Label-free, non-contact determination of resting membrane potential using dielectrophoresis.
Journal
Scientific reports
ISSN: 2045-2322
Titre abrégé: Sci Rep
Pays: England
ID NLM: 101563288
Informations de publication
Date de publication:
09 Aug 2024
09 Aug 2024
Historique:
received:
06
03
2024
accepted:
30
07
2024
medline:
10
8
2024
pubmed:
10
8
2024
entrez:
9
8
2024
Statut:
epublish
Résumé
Measurement of cellular resting membrane potential (RMP) is important in understanding ion channels and their role in regulation of cell function across a wide range of cell types. However, methods available for the measurement of RMP (including patch clamp, microelectrodes, and potential-sensitive fluorophores) are expensive, slow, open to operator bias, and often result in cell destruction. We present non-contact, label-free membrane potential estimation which uses dielectrophoresis to determine the cytoplasm conductivity slope as a function of medium conductivity. By comparing this to patch clamp data available in the literature, we have demonstratet the accuracy of this approach using seven different cell types, including primary suspension cells (red blood cells, platelets), cultured suspension cells (THP-1), primary adherent cells (chondrocytes, human umbilical mesenchymal stem cells), and adherent (HeLa) and suspension (Jurkat) cancer cell lines. Analysis of the effect of ion channel inhibitors suggests the effects of pharmaceutical agents (TEA on HeLa; DMSO and neuraminidase on red blood cells) can also be measured. Comparison with published values of membrane potential suggest that the differences between our estimates and values recorded by patch clamp are accurate to within published margins of error. The method is low-cost, non-destructive, operator-independent and label-free, and has previously been shown to allow cells to be recovered after measurement.
Identifiants
pubmed: 39122771
doi: 10.1038/s41598-024-69000-7
pii: 10.1038/s41598-024-69000-7
doi:
Types de publication
Journal Article
Langues
eng
Sous-ensembles de citation
IM
Pagination
18477Subventions
Organisme : Khalifa University of Science, Technology and Research
ID : FSU-2022-020
Organisme : Khalifa University of Science, Technology and Research,United Arab Emirates
ID : 8474000443
Informations de copyright
© 2024. The Author(s).
Références
Carmeliet, E. From Bernstein’s rheotome to Neher-Sakmann’s patch electrode. The action potential. Physiol. Rep. 7, e1386 (2019).
doi: 10.14814/phy2.13861
Cole, K. S. & Curtis, H. J. Electrical impedance of the squid giant axon during electrical activity. J. Gen. Physiol. 22, 649–670 (1939).
pubmed: 19873125
pmcid: 2142006
doi: 10.1085/jgp.22.5.649
Hodgkin, A. L. & Horowicz, P. The influence of potassium and chloride ions on the membrane potential of single muscle fibres. J. Physiol. 148, 127–160 (1959).
pubmed: 14402240
pmcid: 1363113
doi: 10.1113/jphysiol.1959.sp006278
Goldman, D. E. Potential, impedance, and rectification in membranes. J. Gen. Physiol. 27, 37–60 (1943).
pubmed: 19873371
pmcid: 2142582
doi: 10.1085/jgp.27.1.37
Hodgkin, A. L. & Katz, B. The effect of sodium ions on the electrical activity of the giant axon of the squid. J. Physiol. 108, 37–77 (1949).
pubmed: 18128147
pmcid: 1392331
doi: 10.1113/jphysiol.1949.sp004310
Abdul, K. L., Stacey, M. & Barratt-Jolley, R. Emerging roles of the membrane potential: Action beyond the action potential. Front. Physiol. 21, 1661 (2018).
doi: 10.3389/fphys.2018.01661
Sachs, H. G., Stambrook, P. J. & Ebert, J. D. Changes in membrane potential during the cell cycle. Exp. Cell Res. 83, 362–366 (1974).
pubmed: 4856272
doi: 10.1016/0014-4827(74)90350-4
Stratford, J. P. et al. Electrically induced bacterial membrane-potential dynamics correspond to cellular proliferation capacity. Proc. Nat. Acad. Sci U.S.A. 116, 9552–9557 (2019).
doi: 10.1073/pnas.1901788116
Yang, M. & Brackenbury, W. J. Membrane potential and cancer progression. Front. Physiol. 17, 185 (2013).
Labeed, F. H., Coley, H. M., Thomas, H. & Hughes, M. P. Assessment of multidrug resistance reversal using dielectrophoresis and flow cytometry. Biophys. J. 85, 2028–2034 (2003).
pubmed: 12944315
pmcid: 1303374
doi: 10.1016/S0006-3495(03)74630-X
Kline, D., Jaffe, L. A. & Tucker, R. P. Fertilization potential and polyspermy prevention in the egg of the nemertean, Cerebratulus lacteus. J. Exp. Zool. 236, 45–52 (1985).
pubmed: 4056704
doi: 10.1002/jez.1402360107
Kobayashi, W., Baba, Y., Shimozawa, T. & Yamamoto, T. S. The fertilization potential provides a fast block to polyspermy in lamprey eggs. Dev. Biol. 161, 552–562 (1994).
pubmed: 8314001
doi: 10.1006/dbio.1994.1053
Henslee, E. A. et al. Rhythmic K+ transport regulates the circadian clock in human red blood cells. Nat. Commun. 8, 1978 (2017).
pubmed: 29215003
pmcid: 5719349
doi: 10.1038/s41467-017-02161-4
Beale, A. D. et al. Casein Kinase 1 underlies temperature compensation of circadian rhythms in human red blood cells. J. Biol. Rhythms. 34, 144–153 (2019).
pubmed: 30898060
pmcid: 6458989
doi: 10.1177/0748730419836370
Erndt-Marino, J. & Hahn, M.S. Membrane potential controls macrophage activation. In Proc. 10th World Biomaterials Congress, Montréal, Canada, 17 May–22 May (2016).
MacIntyre, D. E. & Rink, T. J. The role of platelet membrane potential in the initiation of platelet aggregation. Thromb. Haemost. 26, 22–26 (1982).
Hai, A., Shappir, J. & Spira, M. E. Long-term, multisite, parallel, in-cell recording and stimulation by an array of extracellular microelectrodes. J. Neurophysiol. 104, 559–568 (2010).
pubmed: 20427620
doi: 10.1152/jn.00265.2010
Neher, E. & Sakmann, B. Single-channel currents recorded from membrane of denervated frog muscle fibres. Nature 260, 799–802 (1976).
pubmed: 1083489
doi: 10.1038/260799a0
Tasaki, I., Teorell, T. & Spyropoulos, C. S. Movement of radioactive tracers across squid axon membrane. Am. J. Physiol. 200, 11–22 (1961).
pubmed: 13775386
doi: 10.1152/ajplegacy.1961.200.1.11
Adams, D. S. & Levin, M. Measuring resting membrane potential using the fluorescent voltage reporters DiBAC4(3) and CC2-DMPE. Cold Spring Harb. Protoc. 4, 459–464 (2012).
Kulkarni, R. U. & Miller, E. W. Voltage imaging: Pitfalls and potential. Biochemistry 56, 5171–5177 (2017).
pubmed: 28745864
doi: 10.1021/acs.biochem.7b00490
Ortiz, G., Naing, S. H. H., Muller, V. R. & Miller, E. W. Synthesis of sulfonated carbofluoresceins for voltage imaging. J. Am. Chem. Soc. 141, 6631–6638 (2019).
pubmed: 30978010
pmcid: 6546115
doi: 10.1021/jacs.9b01261
McCann, J. T. et al. Flipping the switch: Reverse-demand voltage-sensitive fluorophores. J. Am. Chem. Soc. 144, 13050–13054 (2022).
pubmed: 35834763
pmcid: 9462387
doi: 10.1021/jacs.2c05385
Bullen, A. & Saggau, P. High-speed, random-access fluorescence microscopy: II. Fast quantitative measurements with voltage-sensitive dyes. Biophys. J. 76, 2272–2287 (1999).
pubmed: 10096922
pmcid: 1300200
doi: 10.1016/S0006-3495(99)77383-2
Brinks, S., Klein, A. J. & Cohen, A. E. Two-photon lifetime imaging of voltage indicating proteins as a probe of absolute membrane voltage. Biophys. J. 109, 914–921 (2015).
pubmed: 26331249
pmcid: 4564826
doi: 10.1016/j.bpj.2015.07.038
Lazzari-Dean, J. R., Gest, A. M. & Miller, E. W. Optical estimation of absolute membrane potential using fluorescence lifetime imaging. eLife 8, e44522 (2019).
pubmed: 31545164
pmcid: 6814365
doi: 10.7554/eLife.44522
Saminathan, A. et al. A DNA-based voltmeter for organelles. Nat. Nanotechnol. 16, 96–103 (2021).
pubmed: 33139937
doi: 10.1038/s41565-020-00784-1
Moersdorf, D. et al. Transmembrane potential of red blood cells under low ionic strength conditions. Cell Physiol. Biochem. 31, 875–882 (2013).
pubmed: 23817052
doi: 10.1159/000350105
Pethig, R. Dielectrophoresis: Theory Methodology and Biological Applications (Wiley, 2017).
doi: 10.1002/9781118671443
Hoettges, K. F. et al. Ten-second electrophysiology: Evaluation of the 3DEP Platform for high-speed, high-accuracy cell analysis. Sci. Rep. 9, 19153 (2019).
pubmed: 31844107
pmcid: 6915758
doi: 10.1038/s41598-019-55579-9
Mansoorifar, A., Koklu, A., Ma, S., Raj, G. V. & Beskok, A. Electrical impedance measurements of biological cells in response to external stimuli. Anal. Chem. 90, 4320–4327 (2018).
pubmed: 29402081
doi: 10.1021/acs.analchem.7b05392
Sawai, S., Shukri, N. A., Mohktar, M. S., Safwani, W. & Zaman, W. K. Dielectrophoresis-based discrimination of hepatic carcinoma cells following treatment with cytotoxic agents. Eng. Sci. Technol. Int. J. 25, 100990 (2022).
Wang, H. C., Nguyen, N. V., Lin, R. Y. & Jen, C. P. Characterizing esophageal cancerous cells at different stages using the dielectrophoretic impedance measurement method in a microchip. Sensors (Basel) 17, 1053 (2017).
pubmed: 28481265
doi: 10.3390/s17051053
Graham, K. A. et al. A dielectrophoretic method of discrimination between normal oral epithelium, and oral and oropharyngeal cancer in a clinical setting. Analyst 140, 5198–5204 (2015).
pubmed: 26086875
pmcid: 11226305
doi: 10.1039/C5AN00796H
Gascoyne, P., Pethig, R., Satayavivad, J., Becker, F. F. & Ruchirawat, M. Dielectrophoretic detection of changes in erythrocyte membranes following malarial infection. Biochim. Biophys. Acta 1323, 240–252 (1997).
pubmed: 9042346
doi: 10.1016/S0005-2736(96)00191-5
Vykoukal, J., Vykoukal, D. M., Freyberg, S., Alt, E. U. & Gascoyne, P. R. C. Enrichment of putative stem cells from adipose tissue using dielectrophoretic field-flow fractionation. Lab Chip 8, 1386–1393 (2008).
pubmed: 18651083
pmcid: 2726253
doi: 10.1039/b717043b
Giduthuri, A. T., Theodossiou, S. K., Schiele, N. R. & Srivastava, S. K. Dielectrophoretic characterization of tenogenically differentiating mesenchymal stem cells. Biosensors (Basel) 11, PMC7919818 (2021).
Ismail, A., Hughes, M. P., Mulhall, H. J., Oreffo, R. O. C. & Labeed, F. H. Characterization of human skeletal stem and bone cell populations using dielectrophoresis. J. Tissue Eng. Regen. Med. 9, 162–168 (2015).
pubmed: 23225773
doi: 10.1002/term.1629
Hughes, M. P. et al. Vm-related extracellular potentials observed in red blood cells. Sci. Rep. 11, 19446 (2021).
pubmed: 34593849
pmcid: 8484267
doi: 10.1038/s41598-021-98102-9
Lewis, R. et al. The role of the membrane potential in chondrocyte volume regulation. J. Cell Physiol. 226, 2979–2986 (2011).
pubmed: 21328349
pmcid: 3229839
doi: 10.1002/jcp.22646
Sukhorukov, V. L. et al. Phloretin-induced changes of lipophilic ion transport across the plasma membrane of mammalian cells. Biophys. J. 81, 1006–1013 (2001).
pubmed: 11463642
pmcid: 1301570
doi: 10.1016/S0006-3495(01)75758-X
Hübner, Y., Hoettges, K. F., Kass, G. E. N., Ogin, S. L. & Hughes, M. P. Parallel measurements of drug actions on erythrocytes by dielectrophoresis, using a three-dimensional electrode design. IEE Proc. Bionanotechnol. 4, 21–25 (2005).
Hoettges, K. F. et al. Dielectrophoresis-activated multiwell Plate for label-free high-throughput drug assessment. Anal. Chem. 80, 2063–2068 (2008).
pubmed: 18278948
doi: 10.1021/ac702083g
Broche, L. M., Hoettges, K. F., Ogin, S. L., Kass, G. E. N. & Hughes, M. P. Rapid, automated measurement of dielectrophoretic forces using DEP-activated microwells. Electrophoresis 32, 2393–2399 (2011).
pubmed: 21800330
doi: 10.1002/elps.201100063
Huang, L., Huang, Q. Y. & Huang, H. Q. The evidence of HeLa cell apoptosis induced with tetraethylammonium using proteomics and various analytical methods. J. Biol. Chem. 289, 2217–2229 (2014).
pubmed: 24297172
doi: 10.1074/jbc.M113.515932
Okada, Y., Ogawa, M., Aoki, N. & Izutsu, K. The effect of K+ on the membrane potential in HeLa cells. Biochim. Biophys. Acta 291, 116–126 (1973).
pubmed: 4684603
doi: 10.1016/0005-2736(73)90066-7
Stein, M. A., Mathers, D. A., Yan, H., Baimbridge, K. G. & Finlay, B. B. Enteropathogenic Escherichia coli markedly decreases the resting membrane potential of Caco-2 and HeLa human epithelial cells. Infect. Immun. 64, 4820–4825 (1996).
pubmed: 8890244
pmcid: 174450
doi: 10.1128/iai.64.11.4820-4825.1996
Szabo, I. et al. Formation of anion-selective channels in the cell plasma membrane by the toxin VacA of Helicobacter pylori is required for its biological activity. EMBO J. 18, 5517–5527 (1999).
pubmed: 10523296
pmcid: 1171620
doi: 10.1093/emboj/18.20.5517
Hsu, K., Han, J., Shinlapawittayatorn, K., Deschenes, I. & Marbán, E. Membrane potential depolarization as a triggering mechanism for Vpu-mediated HIV-1 release. Biophys. J. 99, 1718–1725 (2010).
pubmed: 20858415
pmcid: 2941015
doi: 10.1016/j.bpj.2010.07.027
Colden-Stanfield, M. & Gallin, E. K. Modulation of K+ currents in monocytes by VCAM-1 and E-selectin on activated human endothelium. Am. J. Physiol. 275, C267–C277 (1998).
pubmed: 9688858
doi: 10.1152/ajpcell.1998.275.1.C267
Colden-Stanfield, M. & Scanlon, M. VCAM-1-induced inwardly rectifying K1 current enhances Ca21 entry in human THP-1 monocytes. Am. J. Physiol. Cell Physiol. 179, C488–C494 (2000).
doi: 10.1152/ajpcell.2000.279.2.C488
Funabashi, K., Fujii, M., Yamamura, H., Ohya, S. & Imaizumi, Y. Contribution of chloride channel conductance to the regulation of resting membrane potential in chondrocytes. J. Pharmacol. Sci. 113, 94–99 (2010).
pubmed: 20453434
doi: 10.1254/jphs.10026SC
Vasanthakumar, N. Effect of hydrogen sulphide (h2s) on the electrophysiological properties of bovine articular chondrocytes. PhD thesis, Christian Medical College, Vellore (2018).
Zavodnik, I. B., Piasecka, A., Szosland, K. & Bryszewska, M. Human red blood cell membrane potential and fluidity in glucose solutions. Scand. J. Clin. Lab Invest. 57, 59–63 (1997).
pubmed: 9127458
doi: 10.3109/00365519709057819
Jay, A. W. L. & Burton, A. C. Direct measurement of potential difference across the human red blood cell membrane. Biophys. J. 9, 115–121 (1969).
pubmed: 5764221
pmcid: 1367420
doi: 10.1016/S0006-3495(69)86372-1
Harberts, J., Kusch, M., O’Sullivan, J., Zierold, R. & Blick, R. H. A Temperature-controlled patch clamp platform demonstrated on Jurkat T lymphocytes and human induced pluripotent stem cell-derived neurons. Bioengineering (Basel) 7, 46 (2020).
pubmed: 32455868
doi: 10.3390/bioengineering7020046
Harberts, J. et al. Culturing and patch clamping of Jurkat T cells and neurons on Al
pubmed: 35520244
pmcid: 9063011
doi: 10.1039/C8RA05320K
Heubach, J. F. et al. Electrophysiological properties of human mesenchymal stem cells. J. Physiol. 554, 659–672 (2004).
pubmed: 14578475
doi: 10.1113/jphysiol.2003.055806
Li, G. R., Sun, H., Deng, X. & Lau, C. P. Characterization of ionic currents in human mesenchymal stem cells from bone marrow. Stem Cells 23, 371–382 (2005).
pubmed: 15749932
doi: 10.1634/stemcells.2004-0213
Sundelacruz, S., Levin, M. & Kaplan, D. L. Membrane potential controls adipogenic and osteogenic differentiation of mesenchymal stem cells. PLoS ONE 3, e3737 (2008).
pubmed: 19011685
pmcid: 2581599
doi: 10.1371/journal.pone.0003737
Zhang, J. et al. Regulation of cell proliferation of human induced pluripotent stem cell-derived mesenchymal stem cells via ether-à-go-go 1 (hEAG1) potassium channel. Am. J. Physiol. Cell Physiol. 303, C115–C125 (2012).
pubmed: 22357737
doi: 10.1152/ajpcell.00326.2011
Mahaut-Smith, M. P. Patch-Clamp Recordings of Electrophysiological Events in the Platelet and Megakaryocyte. In Platelets and Megakaryocytes. Methods in Molecular Biology Vol. 273 (eds Gibbins, J. M. & Mahaut-Smith, M. P.) (Humana Press, 2004).
Maruyama, Y. A patch-clamp study of mammalian platelets and their voltage-gated potassium current. J. Physiol. 391, 467–485 (1987).
pubmed: 2451010
pmcid: 1192226
doi: 10.1113/jphysiol.1987.sp016750
Hughes, M. P., Fry, C. H. & Labeed, F. H. Cytoplasmic anion/cation imbalances applied across the membrane capacitance may form a significant component of the resting membrane potential of red blood cells. Sci. Rep. 12, 15005 (2022).
pubmed: 36056086
pmcid: 9440063
doi: 10.1038/s41598-022-19316-z
Ponce, A. Expression of voltage dependent potassium currents in freshly dissociated rat articular chondrocytes. Cell Physiol. Biochem. 18, 35–46 (2006).
pubmed: 16914888
doi: 10.1159/000095134
Pelletier, R. P. & Hablitz, J. J. Tetraethylammonium-induced synaptic plasticity in rat neocortex. Cerebral Cortex 6, 771–780 (1996).
pubmed: 8922333
doi: 10.1093/cercor/6.6.771
Nowak, L. M. & MacDonald, R. L. Substance P: Ionic basis for depolarising responses of mouse spinal cord neurons in cell culture. J. Neurosci. 2, 1119–1128 (1982).
pubmed: 6180151
pmcid: 6564281
doi: 10.1523/JNEUROSCI.02-08-01119.1982
Kim, N. et al. Role of K+ channels to resting membrane potential of rabbit middle cerebral arterial smooth muscle cells. Korean J. Physiol. Pharmacol. 3, 547–554 (1999).
Lyklema, J. Fundamentals of Interface and Colloid Science (Academic Press, 1991).
Laszuk, P., Urbaniak, W. & Petelska, A. D. The equilibria in lipid-lipoic acid systems: Monolayers, microelectrophoretic and interfacial tension studies. Molecules 25, 3678 (2020).
pubmed: 32806764
pmcid: 7465766
doi: 10.3390/molecules25163678
Ohki, S. Electrical potential of an asymmetric membrane. J. Coll. Int. Sci. 37, 318–324 (1971).
doi: 10.1016/0021-9797(71)90299-2
Pethig, R. Dielectric properties of Biological Materials (Wiley, 1979).
Strickholm, A. Ionic permeability of K, Na, and Cl in potassium-depolarized nerve. Dependency on pH, cooperative effects, and action of tetrodotoxin. Biophys. J. 35, 677–697 (1981).
pubmed: 7272457
pmcid: 1327556
doi: 10.1016/S0006-3495(81)84820-5
Balach, M. M., Casale, C. H. & Campetelli, A. N. Erythrocyte plasma membrane potential: Past and current methods for its measurement. Biophys Rev. 11, 995–1005 (2019).
pubmed: 31741171
pmcid: 6874941
doi: 10.1007/s12551-019-00603-5
Sperelakis, N. Cell Physiology Sourcebook 4th edn. (Academic Press, 2011).