Cell membrane tension in disease and emerging strategies for its regulation and measurement

Regulators: Three axes governing membrane tension

Membrane tension in living cells emerges from the dynamic interplay of multiple mechanical processes rather than from membrane stretch alone. In cellular systems, membrane tension is primarily regulated by three principal mechanical axes: membrane–cortex attachment, actomyosin contractility, and membrane surface availability.^5,32 These mechanisms determine how mechanical forces are transmitted to plasma membrane and how cells maintain surface tension homeostasis under changing mechanical conditions (Figure 3).OPEN IN VIEWER

Membrane-cortex attachment

The effective mechanical tension at the plasma membrane depends on its mechanical coupling to the underlying actin cortex, as membrane alone cannot sustain stable tension without cytoskeletal support (Figure 3(a)).^14,33 Membrane–cortex attachment is mediated by linker proteins such as the ERM family and spectrin-ankyrin complexes, which physically connect transmembrane proteins and phospholipids to cortical actin filaments, forming a mechanically resilient submembranous scaffold.^14,34 Through this linkage, forces generated within the actin cortex are transmitted directly to the plasma membrane. The degree of membrane–cortex coupling is regulated upstream by the phosphorylation state of ERM proteins and phosphatidylinositol 4,5-bisphosphate (PIP2) levels. ³⁵ ERM activation occurs in a two-step process. PIP2 binding to the Four-point-one, Ezrin, Radixin, Moesin domain (FERM) domain induces a conformational change, followed by C-terminal threonine phosphorylation, stabilizing membrane–cortex attachment and increasing effective membrane tension.^35,36 Conversely, PIP2 depletion via phospholipase C (PLC) promotes ERM dephosphorylation and release from the cortex, weakening membrane–cortex coupling, reducing resistance to membrane deformation, and promoting membrane detachment. ³⁵ Such reduction in membrane–cortex adhesion facilitates protrusive activity and favors bleb-based migration modes. Thus, membrane tension in cells does not simply reflect membrane stretch but rather arises from the degree of mechanical coupling between the plasma membrane and the cortical cytoskeleton. ⁵

Actomyosin contractility

Actomyosin contractility, the force-generating process in which non-muscle myosin II motors slide anti-parallel actin filaments to produce mechanical stress within the cortical network, represents the second major axis regulating membrane tension by generating cortical mechanical stress (Figure 3(b)). ³³ The actin cortex, a thin contractile network underlying the plasma membrane, produces tension through non-muscle myosin II–driven contraction. ¹² Because the plasma membrane is mechanically coupled to the actin cortex via membrane–cortex linkers such as ERM proteins, contractile forces generated within the actin networks are transmitted to the cell surface.^12,14 Thus, increased actomyosin contractility elevates cortical tension, stiffens the cortex, and raises apparent membrane tension.^12,37

Actomyosin contractility is primarily regulated by the RhoA–Rho-associated coiled-coil containing protein kinase (ROCK)–myosin II signaling pathway.^38,39 Activation of RhoA stimulates ROCK, which phosphorylates myosin light chains (MLC) at Ser19, increasing motor activity and promoting cortical tension. ³⁹ Additionally, ROCK simultaneously inhibits myosin light chain phosphatase (MLCP) through phosphorylation of its targeting subunit myosin phosphatase target subunit 1 (MYPT1), thereby sustaining elevated MLC phosphorylation and maintaining contractile force generation. ³⁸ Conversely, inactivation of this pathway dephosphorylates MLC, reduces actomyosin contractility, and lowers membrane tension. Together, RhoA-ROCK-myosin II axis dynamically tunes mechanical state of the cell surface by setting the level of force generation within the actin cortex.

Membrane surface

The third regulatory axis involves membrane area regulation through membrane reservoirs and vesicular trafficking (Figure 3(c)). Because membrane tension rises when surface area becomes limited under mechanical load, cells employ buffering mechanisms that rapidly supply additional membrane. ⁴⁰

Caveolae are flask-shaped invaginations of the plasma membrane enriched with caveolin and cavin proteins that function as membrane reservoirs.^11,41 Under acute mechanical stress, caveolae flatten and release additional membrane surface, thereby buffering sudden increases in membrane tension.^11,42 When buffering capacity of caveolae is exceeded, cells engage vesicular trafficking pathways as a secondary mechanism to regulate membrane area.^11,40 Elevated membrane tension suppresses endocytosis, as the energetic cost of membrane bending rises under high tension, while simultaneously stimulating exocytosis from recycling membrane pools, which expands plasma membrane surface area and relieves tension buildup.^9,43 This bidirectional regulation including reduced endocytosis and enhanced exocytosis acts as a coordinated feedback system that dynamically adjusts membrane area in response to mechanical demand, thereby maintaining tension within a physiological range.

How to modulate membrane tension: Experimental and engineering approaches

Membrane tension can be experimentally manipulated through diverse strategies that target membrane area, membrane reservoirs, cytoskeletal contractility, or genetic regulators.^40,44 These approaches enable controlled perturbation of the mechanical state of the plasma membrane and provide valuable insight into tension-dependent cellular behaviors.

Physical perturbation

Membrane tension can be directly modulated through controlled physical perturbations that alter membrane area or surface stress. Two widely used approaches are osmotic challenge and mechanical stretching (Figure 4(a)).^44,45 Osmotic challenge exploits the relationship between cell volume and membrane surface strain. In practice, cells are exposed to hypotonic solutions prepared by diluting culture medium or PBS with distilled water, which reduces extracellular osmolarity and drives water influx. ⁴⁶ Acute hypotonic swelling increases membrane tension by stretching the plasma membrane as cell volume expands. Whereas hypertonic treatment, achieved by supplementing medium with membrane-impermeant solutes such as sucrose or mannitol, shrinks the cell, introduces membrane slack, and reduces effective membrane tension.^45–48 Membrane tension changes occur within seconds of osmotic shock and closely follow cell volume dynamics.OPEN IN VIEWER

Mechanical stretching of adherent cells or tissues also provides a strategy to elevate membrane tension at larger scales (Figure 4(b)). Cells cultured on flexible silicone membranes can be subjected to uniaxial or biaxial cyclic strain by applying vacuum-driven deformation to the substrate. ¹¹ When external strain is applied, caveolae progressively flatten and disassemble in an actin and ATP-independent manner, reducing membrane slack and shifting the plasma membrane into a mechanically strained state. Once the caveolar reservoir is depleted, additional strain directly elevates effective membrane tension. ⁴⁹ Unlike osmotic challenge, mechanical stretching perturbs membrane tension without directly altering intracellular ion concentrations or osmolarity, making it more suitable for isolating membrane tension-specific effects.

Physical perturbations provide high sensitivity for inducing rapid and robust tension shifts, but they often lack molecular and spatial specificity. Osmotic challenge is easily implemented, yet it alters intracellular ionic strength and volume, which can confound downstream signaling analysis.^46,47 In contrast, mechanical stretching offers precise control over substrate strain but is largely restricted to adherent cultures. ⁴⁹ The requirement for specialized, substrate-dependent hardware severely limits its use in non-adherent cells or complex in vivo environments. In addition, long-term stretching can induce cellular mechanoadaptation, in which the cytoskeleton reinforces itself over time, potentially confounding the interpretation of steady-state tension responses. ⁵⁰

Cytoskeletal manipulation

Membrane tension can be modulated by manipulating cytoskeletal contractility, which directly alters cortical prestress and membrane–cortex mechanical coupling.^33,44 The actin cortex, a thin contractile network underlying the plasma membrane, generates mechanical tension through non-muscle myosin II–driven contraction. Because the plasma membrane is mechanically coupled to this cortex via membrane–cytoskeleton linker proteins, changes in cortical contractility directly influence effective membrane tension.^12,33 A widely used pharmacological approach is inhibition of non-muscle myosin II with blebbistatin, a small-molecule inhibitor that binds the myosin ATPase intermediate state and blocks myosin heads in an actin-detached conformation, thereby preventing cross-bridge cycling and reducing actomyosin-generated cortical prestress (Figure 4(c)). ⁵¹ Treatment with blebbistatin decreases cortical tension by more than 50%, promotes bleb-based protrusions, and reduces effective membrane tension in a dose-dependent manner. ³⁷ Conversely, enhanced actomyosin contractility, achieved by direct RhoA activation or ROCK-mediated myosin light chain phosphorylation, increases cortical tension and strengthens force transmission to the plasma membrane, and suppresses bleb expansion.^12,33,37 At the molecular level, actomyosin contractility is primarily regulated by the RhoA–ROCK–myosin II signaling pathway (Figure 4(d)). When RhoA is activated, it stimulates ROCK-mediated phosphorylation of myosin light chains, increasing myosin motor activity and promoting cortical tension.^52,53 Together, inhibition of RhoA or ROCK with pharmacological agents such as Y-27632 reduces cortical prestress and decreases membrane tension. ⁵⁴ Cytoskeletal manipulation offers advantages of directly targeting the force-generating machinery of the cell, enabling precise modulation of cortical tension without altering cell volume or osmolarity.

Pharmacological modulation of the actomyosin cortex is widely applicable across diverse cell types and exhibits relatively high in vivo compatibility. However, these global perturbations suffer from poor subcellular spatial resolution, making it impossible to investigate localized tension gradients or compartmentalized mechanical responses. ⁵⁵ Moreover, because the actomyosin machinery governs a broad spectrum of cellular functions, from cargo transport to gene expression, the pleiotropic effects of inhibitors like blebbistatin or Y-27632 necessitate careful controls. ⁵⁶ Such measures are essential to ensure that observed phenotypes are directly attributable to specific changes in membrane tension rather than broader cytoskeletal disruption. ⁵⁶

Optogenetic control and genetic engineering of membrane tension

Optogenetics uses genetically encoded light-sensitive proteins to control specific signaling pathways with high spatiotemporal precision, enabling reversible activation or inhibition of target molecules in living cells upon illumination. ⁵⁷ This approach overcomes a key limitation of pharmacological perturbations, which act globally and often have off-target effects, by allowing localized and temporally defined manipulation of cytoskeletal contractility and membrane–cortex coupling. In particular, optogenetic activation of RhoA can provide a direct method to elevate actomyosin contractility and increase cortical tension (Figure 4(e)). Light-induced recruitment of RhoA to the plasma membrane is achieved using engineered light-sensitive dimerization systems, such as Cryptochrome 2–CIB1 N-terminal domain (CRY2–CIBN) or light-oxygen-voltage (LOV) domain-based modules, in which RhoA is fused to a photosensitive domain that undergoes rapid heterodimerization upon blue-light illumination.^57,58 In these systems, for instance, CIBN is anchored to the plasma membrane, while RhoA-fused light-sensitive partner remains cytosolic in the dark. Upon illumination, light-triggered protein–protein interaction drives acute membrane recruitment of RhoA, leading to localized activation of the RhoA–ROCK–myosin II pathway. Light-induced recruitment of the optogenetic guanine nucleotide exchange factor-RhoA construct (optoGEF-RhoA) to the plasma membrane causes a rapid and local increase in cellular traction, intercellular tension, and tissue compaction, while sequestration of the same construct to mitochondria produces the opposite effect, demonstrating bidirectional optogenetic control of contractile forces. ⁵⁷ Spatially restricted activation further demonstrated that local increases in contractility can generate anisotropic tension patterns and drive directed cell deformation. ⁵⁷ These studies demonstrate that optogenetic RhoA systems represent powerful tools for actively manipulating membrane tension in a controlled manner.

Engineering membrane–cortex attachment represents another strategy to modulate effective membrane tension. Membrane tension is genetically modulated by altering the activity of ERM proteins (Figure 4(f)). Morpholino-mediated knockdown of ERM reduced membrane–cortex attachment and lowered effective membrane tension, promoting bleb-based migration. ¹⁰ In contrast, overexpression of constitutively active ERM strengthened membrane–cortex coupling, increased apparent membrane tension, and suppressed blebbing. Enhanced attachment also restricts membrane deformation and stabilizes lamellipodial protrusions. ¹⁰ These findings demonstrate that membrane tension can be modulated through genetic manipulation of membrane–cortex coupling strength. Together, these approaches illustrate how engineered signaling systems enable direct and quantitative control of membrane tension.

Optogenetic tools represent the current standard for spatiotemporal resolution, allowing researchers to manipulate tension at specific subcellular locations with millisecond-scale reversibility. ⁵⁷ Despite these advantages, their sensitivity can be inconsistent due to inherent variations in exogenous protein expression levels across different cells. ⁵⁹ The stringent technical requirements, most notably genetic modification and specialized optical setups, further limit their applicability in primary cells or clinical samples. Similarly, while genetic manipulation of ERM proteins directly targets the molecular interface between the membrane and the cortex, it possesses a chronic nature, typically requiring hours to days for phenotypic expression. ¹³ This slow onset potentially triggers compensatory mechanical pathways, which may mask the primary effects of tension modulation. As a result, these genetic approaches may not capture the immediate mechanical responses that acute perturbation methods can reveal, necessitating careful experimental design when interpreting long-term tension alterations. ⁶⁰

How to measure membrane tension

Accurate quantification of membrane tension is essential for understanding how cells sense and respond to mechanical cues, as it provides a direct readout of the mechanical state of the cell surface and enables mechanistic interpretation of tension-dependent behaviors such as migration, division, and signaling. Membrane tension can be quantified using several complementary approaches, each with distinct advantages and limitations.

Membrane tether pulling with optical tweezers is one of the most widely used force-based methods for measuring membrane tension. ⁶¹ In this method, a micron-sized bead attached to the cell surface is trapped by a focused laser beam and pulled away from the membrane to extract a thin membrane tether.^61–63 The steady-state force required to maintain this tether reflects effective membrane tension, which integrates intrinsic bilayer tension and membrane–cortex adhesion strength.^14,19,63 Because tether formation mechanically isolates a narrow membrane tube from the cell body, this approach provides a sensitive readout of changes in surface mechanical state in response to pharmacological or genetic perturbations. ⁶⁴

Complementary approaches such as micropipette aspiration provide an alternative force-based measurement by applying a defined negative pressure to a portion of the cell membrane. As the membrane is aspirated into the pipette, the relationship between applied pressure and membrane deformation allows the estimation of apparent cortical tension. ⁶⁵ This method was instrumental in establishing cortical tension as a quantifiable mechanical parameter in cell biomechanics, providing the first systematic framework for relating applied force to membrane deformation and laying the conceptual groundwork for subsequent tension measurement approaches. ⁶⁶ It remains particularly well suited for suspended or spherical cells.

In addition to mechanical perturbation methods, fluorescence-based approaches enable non-contact measurement of membrane tension. Mechanosensitive probes such as fluorescent lipid tension reporter (Flipper-TR) insert into the lipid bilayer and exhibit changes in fluorescence lifetime depending on membrane packing and lateral tension. ⁴⁵ These probes contain a planarizable molecular rotor structure whose conformation depends on the mechanical constraints imposed by surrounding lipids. Under low tension, looser lipid packing allows partial twisting of the probe, shortening its fluorescence lifetime. In contrast, increased membrane tension promotes tighter lipid ordering and restricts probe rotation, stabilizing a more planar conformation and prolonging fluorescence lifetime. Therefore, changes in membrane tension are transduced into measurable shifts in fluorescence decay kinetics, which can be quantified using fluorescence lifetime imaging microscopy (FLIM). ⁴⁵ These probes allow real-time monitoring of membrane tension with subcellular spatial resolution and are particularly advantageous for dynamic processes such as migration and division.

While fluorescence-based approaches such as Flipper-TR enable non-contact measurement of membrane tension, they are limited in their ability to directly quantify the specific mechanical forces transmitted via individual receptors. To complement these methods, DNA-based molecular tension probes have emerged as an alternative strategy for mapping mechanical forces at the cell surface. ⁶⁷ In this approach, surface-immobilized DNA hairpin or duplex structures are mechanically unfolded by piconewton-scale forces transmitted through receptors such as integrins, generating fluorescence signals that report the magnitude of forces applied by specific molecules. However, conventional DNA probes, which represent the naturally occurring biological stereoisomer of DNA, are susceptible to nuclease degradation, restricting their application to brief imaging windows. This limitation has been addressed by the development of L-DNA–based tension probes, the non-biological stereoisomer of DNA, which are resistant to enzymatic degradation and enable stable force imaging over extended periods. ⁶⁸ L-DNA–based probes offer critical advantages even during short-term measurements currently used in mechano-profiling of cells, enabling cleaner imaging without the need for extensive background subtraction. Furthermore, to improve quantitative resolution, tandem tension sensors (TTS)-advanced DNA probes incorporating dual force-responsive elements with distinct thresholds-have been developed. These sensors allow self-calibrated measurements and enable the detection of force heterogeneity that single-threshold sensors cannot resolve. In parallel, polarity-sensitive ratiometric fluorescence probes have been introduced to overcome the requirement for fluorescence lifetime imaging microscopy (FLIM). ⁶⁹ These probes detect changes in lipid packing of the plasma membrane and convert tension-dependent changes into intensity-based readouts, thereby enabling real-time imaging of membrane tension using conventional fluorescence microscopy while maintaining subcellular spatial resolution.

Membrane tension can also be inferred from analysis of thermally driven membrane fluctuations. In flicker spectroscopy, high-resolution imaging is used to quantify spontaneous membrane undulations. As membrane tension increases, fluctuation amplitude decreases because the membrane becomes mechanically constrained. ⁷⁰ By analyzing such fluctuation spectra, investigators can estimate relative changes in membrane tension without applying external force. Although indirect, this approach provides insight into baseline mechanical states and is useful for measuring membrane tension in suspended cells.