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1Departments of Biomedical Engineering, 3Internal Medicine (Cardiovascular Division), 4Molecular Physiology and Biological Physics, and the Cardiovascular Research Center, and 2Mechanical, Aerospace and Nuclear Engineering, University of Virginia Health Sciences Center, Charlottesville, Virginia 22908
Submitted 14 March 2003 ; accepted in final form 25 October 2003
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ABSTRACT |
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 TOP ABSTRACT THEORYMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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regulatory volume decrease; cell swelling; phospholemman
Osmolyte efflux is a more intermittent process, as changing conditions necessitate active responses to control cell volume to precise specifications. Changes in extracellular tonicity cause passive and active changes in cell volume (14). If extracellular tonicity is lowered, there is an initial passive increase in cell volume as water enters cells. As cell swelling progresses, critical regulatory mechanisms are activated, which limit or reduce the increase. The secondary response is called the "regulatory volume decrease" (RVD), and it is a common experimental finding (3). RVD returns cells toward normal size and is hypothesized to result from the opening of unidentified osmolyte-selective membrane ion channels activated by cell swelling (29). This allows selective efflux of osmolytes, including cations, anions, and small zwitterionic amino acids. This active solute efflux is accompanied by an obligatory water efflux and a reduction in cell size.
Cell volume has been measured using imaging such as video microscopy (6, 11, 30), fluorescence imaging (1, 9, 13, 27), electrical impedance (19), and Coulter counters (12, 22, 25). Each of these techniques has strengths and limitations. We have devised a new technique that is based on the principle that fluid flow in a rigid system is well determined, particularly for capillaries and other small-diameter tubes. The strategy is to grow cultured cells to line the inner lumen of capillary glass and to measure changes in the pressure gradient across the capillary tube as cell volume varies with changes in extracellular osmotic pressure. Advantages of this new method are that it is independent of cell morphology, records continuous changes in cell volume from large populations of cells, and allows for multiple experimental conditions within a single trial.
To test the ability of this new method to detect kinetic differences under conditions known to enhance RVD, we sought to confirm the finding that phospholemman (PLM) overexpression enhances RVD. PLM is a member of the FYXD family of single transmembrane domain proteins that regulate ion channels and pumps (31). In addition, PLM forms taurine-selective channels in lipid bilayers (5, 17, 21). The work to date does not allow definitive characterization of the cellular role of PLM as a channel or as a regulator but supports the possibility that PLM is involved in the signal transduction pathway responsible for regulation of cell volume. The most direct evidence is that overexpression of stably transfected PLM in cultured kidney cells enhances RVD and taurine efflux (22) and that suppression of PLM expression using anti-sense oligonucleotides reduces taurine efflux in cultured astrocytes (23).
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THEORY |
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TOPABSTRACTTHEORYMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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P) is dissipated across the two capillaries; thus the total pressure drop is equal to the sum of the pressure drops P1 and P2 across each capillary measured with transducers T1 and T2. Because the tubing diameter in the system is many orders of magnitude larger than the capillary diameter, the total resistance of the system is equal to the sum of the resistances R1 and R2 of the two individual capillaries that are connected in series. The volumetric fluid flow through both capillaries is the same, and the flow through the total system is equal to the flow through each capillary. Thus P1/R1 = P2/R2.
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Because the reference capillary obeys Poiseuille's law, the resistance of the capillary is R1 = (8
l1)/(
r14), where r1 is the internal radius of the capillary, is the fluid viscosity, and l1 is the length of the reference capillary tube. Thus the resistance of the capillary lined with cells is
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As the cells swell, we can calculate the resistance of the capillary containing the cells by using the two measured pressures in the system and the constants defining the reference capillary. The length of the capillary, l1, and the radius of the reference capillary, r1, are known. The viscosity, , was empirically measured at 37°C for the isotonic, hypotonic, and hypertonic solutions (9).
Calculation of Reynolds numbers. Our measurement system is based on the assumption that the fluid flow through the capillaries is laminar (2). To verify this assumption, we calculated Reynolds numbers for the system under normal and hypotonic conditions. This calculation characterizes the general flow conditions in the system and estimates the ratio of the inertial force loss to the viscous force loss of the fluid. For fluid flow through a cylinder, Re = Dv
, where Re is the dimensionless Reynolds number, D is the diameter of the tube, v is the estimated velocity of the fluid flow, = 1,000 kg/m3 is the density of the water at 20°C, and is the viscosity of the solution.
We calculated a Reynolds number of Re = 136 for a 244-µm-diameter reference capillary tube with a measured solution velocity of 5.6 x 10-3 m/s. From this low value, we conclude that there is laminar flow through the empty capillary under normal conditions. If the cell monolayer height increases from 2.0 µm under isotonic conditions to 3.0 µm under extreme hypotonic conditions, the Reynolds number changes to 134, reflecting the changes in cell height and solution viscosity. In reality, the cell monolayer surface is not smooth; therefore, the flow may encounter some additional frictional losses against the cells. However, the calculated Reynolds numbers are so low (Re << 1,000) that the occlusions and morphology changes from the cell surface would have to be extreme to induce turbulent flow patterns. We conclude that there should be laminar flow through capillaries under our experimental conditions.
Calculation of expected shear stresses. The shear stress applied to the cells lining the capillary is a function of the pressures in the system, and shear stress calculations will determine whether the fluid flow remains in a physiologically relevant range. The average fluid velocity, v, of our system is found with the use of the previously determined resistance of the capillary, R1 = (8l1)/(r14) and is given by v = P1/R1A = P1r2/8l, where A is area. The shear stress at the edge of the capillary is 
= 4v/r, where r is the radius of the capillary after cells were plated. Thus the shear stress experienced by the cells in the capillary is = P1r/2l. In a typical experiment, the pressure across a single capillary would be no more than 700 Pa (7,000 dyn/cm2) for a 244-µm-diameter capillary that is 40 mm long. This maximum pressure corresponds to very low shear stress, on the order of = 0.133 Pa (0.13 dyn/cm2).
In addition to calculating the shear stresses directly from the measured pressure drop P1, we can theoretically estimate the shear stresses in the system. If we assume laminar fluid flow, the shear stress that the cells on the luminal surface of the tube will experience is the tangential stress of viscous fluid flow (2): = 2umax/r where is the shear force, umax is the maximum velocity of the laminar wavefront, and r is the capillary radius. We used fluorescein to visualize the fluid flow in the system and measured the fluid velocity to be 5.6 x 10-3 m/s for a pressure of 500 Pa (5,000 dyn/cm2) across a single capillary. Thus this estimated maximum shear stress on the cells is also quite low under isotonic conditions: = 0.250 Pa (2.50 dyn/cm2).
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METHODS |
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The experimental device was housed in a Plexiglas isolation chamber. The interior of the chamber was heated to 37°C before experiments. A round incoloy sheath tubular heating element rated at 300 W running off 115 V (Chromalox, Ogden, UT) was chosen for the chamber. It was connected in series with a limit-filled, saturated vapor sensor rated at 5 A (Omega Engineering, Stamford, CT), which acted as an adjustable temperature switch. The temperature switch had a repeatability of 2.88°C. During the experiment, the heating apparatus was turned off, and heat loss was minimized by an insulating cover. We found that the temperature was maintained within 1°C over the 10-min experiments.
Cell culture. cos-7 Were grown in DMEM supplemented with 10% FBS and 1% penicillin-streptomycin (Life Technologies) at 37°C in a 5% CO2 environment. The cells were passaged once per week at a 1-to-10 ratio when cells reached 
75% confluency. We removed cells from the flasks for experiments with 6 ml of an EDTA-buffered 0.05% trypsin solution (Life Technologies), which bathed the cells for 5 min until they became loosely adherent. The cells were then manually dislocated from the flask surface by flushing fresh media over the surface. The cells from an entire flask (>5 x 106) were concentrated into 0.5 ml of supplemented DMEM media. Capillary action was used to fill single-bore, nonfilamented borosilicate glass tubes (A-M Systems, Carlsborg, WA) with the cell mixture. The capillary tubes were placed horizontally and rolled to ensure uniform cell distribution along the luminal surface of the glass.
The cells lining the capillary lumen were assayed for confluency and viability by trypan blue exclusion at the end of each experiment. Healthy living cells exclude 0.05% trypan blue, whereas dead or sick cells take up the dye. Capillary glass tubes containing cells were examined under x60 magnification, and the percent confluency was estimated by counting healthy cells in a given length of the tube. When the cells were plated onto uncoated glass, the cells formed an 40% confluent monolayer. To increase the confluency, an extracellular substrate matrix was coated inside the capillaries to increase cell viability. The matrix proteins provided the conditions necessary to grow and differentiate in vitro. Capillary glass was washed with acetone overnight, rinsed multiple times with deionized H2O, dried overnight at 150°C, and gravity autoclaved for 35 min to sterilize for cell culture. Lyophilized human fibronectin (Becton Dickenson, Bedford, MA) was diluted to 50 µg/ml under sterile conditions. The glass was then placed into the diluted fibronectin, capillary action filled the tubes with the mixture, and the capillaries remained in the mixture for 30-60 min. Once the capillaries were coated internally with the matrix proteins, the capillaries were removed from the mixture and rinsed once with sterile deionized H2O. The capillaries were allowed to air dry at room temperature under sterile conditions. Cell confluency was estimated, using trypan blue exclusion, to be as high as 90% for fibronectin-coated capillaries. Confluencies of 30-75% (as desired) were utilized for experimental recordings.
Cell transfection. The coding region of canine PLM was cloned into the pcDNA3 vector (Stratagene). Lipofectamine was used according to the manufacturer's directions to transfect cos-7 African green monkey kidney cells (American Type Culture Collection, Manassas, VA). We verified transient expression at 1-3 days posttransfection using a fluorescent antibody technique as follows. Cells were grown on fibronectin-coated glass coverslips in Iscove's medium until 50-80% confluent and then transfected in 2 µg of DNA and 15 µg of lipofectamine for 5 h at 37°C in 5% CO2. They were fixed in paraformaldehyde (600 mosM) and incubated in DMEM containing a 1:1,000 dilution of anti-PLM B8 monoclonal antibody for 1 h. After cells were rinsed, they were incubated with 1-µl beads (Dynal) in 2 ml of DMEM for 30 min and then in 1:500 dilution of FITC-conjugated goat anti-mouse IgG for 20 min. After five or more washes, we examined the cells with a fluorescence microscope. For experiments with the device, they were harvested and loaded into capillary tubes by capillary action as described above.
Signal acquisition and processing. We measured the pressure using piezoresistive pressure transducers from the PX series, PX164-010D5V (Omega Engineering, Stanford, CT). The continuous analog differential pressure records were digitized using a 12-bit, 40-kHz AD221 Lab Master DMA daughter board (Scientific Solutions,) with the dynamic range adjusted from 0 to +5 V. The signal was acquired by pCLAMP 5.1 software (Axon Instruments), and the voltage output from each transducer was acquired on a separate analog-to-digital channel and digitized at 1.7 Hz. The P2-to-P1 ratios (P2/P1) of the time series were then calculated offline for each of the experiments. The normalization, which was necessary because of variations in P2/P1 due to subtle differences in capillary tubes or the degree of cell confluency, was performed by 1) subtracting the P2/P1 value under isotonic conditions and then 2) dividing by the largest P2/P1 value, at the peak of the cell swelling or shrinking response. The normalized records were averaged. To compare RVD kinetic differences between cell types, the resistances were normalized to 0 at the beginning of each experiment and 1 at the peak of the passive swelling response.
Solutions and experimental protocols. We tested the effects of three solutions of identical ionic strength but differing osmotic pressures. The hypotonic 250 mosmol/kgH2O saline solution, which caused cultured cos-7 cells to swell, contained (in mM) 100 NaCl, 2.5 MgCl2, 2.5 CaCl2, and 10 HEPES, pH 7.2. The isotonic 300 mosmol/kgH2O solution, which had the same osmotic pressure as the DMEM culture media, had no effect on cos-7 cell size and was adjusted to the final 300 mosmol/kgH2O osmolality by adding mannitol to the hypotonic solution. Hypertonic 350 mosmol/kgH2O solution was also adjusted from the original hypotonic solution with mannitol, and this solution induced the cells to shrink. After the capillary tubes were positioned, records were obtained for 1 min in the isotonic solution, 7 min in either hypotonic or hypertonic test solution, and 2 min in the original isotonic solution.
Measurements of solution viscosity. Bulk solution viscosity was empirically measured by an Oswald-type viscometer (VWR Scientific, West Chester, PA) relative to deionized H2O. Relative viscosity (/0) was calculated according to (/o) = (t/to) where t is the time measured in the viscometer and the subscript 0 indicates measurements on water (20). All viscosity measurements were made at the experimental temperature (37°C) by immersing the viscometer in a temperature-controlled, circulating water bath. The viscosities of the hypotonic, isotonic, and hypertonic solutions were 1.009 ± 0.004, 1.032 ± 0.004, and 1.049 ± 0.004 x 10-5 kg·m-1·s-2 (n = 25, mean ± SE), respectively. Pure water has a viscosity of 1.002 x 10-5 kg·m-1·s-2. This corresponds to an average increase of solution viscosity of 0.7, 3.0, and 4.7% for the hypotonic, isotonic, and hypertonic solutions relative to water.
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RESULTS |
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Verification of Poiseuille's law for system. Poiseuille's fluid flow equation for rigid-wall capillary systems was empirically verified for our recording system by applying a constant pressure across the system and measuring resulting fluid flow through three pairs of capillaries with different radii of 1.15 x 10-4, 2.82 x 10-4, and 4.03 x 10-4 m. The theoretical fluid flow was calculated for each capillary pair and found to be 8.33 x 10-4, 2.08 x 10-3, and 1.00 x 10-2 ml/s, respectively. These calculated values were compared with those empirically measured and found to be, respectively, 4, 2, and 1% lower than the measured values of 8.02 x 10-4, 2.04 x 10-3, and 0.99 x 10-2 ml/s. We concluded that the limiting resistive components of our recording system were the two capillaries in series and that a radius change in the cell capillary would directly dictate changes in the recorded pressures P2 and P1.
Cells contribute to R2. The presence of cultured cells should make a measurable contribution to P2 - P1, the difference in pressure drops across the capillary tubes with and without cells, respectively. We measured this difference in 9 pairs of capillaries containing no cells and in 11 pairs of capillaries containing cells in isotonic conditions. The average ± SD transducer output for empty capillaries was 0.15 ± 0.08 V (n = 9) and 0.63 ± 0.12 V (n = 11) for capillaries containing cells, which translates to 36.7 ± 16.2 Pa (n = 9) and 170.3 ± 32.4 Pa (n = 11) for the empty capillaries and pairs containing cells, respectively (P < 0.001, Student's t-test). This 133.6-Pa difference between the average pressures represents a 66.8-Pa increase for T2 and, compared with the average reading of 675.7 Pa, a 9.8% increase in the signal due to the presence of cells.
Static and dynamic relationship between pressure differences and cell confluency. Given that the presence of cells contributed to the total resistance of the system, we hypothesized that there is a relationship between the percent confluency of the cell monolayer and the peak amplitude of the change in resistance R2 due to cell volume changes. That is, capillaries with more cells should show larger evidence for differences in pressure between the two transducers and a higher peak response for volume changes. We plated capillaries at confluencies of 30, 40, 50, and 60% of the total luminal surface area using sequential dilutions of cells. The cells were perfused with hypertonic solution to induce passive shrinking. The P2 - P1 voltage (V) was calculated for each treatment group as well as for a group of blank capillaries. The blank capillaries had an average signal difference of 0.18 ± 0.06 V (n = 13). The signal differences for the different cell confluencies were corrected for this offset by subtracting 0.18 V from each measured difference. The corrected P2 - P1 values for each treatment were grouped together and plotted as a box plot in Fig. 2A. As shown, the corrected voltage difference between the raw signals P2 and P1 monotonically increases with the confluency of cells lining the capillary.
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In addition to this static difference, we might expect larger dynamic changes when more cells were present. Figure 2B is a plot of the peak amplitude of the P2/P1 passive hypertonic cell shrinkage response as a function of the corrected V, which we used as an estimate of cell number. Because individual capillaries had different confluencies, it was not surprising that the corrected P2 - P1 values had a broad distribution. In keeping with our hypothesis, there was a highly significant correlation of moderate degree between voltage offset and peak signal output (P < 0.001, Spearman's rank order coefficient = 0.5). These two results provide reassurance that the cells in the R2 capillary contribute to both static and dynamic pressure drops that we record experimentally.
Dynamic changes in cell volume detected with the new method. Figure 3 shows recordings from four experimental conditions. The pair of records in Fig. 3A is of P1 and P2 for blank capillary tubes. The voltage signals are constant and demonstrate that there is no significant drift of the signal or significant artifact from the solution switching at minutes 3 and 6. Shown in Fig. 3, C, E, and G, are representative signals from capillaries containing cells. In each panel, there is a larger difference between P1 and P2, as expected from the increase in R2 due to the cells. The records in Fig. 3C were obtained when the test solution was the isotonic solution. As expected, there was no change in the signal as the solutions were changed. The records in Fig. 3E were obtained when the test solution was the hypotonic solution. P2 increases, consistent with the idea that R2 rises as the cells swell. Because the total pressure drop does not change, P1 correspondingly decreases. Note that the two pressures slowly return toward baseline, suggesting that a RVD mechanism is present. The records in Fig. 3G were obtained when the test solution was hypertonic. P2 falls and P1 rises, as expected for a decrease in R2 as the cells shrink. By inspection, we observed the expected directional changes in P1, P2, or both for each experiment in hypotonic and hypertonic solution conditions.
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We calculated P2/P1 and averaged the normalized results from several experiments. The results are shown in Fig. 3, B, D, F, and H. In Fig. 3, B and D, when no osmotic changes in cell volume were expected, the ratios are unchanging. Figure 3F shows a rise in the ratio followed by a gradual fall, in keeping with the expected finding of an increase in cell volume followed by RVD. Figure 3H shows a fall in the ratio with no recovery, as expected if the cells shrank passively and did not activate a regulatory volume increase. We interpret this finding to reflect loss of intracellular solute during the RVD process.
Calculated radius changes based on experimental results. Using the example traces from Fig. 3, E and G, we calculated the approximate radius change in the capillaries due to the cell volume changes. We assumed all cell volume changes reflect only a change in the effective radius of the capillary. We calculated a change in the radius of the capillary containing the cells by using Poiseuille's law, where we know P1/R1 = P2/R2. The resistance of the reference capillary is R1 = (8l)/(r14), where r1 is the internal radius of the capillary. The resistance of the capillary containing the cells is R2 = (8l)/(r24), where r2 is the internal radius of the capillary. The difference r2 - r1 would then give the height of the cell monolayer, and we calculated this height under isotonic and hypertonic conditions after correcting for the characteristic voltage offset of the blank capillaries (Table 1). Our calculations revealed that, under hypotonic conditions, the calculated radius of the capillary containing cells decreased by 0.32 µm at the peak of the hypotonic response, a 17.6% increase in cell height. Similarly, under hypertonic conditions, the radius increased by 0.29 µm at the peak of the passive cell shrinking, a 15.1% decrease in cell height. These calculations are in reasonable agreement with an estimated 20% volume shift as a result of extreme osmotic stress in many cell types (3).
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A cell line overexpressing PLM. We expressed canine PLM in cos-7 cells (Green African monkey kidney cells) using a lipofection technique. Cells were fixed 24 h after transfection and incubated with a mouse IgG2a monoclonal antibody to PLM that attached to expressed or endogenous PLM or PLM-like membrane protein. Cells carrying this primary anti-PLM antibody were detected in two ways. First, they were incubated with microbeads carrying a rat antibody to mouse IgG2a. The presence of a bead on a cell would be interpreted as a demonstration of PLM expression by that cell. To confirm this assumption, the cells were next incubated with a fluorescence-tagged goat anti-mouse antibody. The amount of fluorescence can be taken to be proportional to the amount of PLM expressed. As expected, the beaded cells had increased fluorescence, demonstrating PLM expression (Fig. 4).
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Enhanced volume regulation in cells overexpressing PLM. We used the new device to compare RVD in cells expressing transiently transfected PLM (n = 10 trials) and cells transfected with the vector alone (n = 7 trials). We found that the PLM-overexpressing cells had a faster RVD than the control cells. The normalized P2/P1 are shown in Fig. 5. The straight lines are linear regressions with slopes of -1.95 x 10-3 for sham-transfected and -4.79 x 10-3 s-1 for PLM-expressing cells. This more than twofold increase in RVD in PLM-overexpressing cells is highly significant (z = 5.2 for the regression slopes, P < 0.001).
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DISCUSSION |
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TOPABSTRACTTHEORYMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Comparison with existing methods. There are several methods presently used to measure cell volumes. Video microscopy has been used very successfully to measure changes in cell volume from cardiac myocytes (6, 11, 30). Although useful for geometrically simple cells, this method is dependent on cell morphology and cannot be applied to most cell types. Another widely used technique to measure cell volume uses electronic particle counters, such as the commercially available Coulter counter, to quantitatively measure cell volume changes in populations of cells (12, 22, 25). The practical disadvantage of this method is that volume measurements must be recorded at discrete time points. Our method records continuous cell volume changes and allows for multiple experimental conditions in a single trial. Moreover, the new method allows for measurements under more physiological conditions, i.e., cells attached to a surface as opposed to cells in suspension as in a light-scattering method (26). Fluorescent probes have also been employed to measure cell volume changes (1, 9, 13, 29). Unlike the single cell fluorescent methods, our method records from populations of cells, and there is no danger of phototoxicity. Volume changes have also been recorded by measuring the electrical impedance above a cell monolayer in an enclosed recording chamber (19). This method's major disadvantage is that the resistivity of the solution above the cell monolayer changes with osmolarity. Thus the stimulus to induce cell swelling also alters the recorded parameter. Unlike this method, we are measuring hydrostatic pressure changes that are independent of the perfusing solution's osmotic strength.
Limitations of the method. The new method also has limitations. First, the shear stress that the cells experience varies due to changes in the pressure across the capillary. However, as we showed, the estimated shear stress is low. Second, cells might become detached during the experiment and could result in a drift of the recorded pressures. Our inspection of the capillary tubes, however, suggested that the fibronectin treatment was successful in securing cells to the luminal surface. Moreover, the voltage signal was stable during the initial equilibrium period of the experiments, suggesting that no large numbers of cells were removed by the initial flow. Third, the time response is limited by the rate of flow through the system and is thus slower to report volume changes than a light-scattering method (26). Fourth, the extent of cell confluency is correlated with the peak amplitude of the recorded responses, and these amplitudes vary slightly from trial to trial. If we are interested in comparing the kinetic differences between different cell types or for different experimental conditions, we can simply normalize the data from baseline to peak to correct for amplitude variation as shown previously in Fig. 3, F and H. Additionally, when cells are plated at the same confluency, it is possible to compare raw amplitude responses between experimental conditions, reflecting different magnitude RVD responses. Finally, we noted that the system was sensitive to extraneous factors such as air bubbles in the tubing and switching of the solenoid valve controlling solution flow. Thus meticulous care was required in the experimental technique.
Increased RVD in cells overexpressing PLM. We tested whether the new cell volume measurement strategy could detect changes in RVD kinetics in cell lines with altered volume regulation constituents. Hence, we compared cell lines differing in the amount of expressed PLM, a membrane protein with a single transmembrane domain that is a major substrate for protein kinases in heart and skeletal muscle (24). Two possible physiological functions have been proposed for PLM.
First, electrophysiological evidence has pointed to a possible role as an ion channel that is selective for zwitterionic osmolytes such as taurine (4, 17, 21). In support of this hypothesis are the findings that the level of PLM expression correlates with the degree and kinetics of RVD in cultured cells. Pasantes-Morales and colleagues have shown increased rates of RVD and of taurine efflux in HEK-293 cells stably expressing transfected PLM (22) and decreased rates in astrocytes treated with PLM anti-sense oligonucleotides (23). Kirschner and coworkers (16) more recently found ion channel activity in hepatocytes under hypertonic conditions that was similar to that of reconstituted PLM. Our present findings affirm those of Pasantes-Morales and colleagues and introduce an entirely novel measurement approach (22, 23).
Second, molecular and biochemical evidence suggests a role for PLM as a modulator of membrane-bound ion-motive pumps. PLM, or FXYD1, is a member of a family of single transmembrane proteins named for a segment of conserved amino acid sequence (31). Several members, including PLM, have been demonstrated to associate with, and modify the function of, the Na+-K+-ATPase (7, 8). Because this enzyme causes net loss of osmolytes, it has a role in regulation of cell volume (28).
Both putative functions are compatible with the findings of altered cell volume regulation as PLM expression is changed.
Summary. We have developed a technique for following dynamic changes in cell volume based on resistance changes in capillary tubes coated with cultured cells expressing transfected proteins. The method was used to confirm the findings of Pasantes-Morales and colleagues (22), who used a Coulter counter technique to show that HEK-293 cells overexpressing PLM had enhanced RVD. These findings suggest that PLM expression plays a role in regulation of cell volume but do not clarify its role in the process as an ion channel itself or as a regulator of other ion channels.
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ACKNOWLEDGMENTS |
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TOPABSTRACTTHEORYMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Present address of C. E. Davis: The Charles Stark Draper Laboratory, 555 Technology Square, MS 37, Cambridge, MA 02141 (E-mail: cristina{at}alumni.duke.edu).
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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) varied.
