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Zhu MX, editor. TRP Channels. Boca Raton (FL): CRC Press; 2011.

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Chapter 16Regulation of TRP Channels by Osmomechanical Stress

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16.1.1. Osmomechanical Sensing and Transduction

Most prokaryotic and eukaryotic cells have the ability to sense and respond to alterations in various mechanical stimuli. This is most apparent in higher organisms (eukaryotic cells), which have developed sensory systems with specialized sensory cells for detecting a wide range of stimuli including mechanical distortions of the cell membrane. However, the basic mechanisms of sensing (mechanosensation) and transducing (mechanotransduction) mechanical stimuli are apparent throughout evolutionary biology with bacteria already displaying mechanosensitive channels that are activated in response to osmotic stimuli (hyperosmolarity).13 Mechanosensitive channel opening in these prokaryotes is induced upon the generation of tension within the cell membrane. In eukaryotes, much more elaborate systems of mechanosensation have been developed in association with the evolution of an extensive scaffolding network, the actin cytoskeleton, with sites of tethering to the plasma membrane and to external sites through integrin-extracellular matrix attachments.46 As a result, changes in cell volume, shape, or tension within the cell membranes can rapidly lead to induction of ion channels or biochemical cascades as part of the transduction process.

Most mammalian cells can sense and respond to mechanical stimuli. This is particularly true in systems with continuously changing mechanical stresses, such as those found in the cardiovascular and renal systems. Indeed, in addition to the cardiomyocytes and vascular smooth muscle cells, the vascular endothelial cells and renal tubular epithelial cells are well known to be sensitive to alterations in fluid flow, hydrostatic pressures, contractile forces, and/or cell shape changes.710 Dysfunctional control of the sensing and transduction machinery can lead to numerous pathophysiological states ranging from hypertension, to atherosclerosis, and to altered fluid and electrolyte balance, to name a few. The underlying alteration in osmomechanical stresses is often associated with an early change in calcium signaling and downstream calcium-dependent processes. The source of the mechanosensitive calcium signals and the downstream effector pathways has only recently begun to be elucidated. Increasing evidence points to a key role of calcium-permeable TRP channels in this process.7,10

This chapter will focus on the methods and techniques of applying and studying the effect of osmotic and mechanical stresses on a functional group of TRP channels dubbed the “osmomechanical TRP channels.” This is a group of TRP channels that are known to be activated when exposed to osmotic and mechanical stresses.1012 This chapter is not meant to be an exhaustive review on osmomechanical TRP channels, as many excellent reviews are available that cover various functional aspects of these channels.7,10,1214 The methods outlined below will focus on application of mechanical stresses to whole cells in the cardiovascular and renal systems that will include osmotic stresses, shear stresses, or fluid flow stresses and cell membrane stretch, stresses these cells typically experience on a minute-to-minute basis.

16.1.2. Osmomechanical-Sensitive TRP Channels

Transient receptor potential (TRP) proteins are a family of nonselective cation permeable channels, most of which are permeable to calcium ions.12,15,16 As first described in Drosophila, TRP channels have been shown to be expressed in a multitude of animal cells and tissues. In mammals, the TRP channels expressed belong to six subfamilies: classical (TRPC1-7), vanilloid (TRPV1-6), melastatin (TRPM1-8), polycystin (TRPP1-4), mucolipin (TRPML1-3), and ankyrin (TRPA1). A subunit of TRP channels is composed of cytosolic carboxyl and amino-termini with multiple protein-to-protein interaction sites, six transmembrane spanning segments, and an ion conductive “pore loop” between the fifth and sixth transmembrane segments. Functional channels are believed to be formed by a homo/heterotetramer configuration of TRP proteins. Furthermore, many TRP channels seem to be activated by a broad range of stimuli including osmomechanical stress. The group of TRP channels that are sensitive to osmomechanical stress includes proteins from multiple subfamilies as outlined below (see Table 16.1). However, there is considerable diversity in channel activation. For example, a member of the TRPC subfamily, TRPC1, was demonstrated to be activated when tension was applied to the lipid bilayer.17 Through a series of experiments where TRPC1 was expressed in liposomes and CHO-K1 cells, activation evaluated using the patch-clamp technique showed that the channel is likely gated by bilayer-dependent tension. Alternatively, a member of the TRPV subfamily, TRPV4, has been shown to respond to mechanostimuli delivered by shear/hypotonic stress.17,18 However, while direct activation of TRPV4 by mechanical tension in a bilayer has not been forthcoming, accumulating evidence points to an indirect control of the channel via arachidonic acid and epoxyeicosatrienoic acids, the latter being metabolites of the P450 (CYP) epoxygenase pathway.19

TABLE 16.1

TABLE 16.1

TRP Channels Regulated by Osmomechanical Stresses

16.1.3. Mechanisms of Channel Activation by Osmomechanical Stresses

Mechano- or osmomechano-sensitive channels are believed to have multiple mechanisms of activation.68 The first model of mechano-sensitivity supposes channels to be activated in a lipid bilayer-dependent manner. In this mechanism, tension applied deforms/bends the membrane, causing the membrane lipids to “pull away” from the channel protein. As a result, tension is increased on the hydrophobic sites of the channel (increasing hydrophobic forces), which then leads to a conformational change of the channel and possible activation. A second model supposes channels to be linked to accessory molecules, such as extracellular matrix/cytoskeletal proteins, and that mechanical forces acting on these molecules may produce tension to activate the channel in a lipid bilayer-independent manner.

A third model of channel activation is an indirect model where the channel is not directly activated by mechanical forces producing tension on the channels but rather is activated by processes involving secondary signaling components (lipases, kinases, G-proteins) that are initiated as a result of mechanical stimulus. This is a widespread scenario that underlies the activation of many “mechano-sensitive” processes in cells, for example, osmotic activation of TRPV4 by epoxyeicosatrienoic acid metabolites of the P450 (CYP) epoxygenase pathway.19

Accumulating evidence indicates that osmotic activation of some TRP channels may occur via one of two mechanisms: (1) Swelling-induced activation of PLA2/arachidonic acid and/or the PLC/diacylglycerol/PKC pathway, whose downstream metabolites then activate TRP channels; (2) PDZ scaffolding proteins such as ezrin/radixin/moesin act as the mediators between some TRP channels and F-actin cytoskeleton.2022 Thus, disruption or de-polymerization of actin cytoskeleton can activate some TRP channels via interaction with these PDZ scaffolding proteins.


There are a number of methods available to study the effect of mechanical stresses on TRP channel properties. Not all methods can be utilized for all cells or for studying the effect of specific osmomechanical stresses, as the application of certain stresses requires defined methods and study configurations that may not always allow a particular technique to be employed. In this section, we first outline the most typical experimental approaches for studying the effect of osmomechanical stresses on TRP channels, followed by the methods for studying the application of three types of specific osmomechanical stresses on cells grown on cover slips: osmotic stress, shear stress (fluid flow), and cell membrane stretch.

16.2.1. Assessing the Activity of Calcium-Permeable TRP Channels Patch-Clamp Techniques

The patch-clamp techniques provide the most direct measure of channel activity, but the methods are often the most difficult to apply. These methods allow one to directly measure the ionic current carried by the channel of interest. Typically, the magnitude of channel currents (ionic fluxes) is measured under voltage clamp conditions using the standard patch-clamp technique.23,24 The patch electrode is lowered onto the cell surface, and a high resistance seal (giga-ohm seal) between the pipette tip and the cell membrane is established, usually requiring a brief application of negative pipette pressure (suction). Once a giga-ohm seal is achieved, small unitary currents from single channels in the patched membrane (single-channel currents) or integrated currents from the whole cell (whole-cell currents) can often be measured. Success can be elusive because not all cells can be successfully patched with formation of giga-ohm seals. Success is best achieved when the cells in culture have “clean” cell surfaces with minimal expression of extracellular glycoproteins/glycocalyx.

There are several configurations of the patch-clamp technique that allow one to measure single-channel currents or whole-cell currents, the latter being the integrated sum of all channel currents activated in a cell. Single-channel currents can be measured in cell-attached patches of an intact cell or in excised patches where a small area of the cell membrane that is attached to the patch pipette tip is literally pulled away (excised) from the rest of the cell by rapid pipette withdrawal (with significant mechanical perturbation). Excised patches have the advantage of using a “simplified” membrane preparation with currents flowing across only a small membrane surface that is isolated from the rest of the cell and many of its signaling components. Both inside-out patches (membrane cytoplasmic side facing the bathing media) and outside-out patches (membrane extracellular side facing the bathing media) allow access to both the cytoplasmic face of the membrane and the extracellular face of the membrane, respectively, giving the investigator access to either face of the channel. However, many of the cellular components and signaling cascades may be missing in the excised patch configuration, components that may be essential for the regulation and control of the channel. These need to be evaluated before proceeding. Hence, the basic properties of the channel are typically first evaluated with an on-cell patch configuration, often in the whole-cell mode, before progressing to the excised patch configuration to establish the basic properties of the channel. The details of these various techniques have been described in various reviews23,24 and will not be discussed in depth in this chapter.

Use of the patch-clamp technique to study the effect of osmomechanical stimuli on channel activation, including for TRP channels, has both advantages and disadvantages. Most TRP channels have been successfully evaluated with the various configurations of the patch-clamp technique, with the whole-cell patch configuration being the most widely used method. The use of the patch-clamp technique to study osmomechanical TRP channel activation is limited to the types of stimuli applied to the cells as outlined below.

Membrane Stretch. As heretofore noted, various patch-clamp configurations have been employed to study the effect of stretching the cell membrane upon channel activation. The most direct approach is to apply a hydrostatic pressure pulse, either negative (suction; membrane flexing into the pipette) or positive (pressure; membrane flexing away from the pipette), to the back of the patch pipette to “stretch” the membrane attached to the tip of the patch pipette.23,25 The first stretch-activated channels were identified using this approach. Typical pressures applied are from –30 to +30 mm Hg for mammalian channels. This approach must be utilized with extreme caution and testing, as the pressure pulses often will lead to loss of the seal or generation of “seal leaks” that may appear like mechanosensitive currents. An alternative approach to apply membrane stretch is to induce a mechanical stretch of the whole cell. This can be accomplished by growing cells on elasticized membranes, then inducing “cell stretch” by stretching of the elasticized membrane (see below). This approach is not readily applicable to patch-clamp techniques because the abrupt movement of the cell during stretching will normally lead to loss of the pipette seal with the cell membrane. In this case, calcium imaging may be a more appropriate technique for calcium-permeable TRP channels as discussed below.

Osmotic Stress (Osmotic-Induced Cell Volume Changes). Alterations in the osmolarity of the media surrounding the cell will lead to changes in cell volume as water (and solute) leaves or enters the cell to maintain osmotic balance. As cells shrink or swell, this leads to cell membrane tension at sites of cytoskeletal network attachment and/or sites of extracellular matrix attachment that can result in channel activation. This is typically evaluated by first establishing a giga-ohm seal in a whole-cell configuration and then exposing the cell to an osmotic challenge (typically exposing the cell to a hypoosmotic media, 200–270 mOsm/kg, to induce cell swelling). Again, care must be used to assess for generation of seal leak currents that may be mistaken as tension-induced channels. Further consideration must also be given to the effect of osmolarity itself, independent of cell volume changes, and this should be tested separately.

Shear Stress (Fluid Flow). Application of a defined shear stress using the patch-clamp technique is not generally attempted. Indeed, a shear stress stimulus normally employs a defined stress of known magnitude in an enclosed compartment of known geometry and fluid flow properties. For example, cells grown on cover slips can be inserted into a parallel plate chamber, which includes an upper cover slip or cover over the cells with a known distance between the cells (lower cover slip) and the upper cover slip (typically 100–300 μm) and a known chamber width and length so that shear stress associated with a defined flow through the chamber can be precisely defined (see below). Such configurations are not readily adaptable to patch-clamp analysis because the cells are not easily accessible with the patch-clamp electrode owing to the presence of an upper cover. In some cases, an open-top chamber is employed and fluid flow applied to generate a shear stress at the cell membrane surface, but the magnitude of the shear stress is not defined. Calcium Imaging

Calcium imaging allows for the monitoring of calcium ion dynamics within living cells in real time. The methods do not provide a direct measure of TRP channel activity but rather provide an integrated calcium signal that can reflect all sources of calcium entry into the cytoplasm. This includes both calcium entry from the extracellular space and any release of calcium from internal stores. Hence, care must be taken to determine what component of the calcium signal reflects calcium influx through the TRP channel (or any other calcium-permeable channel) of interest. This can be accomplished by performing the experiment in the presence of extracellular calcium and then, again, in the absence of extracellular calcium to abolish all calcium influx. The difference in intracellular calcium levels between the two conditions is typically used as an index of calcium influx, but not a quantitative index because the absolute rates of calcium influx cannot be determined with this approach.

Typically, intracellular calcium levels are determined with the use of fluorescent molecular probes, which upon binding/interacting with calcium ions, undergo a chemical/structural change leading to an alteration of their fluorescent signal. The growing number of available calcium-sensitive molecular probes can be divided into two groups: genetically encoded calcium indicators26 and small molecule calcium indicators.27 Genetically encoded calcium indicators are produced by the translation of a nucleic acid sequence, typically incorporated into the cell by a gene transfer technique. An advantage of using genetically encoded calcium indicators is that the cellular localization and the expression level of the indicator can be controlled by the incorporation of a signal sequence and a promoter sequence in the cDNA construct. On the other hand, small molecule calcium indicators do not require translation by the cell but rather are introduced into cells by diffusion or disruption of the lipid bilayer of the membrane. Use of these indicators generally provides greater sensitivity, a greater ability to measure rapid calcium dynamics, and no need for transfection. An example of a commonly utilized small molecule calcium indicator is Fura-2. Fura-2 is a useful ratiometric dye because of its ability to undergo an absorption shift from 380 nm in its Ca2+-free state to a shorter wavelength of 340 nm in its Ca2+- bound state. Additionally, it offers the advantages of having a high affinity for Ca2+ (Kd ≈ 225 nM), little affinity for other ions, no detectable binding to membrane, and a wide Ca2+ sensitivity ranging from approximately 20 nM to 10 μM. Furthermore, the attachment of an acetoxymethyl (AM) ester group to the carboxyl tail allows Fura-2/AM to freely diffuse through the plasma membrane, making its loading into cells a simple procedure. Once inside the cell, esterases cleave off the AM tail from Fura-2, which leaves the indicator in its charged form and therefore unable to cross the cellular membranes, i.e., it becomes trapped in the cytosol.

The calcium imaging techniques can generally be applied to study TRP channel activity in a variety of experimental setups. Typically, cells are grown on cover slips or elasticized membranes (silastic membranes that can be stretched) and then attached to the bottom of a flow chamber that can be mounted on an inverted microscope that allows ready imaging of the cells. Cells can then be exposed to the various osmomechanical stresses while simultaneously monitoring intracellular calcium: osmotic stresses, shear stresses (or fluid flow), and cell membrane stretch. The methods are outlined in more detail below for each of the conditions. Channel Abundance and Localization

As complementary approaches to patch-clamp and calcium imaging, Western blotting and immunocytochemistry can be used to study TRP channel regulation under different osmomechanical stimuli. These methods provide measures of channel abundance and localization within the cell, respectively. Whole-cell lysates and plasma membrane proteins are harvested, and standard Western blotting (protocol, AbCam) is performed to characterize and quantify the TRP channel protein of interest. Although the biotin-streptavidin method (protocol, Thermo Scientific) has been used extensively to pull down biotinylated plasma membrane proteins, cytosolic proteins, especially cytoskeleton proteins, readily come down as a contaminant with the plasma membrane proteins. Therefore, one has to be careful with the purity of plasma membrane proteins, depending on the goal of the investigation. The sucrose gradient differential centrifugation method28 is by far the cleanest way to extract plasma membrane proteins; however, the low yield is always a concern. With the internal loading control, for instance, pan cadherin in the plasma membrane preparation, one can perform densitometry (Image J) to quantify the abundance of a TRP channel in the plasma membrane sample.

Quantification of a functional TRP on the plasma membrane is a must to understand TRP channel trafficking and recycling under osmomechanical stimuli. Immunostaining techniques are widely used to study the subcellular distribution of TRP channels in various cell and tissue types. Usually, a primary antibody against a TRP channel and another primary antibody against a possible partner of the TRP channel are carefully selected. Then incubation of the primary antibodies and the matching secondary antibodies with fluorescence tags is performed according to standard immunostaining protocols (Millipore). Confocal microscopy is then utilized to visualize the TRP channel distribution and their relations to other target proteins of interest.

16.2.2. Osmotic Stress/Cell Volume

There have been numerous studies showing that certain TRP channels can be activated by hypoosmotic stress, leading to cell swelling as heretofore noted. Some of the examples are TRPV2,29,30 TRPV4,11,3133 TRPM3,34 TRPM7,35 TRPC1,36 and TRPP237 (see Table 16.1). However, there are only limited studies assessing the effect of hyperosmotic stress on TRP channels, with few channels identified as sensitive to shrinkage. A notable exception is an apparent N-terminal truncated version of TRPV1. Ciura and Bourque have shown that TRPV1 truncated protein is activated by hyperosmotic stress, and TRPV1–/– mice show a reduced drinking response under hypertonic challenge.38

Subsequent to hypoosmotic-induced cell swelling, most cells undergo a regulatory volume decrease (RVD) where the cell volume is returned toward the basal isotonic volume. Although there is little evidence that TRP channels directly activate RVD, it is proposed that TRP channels regulate RVD indirectly by activating the Ca2+-activated K+ channel via calcium influx.17,35,39 Indeed, it has been shown recently that TRPV4 may be involved in volume regulation. For example, CHO cells gain RVD capability when overexpressed with TRPV4.40 Interactions between TRPV4 and other channels like AQP5 and CFTR may also affect RVD.39,41

The application of hypoosmotic stress to cells has been widely used to study cell volume-dependent phenomena for many years. In a typical experiment, hypotonic stress is usually achieved by applying an extracellular bathing solution with an osmolarity typically ranging from 200 to 270 mOsm (with reduced NaCl concentration) to the cells. Figure 16.1a shows the diagram of the expected cell volume changes upon exposure of the cells to a hypotonic media (HYPO). The volume flow of water into the cells can be expressed as

FIGURE 16.1. Cellular responses upon exposure of cortical collecting duct M-1 cells to hypoosmotic stress.


Cellular responses upon exposure of cortical collecting duct M-1 cells to hypoosmotic stress. (a) Diagram of the expected cell volume changes upon exposure to a hypotonic media (HYPO). The volume flow of water into the cells is shown as Jv = σ (more...)

Image ch16_ueq01

where Jv is the volume flow, σ is the reflection coefficient of the membrane (typically 1 for cell membranes), Kf is the membrane filtration coefficient, and Δ[Osm] is the applied osmotic gradient across the cell membrane. Figure 16.1b gives an example of the influence of hypotonic stress on intracellular calcium levels based on ratiometric calcium imaging, and Figure 16.1c gives a representative current-voltage plot (I-V plot) from a whole-cell patch-clamp study demonstrating activation of TRPV4 upon cell swelling (HYPO). Similarly, hypertonic stress is achieved by adding mannitol or NaCl to the extracellular media to make it hypertonic, typically in the range of 310–500 mOsm. The patch-clamp technique is utilized to directly measure TRP channel activity or that of associated channels, i.e., the Ca2+-activated K+ channel, when the bath solution osmolality is changed. Standard patch-clamp techniques for single-channel and whole-cell current recording are described above. Calcium imaging can be used in combination with the patch clamp to monitor the changes in intracellular calcium concentration following activation of the TRP channel during cell volume changes.

Actual measurements of the cell volume are complicated. Scanning laser confocal microscopy (SLCM) has been utilized to estimate the cell volume. Basically, a Z-stack of thin optical slices is obtained using SLCM and a three-dimensional (3D) image of a cell constructed using imaging/morphometric software.4244 However, laser scanning can readily induce photodynamic damage to the biological sample and must therefore be used with care, thus greatly limiting its application for cell volume measurements. Newly developed techniques, particularly scanning probe microscopy (SPM), may be more promising for cell volume measurements. Scanning ion-conductance microscopy (SICM) is one type of SPM with great promise. SICM utilizes a glass micropipette filled with electrolyte solution. The micropipette tip is positioned over, but very close to, the cell surface for scanning. The gap between the tip of the microelectrode probe and the sample strongly affects the ion current through the pipette, which decreases as the gap diminishes. Changes in the ion current are used in a feedback configuration to control the positioning system to keep the distance between the sample surface and microelectrode probe constant. Thus, the tip of the pipette traces the surface of the cell during scanning from which an actual cell volume can be calculated.4548

A more widely used technique for measuring the cell volume uses a cytosolic fluorescent probe, e.g., calcein-AM,49,50 to indirectly monitor the cell volume. Calcein-AM is calcein-acetoxymethyl ester, a cell-permeable calcein derivative that is cleaved and trapped in the cytosol once loaded into the cells, similar to that observed for Fura-2/ AM loading. Hamann and coworkers first described an approach to measure the cell volume by calcein-AM (at μM concentration) fluorescence self-quenching due to collisional quenching and dimerization of calcein molecules within the cell cytoplasm.49 Subsequently, Solenov and coworkers demonstrated that rather than calcein self-quenching, intracellular proteins would effectively quench calcein fluorescence as the protein concentration increased upon hyperosmotic challenge. For example, when cells shrink in response to hyperosmotic media, the intracellular protein concentration increases, which enhances calcein quenching; when cells swell, calcein quenching is reduced due to decreased intracellular protein concentration.50 Calcein can be excited at 488 nm, and its emission between 503 and 530 nm recorded and converted to a digital signal using a photomultiplier detector and a 14-bit analog-to-digital converter (or a digital camera). Both the Hamann group and the Solenov group have shown that the change in calcein fluorescence is approximately linear to the relative changes in the cell volume.49,50 Mitchell et al. have used an indirect approach to distinguish between swelling and shrinkage by monitoring the cell area as an index of the cell volume in calcein-loaded cells.5154 A brief experimental procedure is as the following.

Cells are loaded with 2 μM calcein-AM and 0.2% Pluronic for 30 to 40 min and subsequently superfused with hypertonic or hypotonic solution for 5 to 30 min while acquiring fluorescent data. The cell area is determined by the number of pixels detected above the preset threshold within the region of interest. Because calcein goes through bleaching when excited at 488 nm, one should always take into consideration the best frequency for data acquisition and balance this against the rate of calcein photobleaching.

16.2.3. Shear Stress/Fluid Flow

Many cells of the body are constantly exposed to fluid mechanical forces arising from the flow of fluid over the cell surface, giving rise to shear stress. This is particularly true for the endothelial cells lining blood vessels and the epithelial cells lining renal tubules where cells are continuously exposed both to circumferential stretch due to the effects of hydrostatic pressures and to shear stress, a frictional force.8,9,55 In recent years, it has also become apparent that fluid flow over the cell surface can also induce bending or mechanical tension in immotile cilia (primary cilia) of endothelial and renal epithelial cells.56 The bending of the cilia can lead to mechanical activation of numerous signaling processes, including TRP channels. This section of the chapter will focus on the frictional shear stress that any cell would experience with alternations in fluid flow over the cell surface.

Shear stress is the force per unit area that is generated on the cell surface due to fluid flow over the surface. The force is parallel to the luminal cell surface. Endothelial and epithelial cells are known to respond to such forces with changes in morphology, cell signaling, and gene expression.8,9,55 Studying the effect of these forces on TRP channels in vitro requires application of these forces in a controlled environment where channel properties can be measured. Typical shear stresses on endothelial cells in the vasculature range from near 2–20 dyn/cm2 in capillaries and small vessels but can increase to more than 30–100 dyn/cm2 near arterial branches or regions of sharp wall curvature.57,58 In the renal tubule, where tubular fluid has a much lower viscosity owing to protein-free tubular fluid, shear stresses in the distal nephron and collecting ducts, the most flow-sensitive regions, can also approach high values (due to high flow rates) with typical values ranging from near 0.3 to 25 dyn/ cm2, although higher values beyond this range are likely in various pathophysiological states.18

The effects of shear stress can be studied in vitro using a parallel plate chamber. The most straightforward method to study the effects of shear stress on TRP channel activity is to grow model culture cells in vitro on cover slips that can be mounted to the bottom of a parallel plate chamber. Parallel plate chambers are constructed with defined dimensions that permit calculation of flow rates, velocities, and shear stresses (Figure 16.2). The cover slip forms the bottom portion of the chamber that allows cells to be visualized microscopically and assessed using fluorescence techniques. In the case of calcium-permeable TRP channels, assessment of channel activity can generally be accomplished by fluorescence analysis of changes in intracellular calcium levels, as outlined above, while applying shear stresses of defined magnitudes, simply by altering the rate of fluid flow, changing the fluid viscosity, or altering the temperature. Flow must be laminar, not turbulent, for estimating shear stress. A laminar flow chamber can be readily constructed as a parallel plate chamber of constant width, length, and height, with the cover slip (with the cells attached) forming the bottom wall. Commercial sources are available for purchase of parallel plate chambers (e.g., C&L Instruments, Inc., Hershey, PA, USA; Flexcell International Corp., Hillsborough, NC, USA). Typical dimensions for such a parallel plate chamber are: 250 microns H × 1 cm W × 2–5 cm L (or similar). Fluid enters at one end and is collected at the opposite end. Typical flow rates through the chamber can be varied from 0 to 30 mL/min or more using a peristaltic pump to generate shear stresses of from 0 to upward of 30 dynes/cm2. For a rectangular chamber, the shear stress at the middle of the chamber (laminar flow) can be calculated as

FIGURE 16.2. Application of shear stress to TRPV4-transfected human embryonic kidney 293 cells grown on cover slips.


Application of shear stress to TRPV4-transfected human embryonic kidney 293 cells grown on cover slips. (a) Diagram of the stress chamber for application of shear stress to cells on the cover slip while simultaneously monitoring intracellular calcium (more...)

Image ch16_ueq02

where τ is the shear stress (dyn/cm2), μ is the fluid viscosity (using the fluid viscosity of water: 0.01002 Poise at 20°C and 0.006915 Poise at 37°C; note, 1 Poise = 1 dynes/ cm2), Q is the flow rate (mL/s), b is the chamber width (cm), and h is the chamber height (cm).57,58 With laminar flow, the shear stress is linearly dependent upon the fluid flow rate. The major determinates of the shear stress in this setting are the fluid flow rate, fluid viscosity, and temperature. The effect of temperature is primarily related to the dependence of fluid viscosity on temperature as noted above for the viscosity of water at 20° and 37°C.

An example of the effect of increasing shear stress on calcium signaling in TRPV4 transfected HEK cells is shown in Figure 16.2.18,33 Intracellular calcium levels were obtained using standard ratiometric calcium imaging techniques (see methods above). As shown, at low shear stress, calcium levels are near basal values of 100 nM or less. Upon rapid application of the shear stress to 15 dyn/cm2, intracellular calcium levels transiently rise to peak values of 200–300 nM, then relax to a low pseudo-plateau value over several minutes. While not shown here, removing extracellular calcium or knocking down TRPV4 by siRNA techniques largely abolishes the rise in intracellular calcium, demonstrating the dependence of TRPV4 on shear stress/fluid flow.18,33 Similar shear stress effects have been demonstrated in the cortical collecting duct cell line, M-1, where TRPV4 is endogenously expressed.18

While the effect of a step change in the shear stress has long been studied in vascular and renal cells, it is well known today that vascular cells, in particular, may also be sensitive to pulsatile shear stress, in addition to steady-state shear stress. Endothelial cells of the vasculature typically experience pulsatile flow (and pulsatile shear stress) as blood flow varies during the cardiac cycle. Similar methods as outlined above can be used to study pulsatile and other variations of the shear stress on cells by regulating the perfusion flow rate in a pulsatile manner through the parallel plate chamber. Indeed, complicated shear stress profiles can be generated using computer-controlled perfusion systems and well-designed algorithms for controlling flow (see Flexcell International).

16.2.4. Membrane Stretch/Cell Stretch Stretch-Induced Stresses and Cell Injury

Stretch-induced regulation of vascular and renal cells can be induced by mechanical stress that occurs rapidly, such as in traumatic injuries (brain, spinal cord, etc.), or slowly, such as pulsatile stretch that accompanies blood pressure and flow changes over the course of the normal cardiac cycle or during development of hypertension. Cellular and tissue stretch occurs rapidly, typically over milliseconds, in traumatic injury events and slowly, typically minutes to hours to days, during pulsatile or blood pressure alterations. Because the time periods for the stretch-induced effect can differ greatly, different instrumentation is normally used to permit assessment of different time periods and magnitudes of stretch-induced processes.

During trauma-induced stretching events, cells typically experience rapid alterations in mechanical stress. Such trauma-induced stress is thought to be equivalent to a linear tensile strain applied on a cell, causing a change in its length, i.e., stretching. This type of mechanical stress has been recognized to be a component of many traumatic injuries stemming primarily from generation of high angular acceleration.5961 Many mechanical strain instruments and methods have been described to generate such stress-induced stretch for studying cells and tissues in culture. The strain systems outlined below utilize deformation of a flexible membrane to generate stress on cultured cells. However, it should be noted that these strain units do not predict the strain generated on the cells but rather rely on measurements of strain generated on the flexible membrane the cells are grown on to predict injury to cells. This section will aim to highlight key features of a few in vitro strain systems with the goal of presenting the reader with a broad view of stretch models designed to be mounted on a microscope and facilitate couplings with standard fluorescent imaging techniques. Membrane Deformation Models

The tension generated on the cell by stretching has been described to be distributed either equally in two dimensions (pulling the cell in all directions) or unequally in one dimension (pulling the cell in one direction more than others). The former is known as biaxial stretch injury, while the latter is regarded as uniaxial stretch injury. Both forms of stretch injury have been frequently utilized in studies of traumatic injury.59,61

Uniaxial Stretch. Lusardi et al. describe a strain system by which rapid uniaxial stretch injury is applied to cell cultures62 as a method for application of stress that could be used as a model of traumatic brain injury. In the strain unit, cell cultures are grown on a flexible elastic membrane that is attached to a stainless-steel well, so that the membrane occupies the center of the steel chamber just above the well. A closed chamber is formed around the cells by attaching a cover plate over the steel well. The cover plate is designed with openings for pressure input and pressure measurement. The delivery of a pressure pulse applied into the closed upper portion of the chamber causes the elastic substrate to deform rapidly downward, resulting in stretching of the cells. Furthermore, a steel support plate may be placed below the elastic membrane to restrict the amount of membrane deformation that can occur. The use of underlying support plates, with holes in their center of different geometric shape/dimension, allows for generation of varying degrees of stretch. To induce uniaxial stretching, a support plate with a narrow rectangular shape (length 8 times greater than width) is placed over the central opening, thereby favoring longitudinal stretch. This model allows for a >50% strain of the elastic membrane; strain beyond this point can produce rupture of the elastic membrane.62

Biaxial Stretch. Biaxial strain units are available, or can be built, to allow application of either rapid or slow/static stretching of cells. Ellis and coworkers developed an in vitro model with a Cell Injury Controller unit designed to deliver rapid biaxial stretch to induce trauma on cell cultures.63 The unit is commercially available from Virginia Commonwealth University (Virginia Commonwealth University Medical Center, Department of Radiology, Richmond, VA, USA). We are using this unit in our own laboratory to mimic traumatic brain injury (TBI) of brain microvessel endothelial cells. In this model, stretch injury is induced by downward deformation of a flexible silastic membrane on which cell cultures have been grown (see Figure 16.3). The chamber unit calls for the use of six-well tissue culture flex plates, with silastic membranes, from Flexcell International. A removable plug fits on top of an individual well creating an airtight seal. Downward deformation of the membrane, into a spherical cap, is produced by a burst of gas from above. As the membrane is deformed, a measurement of pressure is taken inside the well that corresponds to the amount of biaxial stretch delivered. Any noncorrosive gas such as nitrogen or air can be utilized to create the pressure burst. The model allows for up to a 72% elastic membrane strain ratio and stretch duration rates of 1–99 ms, allowing for examination of various rapid injury conditions. The parameters of the stretch-induced (mild, moderate, and severe) injury were assessed in this model. Elastic membrane deformation of 5.7, 6.3, and 7.5 mm generates cell injuries typical of mild, moderate, and severe traumatic injury corresponding to membrane stretch of 20, 35, and 55%, respectively.

FIGURE 16.3. Membrane/cell stretch on brain microvessel endothelial cell line, bEnd3 cells.


Membrane/cell stretch on brain microvessel endothelial cell line, bEnd3 cells. (a) Stretch injury unit where a cell monolayer is grown on top of an elastic membrane (a1). A burst of pressure causes the elastic membrane to deform (stretch) and generates (more...)

The rapid strain units outlined above all induce injury via deformation of the elasticized membrane, and therefore, all lack the ability to maintain cells at a continuous focus plane during induction of stretch. This presents a fundamental complication in the ability to gain complete injury data with microscopic tools, such as those used for standard fluorescence imaging. Other stretch models, like that described by Hung and coworkers, produce stretch tension on cells while maintaining cells at a continuous focus plane.64 The model utilizes a polysulfone well designed with two grooves at the bottom (inner and outer) and an open area at the center (cell growth area) for cell visualization. At the bottom of the well, an elastomeric circular membrane, on which a monolayer of cells has been grown, is attached to the outer grove by an O-ring. Cells situated in the growth area of the well then will receive radial stretching as a ring located below the membrane is raised vertically, causing indentation of the membrane into the inner groove of the well. The indentation is designed so that while some membrane is raised vertically, the membrane of the cell growth area remains in the same plane. Ring indentation of 0.1–3.5 mm was calculated to achieve strain on the elastic membrane surface in the range of 0.04–8.0% strain.

It should be mentioned that a growing appreciation exists for the rate of strain induced in stretch models, as several studies have documented that the magnitude of injury is dependent on the strain rate.6567 Thus far, stretch-induced stress models have proven a useful tool in the study of various physiological and pathological states, such as the effects of strain on bone or stretch on cells in traumatic brain injury. Yet in many instances, it is assumed that stretch alone can mimic the physiological stress state, when in reality it is the combination of multiple stress factors. To this end, it seems likely that further evolution of these mechanical models will aim at the combining of osmomechanical stimuli in pursuit of developing a more accurate representation of the physiological stress states facing cells.

Chamber study units are also available for a much slower application of cell stretch to better mimic the pulsatile or static stretch normally present in the mammalian vasculature. Flexcell International has developed elasticized membrane systems in six-well plate formats (Flex I plates) to promote the assessment of pulsatile or static stretch on cells grown on the elasticized membranes, using pressure-pulse methods to apply stretch to the membrane as outlined above for the rapid application models. Such systems allow simple pulsatile stretch of cells but can also be programmed to provide more complex waveforms that include combined static and pulsatile components. The chamber can be attached to a microscope stage to allow application of fluorescent techniques to the cells, such as for measurement of intracellular calcium levels, while undergoing cycles of stretching. Flexcell International has developed both culture plates (Flex I culture plates) and computer-controlled strain systems (e.g., Flexcell 4000 Strain Unit) to perform such analysis. Investigators interested in their systems should visit their Web site for more details (; Flexcell International Corp, Hillsborough, NC, USA).


16.3.1. Patch-Clamp versus Calcium Imaging Methods

This chapter has described the basic methods for studying the effects of osmomechanical stresses on the properties of TRP channels. The methods employed to study the channel properties have limitations owing to the fact that application of mechanical stresses often leads to movement of cells in the in vitro study chambers or requires the use of mechanical units that restrict access to the cells. This is particularly applicable to the two methods of studying TRP channel properties described here: patch-clamp versus ratiometric calcium imaging. The patch-clamp technique can be very limited for this type of study because the application of mechanical stresses will typically lead to movement of the cells and loss of the electrical seal with the patch pipette. Further, some strain units, such as for application of shear stress, have an upper cover that restricts access to the cells with a patch pipette.

In contrast to patch-clamp techniques, calcium imaging can usually be used to study calcium influx in cells that are attached to glass cover slips of the study chambers during the application of stress. While some methods for applying strain, such as with the trauma injury models with cells on elasticized membranes, can lead to limitations in maintaining cells of interest in focus, this can be largely overcome with specialized chambers as outlined above. Alternatively, if continuous imaging is not needed during stretching, one can immediately refocus the microscope over a few seconds after application of injury and continue imaging of the cells, although a small degree of lateral movement may be introduced. We have also tested the use of automatic infrared focus systems on today's newer microscopes, but the elasticized membrane is not compatible with the infrared focusing beam. Hence, the investigator will typically either have to refocus after the stretch stimulus or invest in more sophisticated strain units that reduce the focusing problem (never fully eliminating the focus issue).

16.3.2. Cell Adhesion

A common problem when applying osmomechanical stimuli is maintenance of cell adhesion during and after the stimulus. This is particularly true for elasticized membranes that are physically displaced during the application of strain but can also be an issue with the fluid flow rate or upon cell swelling with cells grown on cover slips. Acid washing the cover slips will improve adhesion (1 N HCl for 4–8 hours followed by extensive washing in very clean distilled water). A more widely used approach for both cover slips and elasticized membranes is to precoat the membranes or cover slips with an adhesive-type material. This can be accomplished by treating the membranes or cover slips with polyamino acids such as poly-L-lysine. The protocol is as follows: First, briefly acid wash the membranes or cover slips, then apply a small droplet (adjust based on the area desired for cells to adhere) of poly-L-lysine (1 mg/mL; use 150 kDa size or larger) to the middle of the cover slip, let stand for at least 30 min, and wash with clean water. This should be followed by rinsing the cover slip with 100% ethanol and then resting it on the edge in an open tissue culture dish in a laminar flow hood until dry. Cells can then be plated in the usual manner. Finally, various extracellular matrix substances, including collagens, laminins, fribronectin, gelatin, and others, can also be pre-plated onto the cover slips and elasticized membranes to both improve adhesion and provide the cells with a more “natural” adhesion substrate. Most manufacturers of the extracellular matrix molecules also provide defined protocols for use of these substrates as adhesion substrates in cell culture (see Sigma-Aldrich).


The current chapter outlines typical methods and instrumentation that can be used to study the effects of osmomechanical stress on mechanically sensitive TRP channels. Methods appropriate for studying TRP channel properties under these conditions are presented including both patch-clamp analysis of channel currents and ratiometric calcium imaging analysis of calcium influx under various states of mechanical stress. Further, the application of various mechanical stresses is outlined with an emphasis on the osmotic and mechanical stresses that are typically experienced by cells of the cardiovascular and renal systems.


This work was supported by the National Institutes of Health awards to R. G. O'Neil, grant numbers DK70950 and R21 DE018522.


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