Flagellar Protein Flhe Influences Motility and Morphology Through Regulation of Assembly in Escherichia Coli

The bacterial flagellar motor is a remarkably complex system, comprising thousands of protein subunits and responsible for motility in numerous species. Of these proteins, FlhE is not universally conserved and is relatively understudied. Usually found cotranscribed with flhB and flhA in the flhBAE operon, the flhE gene encodes a 14 kDa, 130 amino acid protein that is present in several peritrichously flagellated Gram-negative species. Deletion of chromosomal flhE in E. coli results in very unusual cellular morphologies, accompanied by an abrogation in swarming (surface-associated) motility and a severe defect in swimming motility. All of these phenomena are relieved upon introduction of full-length flhE from a plasmid, but not by an N-terminally truncated version of the gene (Δ1-17) lacking the signal sequence that targets FlhE to the periplasm. Defects are complemented even at a basal level of expression suggesting that FlhE is present in low-copy number, though overexpression of the protein harms cells and results in a growth defect. Both the growth and morphology defects are remedied in ΔflhE mutants with the overexpression of FlgM, an anti-σ28 factor responsible for repressing late-stage flagellar components. This suggests that at least one of these components plays a role in the defects seen in ΔflhE cells. These and other observations support a model in which FlhE is responsible for keeping the basal body pushed up against the peptidoglycan (PG) layer, allowing for proper assembly and export of the motor and filament. In the absence of this stabilizing interaction, structures that normally assemble outside the cell (the hook and filament) assemble in the periplasm, leading to distortion of the cell envelope and consequent growth defects.

FLAGELLAR PROTEIN FLHE INFLUENCES MOTILITY AND MORPHOLOGY THROUGH REGULATION OF ASSEMBLY IN ESCHERICHIA COLI by Rachel Katherine Ridge A Senior Honors Thesis Submitted to the Faculty of The University of Utah In Partial Fulfillment of the Requirements for the Honors Degree in Bachelor of Science In The Department of Chemistry Approved: ______________________________ David Blair, PhD Thesis Faculty Supervisor _____________________________ Cynthia Burrows, PhD Chair, Department of Chemistry _______________________________ Thomas Richmond, PhD Honors Faculty Advisor _____________________________ Sylvia D. Torti, PhD Dean, Honors College December 2017 ii ABSTRACT The bacterial flagellar motor is a remarkably complex system, comprising thousands of protein subunits and responsible for motility in numerous species. Of these proteins, FlhE is not universally conserved and is relatively understudied. Usually found cotranscribed with flhB and flhA in the flhBAE operon, the flhE gene encodes a 14 kDa, 130 amino acid protein that is present in several peritrichously flagellated Gram-negative species. Deletion of chromosomal flhE in E. coli results in very unusual cellular morphologies, accompanied by an abrogation in swarming (surface-associated) motility and a severe defect in swimming motility. All of these phenomena are relieved upon introduction of full-length flhE from a plasmid, but not by an N-terminally truncated version of the gene (Δ1-17) lacking the signal sequence that targets FlhE to the periplasm. Defects are complemented even at a basal level of expression suggesting that FlhE is present in low-copy number, though overexpression of the protein harms cells and results in a growth defect. Both the growth and morphology defects are remedied in ΔflhE mutants with the overexpression of FlgM, an anti-σ28 factor responsible for repressing late-stage flagellar components. This suggests that at least one of these components plays a role in the defects seen in ΔflhE cells. These and other observations support a model in which FlhE is responsible for keeping the basal body pushed up against the peptidoglycan (PG) layer, allowing for proper assembly and export of the motor and filament. In the absence of this stabilizing interaction, structures that normally assemble outside the cell (the hook and filament) assemble in the periplasm, leading to distortion of the cell envelope and consequent growth defects. iii TABLE OF CONTENTS ABSTRACT ii INTRODUCTION 1 MATERIALS AND METHODS 8 RESULTS 17 DISCUSSION 31 REFERENCES 36 1 INTRODUCTION The bacterial flagellar motor is a complex system of thousands of protein subunits working in concerted effort to drive the motion of the cell. Species like Escherichia coli and Salmonella typhimurium are peritrichously flagellated, with extracellular helical filaments uniformly dispersed across the cell surface (1). The filaments rotate by means of the rotary motors at their base, spinning clockwise or counterclockwise in response to the surrounding medium (2). Harnessing a chemical gradient of protons (or Na+ ions in some marine species), the helical flagella can rotate at speeds as high as 102,000 rpm (3). Over 50 genes are responsible for the biosynthesis and operation of the motor, and are expressed in a very specific order (Fig. 1). Early flagellar proteins are controlled by master regulator flhDC, for example, while late-stage flagellar genes are activated by the export of FlgM – an anti-σ28 factor that binds and represses late-stage genes until the hook-basal body structure is properly assembled (4). Flagellar assembly begins with the formation of the MS-ring (composed of FliF subunits) in the inner membrane (5). The MS-ring then acts as a scaffold upon which the export apparatus, motor, and switch are built (2). Extracellular components of the growing bacterial flagellum are exported via the type III secretion system (T3SS), used also in the injectisome present in some virulent bacteria. The injectisome allows Gram-negative bacteria to deliver a payload of effector proteins directly into their eukaryotic host cells by way of a short, straight needle (5). Many components of the injectisome have homologs in the flagellar motor, and the two systems share many structural and functional similarities as well as the T3SS for protein export (5). The flagellar type III secretion (T3S) apparatus is comprised of a number of proteins (FlhA, FlhB, FliO, FliP, FliQ, FliR, FliH, FliI, and 2 FliJ) and is built in the membrane and on the cytoplasmic side of the MS-ring (5). Substrate proteins, likely in their unfolded form, are exported through a narrow channel formed primarily by FliP and driven by the proton motive force (PMF) (6). After the secretion apparatus is in place, the rotor/switch complex (also known as the C-ring, composed of FliG, FliM, and FliN) attach to the MS-ring on its cytoplasmic face (5). Motor proteins MotA and MotB are then recruited to the inner membrane to form the stator complex, responsible for the rotation of the flagellar motor by working in concert with the rotor (FliG) to generate torque. FIG. 1. Sequential assembly of flagellar motor in Salmonella enterica. Components of the flagellar motor and their stepwise synthesis are shown in order. Brackets indicate components that are assembled before the type III secretion (T3S) apparatus is employed to export flagellar proteins. Newly-assembled substructures are shown in light gray at each step, and the proteins and genes (shown in italics) responsible for assembly are listed in the order they are recruited. The stators (composed of Mot proteins) are noncovalently attached to the MS ring, and are still present in later stages but not shown. OM, P, and CM stand for Outer Membrane, Peptidoglycan, and Cytoplasmic Membrane, respectively. Figure taken from Macnab 2003. 3 After the rotor, switch, and stator are in place, the T3S apparatus secretes the majority of the remaining components of the flagellar motor which self-assemble at the distal end of the structure as it develops (5). The rod – a hollow, cylindrical structure that traverses the periplasmic space – is built first, followed by the recruitment of the L- and Prings which are exported via the Sec pathway (7). The L- and P- rings are encoded by flgH and flgI and are named for the lipopolysaccharide and peptidoglycan layers they are embedded in, respectively (7). These components – including the MS ring, C-ring, rod, and L- and P- rings – are collectively referred to as the basal body. In the final stage of flagellar assembly, T3S is employed to export proteins that make up the hook (FlgE, FliK) and filament (FliC, also called flagellin) as well as the junctions and caps associated with them (7). The hook cap (FlgD) displaces the rod cap (FlgJ) and the hook begins elongating, stopping at the precise length of 55 nm in wild-type cells controlled by molecular ruler FliK (8). At this point, the hook cap is replaced by hook-associated proteins and hookfilament junction proteins and the cell begins assembling the helical filament (2). The filament, preceded by filament cap (FliD), is composed of as many as 30,000 subunits of the protein FliC and grows up to 10-15 μm long (5). At this point, the flagellum is fully assembled (Fig. 2) and can begin propelling the cell forward through liquid medium. The motor harnesses the electrical (Δψ) and chemical (ΔpH) transmembrane potential of protons to drive mechanical rotation; this is known as the protonmotive force (PMF) (9). The PMF represents the work per unit charge that is required to move a proton (H+) from the outside to the inside of the cell. It is given by the equation RT ∆p = ∆ψ − 2.3 ( ) ΔpH F (Equation 1) 4 where p is the protonmotive force in V, R is the universal gas constant in J·K-1·mol-1, T is the temperature in degrees Kelvin, F is Faraday’s constant in C·mol-1, and ΔpH and Δψ (in V) are the differences in pH and electrical potential, respectively, between the inside and outside of the cell (10). Protons pumped outside the cell by various respiratory processes flow back into the cell, down their concentration gradient. When protons return through the flagellar motor they generate torque by inducing conformational changes in the stator that then drives the motion of the rotor (11). This rotation is then translated up the flagellum to the rigid filament whose helical nature produces thrust (12). Not all bacteria are propelled by a proton-driven motor, however. There are some marine and/or alkalophilic species (e.g. Vibrio spp. and Bacillus spp.) that harness Na+ ions for motility instead; this is referred to as the sodium-motive force (2, 13). This complex pattern of assembly and energy coupling culminates in bacterial motility, allowing these cells to navigate and respond to their environment. Enteric bacteria capable of chemotaxis can control the direction of rotation of the flagellar motor in response to environmental cues. Membrane-associated receptors detect the presence of ligands in the cell’s surrounding environment and transmit this information to the bacterial cell (12). The switch complex (C-ring; FliG/M/N) then responds by changing the direction of rotation between clockwise (CW) and counterclockwise (CCW) rotation (11). When the motor is rotating CCW, the cell exhibits a roughly linear “run” and propels itself through the medium. When the motor switches rotation to CW, it “tumbles” in place and reorients itself in a new direction. Bacterial cells possess a rudimentary form of memory by means of a delayed response to external ligand binding; this allows them to compare present conditions to those a few seconds previous and respond accordingly (12). This 5 phenomenon allows enteric bacteria to bias the rotation of their flagellar motor, resulting in longer “runs” in a favorable direction and ultimately a net migration to a more desirable environment (12). FIG. 2. Structural overview of completed flagellum in Salmonella. This diagram shows the result of proper assembly of the flagellum in Salmonella, similar in many ways to that of Escherichia coli. The membrane-bound basal body provides the anchor that confers rotational torque through the flexible hook (FlgE) by means of rotor-stator interactions. The hook acts as a universal joint that translates this motion to the rigid filament (FliC), allowing cells to propel themselves through liquid media. Dashed boxes indicate proteins that are involved in flagellar type III secretion. During the highly-ordered assembly process, several sets of rings self-assemble and are recruited to their appropriate positions. Notably, the motor is held in place by the MS ring embedded in the inner membrane (IM), the P-ring embedded in the peptidoglycan (PG) layer, and the L-ring embedded in the outer membrane (OM). FlhE is absent from this diagram as we do not yet know where it fits into this picture; this is the subject of the current study. Figure taken from Erhardt et al, 2010. Of these many components that allow bacteria to swim, flagellar motor protein FlhE is understudied and presently not well understood. FlhE, encoded by the flhE gene in the bacterial chromosome of many enterics, is 130 amino acids long. While not universally 6 conserved, an analysis using the Basic Local Alignment Search Tool (BLAST) of the NCBI reveals that flhE is present in many peritrichously-flagellated enterics, γ-proteobacteria, and β-proteobacteria. Following flhA in the flhBAE operon, flhE encodes an 14 kDa protein whose crystal structure has been solved (13). A role for FlhE in regulating proton flow through the export apparatus has been proposed by the Harshey group (14). Evidence for this proposal will be discussed below, as well as more-recent results that we believe supports a different model for FlhE function. To understand how FlhE may play a role in bacterial motility and morphology, one must be familiar with the general structure of the cell envelope. Of the three layers that compose the cell envelope, the lipopolysaccharide (LPS) layer refers to the outermost membrane (OM) that is covered in LPS – a glucosamine disaccharide with a polysaccharide core, six or seven mostly-saturated acyl chains, and a stretch of polysaccharide called the “O-antigen” that is used in classifying pathogenic E. coli strains. The OM is attached to the underlying peptidoglycan (PG) layer by means of murein lipoprotein (Lpp), by far the most abundant protein in E. coli (15). The peptidoglycan layer is a heteropolymer unique to bacterial cell walls, made up of repeating units of alternated N-acetylglucosamine and N-acetylmuramic acid that are cross-linked by pentapeptide side chains (15). Any agent that damages the PG can cause cell lysis due to the high turgor pressure of the cytoplasm; this lysis can be prevented in liquid media of high osmolarity (15). Beneath the PG lies the inner membrane (IM), a phospholipid bilayer which encases the cytoplasm and in which many cellular processes take place (e.g. energy production, transport, lipid biosynthesis, etc.). The periplasm refers to the aqueous compartment between the inner and outer membranes of the bacterial cell. More viscous than the cytoplasm, the periplasm is crowded 7 with densely-packed proteins and allows for the compartmentalization of cellular processes, especially those that may otherwise harm the cell (15). FlhE has a predicted N-terminal signal peptidase I leader sequence, meaning that it is targeted to the periplasm after being synthesized by ribosomes in the cytoplasm (16). Because it is periplasmically targeted, FlhE likely interacts with some entity within the periplasm to carry out its function. Screening of several periplasmic proteins for interaction with FlhE has suggested that FlhE may interact with the periplasmic domain of FliF, though this interaction is the subject of ongoing research. FliF subunits form the MS ring of the flagellar motor, responsible for the framework on which the rotor/switch complex and export apparatus are built during the first steps of flagellar assembly (2). We propose a model for the function of FlhE in which the protein is involved in the promotion of proper flagellar assembly, by holding the inner parts of the basal body in place and preventing the premature export of hook and filament. We hypothesize that in the absence of this interaction, might move away from the peptidoglycan layer to allow assembly of late stage products (e.g. the filament) in the periplasm. We believe that the inappropriate assembly of the flagellum in the periplasmic space leads to the defects observed in ΔflhE cells. Either damage to the peptidoglycan or the presence of the filament in the periplasm could spell disaster for these cells and explain both the grotesque morphologies and cell lysis associated with the ΔflhE phenotype. 8 MATERIALS AND METHODS All plasmids and strains of Escherichia coli and Salmonella enterica used in this study are detailed in Table 1. Transformations and plasmid isolations were performed as previously described (17). Tryptone broth (TB) contained 10 g/L tryptone and 5 g/L NaCl. Luria-Bertani broth (LB) contained 10 g/L tryptone, 5 g/L NaCl, and 5 g/L yeast extract. Swim plates were prepared by adding 0.27% agar to TB, and contained 25 μg/mL chloramphenicol and varying concentrations of sodium salicylate (Na-Sal). Sodium salicylate and IPTG (isopropyl-β-D-1-thiogalactopyranoside), both inducers used in this study, were prepared respectively as 100 mM and 25 mM stocks in water. In liquid media, ampicillin was added to a final concentration of 125 μg/mL and chloramphenicol and kanamycin were both used at a final working concentration of 50 μg/mL. These antibiotics were prepared as 125 mg/mL and 50 mg/mL stocks, respectively, in 95% ethanol. Site Directed Mutagenesis Point mutations in the sequence of wild-type flhE were made using the Quick Change mutagenesis protocol (Stratagene, La Jolla, CA) in either the pKG116 or pRR48 vector and confirmed by sequencing. Functional chromosomal deletions of flhE were performed using λ-red recombination techniques as previously described (18). 9 Table 1. Strains and plasmids used in this study Strain or Plasmid Relevant Characteristica Source or Reference Escherichia coli strains RP437 Escherichia coli str. K-12 subst. MG1655 RP3098 E. coli ΔflhDC MS3367 E. coli ΔflhE (flhE::TetRA) MS3016 E. coli ΔflhE (FRT::flhE::FRT) DB225 E. coli ΔfliG MS2947 E. coli ΔflhE (FRT::flhE::FRT) in DB225 MS371 pKG116 in RP437; chlR MS3501 pKG116 in MS3367; chlR MS3484 pKP545 in MS3367; chlR MS3485 pKP575 in MS3367; chlR MS3486 pKP578 in MS3367; chlR MS3487 pKP577 in MS3367; chlR MS3488 pKP595 in MS3367; chlR MS3489 pKP598 in MS3367; chlR J. S. Parkinson J. S. Parkinson This study This study (Lloyd et al., 1996) This study This study This study This study This study This study This study This study This study Salmonella enterica strains LT2 Salmonella enterica serovar Typhimurium TH12483 S. enterica ΔflhE in LT2 K.T. Hughes K.T. Hughes Plasmids pRR48 pKG116 pHT100 pKP147 pTRC99A pMC64 pKP545 pKP548 pKP552 pKP553 pKP575 pKP577 pKP578 pKP596 pKP597 pKP598 pKP601 J.S. Parkinson J.S. Parkinson (Tang et al., 1996) This study K.T. Hughes K.T. Hughes This study This study This study This study This study This study This study This study This study This study This study a Ptac expression vector; ampR Psalicylate expression vector; ; chlR GST-only control vector; kanR flhDC in pKG116; chlR Psalicylate expression vector; chlR S. enterica FlgM in pTRC99A; chlR flhE in pKG116; chlR flhEcodons 17-30 in pKG116; chlR GST-tagged flhEcodons 17-30 in pHT100; kanR HA-tagged fliFcodons 43-454 in pRR48; ampR flhE.W22D in pKG116; chlR flhE.R34D in pKG116; chlR flhE.L31D in pKG116; chlR flhE.W22D+L31D in pKG116; chlR flhE.W22D+L31D in pKP552; kanR flhE.W22D+R34D in pKG116; chlR flhE.W22D+R34D in pKP552; kanR chlR, chloramphenicol resistant; ampR, ampicillin resistant; kanR, kanamycin resistant 10 Motility Assays To ascertain the motility of certain strains, swimming in soft agar was measured quantitatively as previously described (19). The soft agar in swim plates was prepared using 0.27% agar in tryptone broth (abbr. TB; 10 g/L tryptone, 5 g/L NaCl), and contained appropriate antibiotics and varying levels of inducer. Figure 3 displays the results of an experiment using wild-type E. coli cells (abbr. WT; strain RP437) transformed with control plasmid pKG116 which confers resistance to chloramphenicol (abbr. chl). Using pKG116 as a vector, flhE in varying forms was restored to cells lacking chromosomal flhE. Sodium salicylate (abbr. Na-Sal) was added as an inducer to final concentrations of 0, 2.5, 5, 10 and 20 μM to increase the amount of FlhE expressed from the pKG116 vector. Plates compared the swimming rates of WT cells to ΔflhE strains containing either a control plasmid (pKG116), full-length flhE, or full-length flhE containing one or two point mutations. Frozen stocks were streaked out onto LB plates containing chloramphenicol, and liquid cultures were started from single colonies in triplicate and incubated overnight at 32°C with shaking. The next morning, 3 μL of overnight culture was spotted onto swim plates which were then incubated at 32°C; and the diameter (in mm) was measured and recorded every hour over the course of several hours. Morphological Observations Cells in liquid culture were observed mainly via phase-contrast microscopy, at 600X (Nikon Phase Contrast Microscope; CFW15x Nikon ocular; Fluor 40, Nikon objective lens). Small volumes (≤5 μL) of overnight or mid-log culture were observed and images were recorded using GrabBee USB Video Grabber software (VideoHome 11 Technology Corp, New Taipei City, Taiwan) and a COHU High Performance CCD camera. GST Pulldown Assay To measure the interaction in E. coli between FlhE and the periplasmic domain of FliF (residues 43-454), a GST pulldown assay was performed as described previously (20, 21). Cells lacking master flagellar regulator flhDC (RP3098) were transformed with one of three plasmids: one containing a GST-tagged construct of FlhE cloned into parent vector pHT100 (20), another containing the periplasmic domain of FliF with a hemagglutinin tag (HA-FliF43-454), and the other (pHT100) expressing GST only. A colony of each freshly transformed strain was grown overnight at 32°C in 40 mL of tryptone broth (TB) containing appropriate antibiotics and 100 μM IPTG to induce plasmid expression. The next day, the A600 was measured and recorded for each strain. Cells were pelleted (3000 x g, 5 minutes), and resuspended in 2 mL of buffer (1X PBS, 5 mM EDTA, 0.5 mM PMSF, 0.1% CHAPS) per absorbance unit measured previously (21). Resuspended cells were aliquoted in 0.5 mL portions, and combined appropriately. HA-FliF43-454 was present in both samples; one sample contained the GST-FlhE fusion construct, and the other contained GST only as a negative control. After mixing of cells, lysozyme (from a 50 mg/mL stock in 50% glycerol) and MgCl2 (from a 1M stock in water) were added to final concentrations of 0.926 mg/mL and 0.056 M, respectively. Samples were incubated on ice for 60 minutes, after which cells were subjected to lysis by sonication (Power 3, Duty Cycle 50%, 3 times, 50 sec each). Lysis was confirmed by phase-contrast microscopy to ensure that no phase-bright intact cells remained, and the samples were then pelleted (16,000 x g, 12 40 min, 4°C). After pelleting, the top 50 μL of each sample was removed and set aside (deemed “lysate”), and the remaining supernatant was transferred to fresh Eppendorf tubes. Glutathione-Sepharose 4B beads (150 μL of a 50% slurry, GE Healthcare) were added to each of the two supernatant samples, and allowed to incubate at room temperature for one hour with gentle rotation. Beads were then pelleted with a one minute microcentrifuge spin (room temperature) and washed once with 1X PBS buffer containing 5 mM EDTA and 0.5 mM PMSF. A 5 second spin in the microfuge pelleted the glutathione-Sepharose beads once more, and elution buffer (50 μL of 50 mM reduced glutathione, 50 mM Tris, pH 8.0) was allowed to incubate with the beads for ten minutes (at room temperature, with gentle rotation). Beads were then pelleted, and the supernatant of each sample (deemed “supernatant”, or “S/N”) was collected for analysis by Western blot as previously described (20). Both anti-HA and anti-GST primary antibodies were used in detecting HA-FliF43-454 and GST-FlhE, respectively, on different identical blots. Immunoblotting Relevant samples were loaded alongside a protein standard (PageRuler™ Plus Prestained Protein Ladder, Thermo Scientific) and resolved on 12% SDS-PAGE minigels (Bio-Rad MiniProtean system). Protein was transferred onto nitrocellulose membrane using a semidry transfer apparatus (Bio-Rad), washed, and incubated with primary antibody overnight at 4°C. Primary antibody stocks were diluted 1:1000 in 0.1% gelatin and 0.01% sodium azide and stored at 4°C. After overnight incubation primary antibody was decanted, and blots were washed and then incubated for at least one hour in secondary antibody with gentle shaking. Secondary antibody was added to a final dilution factor of 13 1:10,000; 1 μL of either α-rabbit (Li-Cor IRDye® Goat anti-rabbit; for α-GST primary antibody) or α-mouse (Li-Cor IRDye® Goat anti-mouse; for α-HA, α-His) secondary antibody was incubated in 10 mL of 5% milk in Tris-Buffered Saline (TBS) for at least one hour at room temperature (RT) or overnight (O/N) at 4°C. After washing with TBS, blots were visualized using the Li-Cor Odyssey blot development system. Peptidoglycan Purification Isolation of peptidoglycan was performed as previously described (22) with minor modifications. For this experiment, strain RP3098 lacking flhDC (and thus not expressing any flagellar genes) was grown overnight at 32°C with shaking (1L LB, inoculated from a 5-mL overnight culture of frozen RP3098 stock). Cells were then spun down (Sorvall RC5B Superspeed Centrifuge, GSA rotor; 4,000 x g, 10’, 10°C) and the pellet was washed with 40 mL of 10 mM Tris, pH 7.13. This process was repeated twice, and the final pellet was resuspended in 30 mL of 10 mM Tris buffer. This solution was then added dropwise to 300 mL of boiling 4% SDS and boiled for 60 minutes. The sample was then pelleted by ultracentrifugation (Beckman L5-50B ultracentrifuge, SW41 rotor; 100,000 x g, 80’, 20°C) and the pellet was resuspended in 150 mL of 2M NaCl. After vortexing, the sample was incubated overnight at room temperature with gentle rotation. The next day, the sample was spun down at 1,450 x g for 10 minutes and the supernatant was transferred to a new container. It underwent another ultracentrifuge spin (100,000 x g, 80’, 20°C) and the pellet was washed with 60 mL of double-distilled water and stored at -80°C overnight. The sample was ultracentrifuged and washed with water once more, and the pellet was then resuspended in 20 mL of buffer containing 0.1 mM MgCl2, 25 μg/mL DNAse I, 50 μg/mL 14 RNAse A, and 200 μg/mL α-amylase. This mixture was incubated at 37°C for 90 minutes, after which point Pronase was added to a final concentration of 200 μg/mL. The sample was then incubated at 60°C for one hour, for the end goal of inactivating the Pronase that had been added, but it was later discovered that this step was not harsh enough and residual protease activity remained in the peptidoglycan sample. After the incubation, SDS was added to a final concentration of 8% and the sample was boiled for 15 minutes. Sample was then ultracentrifuged (100,000 x g, 80’, 20°C), supernatant was decanted, and grayish, firm pellet was resuspended in 20 mL of 60°C water. After this spin was repeated and grayish-white pellet was resuspended in 60°C water, the sample was spun down once more (100,000 x g, 80’, 20°C) and resuspended in 2 mL of buffer (25 mM HEPES, 50 mM NaCl, pH 7.51 with KOH). Peptidoglycan FlhE Pulldown Assay Purified FlhE(His)6 and peptidoglycan (PG) were combined in a 100 μL reaction volume and allowed to incubate at room temperature for one hour. Prior to combining, the purified FlhE necessary for that day’s experiment was centrifuged (Madell Technologies TG16A-WS, rotor 3; 9,000 x g, 30’, 4°C) to “clarify” the FlhE sample. Due to the observation that FlhE had a tendency to aggregate and spin out of solution by itself, this step was added to reduce the amount of aggregated protein used in the experiment. After purified FlhE(His)6 was centrifuged, only its supernatant was used as the protein sample to which 1% Na-deoxycholate was added to further discourage aggregation. FlhE and PG in the same buffer (25 mM HEPES, 50 mM NaCl, 1 mM DTT, pH 7.81) were combined in 100 μL reaction volumes. After one hour of incubation at room temperature, 30 μL of 15 sample was removed and set aside (deemed the “unspun” sample). After centrifugation (9,000 x g, 30’, 4°C) the top 30 μL of the supernatant was removed and set aside (deemed “supernatant”, or “S/N”). To maintain the original concentrations of protein/PG across all three samples collected, another 30 μL of sample was removed and the pellet was resuspended with 60 μL of buffer; this was then deemed the “pellet” sample. From there, samples were analyzed via Ponceau Staining and sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) followed by either Coomassie staining (to ascertain all protein products present in the gel) or Western blotting (with α-His monoclonal antibody to monitor the amount of FlhE(His)6 in the gel). Ponceau S Staining To nonspecifically detect the presence of protein in a given sample, Ponceau S staining solution was used on nitrocellulose membranes. This assay is able to detect microgram quantities of protein on a nitrocellulose membrane. A negatively charged stain, Ponceau S binds the positively charged amino groups on proteins and binds non-covalently to nonpolar areas of the protein. The stain was prepared as previously described (23), with 0.1% (w/v) Ponceau S in 1% (v/v) acetic acid. The nitrocellulose membrane was labeled and spotted with 2 μL of samples, and allowed to dry on the bench at room temperature for several minutes. The membrane was then immersed in the Ponceau S staining solution for five minutes at room temperature. After safely disposing of the stain, the membrane was washed twice with 5% (v/v) acetic acid and twice with distilled water, for five minutes each time. Because Ponceau S is very toxic, all solutions containing traces of it were 16 collected in a glass bottle for disposal and passed off to the Occupational and Environmental Health and Safety Department of the University of Utah. Coomassie Staining To examine the total protein content of various samples, gels were stained with Coomassie Brilliant Blue. Coomassie blue is a non-specific protein dye and therefore useful in estimating the presence of a variety of proteins. A twelve percent SDS-PAGE gel was run and used directly by prying apart the glass plates, cutting off the stacking gel (containing the now-empty wells the sample was loaded into), and putting the entire membrane in a container with enough Destain I (50% ethanol, 7% acetic acid) to cover the gel. The container was then covered with saran wrap and vented at one corner before being microwaved for one minute at full power. This was allowed to cool for several minutes at room temperature, after which point Destain I was decanted and replaced with Destain II (5%, 7.5% acetic acid). A few mL of Coomassie R-250 (3.2% w/v in water) was added into the container, which was microwaved once more for 1’ and allowed again to sit at RT for several minutes. Destain II and Coomassie blue were then decanted and replaced with fresh Destain II. The gel was then left to destain overnight with gentle shaking, and imaged the following day after a 30’ soak in ddH2O by using a printer scanner (gel placed on top of a layer of saran wrap to protect scanner). 17 RESULTS When FlhE is deleted from the bacterial chromosome in Escherichia coli and Salmonella enterica, several cellular defects are observed. ΔflhE mutants exhibit severely reduced rates of swimming and swarming motility, slower growth rates, and abnormal morphologies. These effects, especially the morphological defect, is observed more strongly in E. coli than in S. enterica. In soft LB agar (0.27%), E. coli ΔflhE mutants swim a fraction of the rate of wild-type (WT) cells (Fig. 3) – a defect that is easily complementable when flhE is expressed from a plasmid (pflhE) even at a basal level of expression (i.e. not induced). Across all induction levels tested, ΔflhE mutants swam at a rate of 20±2% that of wild-type cells and complemented knockouts (pflhE/ΔflhE) swam at a rate of 78±3% that of WT. Examining the average velocities measured (change in diameter over time) across all induction levels, WT cells outpaced ΔflhE cells by 3.2±0.1 mm/hr and pflhE/ΔflhE cells by only 0.9±0.1 mm/hr. Furthermore, the deletion of flhE in S. enterica completely abrogates the surface-associated motility known as “swarming”. Interestingly enough, this defect is greatly relieved by adding a surfactant (such as TWEEN 80) to the surface of the swarm plates which increases wettability and decreases surface tension (16). The swarming defect is also complementable, meaning that swarming function is restored when flhE is resupplied via plasmid to a cell lacking chromosomal flhE (termed ΔflhE; see Fig. 4). If the first 17 amino acids of the sequence are deleted, thereby removing the signal sequence targeting FlhE to the periplasm, the cells are again incapable of swarming. 18 (b) Diameter (mm) (a) 25 20 15 10 5 0 2.0 3.0 4.0 5.0 Time Elapsed (hours) C/WT AV E/ΔE AV 6.0 C/ΔE AV FIG. 3. ΔflhE Mutants are Complementable. (a) Deletion of chromosomal flhE results in a near abrogation of swimming motility. Supplying flhE on a plasmid, even at a basal level of expression (no added inducer), complements the deletion fairly well. The pflhE/ΔflhE colony swims at a rate closer to that of wild-type cells (control plasmid in RP437). Image shown is of plate spotted from overnight cultures in triplicate and allowed to grow for 5 hours at 32°C. (b) A quantification of this effect. The diameter of each colony is measured over time and plotted here. This data represents the average of three biological replicates of a control plasmid (conferring only antibiotic resistance) in wild-type cells (C/WT), a control plasmid in cells lacking chromosomal flhE (C/ΔE), and plasmid-borne flhE transformed into ΔflhE cells (E/ΔE). The 0.27% swim agar contained 20 μM NaSalicylate to induce expression, but results shown here are representative of the effect seen across all induction levels. Plates were incubated for over 6 hours at 32°C. Con/RP437 Con/ΔflhE pFlhE/ΔflhE pFlhEΔ1-17/ΔflhE FIG. 4. ΔflhE Mutants are Severely Defective in Swarming, but complementable. Plates were incubated at 32°C for 24 hours. A control plasmid bearing only antibiotic resistance (Con) was transformed into both wild-type (RP437) E. coli cells and those lacking chromosomal flhE (ΔflhE). The same plasmid vector, bearing either a full-length or an N-terminally truncated version of flhE, was transformed into the ΔflhE strain to test for complementation. Only the full-length version of flhE complemented the swarming defect; without a signal sequence (amino acids 1-17) targeting FlhE to the periplasm, cells swarm as if they still lacked a copy of flhE. 19 Most notably, E. coli cells that lack flhE take on rather whimsical morphologies – many different shapes are observable in a liquid culture of the ΔflhE cells, from “blobs” to “U-shapes” to corkscrew-like cells (Fig. 5). Nearly all of these bizarre looking cells are capable of swimming, interestingly enough. Though obviously deformed, they still have the ability to careen through liquid media and are fascinating to watch. The morphology defect, seen comparatively infrequently in S. enterica ΔflhE mutants, becomes more pronounced when cells are subjected to osmotic stress in the form of increased NaCl concentration. It is worth noting that, under high salt conditions, water moves out of the cell and decreases internal turgor pressure. This effect prevents lysis in cells whose turgor pressure threatens to burst through any weak points in the cell wall. This may explain why distorted morphologies are not abundant in ΔflhE Salmonella cells, but cell lysis is. FIG. 5. ΔflhE Mutants Display Aberrant Morphology. Deletion of chromosomal flhE results in contorted, abnormal cell morphology in Escherichia coli. Panels i and ii identify and outline commonly observed morphologies. While ΔflhE cells are evidently distorted, most are still capable of motility. Panel iii shows wild-type cells (RP437) under the same conditions, by means of comparison. Figure prepared by Griffin Chure. 20 While the structure of FlhE has been solved by the Harshey group (13), its precise function is still unknown. FlhE is bad for cells in high amounts; cells forced to overexpress FlhE suffer from a severely hampered growth rate (Fig. 6). In its absence, however, cells experience a defect both in growth and swimming motility. Several point mutations (W22D, L31D, and R34D) in FlhE in Escherichia coli have been shown to be critical to the function of the protein in swimming motility assays (Fig. 7). Cells expressing FlhE with these mutations behave as though they had been given no flhE at all, though resupplying a ΔflhE strain with wild-type flhE borne on a plasmid provides near complete complementation (at low induction levels) in swimming and growth assays. In liquid culture, cells lacking chromosomal flhE suffer from an initial growth lag followed by slower overall growth. Both this growth lag and the morphological disturbance inherent to ΔflhE cells are cured by the overexpression of flgM from a plasmid (Fig. 8). flgM encodes for an anti-σ28 factor that is responsible for suppressing late-stage flagellar genes (including those encoding the hook-filament junction, filament, chemotaxis proteins and motor torque generators) (24), which suggests that the defects seen in ΔflhE cells require the expression of one or more of these protein products. A straightforward possibility is that the premature building of the filament in the periplasm in ΔflhE mutants is responsible for the damage and deformation to the cell envelope that results in aberrant morphologies. 21 FIG. 6. Overexpression of FlhE Halts Growth in E. coli Cells. While cells lacking FlhE grow more slowly than WT or ΔflhDC cells, an overexpression of FlhE harms growth as well. Supplying flhE via a plasmid and increasing inducer concentration results in more severe growth defects, even in a ΔflhDC strain (not pictured). Cells were allowed to grow in LB and appropriate antibiotic at 32°C for 9 hours, and induced with 0 μM, 100 μM, or 500 μM IPTG. Data to quantify growth rate is measured by the optical density of the sample at 600 nm. Figure prepared by Griffin Chure. FIG. 7. Several Point Mutations Alter Swimming Motility. Of many candidates tested, three point mutations in particular appear to affect the function of FlhE. When W22D, L31D, or R34D mutations are introduced into the sequence of flhE cells swim much more slowly; this effect is more pronounced if two of these point mutations are present.. Plates above were spotted from overnight culture onto plates containing no inducer and incubated at 32°C for 5 hours. 22 FIG. 8. Overexpression of FlgM Cures Growth Defect of ΔE. Overexpression of FlgM represses late-stage flagellar genes, like those responsible for the hook, rod, and filament. Overexpression of flgM from a plasmid rescues the growth defect observed in ΔflhE as measured by absorbance readings at 600 nm. Wild-type growth rate over time, under the same conditions, is shown for comparison. Figure prepared by Griffin Chure. To study the effects of a chromosomal flhE deletion on the morphology of Salmonella cells, an experiment was conducted in which ΔflhE cells were subjected to growth at increasing levels of osmolarity. Cell lysis can be prevented in liquid media of high osmolarity (in this case, tryptone broth [TB] containing additional NaCl), allowing deformed ΔflhE cells to be observed when they would otherwise lyse (15). As the osmolarity of the liquid medium was raised (in TB, from 0 M – 0.3 M additional NaCl), aberrant cell morphologies became increasingly apparent. Increasing the osmolarity of the surrounding solution did cause some wild-type Salmonella cells to deform, though at a much lower frequency than the ΔflhE cells. Because cells shut down the building of late- 23 stage flagellar genes when subject to high osmotic stress, master flagellar regulator flhDC was transformed into both wild-type and ΔflhE cells to ensure that flagella were assembled under all conditions. Increasing the induction of the flhDC-bearing plasmid, the total time incubated with high-salt medium, and the concentration of additional NaCl resulted in the observation of motile, morphologically disturbed cells (Fig. 9). The lack of documentation in the literature of morphologically “sick” ΔflhE Salmonella cells reflects the need to both increase the osmolarity of the medium and provide cells with flhDC to observe an effect. FIG. 9. Aberrant Morphologies Arise in Salmonella ΔflhE Mutants When Under High Osmotic Stress. When subjected to osmotic stress in the form of added NaCl (either 0, 0.22, or 0.3 M in excess of normal [NaCl]), Salmonella cells begin to show more morphological disturbance. Cells were transformed with pflhDC to promote the expression of flagellar genes, as cells have the tendency to shut down flagellar synthesis when under duress. While the wild-type (WT) Salmonella also begins to exhibit deformities at high salt concentrations, the ΔflhE strain becomes markedly more aberrant and begins to resemble its E. coli equivalent. Figure prepared by Griffin Chure. 24 In ΔflhE mutants in both E. coli and Salmonella, the presence of “sausage link” looking cells led us to investigate whether FlhE has a role in directing flagellar assembly away from the cell’s poles. In some species, genes flhF and flhG are found upstream of flhE (16); in Vibrio alginolyticus FlhF and FlhG are implicated in the regulation of its mono-polar-flagellum (25) . If these homologs are responsible for directing assembly away from the poles, perhaps the absence of flhE could allow flagella to be built at the cell division plane. The filament would be exported, therefore, into the daughter cell and cause the two to be linked together. To test this hypothesis, growing cells were treated with 10 μg/mL Cephalexin and allowed to incubate for two hours. Cephalexin is an antibiotic that prevents cell division and results in long filamentous cells (26). If the morphology defect in ΔflhE cells is due to the building of flagella at the poles, then cephalexin should, by preventing the formation of septa, obviate the defect. ΔflhE cells were compared to a double mutant lacking both flhE and fliG, allowing us to control when the cells built flagella. Without FliG, the switch complex (composed also of proteins FliM and FliN) cannot be completed. The results of the experiment demonstrate that FlhE is likely not involved in directly flagellar assembly away from the poles; ΔflhE cells exposed to Cephalexin still exhibit deformity in the form of long, kinked cells (Fig. 10). Double ΔflhEΔfliG mutants appear like wild-type cells before addition of Cephalexin, and, like the wild type, become long and serpentine after incubation with the antibiotic. 25 FIG. 10. Flagella are not built at cell poles in ΔE Mutants. Addition of Cephalexin, an antibiotic that prevents cell division, sheds light on the flagellation patterns in ΔflhE cells. After incubation with cephalexin, ΔflhE cells become visibly “kinked”, perhaps as a result of peritrichous flagella being built defectively as the cells attempt to divide. Cells lacking fliG, which contributes to the assembly of the switch complex, are not able to build flagellar motors and therefore should appear merely long, not deformed. In the search to find a binding partner for FlhE, pulldown assays, fluorescence spectrophotometry, and the Bacterial II Hybrid System (BACTH) have thus far been employed. In these experiments, point mutants known to have phenotypes were used to test the functional relevance of any interactions demonstrated or suspected. Swim motility assays suggest that FlhE has at least three critical residues that, when mutated and supplied to ΔflhE cells via a plasmid, result in a swimming phenotype closest to that of ΔflhE cells (Fig. 6). These three point mutations (W22D, L31D, and R34D), especially when combined, yield a non-functional version of the protein that is unable to complement an 26 flhE chromosomal deletion. Of several protein candidates assayed, only the periplasmic region of FliF (residues 43-454) suggested a positive interaction per GST pulldown assays, and efforts to reproduce the experiment are currently underway. The three critical point mutations did not appear to disrupt this interaction however, which suggests that FlhE may have another binding partner besides FliF. Once it was surmised that FlhE could be binding to the MS ring and likely has at least one other binding partner, several experiments were conducted to determine what the other entity might be. Because the MS ring is embedded in the cytoplasmic membrane and very near the peptidoglycan layer of the cell, a series of pulldown assays were conducted to determine if peptidoglycan could be the unknown binding partner. In this model, FlhE would be responsible for transiently pushing the MS ring against the peptidoglycan layer of E. coli (thus also pushing the rod against the outer membrane) long enough for the proper assembly and export of late-stage flagellar proteins (Fig. 11). These include the proteins of the hook (fliK), rod, and filament, accompanied by the recruitment of the lipopolysaccharide (L-) and peptidoglycan (P-) rings. These rings are named for the layer they are embedded in and encoded for by flgH and flgI, respectively (5). In the absence of the hypothesized tethering interaction of FlhE, assembly of these normally-exterior structures might occur in the periplasm and lead to cell defects. 27 FIG. 11. A proposed mechanistic model of FlhE function. In this model, FlhE is responsible for attaching the basal body to the peptidoglycan (PG) layer in a transient interaction that pushes the rod of the motor against the outer membrane of the cell. This would allow for the proper recruitment and export of late-stage flagellar elements; in the absence of FlhE the flagellum may continue assembly in the periplasmic space. Figure prepared by Griffin Chure. To test for a possible interaction between FlhE and the peptidoglycan layer of the cell, a simple pulldown assay was performed. Peptidoglycan was isolated from Escherichia coli through a process that used boiling, a barrage of different enzymes to chew up DNA, RNA, and any associated proteins that remained, and separation by high-speed ultracentrifugation. Purified peptidoglycan was added in vitro to purified FlhE protein and allowed to incubate, after which point centrifugation would – in theory – pull down the 28 high-MW peptidoglycan and any protein associated with it. We ran into several problems with this series of experiments; FlhE, for example, appeared to self-aggregate and pellet spontaneously. To discourage FlhE-FlhE aggregation several detergents were tested, of which 1% Na-deoxycholate was found to be sufficient to dissociate E-E aggregates. When purified peptidoglycan was added to a sample of purified FlhE-His, the FlhE signal decreased in intensity (as measured either by a monoclonal α-His antibody or a Coomassie R-250 stain). This was surmised to be the result of a peptidoglycan-FlhE interaction, in which the peptidoglycan pulled down soluble FlhE out of solution (Fig. 11). 250 kDa 15 kDa Pellet S/N PG Only Unsp. Pellet S/N FlhE + PG Unsp. Pellet S/N Unsp. Marker FlhE Only PG FlhE 10 kDa FIG. 11. Presence of peptidoglycan greatly reduces FlhE signal in supernatant. After combining FlhE and PG together (as well as controls containing one or the other), samples were centrifuged and both the supernatant (S/N) and pellet samples were analyzed alongside their pre-spin (Unsp.) counterparts. The signal of FlhE is greatly reduced when combined with PG, especially in the supernatant post-spin. To release the FlhE in the peptidoglycan sacculi (or fragments thereof), samples were treated with lysozyme and run on a gel. A band appeared at the expected size (appx. 14 kDa for FlhE-His) but turned out to be the lysozyme (also appx. 14 kDa and α-Hisdetectable) that samples were treated with. After further investigation, it was found that the 29 purified peptidoglycan sample had residual (Pronase) protease activity that should have been eliminated during the purification process in a rigorous boiling step. This explains why the FlhE signal decreased in intensity when combined with PG, but a BSA sample exposed to the same conditions remained relatively unchanged in most experiments. Further experiments were conducted using a broad-spectrum protease inhibitor (Roche) to suppress any residual protease activity, but have thus far remained inconclusive. Another angle was taken to explore a possible FlhE-PG interaction; fluorescence spectrophotometry was employed to monitor any changes in signal upon mixing of FlhE and PG. Wildtype FlhE has three tryptophan residues, which contribute to a measureable emission spectrum. Tryptophan (abbr. W) has a maximum absorbance at 280 nm and has an emission spectrum dependent on the local environment (27). A shift in the emission spectrum of FlhE (either a change in maximum wavelength and/or intensity) upon addition of purified peptidoglycan fragments may indicate an interaction between the two. Unfortunately, results of this experiment were again inconclusive. After FlhE-PG interaction experiments were abandoned, we began exploring a possible interaction between FlhE and the L- and P-rings. Encoded for by flgH and flgI, respectively, the Land P-rings are embedded in the lipopolysaccharide (outer membrane, OM) and the peptidoglycan (PG) layers of the cell envelope (5). If FlhE is responsible as a type of chaperone to ensure for the proper recruitment of these rings, its absence could result in the building of late-stage flagellar elements before they are properly anchored in the PG and OM. Attempts to increase the likelihood of proper L- and P-ring assembly by overexpression of flgHI were unsuccessful; overexpressing the proteins hurt wild-type cells and all but killed ΔflhE cells (data not shown). 30 Another assay was then employed to explore possible interaction between FlhE and FlgI in a way that doesn’t result in cell death. The Bacterial II Hybrid System (BACTH) is a semi-sensitive assay for protein interaction. In essence, two proteins surmised to potentially interact are cloned into two of four possible vectors. These vectors, pKT25, pKNT25, pUT18, and pUT18c each contain half of the enzyme adenylate cyclase, which restores the synthesis of cAMP when reconstituted. This upregulates, among other things, the maltose and lactose catabolic operons and thus the ability to metabolize both sugars (28). If two proteins do indeed interact, and come close enough to each other to do so, the theory is that the enzyme halves will be close enough to re-form and produce a purple pigment on MacConkey agar plates. Tests ran with FlgI (responsible for the P-ring that penetrates the peptidoglycan layer in the flagellar motor) has yielded no positive results as of yet, though tentative evidence of FlhE-FlhE interaction was observed. This makes some sense, as we have seen in several different assays that FlhE has a tendency to selfaggregate. BACTH will be employed next to ascertain the existence of an interaction between FlhE and the periplasmic domain of FliF (amino acids 43-454). If any positive interactions in the BACTH system are discovered, the next step will be to test the same interactions using FlhE with one (or two) of the critical point mutations found in swim plate studies. 31 DISCUSSION The deletion of flhE from the bacterial chromosome in Escherichia coli and Salmonella enterica causes numerous cellular defects. These defects – including swimming and swarming ability as well as morphology – are relieved fairly well in E. coli when plasmid-borne flhE (pflhE) is supplied to ΔflhE cells even at a basal level of expression. At low levels of induction, pflhE/ΔflhE shows near complete complementation on swim plates, indicating that FlhE is likely present in low copy number as only the “leakiness” of the plasmid’s expression is leading to sufficient FlhE expression. Moreover, the defects in swimming and swarming remain when an N-terminally truncated version of the protein (lacking the first 17 residues which function as a signal sequence). This indicates that not only does FlhE need to be present in the cell, but it must be periplasmically targeted to carry out its function. If flhE is overexpressed, however, cells experience severely hampered growth rates meaning that the overabundance of the protein is somehow toxic to cells. Both the morphology and growth defects of ΔflhE cells are relieved upon the overexpression of protein FlgM. This anti-σ28 factor is responsible for binding and repressing late-stage genes until the hook-basal body structure is properly assembled, at which point it is secreted out of the cell and allows for assembly of the flagellum to complete. When overexpressed, FlgM suppresses the expression of late flagellar genes including the hook-filament junction, the filament, and genes necessary for motility and chemotaxis. These cells are therefore inherently nonmotile, but capable of other cell processes like growth and division. The fact that FlgM overexpression relieves the defects in ΔflhE cells suggests that at least one of these late-stage components is 32 involved in the abnormalities observed, from motility to morphology. We propose that the severe morphological defects could be the result of the improper building of the filament in the periplasm. This structure, comprised of roughly 30,000 FliC subunits and up to 15 μm long, could cause catastrophe if exported into an already-crowded periplasmic space. What, then, could FlhE be doing to prevent this phenomenon? We believe that FlhE plays a role in keeping the basal body pressed up against the peptidoglycan layer (and, by extension, the outer membrane) long enough for proper assembly of the hook, L- and P-rings, and filament. FlhE likely binds to some component of the basal body (perhaps the MS ring) and attaches noncovalently to another structure like the peptidoglycan layer. In its absence the basal body would be allowed to hover beneath the peptidoglycan layer, sometimes building the hook, rings, and filament at the right time and sometimes building them prematurely. If the flexible inner membrane (IM) that houses the MS ring and export apparatus is too far from the peptidoglycan layer when the cell begins the next steps of assembly, then the rod, hook, and filament will be built in the periplasm. This would result in the distortion of the cell envelope and thus the fascinatingly aberrant cell morphologies observed in a ΔflhE strain. Because E. coli and S. enterica are peritrichously flagellated and possess many flagella dispersed across their surfaces, there is the opportunity for motility if cells manage to export their filaments properly. Again, this could occur if assembly begins while the basal body is near enough the PG and OM for assembly to proceed smoothly. This phenomenon would explain why most cells lacking flhE are morphologically disturbed and yet still motile. 33 It is worth noting that the absence of FlhE does not appear to be nearly as harmful to Salmonella cells as it is to E. coli. The deletion of chromosomal flhE in Salmonella results in an abrogation of swarming motility (again complemented by pflhE but not pflhEΔ1-17) and a decrease in swimming motility. Moreover, Salmonella ΔflhE cells show little morphological disturbance especially when compared to their E. coli counterparts. The Harshey group does not report aberrant morphologies of their cells (14), which is not surprising seeing as they studied flhE in the context of Salmonella enterica. Distorted morphology is not observed in ΔflhE Salmonella mutants nearly as frequently as it is in E. coli. This could be due to the fact that Salmonella cells are smaller than E. coli (generally 0.5 μm wide by 2 μm long in Salmonella (29) vs. 1-2 μm wide by 2-5 μm long in E. coli (30)). The osmotic pressure of a whole cell will vary inversely with its volume, albeit only under the assumption that both species have roughly the same concentrations of solutes in the periplasm and cytoplasm (31). If the osmotic pressure in the smaller Salmonella cells is higher than that in E. coli, the basal body might be forced nearer the peptidoglycan layer by default regardless of the presence of FlhE. This would result in a higher frequency of “normal” cells which were able to properly build and export late-stage flagellar proteins. Conversely, with higher osmotic pressure comes the increased likelihood of cell lysis given stress to the cell envelope; this would result in the DNA shedding observed in ΔflhE mutants reported by Lee et al, 2010. Their data appears to be consistent with our model; DNA shedding, for example, was greatly reduced in their study if genes encoding the hook or filament were deleted. Attempts to explore an interaction between FlhE and peptidoglycan have thus far been unsuccessful. The peptidoglycan pulldown experiment looked promising because the 34 signal of FlhE was greatly reduced upon addition of peptidoglycan, but it was later found to be partly explainable by residual protease activity in the purified PG sample. Though rigorous boiling steps during the purification process should have inactivated the Pronase, experiments with BSA and ovalbumin indicated that the purified PG sample was chewing up protein. This effect was shown to be greatly reduced upon the addition of a broad-spectrum protease inhibitor (Roche), but further experiments with proteaseinhibitor-treated PG have not yet been executed. Moreover, FlhE appeared to have the tendency to aggregate and pellet by itself. This phenomenon was relieved upon the addition of small amounts of detergent; 1% sodium deoxycholate was shown to be sufficient to this end. Fluorescence spectrophotometry was used as another approach to monitor the same interaction; if FlhE interacts with PG, its emission spectrum should change either in wavelength or intensity due to the intrinsic fluorescence of FlhE’s three tryptophan residues. Unfortunately, results of this experiment again proved inconclusive. To conserve sample, a narrow cylindrical cuvette was used and held in place by a specially machined metal vessel with windows to allow light to pass through. This method proved to be too sensitive; mere rotation of the cuvette within the holder affected the spectra measured and scans of the same sample could be inconsistent, changing intensities sometimes and flatlining others. With the simple expedient of a more suitable cuvette, experiments using fluorescence spectrophotometry will likely provide an avenue to test for the hypothesized interaction between FlhE and PG. Current study aims to further explore the interaction of FlhE with other binding partners. The existence of point mutations that can knock out function suggests that these critical residues likely play a part in the function of FlhE and are potential binding sites for 35 interaction. Experiments are underway to explore interaction between FlhE and FliF (whose subunits constitute the MS-ring) using either GST pulldown assays or the bacterial two-hybrid system (BACTH) in E. coli. Regardless of what entity it interacts with, FlhE is absolutely essential to normal cell motility and morphology. Its deletion culminates in the abrogation of swarming motility, severe reduction in swimming motility, and morphological aberrancy, especially in E. coli. These phenomena are relieved when cells are supplied with flhE from a plasmid, and appear to be dependent on the building of late-stage flagellar elements. Therefore, we propose that FlhE plays a critical role in the proper assembly of the bacterial flagellar motor by preventing premature export of external flagellar components. 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