This work was supported by National Institutes of Health grant GM118085 and in part by the Robert Welch Foundation (grant F-1811 to R.M.H.).

1. Isolation of suppressor mutants in motility-deficient strains
NOTE: Use this method as a broad &#8216;catch-all&#8217; to identify the general nature of the motility defect.
<ol>
	<li>Soft-agar plate preparation 
	NOTE: Soft-agar, also referred to as motility- or swim-agar, is a low percentage agar (~0.2-0.35% w/v), long used to assay chemotaxis31,32.
	<ol>
		<li>Add 3 g of bacto-agar (0.3% w/v) and 20 g of LB to a 2 L round bottom flask. Add 1 L of ddH2O (double-distilled water) to the flask and evenly mix the suspension using a stir rod and magnetic stirring plate.</li>
		<li>Autoclave for 20 min at 121 &#176;C.</li>
		<li>Allow to cool with gentle agitation using the rod/plate as above. When the temperature reaches approximately 50 &#176;C, pour 25 mL into sterile Petri dishes (100 mm x 15 mm), and allow the molten agar to set with lid in place for at least 1 h, for use within 16 h.</li>
	</ol>
	</li>
	<li>Culture preparation, inoculation, and isolation of suppressor mutants 
	NOTE: E. coli inoculated in the center of nutrient rich media solidified with soft agar consume nutrients locally, creating a nutrient gradient that they follow. As they move outward, defined &#8216;rings&#8217; appear (Figure 1A), which are related to specific chemoattractants the bacteria respond to. Defects in either the chemotaxis system or structural components of the flagella motor can compromise performance in this assay. Often, mutants with a motility advantage arise during screening, and can be seen emerging from single or from multiple points along the periphery of the ring, from where they &#8216;flare&#8217; out (Figure 1C). One will notice that the outermost edge of the swimming front contrasts readily against the uncolonized virgin soft agar.
	<ol>
		<li>Grow overnight cultures of the desired motility-deficient strain in 5 mL of Lennox Broth (LB; 10 g/L tryptone, 5 g/L yeast extract, and 5 g/L NaCl, Table of Materials) at 30 &#176;C with horizontal shaking (220 r.p.m.). The following day, sub-culture (1:100 dilution) in fresh LB, growing under the same conditions to exponential phase (OD600 of 0.6).</li>
		<li>Inoculate 6 &#181;L of the culture into the center of a soft-agar plate (1.1) using a pipette, pushing the loaded sterile tip into the agar to gently expel the contents. Transfer to 30 &#176;C and incubate (Figure 1B), until motility &#8216;flares&#8217; are evident, emanating from the inoculation point or the periphery of the motility rings, typically in 24-36 h (Figure 1C). 
		NOTE: In motility assays, inoculate a wild-type strain alongside the mutant isolates for comparison. This wild-type strain will show the characteristic concentric chemotactic rings (Figure 1A) and fill the plate within 8-10 h.</li>
		<li>Use a sterile wire-loop to lift the cells from the &#8216;flare&#8217; region and streak to purify single colonies onto an LB hard agar plate (LB prepared as above, solidified with 15 g/L bacto-agar).</li>
		<li>Pick single colonies from the streak plate using a sterile wire loop and re-purify by streaking for single colonies to ensure isolation of a &#8216;pure&#8217; colony isolate.</li>
	</ol>
	</li>
	<li>Confirmation, and characterization of suppressor mutants
	<ol>
		<li>Confirm that the isolated suppressor mutant(s) have restored motility. Prepare soft-agar plates (1.1), and cultures for strains of interest (as in 1.2.1), including wild-type and the starting &#8216;motility-deficient&#8217; strain for comparison.</li>
		<li>Inoculate the plates (as in 1.2.2) and incubate at 30 &#176;C for 8-10 h.</li>
		<li>Record the diameter of the outermost ring (edge of the circle) and compare to establish which of the isolates have substantially restored motility. 
		NOTE: It is recommended that plates be photographed throughout the time-course of the experiment. For best results, use a &#8220;bucket of light&#8221; device30, where a digital camera is mounted above a light source for better illumination to measure the diameter of the swim colony and distinguish it from uncolonized agar.</li>
		<li>Subject verified isolates to WGS as required, allowing for sufficient &#8216;sequence coverage&#8217; to positively identify the mutations that restored wild-type function.</li>
	</ol>
	</li>
</ol>
2. Quantifying flagella motor behavior via cell tethering
NOTE: Use this method when normal run-tumble behavior (chemotaxis) appears to be compromised.
<ol>
	<li>Culture preparation and flagella shearing
	<ol>
		<li>Prepare an exponential phase culture of the strain of interest as described in step 1.2.1.</li>
		<li>Pellet 10 mL of cells by centrifugation at 2,000 x g for 3 min before resuspending in 10 mL of filter-sterilized Motility Buffer (MB; 10 mM potassium phosphate buffer [0.0935 M K2HPO4, 0.0065 M KH2PO4, pH 7.0], 0.1 mM EDTA [pH 7.0], 10 mM NaCl, 75 mM KCl). 
		NOTE: MB supports motility, but does not support bacterial growth</li>
		<li>Repeat step 2.1.2 two more times before resuspending the final pellet in 1 mL of MB.</li>
		<li>Transfer the cell suspension into a 1 mL syringe and attach a 23G needle to the end. Assemble an identical syringe/needle apparatus and attach the two together via 6 inches of polyethylene tubing (inner diameter of 0.58 mm) tightly sheathed over each needle tip.</li>
		<li>Shear the flagella (they are fragile and break easily) by gently passing the cell suspension back and forth from one syringe to the other 50x, with 1 min pauses between every 10 passes.</li>
		<li>Centrifuge the sheared cells at 2,000 x g for 3 min and resuspend in a final volume of 500 &#181;L of MB.</li>
	</ol>
	</li>
	<li>Slide preparation and cell tethering
	<ol>
		<li>Prepare a cell fixation chamber by stacking an 18 mm x 18 mm coverslip over a 3 inch x 1 inch x 1 mm glass microscope slide, separated by double-sided tape (Figure S1).</li>
		<li>Flush the chamber with 0.01% (w/v) poly-lysine solution by applying to the top of the chamber. Tilt the bottom edge (the coverslip flush with the microscope slide) onto a task wiper (tissue paper) to help draw solution through the chamber (Figure S1). Then incubate at room temperature for 10 min.</li>
		<li>Wash the chamber three times with 40 &#181;L of MB using as described in step 2.2.2</li>
		<li>Add 40 &#181;L of the sheared cell suspension (prepared above) to the top of the chamber and incubate at room temperature for 10 min to allow cells to attach to the coverslip.</li>
		<li>Gently flush the chamber with 40 &#181;L of MB as in 2.2.3 to remove unattached cells.</li>
	</ol>
	</li>
	<li>Cell rotation recording and quantification
	<ol>
		<li>Transfer the microscope slide loaded with tethered cells to the microscope stage.</li>
		<li>Using phase-contrast microscopy and a 100x objective, scan the population for cells that are fixed in place, and rotating on a single axis, i.e., smooth rotations on a fixed point rather than presenting at an angle wherein the cell moves in and out of focus (Video 1).</li>
		<li>Use a commercial microscope and associated camera. Open the associated software, ensure cells of interest are in focus and click video acquisition to record the cell rotation for one minute (at 10 frames per second or higher).</li>
		<li>From video playback, quantify the number of complete rotations per minute and the number of times the cell changes direction (switching frequency). 
		NOTE: Rotational speeds and switching frequency may be too fast to gauge by eye so it is recommended to use video software that offers slow/fine playback, or adopts an automated software system to quantify rotational patterns33. An alternative would be to increase the viscosity of the MB using methyl cellulose (or similar agent) to help slow and resolve the rotation of faster bacteria or compensate when cameras with low framerates are in use.</li>
		<li>Repeat step 2.3.2 with biological replicates to compile a representation of the population of interest.</li>
	</ol>
	</li>
</ol>
3. Preparation of swarms in a border-crossing assay
NOTE: Use this method for assessing the impact of a mutation or condition on group motility. Swarm-agar refers to agar where the percentage is typically higher than that of soft-agar. In soft agar (0.3 %), cells swim individually inside the agar. In swarm agar (0.5% and above), cells move as a group on the surface. While swarm plates must be used as detailed here, swim plates have a longer shelf life, and may be used for several days. Our personal preference is to use in 1-2 days.
<ol>
	<li>Preparation of swarm agar
	<ol>
		<li>Add 5 g of Eiken agar (0.5% w/v) and 20 g of LB to a 2 L round bottom flask. Add 1 L of ddH2O to the flask and evenly mix the suspension using a stir rod and magnetic stirring plate.</li>
		<li>Autoclave for 20 min at 121 &#176;C.</li>
		<li>Allow to cool with gentle agitation to avoid any air bubbles using the rod/plate as above. When approximately 50 &#176;C, add filter-sterilized glucose for a final concentration of 0.5 %.</li>
		<li>Pour 25 mL into sterile Petri dishes (100 mm x 15 mm) and allow to set at room temperature for at least 14 h and no more than 20 h. Do not store for future use.</li>
	</ol>
	</li>
	<li>Inoculation and incubation of swarm plates
	<ol>
		<li>Inoculate 6 &#181;L of a mid-exponential culture (prepared as in 1.2) by spotting on top of the agar.</li>
		<li>Leave the lid off for 5-10 min and replace when the inoculum has dried into the agar surface.</li>
		<li>Incubate at 30 &#176;C for 8 h. Avoid the temptation to inspect swarm progress by removing the lid, as this will contribute to drying of the agar and impair swarming. 
		NOTE: Incubation time may vary depending on the strain phenotype. Some isolated mutations may hamper swarming ability and will require a reduced percentage of agar or a more prolonged period of incubation.</li>
	</ol>
	</li>
	<li>Preparation of border-crossing assay plates 
	NOTE: This assay utilizes a modified Petri dish, where a plastic divide (border) creates two chambers, rather than one (Figure 2A). Each chamber can be prepared independently of the other, offering differing conditions for swarming, prior to &#8216;connecting&#8217; the two. Depending on experimental design, the first chamber (designated left) can be prepared with either swim agar (0.3% w/v) or swarm agar (0.5% w/v) from where the bacteria can migrate across the border into the right chamber containing swarm agar +/- any required supplement or challenge (e.g., antibiotics). Migration on either agar is typically measured/compared by recording the widest diameter of bacterial colonization (edge to edge) from the original point of inoculation.
	<ol>
		<li>Prepare swim agar as described in 1.1 if required.</li>
		<li>Prepare swarm agar as described in 3.1.</li>
		<li>Pour ~30 mL of swarm agar (with desired supplementation if required) into the right chamber of a dual-compartment Petri dish (100 mm x 15 mm), to the point where it is level with the plastic divider between chambers, but not overflowing into the left (Figure 2B).</li>
		<li>After the agar has hardened, fill the left chamber with ~30 mL of swim or swarm agar, again to the point of contact with the plastic divide (Figure 2C). Before it sets, use a sterile pipette tip to gently drag the agar over the border to connect the two sides with a ~1 mm tall agar bridge that spans the entire length of the divide (Figure 2D).</li>
		<li>Allow the plate to dry at room temperature (3.1). 
		NOTE: An alternative method of creating bridge is allow the left chamber agar to dry and then slowly pipette ~100 &#956;L of molten swarm agar along the plastic divider to bridge the two chambers (3.4). Inoculate the plates on the left chamber as detailed above for swim (1.2.2) or swarm (3.2) agar, before incubating at 30 &#176;C for 12-16 h, or until swarms have made sufficient progress over the right chamber to allow comparisons between strains of interest.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The isolation and characterization of suppressor mutations have successfully contributed to identifying key components of the chemotaxis system35,36,37, as well as the motor machinery itself38,39,40. While using Protocol 1, it is important to include multiple independent replicates to ensure the isolation of a large spectrum of possible mutations that could compensate for the loss of motility. Increasing the number of bacteria by streaking the culture in a line rather than a spot, can improve the odds of generating suppressors41. Isolation of the same mutation (as determined by DNA sequencing) multiple times increases confidence in its authenticity. WGS will invariably reveal the presence of other mutations in the genome. It is therefore important to verify the results by transducing (where possible) the identified mutation back into the original motility-deficient background. The suppressor mutant approach is rooted in restoring the function through a secondary mutation, so a limitation of this method is that if a critical structural gene is deleted, i.e., one that underpins the entire pathway or structure, there may be no scope for compensation. Despite being an old method, our recent work34 demonstrates its continuing utility in elucidation of new pathways that contribute to bacterial motility.
For the quantification of motor output, the cell-tethering approach remains a universally accessible tool requiring only a microscope with a camera attachment. Cell tethering has already been used in a diverse number of bacterial species including Salmonella42, Pseudomonas43, Streptococcus44, and Rhodobacter33. The success of the protocol is largely contingent on proper shearing and attachment of cells. Shearing too aggressively or omitting the pause between shears (2.1.5) tends to promote inconsistent or incomplete shearing of filaments, resulting in non-motile cells or cells tethered on a skewed axis. The enduring relevance of this protocol remains, despite the adoption of the higher-resolution bead assay by many research groups (including ours). The primary limitation of the bead assay comes from the need for the bead to adhere to the filament of the bacteria of interest. This technique has greatly benefitted from studies in E. coli that identified a &#8216;sticky&#8217; flagellin allele, which facilitates adherence45. The sticky variant is also superior in the cell-tethering assay. Such a variant is not yet available for the majority of flagellated bacteria. The situation is complicated further with some organisms possessing multiple flagellin proteins46, and in the case of Vibrio sp., also possessing a membranous sheath47. Cell-tethering can also be performed using a species-specific anti-flagellin antibody or an antibody to an engineered epitope tag.
While bacteria can swim immediately upon introduction into a liquid medium, this is not true of swarming, where cells must be first primed into a swarming state. Surface contact triggers a physiological change required for cells to initiate swarming48,49,50,51, resulting in a lag phase and a buildup of high cell density. Physiological changes include remodeling of the chemotaxis system in E. coli16, and adaptations such as cell elongation and/or hyperflagellation for other bacteria13,14,21. Given the physiological changes required to initiate swarming, we strike a cautionary note about studies that have attempted to mimic select facets of swarming - such as high density or increased cell length &#8211; simply by concentrating planktonic cell cultures to increase density, and/or inducing cell elongation by use of antibiotics that inhibit cell division52,53. If using cell elongation as a marker, we caution also that compared to planktonic cells, there is only a marginal increase in the average length of swarmer cells in Salmonella or E. coli54. Swarm assays are harder to establish than swim assays. Variables include the commercial source of the agar used to solidify the media (the special Eiken agar is essential for swarming in E. coli, whereas the more standard Difco agar supports swarming for all other bacteria), use of rich versus minimal media (E. coli and Salmonella require glucose supplementation), and most critically ambient humidity2,13,14,21. Of these, maintaining optimal humidity can be the most frustrating. [An excellent, methodical, optimization for swarming in an organism of choice (Pseudomonas aeruginosa) has been demonstrated by Morales-Soto and co-workers55.] Swarm media must be sufficiently moist to promote swarming but not so moist as to allow passive spreading/sliding, which can be readily mistaken for active swarming56. It is therefore critical that swarms be checked under a microscope to confirm the distinct patterns of movement associated with this collective motility (Video 2). Temperature is also an important consideration for optimizing the swarming assay. Higher temperatures, for example 37 &#176;C, will dry out the plates sooner than at 30 &#176;C. Using an incubator with humidity control (~70-80%) can help mitigate these issues, including seasonal changes that could affect internal building temperatures and humidity. Once successfully established, protocol 3 provides a powerful way to investigate one of the most interesting aspects of swarming bacteria, elevated resistance to antibiotics17. All protocols described here can be applied to new organisms to identify pathways that specify and control flagella mediated motility.

Bacteria employ many appendages for movement and dispersal in their ecological niches1. Flagella-driven motility is the fastest of these, promoting the colonization of favorable locales in response to environmental signals, and contributing significantly to the pathogenic ability of some species2,3. Flagellated bacteria can swim individually in bulk liquid, or swarm as a collective over a semi-solid surface4. Extracellular flagella attach to and are driven by rotary motors embedded in the membrane, which harness the power of ion gradients to generate torque that causes rotation1,2,4,5,6,7,8. In E. coli, whose motors run at a constant torque9, the motor output can be categorized in terms of rotational speed and switching of the rotor between counter-clockwise (CCW) and clockwise (CW) directions. CCW rotation promotes formation of a coherent flagellar bundle that propels the cell forward (run), while a transient switch in rotational direction (CW) causes the bundle to disassemble either partially or fully10, and the cell to reorient its swimming direction (tumble). E. coli typically run for a second and tumble for a tenth of a second. Switching frequency of the rotor or &#8216;tumble bias&#8217; is controlled by the chemotaxis signaling system, wherein transmembrane chemoreceptors detect external chemical signals and transmit them via phosphorelay to the flagellar motor to extend runs in response to attractants, or suppress them in response to toxic chemicals11,12. Swimming motility is assayed in 0.3% soft agar.
During swarming, bacteria navigate on a semi-solid surface as a dense collective, where packs of bacteria stream in a continuous swirling motion2,13,14,15. E. coli swarms exhibit altered chemosensory physiology (lower tumble bias), higher speeds, and higher tolerance to antimicrobials over cells swimming in bulk liquid16,17. Swarmers vary in their deployment of a plethora of strategies that aid movement, including surfactant production, hyperflagellation, and cell elongation2. Swarming offers bacteria a competitive advantage in both ecological and clinical settings18,19,20. There are two categories of swarming bacteria: temperate swarmers, which can swarm only on media solidified with 0.5-0.8% agar, and robust swarmers, which can navigate across higher agar concentrations21.
A variety of assays exist to interrogate swimming motility and its regulation. When impaired by mutations or environmental conditions, motility itself offers a strong selection for identifying gain-of-function suppressor mutations. These suppressors can be genuine revertants of the original mutation, or pseudo-revertants, where a second mutation restores functionality. Such mutants can be identified by whole genome sequencing (WGS). An alternative to unbiased suppressor selection is a biased targeted mutagenesis strategy (e.g., PCR mutagenesis). These methodologies often shed light on the function or environmental regulation of the motility apparatus. If the goal is to study motor function, then the restoration of wild-type motility as measured in soft agar may not necessarily indicate restoration of wild-type motor output. The cell-tethering assay, in which cells are attached to a glass surface by a single flagellum and rotation of the cell body is subsequently monitored, can be the initial assay of choice for assessing motor behavior. Although more sophisticated methodologies are now available to monitor motor properties, the required high-speed camera set-up and application of software packages for motion analysis limit their widespread use22,23,24,25. The cell-tethering assay requires only that the flagella be sheared, allowing attachment of the short filaments to a glass slide, followed by videotaping the rotation of the cell body. Although the recorded motor speeds are low in this assay because of the high load the cell body exerts on the flagellum, this assay has nonetheless contributed to valuable insights into chemotactic responses26,27,28,29, and remains a valid investigative tool as discussed below.
Swarming motility poses a different set of challenges to researchers. Selection of gain-of-function suppressors only works in swarmers that produce copious surfactants and swarm readily13. Surfactant non-producers such as E. coli are fastidious with respect to the choice of agar, media composition and humidity of the environment2,13,14,21. Once swarming conditions are established, the border-crossing assay17 is a useful methodology to interrogate the ability of a swarm to navigate new/harsh conditions. Though the protocols presented below&#160;relate to E. coli, they can be readily adapted for application in other species.

<table><tbody><tr><td>&lt;strong&gt;Reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Bacto Dehydrated Agar</td><td>Fisher Scientific</td><td>DF0140-15-4</td><td></td></tr><tr><td>EDTA Disodium Salt, Dihydrate</td><td>Fisher Scientific</td><td>02-002-786</td><td></td></tr><tr><td>Eiken agar</td><td>Eiken Chemical Co. Japan</td><td>E-MJ00</td><td>Essential for E. coli swarming</td></tr><tr><td>Glucose D (+)</td><td>Fisher Scientific</td><td>410955000</td><td></td></tr><tr><td>LB (Lennox) Broth</td><td>Fisher Scientific</td><td>BP1427-500</td><td></td></tr><tr><td>Poly-L-lysine Solution (0.1%)</td><td>Sigma-Aldrich</td><td>P8920</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>Fisher Scientific</td><td>18-605-496</td><td></td></tr><tr><td>Potassium Phosphate monobasic (KH2PO4)</td><td>Fisher Scientific</td><td>BP362-500</td><td></td></tr><tr><td>Potassium Phosphate dibasic (K2HPO4)</td><td>Fisher Scientific</td><td>BP363-500</td><td></td></tr><tr><td>Sodium chloride (NaCl)</td><td>Fisher Scientific</td><td>S271-500</td><td></td></tr><tr><td>&lt;strong&gt;Materials and Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>CellSense microscope imaging software (V. 1.6)</td><td>Olympus</td><td></td><td>Or equivalent software for microscope used</td></tr><tr><td>Electron Microscopy Sciences Scotch 666 Doube Sided Tape</td><td>Fisher</td><td>50-285-28</td><td></td></tr><tr><td>Frosted microscope slides 3x1x1mm</td><td>Fisher</td><td>12-550-343</td><td></td></tr><tr><td>Olympus BX53 microscope</td><td>Olympus</td><td>BX53</td><td>Any upright or inverted phase microscope can be used</td></tr><tr><td>Petri dishes (100 mm diameter)</td><td>Fisher Scientific</td><td>FB0875712</td><td>For soft-agar assays</td></tr><tr><td>Polyethylene Nebulizer Capillary Tubing (0.58mm x 99mm 3.0m)</td><td>Perkin Elmer</td><td>9908265</td><td></td></tr><tr><td>Round Petri Dish with 2 Compartments</td><td>VWR</td><td>89200-944</td><td>For border-crossing assays</td></tr><tr><td>Safety Hypodermic Needles (23G)</td><td>Fisher Scientific</td><td>14-826A</td><td></td></tr><tr><td>Sterile Syringe - 1 mL</td><td>Fisher scientific</td><td>14-955-450</td><td></td></tr><tr><td>Task/Tissue wipes</td><td>Fisher scientific</td><td>06-666</td><td>Or equivalent single use tissue wipes</td></tr><tr><td>VWR micro cover-glass 18x18mm</td><td>VWR</td><td>48366205</td><td></td></tr><tr><td>XM10 camera</td><td>Olympus</td><td>XM10</td><td>Or equivalent microscope camera</td></tr></tbody></table>

investigating flagella-driven motility in escherichia coli by applying three established techniques in a series

Motility is crucial to the survival and success of many bacterial species. Many methodologies exist to exploit motility to understand signaling pathways, to elucidate the function and assembly of flagellar parts, and to examine and understand patterns of movement. Here we demonstrate a combination of three of these methodologies. Motility in soft agar is the oldest, offering a strong selection for isolating gain-of-function suppressor mutations in motility-impaired strains, where motility is restored through a second mutation. The cell-tethering technique, first employed to demonstrate the rotary nature of the flagellar motor, can be used to assess the impact of signaling effectors on the motor speed and its ability to switch rotational direction. The &#8220;border-crossing&#8221; assay is more recent, where swimming bacteria can be primed to transition into moving collectively as a swarm. In combination, these protocols represent a systematic and powerful approach to identifying components of the motility machinery, and to characterizing their role in different facets of swimming and swarming. They can be easily adapted to study motility in other bacterial species.

The isolation of pseudo-revertants in an E. coli strain whose motility is impaired by high levels of the signaling molecule c-di-GMP, was detailed in recent work from our lab34. This strain (JP1442) harbored two mutations: &#916;yhjH and &#916;ycgR. YhjH is the most active phosphodiesterase that degrades c-di-GMP in E. coli. Absence of YhjH leads to elevated c-di-GMP levels and inhibition of motility. YcgR is a c-di-GMP effector. In complex with c-di-GMP, YcgR binds to the flagellar rotor to first induce CCW motor rotation and subsequently decrease motor speed. Cell tethering and bead assays showed that motor behavior returned to normal in the double mutant, yet motility in soft agar did not34. So, we deployed step 1 of the protocol to isolate pseudo-revertant flares in the double mutant (Figure 1C). The majority of the mutations mapped by WGS (HiSeq 4000 platform, PE 2 x 150 setup34) to rssB, which codes for a response regulator/adaptor protein that normally directs ClpXP protease to target &#963;S for degradation34. One of these revertants, which displayed motility close to wild-type (AW405, compare Figure 1A,D), was used to generate representative results for step 2 and 3 of the protocol section, using as controls both its double mutant parent (Figure 1B) and isogenic wild-type strain (Figure 1A).
For step 2 of the protocol section, video captures were analyzed to calculate rotations per minute (each 360&#176; complete rotation), and CWBias (the fraction of time motors rotate in a CW direction, or tumble bias). The &#916;yhjH showed fewer rotations per minute and a lower CW bias compared to the wild-type, as expected (Figure 3). Both the &#916;yhjH &#916;ycgR double mutant and its suppressor showed motor behavior similar to wild-type, observations supported by a previous analysis using the higher-resolution &#8216;bead&#8217; assay detailed in the introduction above in previous work34.
For step 3 of the protocol section, the border-crossing assay (Figure 2) was used to compare the abilities of the wild-type and the suppressor isolate, first to swarm, and then to move across the border and swarm on agar supplemented with kanamycin. Results show that both strains reached the border at a similar time (data not shown) indicating similar rates of swarming from an identical inoculation point. However, cross-over of the swarm to the right (antibiotic) chamber was marginally, but consistently greater for the wild-type than the suppressor at 20 &#181;g/mL kanamycin (Figure 4). The difference between the two strains was more pronounced at 40 &#181;g/mL kanamycin. Together, these data suggest that the mutations in rssB that restored motility on soft-agar plates (Figure 1D), negatively impact the antibiotic resistance of the suppressor strain during swarming (Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61364/61364fig01.jpg" /> 
Figure 1: Soft-agar motility assays and emergence of suppressor flares. 
The plates contain LB solidified with 0.3 % w/v agar. E. coli strains were inoculated in the center of each plate and incubated at 30&#176;C for 8 h, except for C, which was incubated for 16 h. (A) Wild-type E. coli (AW405). (B) Motility-deficient variant &#8710;yhjH &#8710;ycgR (JP1442). (C) As in B, except longer incubation times. Arrows point to faster moving &#8216;flares&#8217; emerging at the peripheral ring of the expanding swim colony. (D) A suppressor isolated from a flare in C. <a href="https://www.jove.com/files/ftp_upload/61364/61364fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61364/61364fig02.jpg" /> 
Figure 2: Schematic for setting up a Border-crossing plate assay. 
(A) Pour ~30 mL of swarm agar (with desired antibiotic) into the right chamber of a divided petri dish until level with the plastic divider and allow to set with lid closed. (B) Fill the left chamber with ~30 mL of swim or swarm agar to the point of contact with the top of the plastic divider. (C) Use a sterile pipette tip to gently drag the molten swarm agar over the border, thereby connecting the two sides with a ~1 mm tall agar bridge and allow to set with lid closed. (D) Allow the plate to dry further at room temperature overnight before inoculating the left chamber with the desired strain, and incubating at 30 &#176;C. <a href="https://www.jove.com/files/ftp_upload/61364/61364fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61364/61364fig03.jpg" /> 
Figure 3: Motor properties of various strains as measured by the cell-tethering technique. 
Wild-type (AW405), &#8710;yhjH (VN133), &#8710;ycgR &#8710;yhjH (JP1442), and its suppressor (JP1836) were grown in LB at 30 &#176;C to mid-exponential phase prior to tethering. (A) Rotations per minute (completed 360&#176; turns), and (B) CWbias (fraction of time motors rotate in a CW direction). Standard deviation of the mean (&#177;). 20 tethered cells were observed for 60 sec in each strain. <a href="https://www.jove.com/files/ftp_upload/61364/61364fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61364/61364fig04.jpg" /> 
Figure 4: Border-crossing assays. 
Mid-exponential phase cultures of wild-type E. coli (AW405) and the suppressor mutant (JP1836) were inoculated at the indicated position (*) in the left compartment of the divided plate containing swarm media, and incubated at 30 &#176;C. They reached the border at comparable times. The plates were incubated for a further 6 h, during which the swarm crossed over to the right chamber, in which the media was supplemented with kanamycin (Kan; numbers indicate &#181;g/mL). Plates are representative of three biological replicates each carried out in triplicate. <a href="https://www.jove.com/files/ftp_upload/61364/61364fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Preparation of a chamber slide for cell tethering. (A) Lay down two pieces of double-sided tape before (B) using a razor blade to trim away the excess. (C) Peel away the top layer to expose the adhesive before (D) affixing a coverslip and gently pressing it into position (indicated by [ ), ensuring all air is pushed out of the interface between the coverslip and tape below. (E) Load sample (shown here with DNA loading dye [30% v/v glycerol, 0.25% w/v bromophenol blue, and 0.25% w/v xylene cyanol] added to aid visualization) into the top of the created channel (arrow) while (F) angling onto clean, tissue task wipe to help draw the solution through the chamber as the tissue absorbs the liquid (arrow) in the channel and draws it through. <a href="https://cloudflare.jove.com/files/ftp_upload/61364/61364supfig01.jpg" target="_blank">Please click here to download this figure.</a>
Video 1: Rotation of tethered E. coli cells. <a href="https://cloudflare.jove.com/files/ftp_upload/61364/Video_1.m4v" target="_blank">Please click here to download this video.</a>
Video 2: An active E. coli swarm filmed under 60x magnification, demonstrating its characteristic swirling motion behind the edge of the moving front. <a href="https://cloudflare.jove.com/files/ftp_upload/61364/Video_2.m4v" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about Investigating Flagella-Driven Motility in Escherichia coli by Applying Three Established Techniques in a Series at JoVE.com

Investigating Flagella-Driven Motility in Escherichia coli by Applying Three Established Techniques in a Series

Many bacteria use flagella-driven motility to navigate their environment and colonize favorable surroundings both individually and as a collective. Demonstrated here is the use of three established methods that exploit motility as a selection tool to identify components/pathways contributing to swimming and swarming motility.

Investigating Flagella-Driven Motility in Escherichia coli by Applying Three Established Techniques in a Series

Motility is crucial to the survival and success of many bacterial species. Many methodologies exist to exploit motility to understand signaling pathways, ...

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Nanyang Technological University

Research

JoVE Journal

Biology

7.5K Views.  The University of Texas at Austin. Many bacteria use flagella-driven motility to navigate their environment and colonize favorable surroundings both individually and as a collective. Demonstrated here is the use of three established methods that exploit motility as a selection tool to identify components/pathways contributing to swimming and swarming motility.

Video: Investigating Flagella-Driven Motility in Escherichia coli by Applying Three Established Techniques in a Series

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

A Microfluidic Device for Quantifying Bacterial Chemotaxis in Stable Concentration Gradients

Chemotaxis allows bacteria to approach sources of attractant chemicals or to avoid sources of repellent chemicals. Bacteria constantly monitor the concentration of specific chemoeffectors by comparing the current concentration to the concentration detected a few seconds earlier. This comparison determines the net direction of movement. Although multiple, competing gradients often coexist in nature, conventional approaches for investigating bacterial chemotaxis are suboptimal for quantifying migration in response to concentration gradients of attractants and repellents. Here, we describe the development of a microfluidic chemotaxis model for presenting precise and stable concentration gradients of chemoeffectors to bacteria and quantitatively investigating their response to the applied gradient. The device is versatile in that concentration gradients of any desired absolute concentration and gradient strength can be easily generated by diffusive mixing. The device is demonstrated using the response of Escherichia coli RP437 to gradients of amino acids and nickel ions.

This protocol describes the development of a microfluidic device for investigating bacterial chemotaxis in stable concentration gradients of chemoeffectors.

Chemotaxis allows bacteria to approach sources of attractant chemicals or to avoid sources of repellent chemicals. Bacteria constantly monitor the ...

Quantifying Bacterial Surface Swarming Motility on Inducer Gradient Plates

Bacterial swarming motility is a common microbiological phenotype that bacterial communities use to migrate over semisolid surfaces. In investigations of induced swarming motility, specific concentration&#160;of an inducer may not be able to report events occurring within the optimal concentration range to elicit the desired responses from a species. Semisolid plates containing multiple concentrations are commonly used to investigate the response within an inducer concentration range. However, separate semisolid plates increase variations in medium viscosity and moisture content within each plate due to nonuniform solidification time.
This paper describes a one-step method to simultaneously test surface swarming motility on a single gradient plate, where the isometrically arranged test wells allow the simultaneous acquisition of multiconcentration responses. In the present work, the surface swarming of Escherichia coli K12 and Pseudomonas aeruginosa PAO1 were evaluated in response to a concentration gradient of inducers such as resveratrol and arabinose. Periodically, the swarm morphologies were imaged using an imaging system to capture the entire surface swarming process.
The quantitative measurement of the swarm morphologies was acquired using ImageJ software, providing analyzable information of the swarm area. This paper presents a simple gradient swarm plate method that provides qualitative and quantitative information about the inducers&#39; effects on surface swarming, which can be extended to study the effects of other inducers on a broader range of motile bacterial species.

Here, we describe the use of inducer gradient plates to evaluate bacterial swarming motility while simultaneously obtaining multiple concentration responses.

Bacterial swarming motility is a common microbiological phenotype that bacterial communities use to migrate over semisolid surfaces. In investigations of ...

Biochemistry

Visualization of Twitching Motility and Characterization of the Role of the PilG in Xylella fastidiosa

Xylella fastidiosa is a Gram-negative non-flagellated bacterium that causes a number of economically important diseases of plants. The twitching motility provides X. fastidiosa a means for long-distance intra-plant movement and colonization, contributing toward pathogenicity in X. fastidiosa. The twitching motility of X. fastidiosa is operated by type IV pili. Type IV pili of Xylella fastidiosa are regulated by pilG, a chemotaxis regulator in Pil-Chp operon encoding proteins that are involved with signal transduction pathways. To elucidate the roles of pilG in the twitching motility of X. fastidiosa, a pilG-deficient mutant Xf&#916;pilG and its complementary strain Xf&#916;pilG-C containing native pilG were developed. A microfluidic chambers integrated with a time-lapse image recording system was used to observe twitching motility in Xf&#916;pilG, Xf&#916;pilG-C and its wild type strain. Using this recording system, it permits long-term spatial and temporal observations of aggregation, migration of individual cells and populations of bacteria via twitching motility. X. fastidiosa wild type and complementary Xf&#916;pilG-C strain showed typical twitching motility characteristics directly observed in the microfluidic flow chambers, whereas mutant Xf&#916;pliG exhibited the twitching deficient phenotype. This study demonstrates that pilG contributes to the twitching motility of X. fastidiosa. The microfluidic flow chamber is used as a means for observing twitching motility.

Visualization of Twitching Motility and Characterization of the Role of the PilG in Xylella fastidiosa

In this study, a nano-microfluidic flow chamber was employed to visualize and functionally characterize the twitching motility of Xylella fastidiosa, a bacterium that causes Pierce&#39;s disease in grapevine.

Xylella fastidiosa is a Gram-negative non-flagellated bacterium that causes a number of economically important diseases of plants. The twitching motility ...

Immunology and Infection

Single-Molecule Analysis of Sf9 Purified Superprocessive Kinesin-3 Family Motors

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved single-molecule studies and has gained immense popularity in detecting and understanding the dynamic behavior of fluorescent-tagged molecules. Here, we describe a detailed method for&#160;in vitro single-molecule studies of kinesin-3 family motors using Total Internal Reflection Fluorescence (TIRF) microscopy. Kinesin-3 is a large family that plays critical roles in cellular and physiological functions ranging from intracellular cargo transport to cell division to development. We have shown previously that constitutively active dimeric kinesin-3 motors exhibit fast and superprocessive motility with high microtubule affinity at the single-molecule level using cell lysates prepared by expressing motor in mammalian cells. Our lab studies kinesin-3 motors and their regulatory mechanisms using cellular, biochemical and biophysical approaches, and such studies demand purified proteins at a large scale. Expression and purification of these motors using mammalian cells would be expensive and time-consuming, whereas expression in a prokaryotic expression system resulted in significantly aggregated and&#160;inactive protein. To overcome the limitations posed by bacterial purification systems and mammalian cell lysate, we have established a robust Sf9-baculovirus expression system to express and purify these motors. The kinesin-3 motors are C-terminally tagged with 3-tandem fluorescent proteins (3xmCitirine or 3xmCit) that provide enhanced signals and decreased photobleaching. In vitro single-molecule and multi-motor gliding analysis of Sf9 purified proteins demonstrate that kinesin-3 motors are fast and superprocessive akin to our previous studies using mammalian cell lysates. Other applications using these assays include detailed knowledge of oligomer conditions of motors, specific binding partners paralleling biochemical studies, and their kinetic state.

This study details purification of KIF1A(1-393LZ), a member of kinesin-3 family, using Sf9-baculovirus expression system. In vitro single-molecule and multi-motor gliding analysis of these purified motors exhibited robust motility properties comparable to motors from mammalian cell lysate. Thus, Sf9-baculovirus system is amenable to express and purify motor protein of interest.

A complex cellular environment poses challenges for single-molecule motility analysis. However, advancement in imaging techniques have improved ...

Continuous Measurement of Biological Noise in Escherichia Coli Using Time-lapse Microscopy

The protocol developed here offers a tool to enable computer tracking of Escherichia coli division and fluorescent levels over several hours. The process starts by screening for colonies that survive on minimal media, assuming that only Escherichia coli harboring the correct plasmid will be able to thrive in the specific conditions. Since the process of building large genetic circuits, requiring the assembly of many DNA parts, is challenging, circuit components are often distributed between multiple plasmids at different copy numbers requiring the use of several antibiotics. Mutations in the plasmid can destroy transcription of the antibiotic resistance genes and interject with resources management in the cell leading to necrosis. The selected colony is set on a glass-bottom Petri dish and a few focus planes are selected for microscopy tracking in both bright field and fluorescent domains. The protocol maintains the image focus for more than 12 hours under initial conditions that cannot be regulated, creating a few difficulties. For example, dead cells start to accumulate in the lenses' field of focus after a few hours of imaging, which causes toxins to buildup and the signal to blur and decay. Depletion of nutrients introduces new metabolic processes and hinder the desired response of the circuit. The experiment's temperature lowers the effectivity of inducers and antibiotics, which can further damage the reliability of the signal. The minimal media gel shrinks and dries, and as a result the optical focus changes over time. We developed this method to overcome these challenges in Escherichia coli, similar to previous works developing analogous methods for other micro-organisms. In addition, this method offers an algorithm to quantify the total stochastic noise in unaltered and altered cells, finding that the results are consistent with flow analyzer predictions as shown by a similar coefficient of variation (CV).

Continuous Measurement of Biological Noise in Escherichia Coli Using Time-lapse Microscopy

This work presents a microscopy method that allows live imaging of a single cell of Escherichia coli for analysis and quantification of the stochastic behavior of synthetic gene circuits.

The protocol developed here offers a tool to enable computer tracking of Escherichia coli division and fluorescent levels over several hours. The process ...

Bioengineering

Generation of Null Mutants to Elucidate the Role of Bacterial Glycosyltransferases in Bacterial Motility

The study of glycosylation in prokaryotes is a rapidly growing area. Bacteria harbor different glycosylated structures on their surface whose glycans constitute a strain-specific barcode. The associated glycans show higher diversity in sugar composition and structure than those of eukaryotes and are important in bacterial-host recognition processes and interaction with the environment. In pathogenic bacteria, glycoproteins have been involved in different stages of the infectious process, and glycan modifications can interfere with specific functions of glycoproteins. However, despite the advances made in the understanding of glycan composition, structure, and biosynthesis pathways, understanding of the role of glycoproteins in pathogenicity or interaction with the environment remains very limited. Furthermore, in some bacteria, the enzymes required for protein glycosylation are shared with other polysaccharide biosynthetic pathways, such as lipopolysaccharide and capsule biosynthetic pathways. The functional importance of glycosylation has been elucidated in several bacteria through mutation of specific genes thought to be involved in the glycosylation process and the study of its impact on the expression of the target glycoprotein and the modifying glycan. Mesophilic Aeromonas have a single and O-glycosylated polar flagellum. Flagellar glycans show diversity in carbohydrate composition and chain length between Aeromonas strains. However, all strains analyzed to date show a pseudaminic acid derivative as the linking sugar that modifies serine or threonine residues. The pseudaminic acid derivative is required for polar flagella assembly, and its loss has an impact on adhesion, biofilm formation, and colonization. The protocol detailed in this article describes how the construction of null mutants can be used to understand the involvement of genes or genome regions containing putative glycosyltransferases in the biosynthesis of a flagellar glycan. This includes the potential to understand the function of the glycosyltransferases involved and the role of the glycan. This will be achieved by comparing the glycan deficient mutant to the wild-type strain.

Here, we describe the construction of null mutants of Aeromonas in specific glycosyltransferases or regions containing glycosyltransferases, the motility assays, and flagella purification performed to establish the involvement and function of their encoded enzymes in the biosynthesis of a glycan, as well as the role of this glycan in bacterial pathogenesis.

The study of glycosylation in prokaryotes is a rapidly growing area. Bacteria harbor different glycosylated structures on their surface whose glycans ...

Preparation, Imaging, and Quantification of Bacterial Surface Motility Assays

Bacterial surface motility, such as swarming, is commonly examined in the laboratory using plate assays that necessitate specific concentrations of agar and sometimes inclusion of specific nutrients in the growth medium. The preparation of such explicit media and surface growth conditions serves to provide the favorable conditions that allow not just bacterial growth but coordinated motility of bacteria over these surfaces within thin liquid films. Reproducibility of swarm plate and other surface motility plate assays can be a major challenge. Especially for more &#8220;temperate swarmers&#8221; that exhibit motility only within agar ranges of 0.4%-0.8% (wt/vol), minor changes in protocol or laboratory environment can greatly influence swarm assay results. &#8220;Wettability&#8221;, or water content at the liquid-solid-air interface of these plate assays, is often a key variable to be controlled. An additional challenge in assessing swarming is how to quantify observed differences between any two (or more) experiments. Here we detail a versatile two-phase protocol to prepare and image swarm assays. We include guidelines to circumvent the challenges commonly associated with swarm assay media preparation and quantification of data from these assays. We specifically demonstrate our method using bacteria that express fluorescent or bioluminescent genetic reporters like green fluorescent protein (GFP), luciferase (lux operon), or cellular stains to enable time-lapse optical imaging. We further demonstrate the ability of our method to track competing swarming species in the same experiment.

Swarming motility is influenced by physical and environmental factors. We describe a two-phase protocol and guidelines to circumvent the challenges commonly associated with swarm assay preparation and data collection. A macroscopic imaging technique is employed to obtain detailed information on swarm behavior that is not provided by current analysis techniques.

Bacterial surface motility, such as swarming, is commonly examined in the laboratory using plate assays that necessitate specific concentrations of agar ...

Biophysical Characterization of Flagellar Motor Functions

The role of flagellar motors in bacterial motility and chemotaxis is well-understood. Recent discoveries suggest that flagellar motors are able to remodel in response to a variety of environmental stimuli and are among the triggers for surface colonization and infections. The precise mechanisms by which motors remodel and promote cellular adaptation likely depend on key motor attributes. The photomultiplier-based bead-tracking technique presented here enables accurate biophysical characterization of motor functions, including adaptations in motor speeds and switch-dynamics. This approach offers the advantage of real-time tracking and the ability to probe motor behavior over extended durations. The protocols discussed can be readily extended to study flagellar motors in a variety of bacterial species.

Recent findings suggest that bacterial flagellar motors sense a variety of environmental signals and remodel in response. The bead-assays discussed here are expected to help explain the role of remodeling in cellular adaptation to environmental stressors.

The role of flagellar motors in bacterial motility and chemotaxis is well-understood. Recent discoveries suggest that flagellar motors are able to remodel ...

Visualizing Bacterial Motility Based on a Color Reaction

Bacterial motility is crucial for bacterial pathogenicity, biofilm formation, and drug resistance. Bacterial motility is crucial for the invasion and/or dissemination of many pathogenic species. Therefore, it is important to detect bacterial motility. Bacterial growth conditions, such as oxygen, pH, and temperature, can affect bacterial growth and the expression of bacterial flagella. This can lead to reduced motility or even loss of motility, resulting in the inaccurate evaluation of bacterial motility. Based on the color reaction of 2,3,5-triphenyl tetrazolium chloride (TTC) by intracellular dehydrogenases of living bacteria, TTC was added to traditional semisolid agar for bacterial motility detection. The results showed that this TTC semisolid agar method for the detection of bacterial motility is simple, easy to operate, and does not involve large and expensive instruments. The results also showed that the highest motility was observed in semisolid medium prepared with 0.3% agar. Compared with the traditional semisolid medium, the results are easier to evaluate and more accurate.

Here, we present a protocol to detect bacterial motility based on a color reaction. Key advantages of this method are that it is easy to evaluate and more accurate, and does not require specialized equipment.

Bacterial motility is crucial for bacterial pathogenicity, biofilm formation, and drug resistance. Bacterial motility is crucial for the invasion and/or ...

Investigating Flagella-Driven Motility in Escherichia coli by Applying Three Established Techniques in a Series

Summary

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Chapters in this video

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

A Microfluidic Device for Quantifying Bacterial Chemotaxis in Stable Concentration Gradients

Quantifying Bacterial Surface Swarming Motility on Inducer Gradient Plates

Visualization of Twitching Motility and Characterization of the Role of the PilG in Xylella fastidiosa

Single-Molecule Analysis of Sf9 Purified Superprocessive Kinesin-3 Family Motors

Continuous Measurement of Biological Noise in Escherichia Coli Using Time-lapse Microscopy

Generation of Null Mutants to Elucidate the Role of Bacterial Glycosyltransferases in Bacterial Motility

Preparation, Imaging, and Quantification of Bacterial Surface Motility Assays

Biophysical Characterization of Flagellar Motor Functions

Visualizing Bacterial Motility Based on a Color Reaction