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...

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>Reagents</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>Materials and Equipment</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 b...

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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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

An Assay for Measuring the Activity of Escherichia coli Inducible Lysine Decarboxyase

Escherichia coli is an enteric bacterium that is capable of growing over a wide range of pH values (pH 5 - 9)1 and, incredibly, is able to survive extreme acid stresses including passage through the mammalian stomach where the pH can fall to as low as pH 1 - 22. To enable such a broad range of acidic pH survival, E. coli possesses four different inducible amino acid decarboxylases that decarboxylate their substrate amino acids in a proton-dependent manner thus raising the internal pH. The decarboxylases include the glutamic acid decarboxylases GadA and GadB3, the arginine decarboxylase AdiA4, the lysine decarboxylase LdcI5, 6 and the ornithine decarboxylase SpeF7. All of these enzymes utilize pyridoxal-5'-phospate as a co-factor8 and function together with inner-membrane substrate-product antiporters that remove decarboxylation products to the external medium in exchange for fresh substrate2. In the case of LdcI, the lysine-cadaverine antiporter is called CadB. Recently, we determined the X-ray crystal structure of LdcI to 2.0 &#197;, and we discovered a novel small-molecule bound to LdcI the stringent response regulator guanosine 5'-diphosphate,3'-diphosphate (ppGpp) 14. The stringent response occurs when exponentially growing cells experience nutrient deprivation or one of a number of other stresses9. As a result, cells produce ppGpp which leads to a signaling cascade culminating in the shift from exponential growth to stationary phase growth10. We have demonstrated that ppGpp is a specific inhibitor of LdcI 14. Here we describe the lysine decarboxylase assay, modified from the assay developed by Phan et al.11, that we have used to determine the activity of LdcI and the effect of pppGpp/ppGpp on that activity. The LdcI decarboxylation reaction removes the &alpha;-carboxy group of L-lysine and produces carbon dioxide and the polyamine cadaverine (1,5-diaminopentane)5. L-lysine and cadaverine can be reacted with 2,4,6-trinitrobenzensulfonic acid (TNBS) at high pH to generate N,N'-bistrinitrophenylcadaverine (TNP-cadaverine) and N,N&#8242;-bistrinitrophenyllysine (TNP-lysine), respectively11. The TNP-cadaverine can be separated from the TNP-lysine as the former is soluble in organic solvents such as toluene while the latter is not (See Figure 1). The linear range of the assay was determined empirically using purified cadaverine.

An Assay for Measuring the Activity of Escherichia coli Inducible Lysine Decarboxyase

The activity of the inducible lysine decarboxylase is monitored by reacting the substrate L-lysine and the product cadaverine with 2,4,6-trinitrobenzensulfonic acid to form adducts that have differential solubility in toluene.

Escherichia coli is an enteric bacterium that is capable of growing over a wide range of pH values (pH 5 - 9)1 and, incredibly, is able to survive extreme ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted into fuel alcohols1. The potential for enzymatic hydrolysis of cellulosic biomass to provide renewable energy has intensified efforts to engineer cellulases for economical fuel production2. Of particular interest are fungal cellulases3-8, which are already being used industrially for foods and textiles processing.
Identifying active variants among a library of mutant cellulases is critical to the engineering process; active mutants can be further tested for improved properties and/or subjected to additional mutagenesis. Efficient engineering of fungal cellulases has been hampered by a lack of genetic tools for native organisms and by difficulties in expressing the enzymes in heterologous hosts. Recently, Morikawa and coworkers developed a method for expressing in E. coli the catalytic domains of endoglucanases from H. jecorina3,9, an important industrial fungus with the capacity to secrete cellulases in large quantities. Functional E. coli expression has also been reported for cellulases from other fungi, including Macrophomina phaseolina10 and Phanerochaete chrysosporium11-12. 
We present a method for high throughput screening of fungal endoglucanase activity in E. coli. (Fig 1) This method uses the common microbial dye Congo Red (CR) to visualize enzymatic degradation of carboxymethyl cellulose (CMC) by cells growing on solid medium. The activity assay requires inexpensive reagents, minimal manipulation, and gives unambiguous results as zones of degradation (&#x201C;halos&#x201D;) at the colony site. Although a quantitative measure of enzymatic activity cannot be determined by this method, we have found that halo size correlates with total enzymatic activity in the cell. Further characterization of individual positive clones will determine , relative protein fitness. 
Traditional bacterial whole cell CMC/CR activity assays13 involve pouring agar containing CMC onto colonies, which is subject to cross-contamination, or incubating cultures in CMC agar wells, which is less amenable to large-scale experimentation. Here we report an improved protocol that modifies existing wash methods14 for cellulase activity: cells grown on CMC agar plates are removed prior to CR staining. Our protocol significantly reduces cross-contamination and is highly scalable, allowing the rapid screening of thousands of clones. In addition to H. jecorina enzymes, we have expressed and screened endoglucanase variants from the Thermoascus aurantiacus and Penicillium decumbens (shown in Figure 2), suggesting that this protocol is applicable to enzymes from a range of organisms.

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

We describe a low cost, high throughput method to screen for fungal endoglucanase activity in E. coli. The method relies on a simple visual readout of substrate degradation, does not require enzyme purification, and is highly scalable. This allows for the rapid screening of large libraries of enzyme variants.

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted ...

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Phenotypes are determined by a complex series of physical (e.g. protein-protein) and functional (e.g. gene-gene or genetic) interactions (GI)1. While physical interactions can indicate which bacterial proteins are associated as complexes, they do not necessarily reveal pathway-level functional relationships1. GI screens, in which the growth of double mutants bearing two deleted or inactivated genes is measured and compared to the corresponding single mutants, can illuminate epistatic dependencies between loci and hence provide a means to query and discover novel functional relationships2. Large-scale GI maps have been reported for eukaryotic organisms like yeast3-7, but GI information remains sparse for prokaryotes8, which hinders the functional annotation of bacterial genomes. To this end, we and others have developed high-throughput quantitative bacterial GI screening methods9, 10.
Here, we present the key steps required to perform quantitative E. coli Synthetic Genetic Array (eSGA) screening procedure on a genome-scale9, using natural bacterial conjugation and homologous recombination to systemically generate and measure the fitness of large numbers of double mutants in a colony array format. Briefly, a robot is used to transfer, through conjugation, chloramphenicol (Cm) - marked mutant alleles from engineered Hfr (High frequency of recombination) 'donor strains' into an ordered array of kanamycin (Kan) - marked F- recipient strains. Typically, we use loss-of-function single mutants bearing non-essential gene deletions (e.g. the 'Keio' collection11) and essential gene hypomorphic mutations (i.e. alleles conferring reduced protein expression, stability, or activity9, 12, 13) to query the functional associations of non-essential and essential genes, respectively. After conjugation and ensuing genetic exchange mediated by homologous recombination, the resulting double mutants are selected on solid medium containing both antibiotics. After outgrowth, the plates are digitally imaged and colony sizes are quantitatively scored using an in-house automated image processing system14. GIs are revealed when the growth rate of a double mutant is either significantly better or worse than expected9. Aggravating (or negative) GIs often result between loss-of-function mutations in pairs of genes from compensatory pathways that impinge on the same essential process2. Here, the loss of a single gene is buffered, such that either single mutant is viable. However, the loss of both pathways is deleterious and results in synthetic lethality or sickness (i.e. slow growth). Conversely, alleviating (or positive) interactions can occur between genes in the same pathway or protein complex2 as the deletion of either gene alone is often sufficient to perturb the normal function of the pathway or complex such that additional perturbations do not reduce activity, and hence growth, further. Overall, systematically identifying and analyzing GI networks can provide unbiased, global maps of the functional relationships between large numbers of genes, from which pathway-level information missed by other approaches can be inferred9.

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Systematic, large-scale synthetic genetic (gene-gene or epistasis) interaction screens can be used to explore genetic redundancy and pathway cross-talk. Here, we describe a high-throughput quantitative synthetic genetic array screening technology, termed eSGA that we developed for elucidating epistatic relationships and exploring genetic interaction networks in Escherichia coli.

Phenotypes  are determined by a complex series of physical (e.g. protein-protein) and  functional (e.g. gene-gene or genetic) interactions (GI)1. While ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has become the method of choice for introducing DNA into prokaryotes that are not naturally competent. Electroporation is a rapid, efficient, and streamlined transformation method that, in addition to purified DNA and competent bacteria, requires commercially available gene pulse controller and cuvettes. In contrast to the pulsing step, preparation of electrocompetent cells is time consuming and labor intensive involving repeated rounds of centrifugation and washes in decreasing volumes of sterile, cold water, or non-ionic buffers of large volumes of cultures grown to mid-logarithmic phase of growth. Time and effort can be saved by purchasing electrocompetent cells from commercial sources, but the selection is limited to commonly employed E. coli laboratory strains. We are hereby disseminating a rapid and efficient method for preparing electrocompetent E. coli, which has been in use by bacteriology laboratories for some time, can be adapted to V. cholerae and other prokaryotes. While we cannot ascertain whom to credit for developing the original technique, we are hereby making it available to the scientific community.

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation is a commonly employed method for introducing DNA into bacteria in a process known as transformation. Traditional protocols for the preparation of electrocompetent cells are time consuming and labor intensive. This article describes an alternate, rapid, and efficient method for the preparation of electrocompetent cells presently employed by some laboratories.

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has ...

Detection of Live Escherichia coli O157:H7 Cells by PMA-qPCR

A unique open reading frame (ORF) Z3276 was identified as a specific genetic marker for E. coli O157:H7. A qPCR assay was developed for detection of E. coli O157:H7 by targeting ORF Z3276. With this assay, we can detect as low as a few copies of the genome of DNA of E. coli O157:H7. The sensitivity and specificity of the assay were confirmed by intensive validation tests with a large number of E. coli O157:H7 strains (n = 369) and non-O157 strains (n = 112). Furthermore, we have combined propidium monoazide (PMA) procedure with the newly developed qPCR protocol for selective detection of live cells from dead cells. Amplification of DNA from PMA-treated dead cells was almost completely inhibited in contrast to virtually unaffected amplification of DNA from PMA-treated live cells. Additionally, the protocol has been modified and adapted to a 96-well plate format for an easy and consistent handling of a large number of samples. This method is expected to have an impact on accurate microbiological and epidemiological monitoring of food safety and environmental source.

Detection of Live Escherichia coli O157:H7 Cells by PMA-qPCR

A qPCR assay was developed for detection of Escherichia coli O157:H7 targeting a unique genetic marker, Z3276. The qPCR was combined with propidium monoazide (PMA) treatment for live cell detection. This protocol has been modified and adapted to a 96-well plate format for easy and consistent handling of numerous samples

A unique open reading frame (ORF) Z3276 was identified as a specific genetic marker for E. coli O157:H7. A qPCR assay was developed for detection of E. ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

We report here that the expression of protein complexes in vivo in Escherichia coli can be more convenient than traditional reconstitution experiments in vitro. In particular, we show that the poor solubility of Escherichia coli DNA polymerase III &#949; subunit (featuring 3&#8217;-5&#8217; exonuclease activity) is highly improved when the same protein is co-expressed with the &#945; and &#952; subunits (featuring DNA polymerase activity and stabilizing &#949;, respectively). We also show that protein co-expression in E. coli can be used to efficiently test the competence of subunits from different bacterial species to associate in a functional protein complex. We indeed show that the &#945; subunit of Deinococcus radiodurans DNA polymerase III can be co-expressed in vivo with the &#949; subunit of E. coli. In addition, we report on the use of protein co-expression to modulate mutation frequency in E. coli. By expressing the wild-type &#949; subunit under the control of the araBAD promoter (arabinose-inducible), and co-expressing the mutagenic D12A variant of the same protein, under the control of the lac promoter (inducible by isopropyl-thio-&#946;-D-galactopyranoside, IPTG), we were able to alter the E. coli mutation frequency using appropriate concentrations of the inducers arabinose and IPTG. Finally, we discuss recent advances and future challenges of protein co-expression in E. coli.

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

Protein co-expression is a powerful alternative to the reconstitution in vitro of protein complexes, and is of help in performing biochemical and genetic tests in vivo. Here we report on the use of protein co-expression in Escherichia coli to obtain protein complexes, and to tune the mutation frequency of cells.

We report here that the expression of protein complexes in vivo in Escherichia coli can be more convenient than traditional reconstitution experiments in ...

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

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

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An Assay for Measuring the Activity of Escherichia coli Inducible Lysine Decarboxyase

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

Mapping Bacterial Functional Networks and Pathways in Escherichia Coli using Synthetic Genetic Arrays

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Detection of Live Escherichia coli O157:H7 Cells by PMA-qPCR

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The Multifaceted Benefits of Protein Co-expression in Escherichia coli

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments