This work was funded by the National Heart, Lung, and Blood Institute (R01 HL073965, IL) and (T32 HL007829-24, ISF). The authors would also like to acknowledge the Scientific Machine Shop at the University of Illinois at Chicago for generating our latest MPP flow chambers.

The use of animals in our studies is approved by the University of Illinois at Chicago Animal Care Committee (#16-183).
1. Assembly of the Modified Parallel Plate Flow Chamber
NOTE: Please refer to Table 1 and Figure 1 for MPP flow chamber piece IDs. Please refer to Figure 1 for a schematic detailing the orientation of chamber pieces for assembly.
<ol>
	<li>To adhere the rectangular cover glass, piece D, to the bottom of piece C, first make the silicone elastomer solution by thoroughly mixing 500 &#181;L silicone elastomer curing agent into 5 mL of silicone elastomer base.</li>
	<li>Apply a thin layer of the silicone elastomer solution around the edges of the rectangular space of piece C and gently place the rectangular cover glass piece D directly on the elastomer solution such that piece D completely covers the open rectangular space of piece C. Carefully wipe away excess silicone elastomer solution.</li>
	<li>Repeat step 1.2 for adhering the rectangular cover glass, piece F, to the bottom of piece E and allow the silicone elastomer solution to cure over night at room temperature. 
	NOTE: Once cured, the rectangular cover glass will remain adhered for up to six months before needing to be replaced.</li>
	<li>Beginning with the bottom chamber piece, piece E, assemble the MPP flow chamber by sequentially placing each piece on top of the previous in the following order: piece E (bottom), piece C, piece B, piece A (top).</li>
	<li>Align the screw holes of each piece at the corners and tightly screw the pieces together to prevent leaks from occurring while administering flow to the MPP flow chamber.</li>
</ol>
2. Cell Preparation and Seeding into the MPP Flow Chamber
NOTE: Follow steps 2.1&#8722;2.7 for cultured endothelial cells. Follow the method detailed in steps 2.8&#8722;2.14 for isolating endothelial cells from the mouse mesenteric arterial arcade and preparation of freshly isolated endothelial cells.
<ol>
	<li>In a 6-well plate, place four to five 12 mm cover glass circles/well and seed cells between 10% and 30% confluency such that single cells can be accessed for electrophysiological recordings.</li>
	<li>Incubate cells under standard culture conditions (5% CO2, 37 &#176;C) for no less than 2 h to allow cells to adhere and no more than 24 h as endothelial cells in authors&#8217; experience become very flat and difficult to patch when seeded at sub-confluency for more than 24 h.</li>
	<li>Remove a cover glass containing adhered cells from a well of the 6-well plate, quickly rinse in phosphate-buffered saline (PBS), and transfer to a 35 mm Petri dish containing 2 mL electrophysiological bath solution (Table 2) prior to transfer to the MPP flow chamber.</li>
	<li>Transfer the cover glass circle to the rectangular cover glass, piece D, which is adhered to piece C of the MPP flow chamber being sure that adequate solution stays on the cover glass so that cells to not become exposed to air. Add the desired bath solution (~250 &#181;L) to the cells so that the cover glass circle and cells are completely submerged in solution.</li>
	<li>Position the cover glass circle such that it rests in the half closest to the vacuum reservoir side so that cells will be in line with the slit openings of piece B. Ensure that the glass cover circle adheres to piece D through solution-glass adhesion so that application of flow to the chamber does not disrupt the position of the cover glass circle.</li>
	<li>Assemble the MPP flow chamber by screwing the pieces together in the appropriate order, as described in steps 1.4 and 1.5 and as shown in Figure 1. Transfer the chamber to the microscope stage and immediately perfuse the chamber with bath solution such that solution reaches the vacuum reservoir for aspiration (~10 mL).</li>
	<li>Identify a healthy cell for the experiment by identifying a cell with a dark border and obvious nucleus. Avoid cells that appear to be blebbing or cells that are in contact with other cells. 
	NOTE: In the authors&#8217; laboratory, human aortic endothelial cells and primary mouse mesenteric endothelial cells in culture are used. However, any other type of adherent cell that is of interest to specific research needs can be used in the same way.</li>
	<li>Wash the isolated arterial arcade in dissociation solution (Table 2). Transfer the arcade to a 2 mL centrifuge tube containing 2 mL of pre-warmed (37 &#176;C) dissociation solution (recipe for dissociation solution shown in Table 2) containing neutral protease (0.5 mg/mL) and elastase (0.5 mg/mL). Incubate for 1 h at 37 &#176;C with gentle shaking every 10 min.</li>
	<li>Remove 1 mL of the neutral protease/elastase dissociation solution and add 1 mg/mL collagenase type 1. Return the collagenase solution to the solution containing the arteries for a final collagenase type 1 concentration of 0.5 mg/mL. Incubate for 2&#8722;3 min at 37 &#176;C.</li>
	<li>Using 5 grade forceps, quickly move the arcade onto a 35 mm diameter Petri dish containing a 750 &#181;L drop of fresh, chilled dissociation solution. Further dissociate the tissue by using two 20 G syringe needles to mechanically liberate endothelial cells from enzymatically digested arteries.</li>
	<li>Using a 9&#8221; disposable Pasteur glass pipet, triturate the cell solution 10x before transferring the cells to a new 1.5 mL centrifuge tube using the glass pipet.</li>
	<li>Wash the Petri dish with another 750 &#181;L of dissociation solution and transfer to the same tube. Using the glass pipette, further mechanically disperse the cells by pipetting at least 10x to create a single cell suspension being careful not to introduce bubbles that may damage endothelial cell integrity.</li>
	<li>Add 750 &#181;L of the endothelial cell suspension (~500&#8722;1,000 cells) directly to piece D of the MPP flow chamber on the half closest to the reservoir vacuum side. Allow the endothelial cells to adhere between 45 min and 1 h at room temperature.</li>
	<li>Assemble the MPP flow chamber by screwing the pieces together in the appropriate order as described in steps 1.4 and 1.5 and as shown in Figure 1. Transfer the chamber to the microscope stage and immediately perfuse the chamber with bath solution such that solution reaches the vacuum aspiration (~10 mL). Identify accessible endothelial cells by their rough and round phenotype19,20. 
	NOTE: A variety of digestion methods and enzyme cocktails have been used to isolate endothelial cells from different arterial beds. See Table 3 for detailed descriptions of protocols that have been used by a variety of investigators to isolate endothelial cells for patch clamp electrophysiology of mechanosensitive ion channels. These methods are likely suitable for use in combination with the MPP flow chamber.</li>
</ol>
3. Controlling Shear Stress to the MPP Flow Chamber for Electrophysiological Recordings of Shear-activated Mechanosensitive Ion Channels
<ol>
	<li>Set-up a gravity perfusion system by connecting a 30 mL graduated syringe cylinder to a 3-way luer lock fitted with microbore tubing (internal diameter: 0.05 inch, outer diameter: 0.09 inch) suited for insertion into the 3 mm diameter inlet hole of piece A of the MPP chamber.</li>
	<li>Attach the graduated cylinder to the outer face of the Faraday cage surrounding the electrophysiology rig (Figure 2) using double-sided tape. Prior to inserting the tubing in the MPP chamber, pre-fill the syringe and tubing with bath solution (see Table 2 for bath solution used for investigating inwardly rectifying K+ channels in endothelial cells). Insert the tubing into the MPP flow chamber inlet hole of piece A.</li>
	<li>Pre-fill the MPP flow chamber with solution such that solution is being removed in the vacuum reservoir. Stop flow to the chamber and refill the graduated cylinder to the top mark. Calculate flow rates manually by allowing the solution to flow through the chamber and using a stop-watch to calculate mL/s at a given syringe cylinder height.</li>
	<li>Raise or lower the syringe to alter flow, and thus shear in the chamber, and continue this process until a desired level of shear stress is found.</li>
	<li>Calculate shear stress in a parallel chamber using the following equation21: 
	&#964; = 6&#181;Q/h2w 
	where &#181; = fluid viscosity (g/cm&#183;s), Q = flow rate (mL/s), and parallel plate chamber width (w = 2.2 cm) and height (h = 0.1 cm). 
	NOTE: In the current gravity perfusion system, at a syringe cylinder height (as measured from the top of the cylinder) of 57 cm above the microscope stage, the flow rate is 0.3 mL/s. The shear calculated in the chamber at this syringe cylinder height and flow rate is 0.7 dyn/cm2. It should also be noted that other perfusion systems, such as a peristaltic pump, can be used to control flow to the MPP flow chamber. However, these devices may add unwanted turbulence and influence stability of the electrophysiology measurements under flow, therefore, using the gravity perfusion system described here is recommended.</li>
	<li>Transfer an assembled chamber containing adhered cells to the microscope stage of the electrophysiology rig and insert the tubing pre-filled with bath solution into the hole of piece A. Simultaneously fill the chamber and wash the cells with 10 mL of bath solution by turning the 3-way luer lock such that solution flows to the chamber.</li>
	<li>Once the desired patch configuration is successfully obtained allow channel currents to stabilize in a static bath at room temperature. Once currents have stabilized, apply shear in a step-wise fashion allowing increases in current to stabilize prior to the next step increase in shear stress. 
	NOTE: The authors find the most success with the perforated patch configuration when studying mechanoactivated ion channels in endothelial cells. To perform perforated whole-cell patch configurations, add 5 &#181;L of a 60 mg/mL stock amphotericin B in dimethyl sulfoxide (DMSO) to 1 mL of 0.2 &#181;m sterile filtered pipette solution. After generating a giga-ohm seal in the cell-attached configuration, perforated whole-cell patches form within 2&#8722;5 min.</li>
	<li>Remove shear exposure to the cells by stopping flow to the chamber allowing mechanosensitive channel currents to return to baseline currents observed in the static bath.</li>
	<li>Isolate mechanosensitive ion channel currents of interest by altering solution valence (e.g., 60 mM K+ in bath solution with 0 Ca2+ in pipette solution to study inwardly rectifying K+ channels. Table 2 shows example solution recipes) and/or pharmacological inhibition of potentially contaminating current sources.</li>
</ol>

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The authors have nothing to disclose.

The vascular system is constantly exposed to active hemodynamic forces, which activate mechanosensitive ion channels3,22 but the physiological roles of these channels in shear stress-induced mechanotransduction is only starting to emerge4,6,8. The mechanisms responsible for the mechanosensitivity of shear stress-activated channels remain unknown. The protocol detailed he...

Hemodynamic forces generated by the blood flow are well known to play major roles in endothelial and vascular function1,2. It is also well known that several types of ion channels acutely respond to changes in shear stress3,4,5 leading to the hypothesis that ion channels can be primary shear stress sensors. More recently, we and others showed that mechanosensitive ion channels play critical roles in several shear-stress sensitive vascular functions, including the vasoactive response to shear stress6,7,8, and developmental angiogenesis9. The mechanisms of the shear-stress sensitivity of ion channels, however, are almost totally unknown. This gap of knowledge is likely to be due to the technical difficulty of performing electrophysiological recordings under well-defined shear stress. In this article, therefore, we provide a step by step detailed protocol routinely performed in our lab to achieve this goal6,7,10,11.
The overall goal of this method is to allow the real-time investigation of ion channel mechanoactivation under well-defined shear stress in the physiological range. This is achieved by modifying a standard parallel plate flow chamber to allow an electrophysiological pipette to be lowered into the chamber and access cells grown on the bottom plate during the real time exposure to flow, providing a unique approach to achieve this goal6,7,11. In contrast, standard parallel plate flow chambers, described in prior publications can be used for the real time imaging analysis of cells exposed to shear forces12 or other non-invasive approaches13,14 but not for electrophysiology. Similarly, the cone and plate apparatus, another powerful approach to expose cells to shear stress15,16 is also not suitable for electrophysiological recordings. Thus, these flow devices do not allow the investigation of shear stress sensitivity of ion channels. The difficulty in performing electrophysiological recordings under flow is the main reason for the paucity of information about the mechanisms responsible for these crucial effects.
In terms of the alternative approaches to achieve the same goal, there are none that are as accurate or controlled. Some earlier studies attempted to record ion channel activity under flow by exposing cells to a stream of liquid coming from another pipette brought to the vicinity of a cell from above17,18. This is highly non-physiological, as the mechanical forces generated under these conditions have little in common with the physiological profiles of shear stress in the blood vessels. Similar concerns apply to the attempts to simulate physiological shear stress by perfusion of open chambers. As discussed in detail in our earlier study10, an open liquid-air interface creates multiple disturbances and recirculation, which are non-physiological. To address all these concerns, we have designed a modified parallel plate (MPP) flow chamber, also referred to as the &#8220;minimally invasive flow device&#8221; in our earlier studies6,7,10,11, made from acrylic and extensively used in our lab. However, in spite of the fact that the original description of the design has been published almost 20 years ago and is the only flow device that allows performing electrophysiological recordings under well-defined shear stress, this methodology has not been adopted by other labs and there are only very few studies that attempt to record currents under flow. We believe, therefore, that providing a detailed description for using the MPP flow chamber will be of great help to researches who are interested in mechanosensitive ion channels and vascular biology.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 &#181;m sterile syringe filters</td>
			<td>VWR</td>
			<td>28145-501</td>
			<td>Used for filtering electrophysiolgoical pipette solution</td>
		</tr>
		<tr>
			<td>5 grade forceps</td>
			<td>Fine Scientific Tools</td>
			<td>1252-30</td>
			<td>Used for transferring digested arteries to fresh solution</td>
		</tr>
		<tr>
			<td>9&#34; Pasteur Pipet</td>
			<td>Fisher Scientifc</td>
			<td>13-678-20D</td>
			<td>Used for mechanically disrupting digested arteries and transferring freshly isolated endohtelial cells&#160;</td>
		</tr>
		<tr>
			<td>12 mm diameter Cover glass circles</td>
			<td>Fisher Scientifc</td>
			<td>12-545-80</td>
			<td>For use with studies involving cultured cells and multiple treatments. Cells adhered to the cover glass are used for patch clamp analyses</td>
		</tr>
		<tr>
			<td>24 x 40 mm Rectangluar Cover glass</td>
			<td>Sigma-Aldrich</td>
			<td>CLS2975224</td>
			<td>Cover glass to be added to MPP flow chamber pieces C (Figure 1)</td>
		</tr>
		<tr>
			<td>24 x 50 mm Rectangluar Cover glass</td>
			<td>Sigma-Aldrich</td>
			<td>CLS2975245</td>
			<td>Cover glass to be added to MPP flow chamber E (Figure 1)</td>
		</tr>
		<tr>
			<td>20 gauge syringe needles</td>
			<td>Becton Dickinson and Co</td>
			<td>305175</td>
			<td>For use in mechanical disruption of digested mesenteric arteries</td>
		</tr>
		<tr>
			<td>35 mm Petri dish</td>
			<td>Genesee Scientific</td>
			<td>32-103</td>
			<td>For use in mechanical disruption of digested mesenteric arteries</td>
		</tr>
		<tr>
			<td>Amphotericin B solubilized</td>
			<td>Sigma-Aldrich</td>
			<td>A9528-50MG</td>
			<td>Used for generating the perforated whole-cell patch configuration.</td>
		</tr>
		<tr>
			<td>collagenase, type I</td>
			<td>Worthington Biochemical</td>
			<td>100 mg - LS004194</td>
			<td>Enzyme used in our laboratory as a brief digestion following the initial cocktail of neutral protease and elastase</td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide (DMSO)</td>
			<td>Fisher Scientifc</td>
			<td>67-68-5</td>
			<td>Solvent for Amphotericin B used in perforated whole-cell patch clamp</td>
		</tr>
		<tr>
			<td>elastase, lyophilized</td>
			<td>Worthington Biochemical</td>
			<td>25 mg - LS002290&#160;</td>
			<td>Enzyme used in our laboratory in a cocktail with neutral protease/dispase to begin digestion of arteries for endothelial cell isolation.</td>
		</tr>
		<tr>
			<td>Falcon Tissue culture Plate, 6-well, Flat Bottom with Low Evaporation Lid&#160;</td>
			<td>Corning</td>
			<td>353046</td>
			<td>For use with studies involving cultured cells and multiple treatments</td>
		</tr>
		<tr>
			<td>neutral protease/dispase</td>
			<td>Worthington Biochemical</td>
			<td>10 mg- LS02100 50 mg - LS02104</td>
			<td>Enzyme used in our laboratory in a cocktail with elastase to begin digestion of arteries for endothelial cell isolation</td>
		</tr>
		<tr>
			<td>SylGard&#160;</td>
			<td>World Precision Instruments</td>
			<td>SYLG184</td>
			<td>Silicone elastomer for adhering the rectangular cover slip to the MPP flow chamber pieces C and E (Figure 1)</td>
		</tr>
		<tr>
			<td>Tygon ND 10-80 tubing</td>
			<td>Microbore Tubing</td>
			<td>AAQ04133</td>
			<td>ID: 0.05 in, OD: 0.09 in, inlet perfusion tubing for adminsitering flow to the chamber</td>
		</tr>
	</tbody>
</table>

electrophysiological recordings of single-cell ion currents under well-defined shear stress

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood flow triggers a signaling cascade resulting in vasodilation thereby alleviating mechanical stress on the vascular wall. The pattern of shear stress is also well known to be a critical factor in the development of atherosclerosis with laminar shear stress being atheroprotective and disturbed shear stress being pro-atherogenic. While we have a detailed understanding of the various intermediate cell signaling pathways, the receptors that first translate the mechanical stimulus into chemical mediators are not completely understood. Mechanosensitive ion channels are critical to the response to shear and regulate shear-induced cell signaling thereby controlling the production of vasoactive mediators. These channels are among the earliest activated signaling components to shear and have been linked to shear-induced vasodilation through promoting nitric oxide production (e.g., inwardly rectifying K+ [Kir] and transient receptor potential [TRP] channels) and endothelium hyperpolarizing factor (e.g., Kir and calcium-activated K+ [KCa] channels) and shear-induced vasoconstriction through an undetermined mechanism that involves piezo channels. Understanding the biophysical mechanism by which these channels are activated by shear forces (i.e., directly or through a primary mechano-receptor) could provide potential new targets to resolve the pathophysiology associated with endothelial dysfunction and atherogenesis. It is still a major challenge to record flow-induced activation of ion channels in real time using electrophysiology. The standard methods to expose cells to well-defined shear stress, such as the cone and plate rheometer and closed parallel plate flow chamber do not allow real time study of ion channel activation. The goal of this protocol is to describe a modified parallel plate flow chamber that allows real time electrophysiological recording of mechanosensitive ion channels under well-defined shear stress.

Multiple photographs showing different views of the MPP flow chamber on the microscope stage (upper panel) and a schematic representation of the MPP flow chamber (bottom panel) are shown in Figure 1. The schematic details the dimensions of the entire device and flow chamber. Figure 2 shows a photograph of the gravity perfusion system to the MPP flow chamber in our laboratory (upper panel). Also shown is a schematic representation...

Watch this Scientific Journal Video about Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress at JoVE.com

Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

The goal of this protocol is to describe a modified parallel plate flow chamber for use in investigating real time activation of mechanosensitive ion channels by shear stress.

Fluid shear stress is well known to play a major role in endothelial function. In most vascular beds, elevated shear stress from acute increases in blood ...

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Immunology and Infection

6.1K Views.  University of Illinois at Chicago. The goal of this protocol is to describe a modified parallel plate flow chamber for use in investigating real time activation of mechanosensitive ion channels by shear stress.

Video: Electrophysiological Recordings of Single-cell Ion Currents Under Well-defined Shear Stress

Introducing Shear Stress in the Study of Bacterial Adhesion

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the initial and determining step of the pathogenesis. Although experimentally adhesion is mostly studied in static conditions adhesion actually takes place in the presence of flowing liquid. First encounters between bacteria and their host often occur at the mucosal level, mouth, lung, gut, eye, etc. where mucus flows along the surface of epithelial cells. Later in infection, pathogens occasionally access the blood circulation causing life-threatening illnesses such as septicemia, sepsis and meningitis. A defining feature of these infections is the ability of these pathogens to interact with endothelial cells in presence of circulating blood. The presence of flowing liquid, mucus or blood for instance, determines adhesion because it generates a mechanical force on the pathogen. To characterize the effect of flowing liquid one usually refers to the notion of shear stress, which is the tangential force exerted per unit area by a fluid moving near a stationary wall, expressed in dynes/cm2. Intensities of shear stress vary widely according to the different vessels type, size, organ, location etc. (0-100 dynes/cm2). Circulation in capillaries can reach very low shear stress values and even temporarily stop during periods ranging between a few seconds to several minutes 1. On the other end of the spectrum shear stress in arterioles can reach 100 dynes/cm2 2. The impact of shear stress on different biological processes has been clearly demonstrated as for instance during the interaction of leukocytes with the endothelium 3. To take into account this mechanical parameter in the process of bacterial adhesion we took advantage of an experimental procedure based on the use of a disposable flow chamber 4. Host cells are grown in the flow chamber and fluorescent bacteria are introduced in the flow controlled by a syringe pump. We initially focused our investigations on the bacterial pathogen Neisseria meningitidis, a Gram-negative bacterium responsible for septicemia and meningitis. The procedure described here allowed us to study the impact of shear stress on the ability of the bacteria to: adhere to cells 1, to proliferate on the cell surface 5and to detach to colonize new sites 6 (Figure 1). Complementary technical information can be found in reference 7. Shear stress values presented here were chosen based on our previous experience1 and to represent values found in the literature. The protocol should be applicable to a wide range of pathogens with specific adjustments depending on the objectives of the study.

During the infection process, a key step is the adhesion of pathogens with host cells. In most instances this adhesion step occurs in the presence of mechanical stress generated by flowing liquid. We describe a technique that introduces shear stress as an important parameter in the study of bacterial adhesion.

During bacterial infections a sequence of interactions occur between the pathogen and its host. Bacterial adhesion to the host cell surface is often the ...

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly constituted by proteins and exopolysaccharides, among other factors 7-10. The microbial community encased within the biofilm often shows the differentiation of distinct subpopulation of specialized cells 11-17. These subpopulations coexist and often show spatial and temporal organization within the biofilm 18-21.
Biofilm formation in the model organism Bacillus subtilis requires the differentiation of distinct subpopulations of specialized cells. Among them, the subpopulation of matrix producers, responsible to produce and secrete the extracellular matrix of the biofilm is essential for biofilm formation 11,19. Hence, differentiation of matrix producers is a hallmark of biofilm formation in B. subtilis.
We have used fluorescent reporters to visualize and quantify the subpopulation of matrix producers in biofilms of B. subtilis 15,19,22-24. Concretely, we have observed that the subpopulation of matrix producers differentiates in response to the presence of self-produced extracellular signal surfactin 25. Interestingly, surfactin is produced by a subpopulation of specialized cells different from the subpopulation of matrix producers 15.
We have detailed in this report the technical approach necessary to visualize and quantify the subpopulation of matrix producers and surfactin producers within the biofilms of B. subtilis. To do this, fluorescent reporters of genes required for matrix production and surfactin production are inserted into the chromosome of B. subtilis. Reporters are expressed only in a subpopulation of specialized cells. Then, the subpopulations can be monitored using fluorescence microscopy and flow cytometry (See Fig 1).
The fact that different subpopulations of specialized cells coexist within multicellular communities of bacteria gives us a different perspective about the regulation of gene expression in prokaryotes. This protocol addresses this phenomenon experimentally and it can be easily adapted to any other working model, to elucidate the molecular mechanisms underlying phenotypic heterogeneity within a microbial community.

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Microbial biofilms are generally constituted by distinct subpopulations of specialized cells. Single-cell analysis of these subpopulations requires the use of fluorescent reporters. Here we describe a protocol to visualize and monitor several subpopulationswithin B. subtilis biofilms using fluorescence microscopy and flow cytometry.

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly ...

Quantitative High-throughput Single-cell Cytotoxicity Assay For T Cells

Cancer immunotherapy can harness the specificity of immune response to target and eliminate tumors. Adoptive cell therapy (ACT) based on the adoptive transfer of T cells genetically modified to express a chimeric antigen receptor (CAR) has shown considerable promise in clinical trials1-4. There are several advantages to using CAR+ T cells for the treatment of cancers including the ability to target non-MHC restricted antigens and to functionalize the T cells for optimal survival, homing and persistence within the host; and finally to induce apoptosis of CAR+ T cells in the event of host toxicity5.
Delineating the optimal functions of CAR+ T cells associated with clinical benefit is essential for designing the next generation of clinical trials. Recent advances in live animal imaging like multiphoton microscopy have revolutionized the study of immune cell function in vivo6,7. While these studies have advanced our understanding of T-cell functions in vivo, T-cell based ACT in clinical trials requires the need to link molecular and functional features of T-cell preparations pre-infusion with clinical efficacy post-infusion, by utilizing in vitro assays monitoring T-cell functions like, cytotoxicity and cytokine secretion. Standard flow-cytometry based assays have been developed that determine the overall functioning of populations of T cells at the single-cell level but these are not suitable for monitoring conjugate formation and lifetimes or the ability of the same cell to kill multiple targets8.
Microfabricated arrays designed in biocompatible polymers like polydimethylsiloxane (PDMS) are a particularly attractive method to spatially confine effectors and targets in small volumes9. In combination with automated time-lapse fluorescence microscopy, thousands of effector-target interactions can be monitored simultaneously by imaging individual wells of a nanowell array. We present here a high-throughput methodology for monitoring T-cell mediated cytotoxicity at the single-cell level that can be broadly applied to studying the cytolytic functionality of T cells.

We describe a single-cell high-throughput assay to measure cytotoxicity of T cells when incubated with tumor target cells. This method employs a dense, elastomeric array of sub-nanoliter wells (~100,000 wells/array) to spatially confine the T cells and target cells at defined ratios and is coupled to fluorescence microscopy to monitor effector-target conjugation and subsequent apoptosis.

Cancer immunotherapy can harness the specificity of  immune response to target and eliminate tumors. Adoptive cell therapy (ACT)  based on the adoptive ...

Single Cell Measurements of Vacuolar Rupture Caused by Intracellular Pathogens

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed compartment allows bacteria to move within the cytosol, proliferate and further invade neighboring cells. Mycobacterium tuberculosis is phagocytosed by immune cells, and has recently been shown to rupture phagosomal membrane in macrophages. We developed a robust assay for tracking phagosomal membrane disruption after host cell entry of Shigella flexneri or Mycobacterium tuberculosis. The approach makes use of CCF4, a FRET reporter sensitive to &beta;-lactamase that equilibrates in the cytosol of host cells. Upon invasion of host cells by bacterial pathogens, the probe remains intact as long as the bacteria reside in membrane-enclosed compartments. After disruption of the vacuole, &beta;-lactamase activity on the surface of the intracellular pathogen cleaves CCF4 instantly leading to a loss of FRET signal and switching its emission spectrum. This robust ratiometric assay yields accurate information about the timing of vacuolar rupture induced by the invading bacteria, and it can be coupled to automated microscopy and image processing by specialized algorithms for the detection of the emission signals of the FRET donor and acceptor. Further, it allows investigating the dynamics of vacuolar disruption elicited by intracellular bacteria in real time in single cells. Finally, it is perfectly suited for high-throughput analysis with a spatio-temporal resolution exceeding previous methods. Here, we provide the experimental details of exemplary protocols for the CCF4 vacuolar rupture assay on HeLa cells and THP-1 macrophages for time-lapse experiments or end points experiments using Shigella flexneri as well as multiple mycobacterial strains such as Mycobacterium marinum, Mycobacterium bovis, and Mycobacterium tuberculosis.

We describe a method for tracking the endomembrane rupture elicited by the intracellular bacteria Shigella flexneri and Mycobacterium tuberculosis upon host cell invasion. Our assay makes use of CCF4, a host cytoplasmic FRET probe in live or fixed cells. This reporter is degraded by an enzyme activity present on the bacterial surface.

Shigella flexneri are pathogenic bacteria that invade host cells entering into an endocytic vacuole. Subsequently, the rupture of this membrane-enclosed ...

A Flow Adhesion Assay to Study Leucocyte Recruitment to Human Hepatic Sinusoidal Endothelium Under Conditions of Shear Stress

Leucocyte infiltration into human liver tissue is a common process in all adult inflammatory liver diseases. Chronic infiltration can drive the development of fibrosis and progression to cirrhosis. Understanding the molecular mechanisms that mediate leucocyte recruitment to the liver could identify important therapeutic targets for liver disease. The key interaction during leucocyte recruitment is that of inflammatory cells with endothelium under conditions of shear stress. Recruitment to the liver occurs within the low shear channels of the hepatic sinusoids which are lined by hepatic sinusoidal endothelial cells (HSEC). The conditions within the hepatic sinusoids can be recapitulated by perfusing leucocytes through channels lined by human HSEC monolayers at specific flow rates. In these conditions leucocytes undergo a brief tethering step followed by activation and firm adhesion, followed by a crawling step and subsequent transmigration across the endothelial layer. Using phase contrast microscopy, each step of this &#39;adhesion cascade&#39; can be visualized and recorded followed by offline analysis. Endothelial cells or leucocytes can be pretreated with inhibitors to determine the role of specific molecules during this process.

Leucocyte recruitment to the liver occurs within the specialized channels of the hepatic sinusoids which are lined by unique hepatic sinusoidal endothelial cells. Phase contrast microscopy of leucocyte recruitment across human hepatic sinusoidal endothelium under conditions of physiological shear stress can facilitate the elucidation of the molecular mechanisms which underlie this process.

Leucocyte infiltration into human liver tissue is a common process in all adult inflammatory liver diseases. Chronic infiltration can drive the ...

Preparation of Single-cell Suspensions for Cytofluorimetric Analysis from Different Mouse Skin Regions

The skin is a barrier organ that interacts with the external environment. Being continuously exposed to potential microbial invasion, the dermis and epidermis home a variety of immune cells in both homeostatic and inflammatory conditions. Tools to obtain skin cell release for cytofluorimetric analyses are, therefore, very useful in order to study the complex network of immune cells residing in the skin and their response to microbial stimuli. Here, we describe an efficient methodology for the digestion of mouse skin to rapidly and efficiently obtain single-cell suspensions. This protocol allows maintenance of maximum cell viability without compromising surface antigen expression. We also describe how to take and digest skin samples from different anatomical locations, such as the ear, trunk, tail, and footpad. The obtained suspensions are then stained and analyzed by flow cytometry to discriminate between different leukocyte populations.

The skin is home to a complex immune cell network. We describe an efficient methodology for the digestion of mouse skin, from different parts of the animal's body, in order to obtain a single-cell suspension and analyze the different leukocyte populations resident in the skin by flow cytometry.

The skin is a barrier organ that interacts with the external environment. Being continuously exposed to potential microbial invasion, the dermis and ...

Assay of Adhesion Under Shear Stress for the Study of T Lymphocyte-Adhesion Molecule Interactions

Overall, T cell adhesion is a critical component of function, contributing to the distinct processes of cellular recruitment to sites of inflammation and interaction with antigen presenting cells (APC) in the formation of immunological synapses. These two contexts of T cell adhesion differ in that T cell-APC interactions can be considered static, while T cell-blood vessel interactions are challenged by the shear stress generated by circulation itself. T cell-APC interactions are classified as static in that the two cellular partners are static relative to each other. Usually, this interaction occurs within the lymph nodes. As a T cell interacts with the blood vessel wall, the cells arrest and must resist the generated shear stress.1,2 These differences highlight the need to better understand static adhesion and adhesion under flow conditions as two distinct regulatory processes. The regulation of T cell adhesion can be most succinctly described as controlling the affinity state of integrin molecules expressed on the cell surface, and thereby regulating the interaction of integrins with the adhesion molecule ligands expressed on the surface of the interacting cell. Our current understanding of the regulation of integrin affinity states comes from often simplistic in vitro model systems. The assay of adhesion using flow conditions described here allows for the visualization and accurate quantification of T cell-epithelial cell interactions in real time following a stimulus. An adhesion under flow assay can be applied to studies of adhesion signaling within T cells following treatment with inhibitory or stimulatory substances. Additionally, this assay can be expanded beyond T cell signaling to any adhesive leukocyte population and any integrin-adhesion molecule pair.

This flow adhesion assay provides a simple, high impact model of T cell-epithelial cell interactions. A syringe pump is used to generate shear stress, and confocal microscopy captures images for quantification. The goal of these studies is to effectively quantify T cell adhesion using flow conditions.

Overall, T cell adhesion is a critical component of function, contributing to the distinct processes of cellular recruitment to sites of inflammation and ...

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

Salmonella typhimurium is a facultative intracellular bacterium that causes gastroenteritis in humans. After invasion of the lamina propria, S. typhimurium bacteria are quickly detected and phagocytized by macrophages, and contained in vesicles known as phagosomes in order to be degraded. Isolation of S. typhimurium-containing phagosomes have been widely used to study how S. typhimurium infection alters the process of phagosome maturation to prevent bacterial degradation. Classically, the isolation of bacteria-containing phagosomes has been performed by sucrose gradient centrifugation. However, this process is time-consuming, and requires specialized equipment and a certain degree of dexterity. Described here is a simple and quick method for the isolation of S. typhimurium-containing phagosomes from macrophages by coating the bacteria with biotin-streptavidin-conjugated magnetic beads. Phagosomes obtained by this method can be suspended in any buffer of choice, allowing the utilization of isolated phagosomes for a broad range of assays, such as protein, metabolite, and lipid analysis. In summary, this method for the isolation of S. typhimurium-containing phagosomes is specific, efficient, rapid, requires minimum equipment, and is more versatile than the classical method of isolation by sucrose gradient-ultracentrifugation.

Isolation of Salmonella typhimurium-containing Phagosomes from Macrophages

We describe here a simple and quick method for the isolation of Salmonella typhimurium-containing phagosomes from macrophages by coating the bacteria with biotin and streptavidin.

Salmonella typhimurium is a facultative intracellular bacterium that causes gastroenteritis in humans. After invasion of the lamina propria, S. ...

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

Analysis of Shear Flow-induced Migration of Murine Marginal Zone B Cells In Vitro

Marginal zone B cells (MZBs) are a population of B cells that reside in the mouse splenic marginal zones that envelop follicles. To reach the follicles, MZBs must migrate up the shear force of blood flow. We present here a method for analyzing this flow-induced MZB migration in vitro. First, MZBs are isolated from the mouse spleen. Second, MZBs are settled on integrin ligands in flow chamber slides, exposed to shear flow, and imaged under a microscope while migrating. Third, images of the migrating MZBs are processed using the MTrack2 automatic cell tracking plugin for ImageJ, and the resulting cell tracks are quantified using the Ibidi chemotaxis tool. The migration data reveal how fast the cells move, how often they change direction, whether the shear flow vector affects their migration direction, and which integrin ligands are involved. Although we use MZBs, the method can easily be adapted for analyzing migration of any leukocyte that responds to the force of shear flow.

Marginal zone B cells (MZBs) respond to the force of shear flow by re-orienting their migration path up the flow. This protocol shows how to record and analyze the migration using a fluidics unit, pump, microscope imaging system, and free software.

Marginal zone B cells (MZBs) are a population of B cells that reside in the mouse splenic marginal zones that envelop follicles. To reach the follicles, ...