it’s noted that the changes occurring to EF-P are probably not linked to that of the Clp proteases (39). Growth curves were performed in LB with and without glucose as the carbon source- glycerol was supplemented instead of glucose- as well as in M9 minimal media with and without glucose as the carbon source. Cells were then grown and collected at 8 different times through their growth curve, lysed, and the lysate separated on an IEF gel. It was discovered that there is a characteristic shift in the ratios of modified to unmodified EF-P.

The same experiment was run for wild type EF-P grown in LB and minimal media, as well as the K32A mutant grown in LB and in minimal media (Figure 9). The K32A analysis shows no change in ratios of modified vs. unmodified EF-P, however there are

two EF-P bands detected. This raises the question of whether or not EF-P has a second modification, such as a phosphorylation. Because figure 7 establishes that the K32A mutant focuses at a charge state that accounts for both charge lost due to the loss of the charge on lysine as well as the charge lost due to the missing modification, the second band seen on the K32A profile can be attributed to something other than the modification of Lys. An extra band was also seen upon performing this experiment with WT B. subtilis. Three bands were detected rather than only two, one of which disappeared after an OD600= 0.5. It is possible that B. subtilis EF-P has a second modification on it, such as phosphorylation or amination. Phosphorylation of eIF5A is essential for function (44), and essential phosphorylations in prokaryotic transcription factors, such as EF-Tu have also been observed (45). Further experiments are required to confirm whether or not a second modification is present, and could be done by mass spectrometry.

Figure 9. The change of the modification status of EF-P as a function of OD600

3.6) EF-P in P. aeruginosa

Mass spectrometry and site directed mutagenesis were also used to probe the modification on P. aeruginosa. Two mutants of P. aeruginosa were made; the conserved

R32 residue was mutated to both an alanine and a lysine. Growth curves performed from these mutants confirmed the importance of the R32 residue as both the alanine as well as the lysine mutant displayed impaired growth.

3.7) Glycoprotein Staining

For the first attempt to confirm that a glycosylation was the modification on P. aeruginosa EF-P, a Pro-Q Emerald 488 Glycoprotein Gel and Blot Stain Kit were used.

This kit allows in gel detection of glycoproteins by first oxidizing and then tagging them with a fluorescent dye. Once EF-P was run on an SDS Page gel, the gel was fixed with methanol and glacial acetic acid. Following fixing, periodic acid was used to oxidize any carbohydrates, opening the sugar ring and leaving a ketone. Finally, a fluorescent dye was attached by reacting the carbohydrate with an amine group bonded to the probe

(Figure 10).

While promising in theory, this technique proved ineffective for a variety of reasons. While it was possible to detect the CandyCane glycoprotein molecular weight standards (Life Technologies), which contained a mixture of both glycosylated and non- glycosylated proteins, these are all heavily glycosylated proteins, whereas EF-P only has one glycosylation. The background for these assays was also very high, potentially because of side reactions with the backbone of EF-P.

+ ProQ Emerald Reagent

Figure 10.The glycoprotein staining reaction. Adapted from Life Technologies Molecular Probes: the Handbook.

3.8) Lectin-Binding Assays

In order to probe further into whether or not the modification on P. aeruginosa was indeed a sugar molecule, lectin staining assays were performed. A lectin is a carbohydrate binding protein. This particular lectin-binding assay uses a biotinylated lectin to bind carbohydrates, and then takes advantage of biotin avidin interaction for detection. Once the lectin is bound to the carbohydrate, the Vectastain Elite ABC kit was

used for detection. The ABC kit is comprised of two components; avidin and biotinylated horseradish peroxidase. Once the biotinylated lectin binds to the carbohydrate, the avidin will bind to the biotinylated lectin, as well as to the biotinylated horseradish peroxidase.

The horseradish peroxidase was then detected using p-coumaric acid, luminol, and hydrogen peroxide. With the knowledge that the mass of the modification on P. aeruginosa EF-P was about 164 Da, the two options for possible modifications were either fucose or rhamnose. The available fucose-binding lectin was Ulex Europaeus agglutinin 1 (UEA). While twenty-one different lectins were tested for binding to EF-P, no binding was detected. Upon closer inspection, this assay failed because the specificity of the lectins was not properly taken into account. While UEA binds fucose, it only binds to fucose when the fucose is in the specific motif of Fucα1-2Gal-R

(http://www.interchim.fr/ft/M/MS902z.pdf).

3.9) [14C] Rhamnose Synthesis

With the knowledge that the mass of the modification on P. aeruginosa EF-P was about 164 Da, as well as the similarity of EarP to a glycosyl transferase, the identity of the modification could be narrowed down to a couple of options. After investigation of P. aeruginosa’s metabolic pathways, the identity of the modification was narrowed down to either fucose or rhamnose. This knowledge prompted performance of in vitro analyses with P. aeruginosa EF-P and rhamnose.

Due to lack of P. aeruginosa EF-P antibody, radioactivity was the method best suited to performing in-vitro reactions. Unfortunately, dTDP-[14C]Rhamnose is not

available to purchase, so it was enzymatically synthesized. The RmlC pathway in P. aeruginosa is the pathway that procures the rhamnose to be used to modify EF-P. The first step of this synthesis is for RmlA to convert α-D-glucose-1-P and dTTP into dTDP- glucose and inorganic phosphate. Next RmlB converts dTDP-glucose to dTDP-4-keto-6- deoxyglucose. RmlC converts 4-keto-6-deoxyglucose into dTDP-4-keto-rhamnose, and with the aid of NADPH, RmlD converts dTDP-4-keto-rhamnose into dTDP-rhamnose

(figure 11). This reaction was performed as stated in the methods section to generate dTDP-[14C]Rhamnose.

Figure 11. The synthesis pathway for dTDP- rhamnose. Adapted from (46)

Chapter 4: Future Work

4.1) Identifying potential modifying enzyme in B. subtilis

As in the other EF-P modification pathways, it is possible that B. subtilis has a protein responsible for attaching the modification to EF-P. Identifying whether or not B. subtilis has a protein that catalyzes the modification of EF-P, or if the modification is spontaneous is important to further understanding EF-P. Because FLAG-tagged EF-P has already been purified and sent for mass spectrometry, looking back through the mass spectrometry data for binding partners could lead to clues as to what this protein is.

If mass spectrometry data doesn’t reveal anything about identity of this protein, then there are several other approaches that can be taken to identify this pathway. One approach is to use ammonium sulfate fractionation on ∆EF-P B. subtilis lysate and recombinant EF-P to identify the fraction that this protein is in. This can be made detectable by cloning a Protein Kinase A (PKA) site into B. subtilis EF-P, followed by labeling with 32P. ∆EF-P lysate from B. subtilis can be fractionated by ammonium sulfate and then the fractions can be mixed with unmodified PKA EF-P. After incubating the mixture for 30 minutes at 37˚C, endpoints will be taken and the mixtures will be run on an IEF gel to separate modified from unmodified. The gels will then be dried and

exposed on phosphorimaging screens. Currently, the PKA site has been cloned into B. subtilis EF-P, but the rest of the project has yet to be carried out.

Another option is to approach this problem from a genetics standpoint. Assuming that EF-P in B. subtilis performs the same function that it does in E. coli and P.

Aeruginosa, A GFP-mCherry reporter system can be used, placing a poly-proline stretch in front of the GFP gene so that a functional EF-P is needed to translate it. Random mutagenesis in the B. subtilis genome can then be performed, with the hope that one or more of the resulting mutations create a critical mutation in the modifying enzyme. While this mutagenesis could create other mutations that affect the production and therefore florescence of GFP, the cells that show a decrease in GFP fluorescence but normal mCherry fluorescence are the target mutants to further pursue. From there, cell sorting will be used to collect the target cells and either deep sequencing or RAPD PCR will be used to pinpoint where the mutations are. RAPD PCR is PCR with arbitrary, 8-10 nucleotide primers with genomic DNA as the template. From there, the common mutants will be further studied to identify which mutations cause EF-P to remain unmodified.

While IEF and electroblotting techniques show that there is the possibility of B. subtilis EF-P being regulated, more work needs to be done to confirm whether this is actually true. However before the question of whether or not it’s regulated with glucose can be answered, several other questions need to be addressed. Glucose starvation of B. subtilis needs to be better quantified, and tests need to be performed to confirm that the point at which glucose starvation occurs in B. subtilis correlates with the point at which changes in the modification status occur. Glucose levels in the cell can be ascertained by

the use of a glucose uptake assay kit. Another important experiment to be performed is to see if adding glucose to growing cells can rescue the modification status. This will address the question of whether or not the modification status of EF-P will revert back to the ratio of modified and unmodified EF-P that the cell was producing before it was lacking in glucose levels.

Once these questions have been addressed, methods other than IEF can be used to confirm and support what the IEF gels show. Mass spectrometry data from different points in the cell’s growth curve can be used to show the different ratios of modified and unmodified EF-P, or a reporter construct, like the already mentioned GFP-mCherry construct could be used. If a poly-proline stretch is placed before gfp, then fluorescence levels of GFP compared to mCherry can be used to monitor the relative amount of EF-P throughout the cells growth.

References:

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