This project was funded by the National Institutes of Health under GM018568. This protocol was performed in part at the Center for Nanoscale Systems (CNS), a member of the National Nanotechnology Coordinated Infrastructure Network (NNCI), which is supported by the National Science Foundation under NSF award no. 1541959. CNS is part of Harvard University. Many thanks are due to Thomas Norman and Nathan Lord for their work in conceiving and fabricating the master template used for the devices shown here and in building the original version of the apparatus. We also thank Johan Paulsson for his valuable collaborative advice and thank members of his lab for their advice and continued improvements to bacterial microfluidic apparatuses.

1. PDMS Device Casting
<ol>
	<li>In a dust-free environment, mix PDMS polymer and curing agent at a 10:1 ratio and degas under vacuum for at least 10 min.</li>
	<li>Place a silicon master (or an epoxy or polyurethane replica thereof) on a piece of unbroken aluminum foil in a polystyrene Petri plate. Pour the degassed PDMS mixture over the master to a depth of approximately 5 mm. Degas the Petri plate for at least 10 min. Use a gentle stream of air to break surface bubbles. 
	NOTE: The dimensions of the two-tiered device used are provided in the accompanying CAD file5 (see Supplementary File 1). A plastic replica master may partially float during degassing; if this occurs, push down the replica with a clean plastic applicator.</li>
	<li>Cure PDMS for at least 1 h in an oven at 60 &#176;C and cool to room temperature. Remove the aluminum foil containing the master and cured PDMS from the Petri plate. Carefully peel off the aluminum foil from the back of the master, then carefully peel the PDMS from the master. 
	NOTE: Alternatively, PDMS may be cured overnight at room temperature. The relative fragility of a silicon-wafer master may be overcome by using epoxy adhesive to permanently bond the wafer to a 1/16 inch-thick aluminum disc of the same size as the wafer.</li>
	<li>As a wafer typically includes several microfluidic devices with slightly different dimensions (see Supplementary File 1), cut a single device to size using a clean, sharp scalpel. 
	NOTE: For routine B. subtilis work, use devices with 1.0-&#181;m-wide cell trenches, which are marked with a C or F in the accompanying CAD file.</li>
</ol>
2. Device Punching and Bonding
<ol>
	<li>Place a piece of translucent office tape (see Table of Materials) over the patterned side of the device to enhance the visibility of the patterned features. The inlet and outlet ports are identified as circular areas at opposite ends of the visible fluidic channels (Figure 1). To improve their visibility when punching holes in the device for the inlet and outlet pins, mark the inlets and outlets with dots using a fine-tipped marker. 
	NOTE: To ensure that the inlet and outlet pins are not crowded, every other channel is punched, beginning with the channel just beneath the alphabetical designator for the device.</li>
	<li>Flip the PDMS device so that the patterned side is down, and punch through the device to the marked dots using an 0.75-mm biopsy punch; discard the punchings. 
	NOTE: The devices are punched with the patterned side down because the hole generated by the biopsy punch is typically slightly flared where the punch enters the polymer. Punching the device in an inverted orientation ensures that the hole on the patterned side has a sharp border.</li>
	<li>Using a stereomicroscope, ensure that the punched holes overlap with the inlet and outlet areas of the device. Use a fine-tipped tweezers to remove any extraneous fragments of PDMS from the punched holes.</li>
	<li>Use adhesive tape to remove dust from both sides of the device and place into a clean polystyrene Petri plate. Prepare a 22 mm x 40 mm cover glass by wetting with 100% isopropyl alcohol and rubbing with a cleanroom wipe; dry with a stream of dust-free air or nitrogen.</li>
	<li>Place the punched PDMS device feature side up together with the cleaned cover glass into an oxygen plasma cleaner.
	<ol>
		<li>Plasma-treat the device with the following settings: vacuum, 70 mTorr; O2 pressure, 200 mTorr; duration, 15 s; power, 30 W. Immediately after the plasma treatment period finishes, remove the device and cover glass from the plasma cleaner.</li>
		<li>Invert the PDMS and place it on the center of the cover glass so that the patterned side contacts the glass; ensure that the feeding channels (running between the inlet and outlet areas) are aligned with the long axis of the cover glass. Press down very gently to ensure that the PDMS seals against the glass.</li>
	</ol>
	</li>
	<li>Bake the assembled device in an oven for at least 1 h at 60 &#176;C. After baking, manually verify that the PDMS is bonded to the glass by pushing gently on one corner of the device. 
	NOTE: Once the device is bonded, it should be used within 24 h, as the polymer will become increasingly hydrophobic with increasing time after plasma treatment.</li>
</ol>
3. Microfluidic Plumbing Preparation
<ol>
	<li>Cut lengths of polymer tubing (inner diameter (ID) 0.02 inch, outer diameter (OD) 0.06 inch) to appropriate lengths to reach from the syringe pump to the switching apparatus (if used) and the device when mounted on the microscope. Cut as many lengths as there are microfluidic lanes.</li>
	<li>Assemble Y junctions for pinch valves (if using) by cutting and attaching 2-cm lengths of flexible silicone tubing (0.03 inch ID, 0.065 inch OD) to the top barbs of the &#34;Y&#34; and 1-cm lengths of silicone tubing to the bottom barb of the &#34;Y.&#34;</li>
	<li>Cut 10-cm (or appropriate) lengths of polymer tubing; connect them at one end to the switch junction by inserting them into the 1-cm silicone tubing segments on the bottom barb of the &#34;Y.&#34;&#160;Connect them at the other end to 21G needles that have been removed from their plastic syringe adaptors and bent approximately 90&#176; approximately 9 mm from the needle end.</li>
	<li>Connect 21G blunt needles to one end of the tubing segments that will run from the syringes to the switch apparatus by inserting the needles into the tubing. Insert the other ends of each length of tubing into the silicone tubing segments on the inlet barbs of the Y junctions. 
	NOTE: Use some sort of apparatus to hold the pinch valves and junctions in place; this can easily be made from a micropipette tip box (Figure 2D).</li>
	<li>Cut lengths of polymer tubing to carry waste from the outlet of the device to a waste beaker. Connect them at one end to bent 21G needles prepared as described above. Tape the other end of the tubes into a waste beaker.</li>
	<li>Prepare appropriate volumes of media containing 0.1 mg/mL bovine serum albumin (BSA) and fill the desired size and number of syringes. Connect the BSA-loaded syringes to the prepared 21G needles attached to the inlet tubing and load syringes into syringe pumps. 
	NOTE: For typical experiments 5 channels of the device are used, each with a 20-mL syringe. An experiment in which the medium is switched will require 2 banks of 5 syringes each. Here, the pump containing the first bank is referred to as the phase-1 pump, and the pump containing the second bank, with the post-switch medium, is referred to as the phase-2 pump.</li>
	<li>Purge air from the inlet plumbing by running the syringe pumps at a high flow rate (&#62;500 &#181;L/min), beginning by orienting the syringe pumps vertically and tapping the syringes to bring air bubbles to the top. Once air has been purged from the syringes, place the syringe pump horizontally and progressively tap or flick the polymer lines from the syringes up through the switch apparatus to dislodge and purge air bubbles. Repeat this process for the second bank of syringes (if switching media).</li>
	<li>After air bubbles are purged, set the phase-2 syringe pump (i.e., with the medium that will be used second, after the switch) to 1.5 &#181;L/min, then pause the flow. Place small binder clips onto the segments of flexible tubing on the branch of the Y junctions corresponding to the second medium phase. Continue to run the phase-1 syringe pump at a modest flow rate (e.g., 10 &#181;L/min) for at least 30 min to purge the second medium from the inlet tubing downstream of the Y junction.</li>
</ol>
4. Cell Culture Preparation
<ol>
	<li>Grow a preculture of the strain of interest in the desired medium overnight, to stationary phase. The bacterial strain should contain a mutation that renders the cells immobile, so that the cells do not swim out of the channels of the device. 
	NOTE: In this protocol, the bacterial strain is B. subtilis 3610 containing a hagA233V substitution (this mutation causes the flagellum to be straight rather than helical, preventing cell motility), a PrsbV-mNeonGreen reporter for cell stress, and a constitutive Phyperspank-mNeptune (red) reporter to highlight cells. LB Lennox medium is used in the example protocol. Typical strains for microfluidics experiments contain one or more fluorescent transcriptional reporters and a constitutively produced, cytoplasmic fluorescent protein to facilitate automated detection of cells. Growth defects were not observed when LB Lennox rich medium was used, but B. subtilis growth in minimal media may be inhibited under microfluidic conditions because secreted siderophores are washed away by the medium flow. Under such circumstances, growth may be restored by the addition of sodium citrate and ferric chloride to the medium, as reported for S750 medium11.</li>
	<li>The morning of the experiment, dilute the B. subtilis cells 1:50 into a baffled 250-mL flask containing 25 mL of growth medium. Grow the cells for 5 - 6 h in shaking culture at 37 &#176;C to an optical density of greater than 1. 
	NOTE: A dense cell culture is preferable because the stationary-phase B. subtilis cells are smaller and less often found in cell chains, facilitating device loading. Different bacterial species may require other medium or growth conditions for optimal loading.</li>
	<li>Using a syringe fitted with a 5-&#181;m pore-size filter, filter approximately 15 mL of cell culture into a 15-mL conical tube to remove B. subtilis cell chains (it is unnecessary for E. coli and may not be necessary with other species). 
	NOTE: Relatively few cells should be removed; the filtrate should be turbid.</li>
	<li>Centrifuge the filtered culture for 10 min at 4,000 x g at room temperature. Pour off the supernatant and resuspend the cell pellet in the residual supernatant fluid remaining in the tube, adding approximately 500 &#181;L of fresh medium if necessary to facilitate pipetting the cell suspension.</li>
</ol>
5. Device Loading
<ol>
	<li>Gently place empty thin gel-loading micropipette tips (see Table of Materials) into the outlet holes of the microfluidic device.</li>
	<li>Using thin gel-loading tips with a P200 micropipette, passivate the device by injecting medium containing 1 mg/mL BSA into the inlet holes, observing the tips in the outlet holes to monitor filling of the microfluidic channels (Figure 2A). Incubate the device at room temperature for approximately 5 min. 
	NOTE: BSA is commonly used as a blocking or passivation agent that will bind to the hydrophobic surface of the PDMS polymer, increasing its hydrophilicity and reducing the binding of other proteins or of cells (via cell surface molecules).</li>
	<li>Using thin gel-loading tips, load each channel with the resuspended cells (from Section 4), using the tips in the outlet channel to monitor the progress of the cells through the device (Figure 2B). 
	NOTE: Loading is facilitated by setting the P200 micropipette to its maximum 200-&#181;L volume and then aspirating a cushion of air before aspirating a small volume (approximately half the volume of the thin section of the pipette tip) of cells. The volume of resuspended cells need not be precise, as the volume of the PDMS device is small relative to that of the micropipette. We know of no upper limit to the density of the resuspended cells, as long as the cell suspension can be pipetted into the device. Cell clumps can clog the device and should be avoided by thorough pipetting and/or vortex mixing.</li>
	<li>Gently remove the gel-loading tips from the outlet holes, working one at a time to prevent damage to the device.</li>
	<li>Centrifuge the device in a bench-top microcentrifuge in an appropriate rotor adaptor at approximately 6,000 x g for 10 min (Figure 2C). 
	NOTE: A custom-machined aluminum rotor adaptor that was designed to fit into a microcentrifuge rotor (Figure 2C) was used; different centrifuge models may be used with appropriate rotor adaptors. The important feature of the adaptor is that it provides lateral force in the direction of the closed ends of the cell trenches, thereby forcing cells into the side channels.</li>
	<li>Verify successful loading under the microscope (Figure 3) but without affixing the device to the slide holder/stage insert. 
	NOTE: In this protocol, a 60X, 1.4 NA Ph3 oil-immersion objective is used for both verification at this stage and for subsequent imaging.</li>
</ol>
6. Device Assembly, Equilibration, and Mounting
<ol>
	<li>Carefully mount the loaded device onto a stage insert by taping the cover glass on either side of the PDMS to the bottom of the stage insert (i.e., invert the stage insert before attaching the device). Invert the stage insert-device assembly so that the PDMS is facing up and place on a soft surface, such as a dust-free wipe (Figure 2D).</li>
	<li>Adjust the flow rate of the phase-1 syringe pump to 35 &#181;L/min. Working with one lane at a time, insert the inlet needle and then the outlet needle (i.e., running to the waste beaker; Figure 2E).</li>
	<li>Wipe away any excess medium with a clean dust-free wipe and visually inspect the device for leaks. Examine the outlet tubing for the appearance of excess cells (typically appearing as a turbid stripe) and the medium meniscus, which will slowly move toward the waste beaker.</li>
	<li>Permit the device to run at 35 &#181;L/min for approximately 15 - 30 min, until all the connected lanes are draining into the waste beaker.</li>
	<li>Set the phase-1 syringe pump to 1.5 &#181;L/min and pause the flow. Bring the entire pump and device apparatus to the microscope (this process is aided by placing it all on a rolling cart) and restart the phase-1 medium flow at 1.5 &#181;L/min. Carefully mount the stage insert with the device onto an inverted fluorescence microscope, using tape as necessary to route the inlet and outlet tubing.</li>
	<li>Locate desired positions on the device for imaging and begin imaging as desired. Note that cells typically require a few hours (approximately 10 generations) to reach steady-state exponential-phase growth, and the onset of image acquisition may be delayed as desired so that imaging begins after the equilibration period. 
	NOTE: The steady-state growth of cells held in exponential phase should be verified during subsequent image analysis by tracking cell division times and ensuring that they have reached a constant minimum value.</li>
</ol>
7. Medium Switching
<ol>
	<li>Between successive rounds of imaging, pause the flow of the phase-1 syringe pump. Carefully unclip the binder clips from the phase-2 silicone tubing, moving each to the silicone tubing on the phase-1 branch of the Y junction. Un-pause the flow of the phase-2 syringe pump (which was set to 1.5 &#181;L/min in step 3.8) and continue imaging. 
	NOTE: At 1.5 &#181;m/min with a 10-cm length of polymer tubing downstream of the Y junctions, the second-phase medium reaches the device in approximately 50 min. The time to the device can be empirically tested using media that contain fluorescent tracer particles.</li>
</ol>

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The authors have nothing to disclose.

This microfluidics protocol is flexible in that many of the steps may be modified to optimize its use with a particular species or strain or for a specific purpose. Indeed, in this protocol we have made modifications to the original &#34;mother machine&#34; concept4 to optimize its use with B. subtilis. Often, the trenches in which the cells are confined constitute a single-layer feature, whereas in this protocol we use two-layer cell trenches, with a shallow channel surrounding the cells...

One of the most striking features of life on Earth is its great resilience and variety. A central goal of molecular biology is to understand the logic by which cells use genes and proteins to maximize their growth and fitness under a wide variety of environmental conditions. To achieve this goal, scientists must be able to confidently observe how individual cells grow, divide, and express their genes under a given set of conditions, noting how cells respond to subsequent changes in their environment. However, traditional analytical methods in microbiology have technical limitations that affect the types of questions that can be addressed. For example, bulk culture-based analyses have been very useful over the years, yet they offer only population-level data that can mask meaningful cell-to-cell variations or the behaviors of smaller sub-populations of cells in the total population. Single-cell analyses of living bacteria based on light microscopy reveal single-cell behavior but are also technically limited. Bacteria are typically immobilized on agarose pads containing growth medium, but cell growth and division crowds the microscopic view and depletes the available nutrients after just a few cell cycles, substantially limiting the observation time1,2. Moreover, the local depletion of nutrients and the concomitant buildup of metabolic byproducts due to cell growth are constantly changing the local cell growth environment in ways that are difficult to measure or predict. Such environmental changes using agarose pads pose a challenge to studies of steady-state behaviors or of cellular responses to specific changes in growth conditions3.
Microfluidic technology, in which a liquid medium is continuously flowed through microfabricated devices, offers a solution to classic experimental limitations. A microfluidic device can keep individual cells in position for live-cell microscopy while the flow of growth medium constantly provides cells with fresh nutrients and washes away metabolic byproducts and excess cells, thereby creating a highly uniform growth environment. Under constant growth conditions, cell behaviors can be observed in isolation from the influence of environmental factors, permitting an unimpeded view of the internal logic of cells. As the fluid flow prevents the microfluidic device from becoming crowded with cells, observation of single cell lineages for tens or hundreds of generations becomes possible4,5. Such long observation times permit the detection of otherwise undetectable long-term or rare cell behaviors. Finally, the composition of the medium that flows through the device can be altered at will, allowing cells to be observed as they respond to the onset of a stress or to the introduction or removal of a particular compound.
Microfluidics has already enjoyed a number of important applications. For instance, it has been used in tissue-, organ-, or body-on-a-chip devices, in which multiple human cell types are co-cultured to simulate an in vivo condition6; for the study of nematode movement in microstructured environments7; to examine interactions among bacterial biofilms (e.g., 8); and for the encapsulation and manipulation of tiny volumes of cells or chemicals (e.g., 9). Microfluidic devices have also become increasingly popular in the field of microbiology (for excellent reviews, see 10 and 11), especially as their physical and flow properties are well-matched to natural microbial niches12. For instance, microfluidics has been recently employed by microbiologists for such purposes as precisely measuring cell growth and division13,14,15, analyzing pathogen movement16, monitoring quorum sensing17 and physiological transitions18, and for protein counting19, among many other examples. The method presented here is specifically designed for the analysis of single bacterial cell lineages rather than combinations of strains or species. The microfluidic device demonstrated here utilizes one variation of the &#34;mother machine&#34; design4, in which cells are grown single-file within a microfluidic trench with one closed end and one open end; cell growth and division pushes progeny cells up and out of the open end into the fluid flow. Our analyses typically focus only on the &#34;mother&#34; cell that is confined at the closed end of the trench. We consider this method as an advancement over previous light microscopy-based single-cell analytical techniques, such as cell immobilization on agarose pads. While B. subtilis is used as a model here, the method is also applicable to other bacterial species (Escherichia coli is another common model; some species with different cell sizes or morphologies may require the fabrication of new devices with different dimensions). The use of fluorescent reporters to mark cells and to visualize changes in gene expression requires the use of genetically tractable species; however, analyses of cell growth and morphology are possible even without fluorescent markers.
The present protocol excludes the process of fabricating the silicon master using photolithography, which has been extensively described elsewhere5; masters can also be easily outsourced from microfabrication facilities. It includes the molding of a PDMS device from a reinforced silicon master; bonding the device to a piece of cover glass; assembling the microfluidic inlet and outlet plumbing, including pinch valves to permit medium switching; passivating the device, preparing bacterial cells, and loading the device with bacterial cells; attaching the plumbing to the device and equilibrating the cells; and loading the device onto a fluorescence microscope for imaging. Because many different image acquisition and processing software tools can be used to visualize and analyze different data of interest4,5, example images are shown, but image-capture methods are not included in this protocol.

<table border="0" cellpadding="0" cellspacing="0" width="580">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Sylgard 184 Silicone Elastomer Kit</td>
			<td style="width:71px;">Dow Corning</td>
			<td style="width:119px;">(240)4019862</td>
			<td style="width:167px;">This is referred to as PDMS in the protocol. There are 2 components that are mixed at a 10:1 ratio</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:223px;">0.75-mm biopsy punch</td>
			<td style="width:71px;">World Precision Instruments</td>
			<td align="right" style="width:119px;">504529</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">22 x 40 mm No. 1.5 cover glass</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">48393 172</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Plasma Etch PE-50</td>
			<td style="width:71px;">Plasma Etch Inc.</td>
			<td style="width:119px;">PE-50</td>
			<td>Instrument used to bond PDMS to glass using oxygen plasma treatment</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:223px;">Tygon flexible tubing ID 0.02&#34;, OD 0.06&#34;</td>
			<td style="width:71px;">Saint-Gobain PPL Corp.</td>
			<td>AAD04103</td>
			<td>For the main part of the tubing, e.g. attached to the needles</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:223px;">Silicone Tubing, 0.04&#34; ID, 0.085&#34; OD</td>
			<td style="width:71px;">HelixMark Standard Silicone Tubing</td>
			<td style="width:119px;">60-795-05</td>
			<td>For pinch valves and Y-junction connection</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Bovine Serum Albumin</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">A7906-100G</td>
			<td>Used as passivation agent</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:223px;">ART Gel Pipet Tips (P200)</td>
			<td style="width:71px;">Molecular BioProducts</td>
			<td align="right" style="width:119px;">2155</td>
			<td>For loading the device with medium and cells</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Acrodisc 32-mm syringe filter with 5-&#956;m Supor membrane</td>
			<td style="width:71px;">Pall Life Sciences</td>
			<td align="right" style="width:119px;">4560</td>
			<td>For filtering cultures before device loading</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">21-ga blunt needles, 1&#34;</td>
			<td style="width:71px;">McMaster-Carr</td>
			<td style="width:119px;">75165A681</td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:223px;">Y connector with 200-series barbs, 1/16&#34; ID tubing, polypropylene</td>
			<td style="width:71px;">Nordson Medical</td>
			<td style="width:119px;">Y210-6005</td>
			<td>The company is AKA Value Plastics</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Eclipse Ti-E inverted microscope</td>
			<td style="width:71px;">Nikon</td>
			<td style="width:119px;"></td>
			<td>This unit has been discontinued by the manufacturer and is replaced by the Ti 2 E.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">LB Lennox</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">L3022</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Microfuge 18</td>
			<td style="width:71px;">Beckman Coulter</td>
			<td align="right">367160</td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:223px;">Bacillus subtilis strain NCIB3610</td>
			<td style="width:71px;">Bacillus Genetic Stock Center</td>
			<td style="width:119px;">3A1</td>
			<td>wild-type parental strain modified for use in the experiments shown in this protocol</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Gravity convection oven</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">414005-106</td>
			<td>for curing PDMS and baking assembled devices</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Scotch-Weld Epoxy Kit</td>
			<td style="width:71px;">3M</td>
			<td style="width:119px;">2216 B/A</td>
			<td>may be used to bond a silicon wafer to an aluminum backing plate</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Scotch Magic tape, 3/4&#34; width</td>
			<td style="width:71px;">3M</td>
			<td style="width:119px;"></td>
			<td>to clean dust from PDMS devices and to place over features to increase visibility (denoted &#34;office tape&#34; or &#34;adhesive tape&#34; in protocol)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Stereomicroscope</td>
			<td style="width:71px;">Nikon</td>
			<td style="width:119px;">SMZ800N</td>
			<td>listed here as an example; nearly any stereomicroscope will do</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Isopropyl alcohol</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">W292907</td>
			<td>To clean cover glass</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:223px;">Syring pump, 6 channel</td>
			<td style="width:71px;">New Era Pump Systems Inc.</td>
			<td style="width:119px;">NE-1600</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Kimwipes</td>
			<td style="width:71px;">Kimberly-Clark</td>
			<td align="right" style="width:119px;">34120</td>
			<td>a brand of dust-free wipes</td>
		</tr>
	</tbody>
</table>

single-cell microfluidic analysis of bacillus subtilis

Microfluidic technology overcomes many of the limitations to traditional analytical methods in microbiology. Unlike bulk-culture methods, it offers single-cell resolution and long observation times spanning hundreds of generations; unlike agarose pad-based microscopy, it has uniform growth conditions that can be tightly controlled. Because the continuous flow of growth medium isolates the cells in a microfluidic device from unpredictable variations in the local chemical environment caused by cell growth and metabolism, authentic changes in gene expression and cell growth in response to specific stimuli can be more confidently observed. Bacillus subtilis is used here as a model bacterial species to demonstrate a &#34;mother machine&#34;-type method for cellular analysis. We show how to construct and plumb a microfluidic device, load it with cells, initiate microscopic imaging, and expose cells to a stimulus by switching from one growth medium to another. A stress-responsive reporter is used as an example to reveal the type of data that may be obtained by this method. We also briefly discuss further applications of this method for other types of experiments, such as analysis of bacterial sporulation.

Successful initial cell loading, as assessed by phase-contrast microscopy before attaching the microfluidic plumbing to the device, would be considered as having all or nearly all of the microfluidic side channels containing one or more bacterial cells (Figure 3A). Optimal loading would show several cells in each channel, but channels will nonetheless fill with cells due to cell growth during the equilibration period (Figure 3B)....

Watch this Scientific Journal Video about Single-cell Microfluidic Analysis of Bacillus subtilis at JoVE.com

Single-cell Microfluidic Analysis of Bacillus subtilis

We present a method for the microfluidic analysis of individual bacterial cell lineages using Bacillus subtilis as an example. The method overcomes shortcomings of traditional analytical methods in microbiology by allowing observation of hundreds of cell generations under tightly controllable and uniform growth conditions.

Single-cell Microfluidic Analysis of Bacillus subtilis

Microfluidic technology overcomes many of the limitations to traditional analytical methods in microbiology. Unlike bulk-culture methods, it offers ...

single-cell-microfluidic-analysis-of-bacillus-subtilis

Research

JoVE Journal

Biology

11.9K Views.  Oklahoma State University. We present a method for the microfluidic analysis of individual bacterial cell lineages using Bacillus subtilis as an example. The method overcomes shortcomings of traditional analytical methods in microbiology by allowing observation of hundreds of cell generations under tightly controllable and uniform growth conditions.

Video: Single-cell Microfluidic Analysis of Bacillus subtilis

Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis

Both prokaryotes and eukaryotes respond to DNA damage through a complex set of physiological changes. Alterations in gene expression, the redistribution of existing proteins, and the assembly of new protein complexes can be stimulated by a variety of DNA lesions and mismatched DNA base pairs. Fluorescence microscopy has been used as a powerful experimental tool for visualizing and quantifying these and other responses to DNA lesions and to monitor DNA replication status within the complex subcellular architecture of a living cell. Translational fusions between fluorescent reporter proteins and components of the DNA replication and repair machinery have been used to determine the cues that target DNA repair proteins to their cognate lesions in vivo and to understand how these proteins are organized within bacterial cells. In addition, transcriptional and translational fusions linked to DNA damage inducible promoters have revealed which cells within a population have activated genotoxic stress responses. In this review, we provide a detailed protocol for using fluorescence microscopy to image the assembly of DNA repair and DNA replication complexes in single bacterial cells. In particular, this work focuses on imaging mismatch repair proteins, homologous recombination, DNA replication and an SOS-inducible protein in Bacillus subtilis. All of the procedures described here are easily amenable for imaging protein complexes in a variety of bacterial species.

Imaging Mismatch Repair and Cellular Responses to DNA Damage in Bacillus subtilis

A detailed protocol is described for imaging the real time formation of DNA repair complexes in Bacillus subtilis cells.

Both prokaryotes and eukaryotes respond to DNA damage through a complex set of physiological changes. Alterations in gene expression, the redistribution ...

Single Cell Transfection in Chick Embryos

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a thorough analysis of the fates of single cells under normal and experimental conditions. To this end, transfection methods that target single progenitors are a prerequisite. We describe here a technically straightforward method for transfecting single cells in chicken tissues in-ovo, allowing reliable lineage tracing as well as genetic manipulation. Specific tissue domains are targeted within the somite or neural tube, and DNA is injected directly into the epithelium of interest, resulting in sporadic transfection of single cells. Using reporters, clonal populations may consequently be traced for up to three days, and behavior of genetically manipulated clonal populations can be compared with that of controls. This method takes advantage of the accessibility of the chick embryo along with emerging tools for genetic manipulation. We compare and discuss its advantages over the widely-used electroporation method, and possible applications and use in additional in-vivo models are also suggested. We advocate the use of this method as a significant addition and complement for existing lineage tracing and genetic interference tools.

Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations.

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Analysis of Single-cell Gene Transcription by RNA Fluorescent In Situ Hybridization (FISH)

Adhesion of Plasmodium falciparum infected erythrocytes (IE) to human endothelial receptors during malaria infections is mediated by expression of PfEMP1 protein variants encoded by the var genes.
The haploid P. falciparum genome harbors approximately 60 different var genes of which only one has been believed to be transcribed per cell at a time during the blood stage of the infection. How such mutually exclusive regulation of var gene transcription is achieved is unclear, as is the identification of individual var genes or sub-groups of var genes associated with different receptors and the consequence of differential binding on the clinical outcome of P. falciparum infections. Recently, the mutually exclusive transcription paradigm has been called into doubt by transcription assays based on individual P. falciparum transcript identification in single infected erythrocytic cells using RNA fluorescent in situ hybridization (FISH) analysis of var gene transcription by the parasite in individual nuclei of P. falciparum IE1.
Here, we present a detailed protocol for carrying out the RNA-FISH methodology for analysis of var gene transcription in single-nuclei of P. falciparum infected human erythrocytes. The method is based on the use of digoxigenin- and biotin- labeled antisense RNA probes using the TSA Plus Fluorescence Palette System2 (Perkin Elmer), microscopic analyses and freshly selected P. falciparum IE. The in situ hybridization method can be used to monitor transcription and regulation of a variety of genes expressed during the different stages of the P. falciparum life cycle and is adaptable to other malaria parasite species and other organisms and cell types.

Analysis of Single-cell Gene Transcription by RNA Fluorescent In Situ Hybridization FISH

Fluorescent in situ hybridization (FISH) to identify mRNA transcripts in individual cells allows analysis of polygenic activity such as the simultaneous transcription of more than one member of the var multigene family in Plasmodium falciparum infected erythrocytes 1. The technique is adaptable and can be used on different types of genes, cells and organisms.

Adhesion of Plasmodium falciparum infected erythrocytes (IE) to human endothelial receptors during malaria infections is mediated by expression of PfEMP1 ...

Single-cell Microinjection for Cell Communication Analysis

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a hemichannel by each neighboring cell to form the gap junction. In mammalian cells, the hemichannel is formed by six connexins, monomers with four transmembrane domains and a C and N terminal within the cytoplasm. Gap junctions permit the exchange of ions, second messengers, and small metabolites. In addition, they have important roles in many forms of cellular communication within physiological processes such as synaptic transmission, heart contraction, cell growth and differentiation. We detail how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells. It is expected that in functional gap junctions, the dye will diffuse from the loaded cell to the connected cells. It is a very useful technique to study gap junctions since you can evaluate the diffusion of the fluorescence in real time. We discuss how to prepare the cells and the micropipette, how to use a micromanipulator and inject a low molecular weight fluorescent dye in an epithelial cell line.

We describe here how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells, and provide some useful tips. We expect that this paper will help everyone to evaluate the degree of cellular coupling due to functional gap junctions. Everything described here could be, in principle, adapted to other fluorescent dyes with molecular weight below 1,000 Daltons.

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a ...

Analysis of Global RNA Synthesis at the Single Cell Level following Hypoxia

Hypoxia or lowering of the oxygen availability is involved in many physiological and pathological processes. At the molecular level, cells initiate a particular transcriptional program in order to mount an appropriate and coordinated cellular response. The cell possesses several oxygen sensor enzymes that require molecular oxygen as cofactor for their activity. These range from prolyl-hydroxylases to histone demethylases. The majority of studies analyzing cellular responses to hypoxia are based on cellular populations and average studies, and as such single cell analysis of hypoxic cells are seldom performed. Here we describe a method of analysis of global RNA synthesis at the single cell level in hypoxia by using Click-iT RNA imaging kits in an oxygen controlled workstation, followed by microscopy analysis and quantification.&nbsp; Using cancer cells exposed to hypoxia for different lengths of time, RNA is labeled and measured in each cell. This analysis allows the visualization of temporal and cell-to-cell changes in global RNA synthesis following hypoxic stress.

We describe a technique for analysis of global RNA synthesis in hypoxia using imaging. Click-chemistry labeling of RNA has not previously been performed under hypoxia and allows visualization of global RNA changes at the single cell level. This approach complements the existing averaged RNA techniques, allowing direct visualization of cell-to-cell changes in global RNA synthesis.

Hypoxia or lowering of the oxygen availability is involved in many physiological and pathological processes. At the molecular level, cells initiate a ...

Applications of the Single-probe: Mass Spectrometry Imaging and Single Cell Analysis under Ambient Conditions

Mass spectrometry imaging (MSI) and in-situ single cell mass spectrometry (SCMS) analysis under ambient conditions are two emerging fields with great potential for the detailed mass spectrometry (MS) analysis of biomolecules from biological samples. The single-probe, a miniaturized device with integrated sampling and ionization capabilities, is capable of performing both ambient MSI and in-situ SCMS analysis. For ambient MSI, the single-probe uses surface micro-extraction to continually conduct MS analysis of the sample, and this technique allows the creation of MS images with high spatial resolution (8.5 &#181;m) from biological samples such as mouse brain and kidney sections. Ambient MSI has the advantage that little to no sample preparation is needed before the analysis, which reduces the amount of potential artifacts present in data acquisition and allows a more representative analysis of the sample to be acquired. For in-situ SCMS, the single-probe tip can be directly inserted into live eukaryotic cells such as HeLa cells, due to the small sampling tip size (&lt; 10 &#181;m), and this technique is capable of detecting a wide range of metabolites inside individual cells at near real-time. SCMS enables a greater sensitivity and accuracy of chemical information to be acquired at the single cell level, which could improve our understanding of biological processes at a more fundamental level than previously possible. The single-probe device can be potentially coupled with a variety of mass spectrometers for broad ranges of MSI and SCMS studies.

Here, we present protocols to perform both ambient mass spectrometry imaging (MSI) of tissues and in-situ live single cell MS (SCMS) analysis using the single-probe, which is a miniaturized multifunctional device for MS analysis.

Mass spectrometry imaging (MSI) and in-situ single cell mass spectrometry (SCMS) analysis under ambient conditions are two emerging fields with great ...

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

Plasma sterilization is a promising alternative to conventional sterilization methods for industrial, clinical, and spaceflight purposes. Low pressure plasma (LPP) discharges contain a broad spectrum of active species, which lead to rapid microbial inactivation. To study the efficiency and mechanisms of sterilization by LPP, we use spores of the test organism Bacillus subtilis because of their extraordinary resistance against conventional sterilization procedures. We describe the production of B. subtilis spore monolayers, the sterilization process by low pressure plasma in a double inductively coupled plasma reactor, the characterization of spore morphology using scanning electron microscopy (SEM), and the analysis of germination and outgrowth of spores by live cell microscopy. A major target of plasma species is genomic material (DNA) and repair of plasma-induced DNA lesions upon spore revival is crucial for survival of the organism. Here, we study the germination capacity of spores and the role of DNA repair during spore germination and outgrowth after treatment with LPP by tracking fluorescently-labelled DNA repair proteins (RecA) with time-resolved confocal fluorescence microscopy. Treated and untreated spore monolayers are activated for germination and visualized with an inverted confocal live cell microscope over time to follow the reaction of individual spores. Our observations reveal that the fraction of germinating and outgrowing spores is dependent on the duration of LPP-treatment reaching a minimum after 120 s. RecA-YFP (yellow fluorescence protein) fluorescence was detected only in few spores and developed in all outgrowing cells with a slight elevation in LPP-treated spores. Moreover, some of the vegetative bacteria derived from LPP-treated spores showed an increase in cytoplasm and tended to lyse. The described methods for analysis of individual spores could be exemplary for the study of other aspects of spore germination and outgrowth.

Investigating the Detrimental Effects of Low Pressure Plasma Sterilization on the Survival of Bacillus subtilis Spores Using Live Cell Microscopy

This protocol illustrates the important consecutive steps required to assess the relevance of monitoring vitality parameter and DNA repair processes in reviving Bacillus subtilis spores after treatment with low pressure plasma by tracking fluorescence-labelled DNA repair proteins via time-resolved confocal microscopy and scanning electron microscopy.

Plasma sterilization is a promising alternative to conventional sterilization methods for industrial, clinical, and spaceflight purposes. Low pressure ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to ...

Single-Cell Analysis of the Expression of Pseudomonas syringae Genes within the Plant Tissue

A plethora of pathogenic microorganisms constantly attack plants. The Pseudomonas syringae species complex encompasses Gram-negative plant-pathogenic bacteria of special relevance for a wide number of hosts. P. syringae enters the plant from the leaf surface and multiplies rapidly within the apoplast, forming microcolonies that occupy the intercellular space. The constitutive expression of fluorescent proteins by the bacteria allows for visualization of the microcolonies and monitoring of the development of the infection at the microscopic level. Recent advances in single-cell analysis have revealed the large complexity reached by clonal isogenic bacterial populations. This complexity, referred to as phenotypic heterogeneity, is the consequence of cell-to-cell differences in gene expression (not linked to genetic differences) among the bacterial community. To analyze the expression of individual loci at the single-cell level, transcriptional fusions to fluorescent proteins have been widely used. Under stress conditions, such as those occurring during colonization of the plant apoplast, P. syringae differentiates into distinct subpopulations based on the heterogeneous expression of key virulence genes (i.e., the Hrp type III secretion system). However, single-cell analysis of any given P. syringae population recovered from plant tissue is challenging due to the cellular debris released during the mechanical disruption intrinsic to the inoculation and bacterial extraction processes. The present report details a method developed to monitor the expression of P. syringae genes of interest at the single-cell level during the colonization of Arabidopsis and bean plants. The preparation of the plants and the bacterial suspensions used for inoculation using a vacuum chamber are described. The recovery of endophytic bacteria from infected leaves by apoplastic fluid extraction is also explained here. Both the bacterial inoculation and bacterial extraction methods are empirically optimized to minimize plant and bacterial cell damage, resulting in bacterial preparations optimal for microscopy and flow cytometry analysis.

Single-Cell Analysis of the Expression of Pseudomonas syringae Genes within the Plant Tissue

The present protocol describes a method that allows single-cell gene expression analysis on Pseudomonas syringae populations grown within the plant apoplast.

A plethora of pathogenic microorganisms constantly attack plants. The Pseudomonas syringae species complex encompasses Gram-negative plant-pathogenic ...