Metabolic reprogramming in response to cell mechanics

Much attention has been dedicated to understanding how cells sense and respond to mechanical forces. The types of forces cells experience as well as the repertoire of cell surface receptors that sense these forces have been identified. Key mechanisms for transmitting that force to the cell interior have also emerged. Yet, how cells process mechanical information and integrate it with other cellular events remains largely unexplored. Here we review the mechanisms underlying mechanotransduction at cell‐cell and cell‐matrix adhesions, and we summarize the current understanding of how cells integrate information from the distinct adhesion complexes with cell metabolism.


INTRODUCTION
All cells experience and exert physical forces that regulate biological function. These forces are first experienced by cells in developing tissues, continue throughout their lifetimes, and are a critical determinant of function and human health. Additionally, human pathologies are strongly linked to alterations in the mechanical properties of cells and their response mechanical stimuli. For example, atherosclerotic, fibrotic, and tumorigenic tissues are stiffer than their non-diseased counterparts and counter external forces using altered responses (reviewed in Ingber, 2003). Thus, a better understanding of how cells sense and respond to these forces may help identify how cells fail to adapt in disease, and ulti-mately serve as targets for therapeutics that attenuate the mechanical properties of cells.
Forces can be internally generated or externally applied ( Figure 1). Endothelial cells, which line the vasculature as well as epithelial cells which line the major organs experience an external force in the form of shear stress (Figure 1a). The origin of shear stress experienced is dependent on tissue localization. Indeed, endothelial cells experience shear from blood flow (Davies, 1995) whereas ductal epithelial cells lining the kidneys (Essig & Friedlander, 2003) or mammary gland experience shear from the flow of urine or milk. In contrast, epithelial cells lining the bronchi and alveoli experience shear from the flow of air (Waters et al., 2012), and corneal epithelial cells experience shear from eye blinking or rubbing (Masterton & Ahearne, 2018).
In addition to shear, cells experience other external forces, such as stretching and compression (Figure 1b,c). Cardiomyocytes, endothelial cells, smooth muscle cells, and osteocytes, are constantly stretched or compressed (Williams, 1998). This phenomenon is well understood in cardiomyocytes. During the development of the heart, pre-cardiac cells are first strained as they migrate through the primitive streak, a transient embryonic structure that delineates bilateral symmetry (reviewed in Happe & Engler, 2016). During the Compression occurs when a pushing force presses the cell inward causing it to be compacted. (c) Cells are surrounded by matrix proteins. Throughout development and disease progression, the density and composition of the matrix can change, producing a stiffer matrix; cells respond to changes in matrix stiffness by tuning their internal contractility. (e) A protrusive force is generated by the polymerization of actin (red box) at the leading edge. At the leading edge, actin monomers are added at the plus end which is oriented towards the plasma membrane. The addition of monomeric actin is accompanied by ATP hydrolysis, but only a small amount of the free energy of nucleotide hydrolysis is required (Dmitrieff & Nédélec, 2016). The remaining free energy is used to generate a protrusive force that pushes the cell membrane forward. (f) Cells forming adhesions with adjacent cells can experience a tugging force from increased tension in the circumferential actin belt (purple cables) that connects the cells. (g) Increased internal contractility of cells is derived from the molecular motor myosin (blue) binding and pulling the actin filaments in opposite directions. This increased internal contractility is transmitted to the cell substrate at sites where the cell adheres to the matrix, thereby generating traction forces. continued development of the heart, pre-cardiomyocyte structures are stretched and compressed during morphogenesis, and cells are strained again when they first start to spontaneously contract. Similarly, cells tune their mechanical properties to respond to external cues from their environments. Changes in the composition and density of extracellular matrix proteins during aging and disease produce a stiff environment. Cells respond to these alterations by tuning their internal contractility ( Figure 1d) (Discher et al., 2005;Engler et al., 2006).
Forces can also be generated internally from the cytoskeleton and generally fall into two categories: traction forces and protrusive forces. Protrusive forces are generated via the polymerization of actin at the leading edge of cells ( Figure 1e) (Pollard & Borisy, 2003). The addition of monomeric actin is accompanied by ATP hydrolysis, but only a small amount of the free energy of nucleotide hydrolysis is required (Dmitrieff & Nédélec, 2016). The remaining free energy is used to generate a protrusive force that pushes the cell membrane forward (Dmitrieff & Nédélec, 2016). Additionally, through this actomyosin-generated tension, cells exert a tug-ging force on their neighbors, and this force is critical to regulating the size and strength of cell-cell junctions ( Figure 1f) . To migrate over a surface, cells must generate protrusions in the direction of chemotactic stimuli and apply mechanical or traction forces on the extracellular matrix. These traction forces are generated via the actions of the molecular motor myosin generating contractility by walking along actin filaments and are transmitted to the extracellular matrix ( Figure 1g) (Meili et al., 2010).
In addition to experiencing different types of forces, the magnitude of the force can vary greatly. Corneal epithelial cells experience between 0.05 to 14 dynes/cm 2 of force during blinking (Ben-Eli et al., 2019;Hampel et al., 2018;Molladavoodi et al., 2017). This is also the approximate amplitude of force experienced by endothelial cells from blood flow or lung epithelia from airflow (Mahto et al., 2014;Satcher et al., 1997;Trepat et al., 2004). However, cells can experience far greater amplitudes of force. For example, vigorous breathing following exercise applies approximately 60 dynes/cm 2 of shear stress to the lung epithelium (Regmi et al.,FIGURE 2 Mechanisms for increased contractility at sites of cell-cell and cell-matrix adhesion. In epithelial cells, shown on the left, E-cadherin (royal blue structures) senses and responds to changes in tension by recruiting proteins, such as αand β-catenin. α-Catenin undergoes conformational changes that expose binding sites for other proteins, such as vinculin. Tension also stimulates vinculin phosphorylation (blue circle) at Y822 by Abelson tyrosine kinase (Abl). The phosphorylation of vinculin at Y822 is required for increased activation of the small GTPase, RhoA, which in turn stimulates a signal transduction cascade that culminates in increased myosin II contractility and cytoskeletal reinforcement. The cell on the right depicts the response of cell-matrix adhesions to changes in tension. Heterodimeric α and β integrin subunits (light blue structures) form bonds with extracellular matrix components. In response to force, integrins undergo a conformational change from an inactive, bent conformation to an active, stood-up conformation. Additionally, talin unfolds revealing vinculin binding sites. Vinculin is recruited and binds actin (purple filaments). Force on integrins also activates the RhoA guanine nucleotide exchange factors, GEF-H1 and LARG, which leads to the activation of the small GTPase RhoA. Activation of the RhoA pathway leads to the increased contractility. 2017). In contrast, during coughing, the lung epithelia can experience up to 1700 dynes/cm 2 (Button & Button, 2013).
While the types and amplitudes of force experienced by cells are recognized, the mechanisms cells employ to respond to these forces are still emerging. The cell surface receptors that sense the changes in tension and many of the signaling pathways that are activated in response to different forms of force have been identified, yet little is known with respect to how these mechanically activated pathways are integrated with other cellular processes. In this review, we will focus on: (1) how different forces are sensed and transmitted to the cell interior and (2) how this response to force is integrated with cellular metabolism.

Cell-cell junctions
External and internal forces are sensed by numerous cell surface receptors. At the apical surface of polarized epithelial and endothelial cells are intercellular junctions, known as adherens junctions or cell-cell junctions (Figure 2a). These intercellular junctions are highly enriched in cadherins. In epithelia, this cadherin is epithelial or E-cadherin, whereas and in endothelia, it is vascular endothelial or VE-cadherin. Eand VE-cadherin are both composed of a N-terminal ectodomain, a transmembrane region, and an intra-cellular tail (Van Roy & Berx, 2008). The extracellular domains of cadherins bind calcium and cadherin extracellular domains on neighboring cells forming a rod-like conformation to allow for cell-cell adhesion.
In response to force, cadherins transmit the force to the cell interior. Mechanical tension across both E-and VE-cadherin have been observed using FRET probes (Borghi et al., 2012;Conway et al., 2013). Cadherins also recruit numerous binding partners such as alphacatenin, beta-catenin, and vinculin (Leckband & De Rooij, 2014;Ozawa et al., 1989). Beta-catenin directly binds to the cadherin tail while also interacting with alpha-catenin and vinculin directly (Ozawa & Kemler, 1992;Shapiro & Weis, 2009). Alpha-catenin binds vinculin only weakly in the absence of force (Peng et al., 2012;Yonemura et al., 2010). The application of force causes an alpha-catenin conformational change that exposes a binding site for vinculin (Yonemura et al., 2010). Vinculin binding prevents alpha-catenin from returning to an autoinhibited state (Yao et al., 2014;Yonemura, 2011). These events are critical for cell-cell adhesion during collective cell migration (Seddiki et al., 2018).
Like alpha-catenin, vinculin undergoes conformational changes in response to the force (Chen et al., 2005). In the unfurled conformation, vinculin stimulates actin polymerization and recruits actin-remodeling proteins, such as VASP and vinexin, to cell-cell junctions (Brindle et al., 1996;Carisey & Ballestrem, 2011;Kioka et al., 1999). Additionally, vinculin is preferentially phosphorylated at tyrosine 822 on vinculin in response to force applied to cadherins (Bays et al., 2014). This phosphorylation event is critical to stimulating downstream activation of the RhoA and increased contractility (Bays et al., 2014).
Vinculin and alpha-catenin bind to F-actin. The proteins critical for dissipating the force cadherins experience onto the actin cytoskeleton has been the subject of much debate. Alpha-and beta-catenin form heterodimers that have been reported to only weakly bind actin Miller et al., 2013;Yamada et al., 2005). More recent evidence suggests that force strengthens the interaction of these heterodimers with actin (Buckley et al., 2014). Other work has identified vinculin, also an actin-binding protein, as the critical protein in transmitting force between E-cadherin and actin (Le Duc et al., 2010). Thus, it is likely that these proteins and other E-cadherin-associated proteins, such as eplin, play a critical role in transmitting the force experienced by E-cadherin onto the actin cytoskeleton (Abe & Takeichi, 2008).

Cell-matrix adhesions
In addition to having contact with other neighboring cells, cells form junctions to the extracellular matrix (ECM). At these cell-ECM junctions otherwise known was focal adhesions, integrins are the primary mechanoreceptor ( Figure 2b). The integrin family are transmembrane proteins composed of noncovalently linked alpha and beta subunits and serve as a connection from inside the cell to the extracellular matrix (reviewed in Hynes, 2002;Martino et al., 2018;Michael & Parsons, 2020;Ross et al., 2013;Schwartz et al., 1995;Sun et al., 2016). The various integrin alpha and beta subunits associate to form 24 different heterodimers, which can be grouped based on ligand binding or the identity of their subunits. Integrins heterodimers adopt an inactive bent confirmation, and upon sensing force adopt an upright confirmation. The force-stimulated conformational change in integrins is supported by the binding of talin to the integrin cytoplasmic tail (Calderwood et al., 1999;Tadokoro et al., 2003). This conformational change allows for integrin to associate with the extracellular matrix on the outside of the cell and increases its affinity for actin-binding proteins, thereby creating a conduit to dissipate the force experienced by integrins onto the actin cytoskeleton.
Talin is one of the many actin-binding proteins that are recruited to integrin cytoplasmic domains to form focal adhesions (Partridge & Marcantonio, 2006;Pasapera et al., 2010) (reviewed in Chakraborty et al., 2019;Zhao et al., 2022). Like the integrins, talin itself undergoes a conformational change in response to force. This conformational change exposes binding sites for other proteins, such as vinculin and the actin cytoskeleton.
This allows for the recruitment of vinculin to focal adhesions in response to force and further transmission of force (Yao et al., 2014).
In addition to recruiting actin-binding proteins, integrins stimulate signal transduction pathways on the inside of the cell. For example, force on integrins activates the RhoA guanine nucleotide exchange factors, GEF-H1 and LARG (Guilluy et al., 2011). The GEFs then simulate the exchange of GDP for GTP, which promotes the activation of RhoA and subsequent phosphorylation of the myosin light chain (MLC) (reviewed in DeWane et al., 2021;Leckband & De Rooij, 2014;Salvi & DeMali, 2018;Sun et al., 2016). This activation of MLC increases actomyosin contractility which ultimately leads to the reinforcement the actin cytoskeleton necessary for cells to withstand the force (reviewed in DeWane et al., 2021;Leckband & De Rooij, 2014;Salvi & DeMali, 2018;Sun et al., 2016).

Metabolic pathways
Emerging evidence suggests that the response of cells to force is coupled with changes in cell metabolism. In this section, we present a brief overview of the some of the major metabolic pathways utilized by cells and then review how these metabolic pathways are coupled to the response of cell-cell and cell-matrix adhesions to force. This discussion is not comprehensive and is limited to those pathways which have been linked cell mechanics.

Glycolysis
It has been demonstrated that in response to various forces, cells upregulate the transport of glucose into the cell (Bays et al., 2017). The glucose is transported into the cells via high-affinity glucose transporters that have different kinetic properties based on their tissue distribution. In the cytoplasm, glucose then oxidized via the ten-step process of glycolysis (reviewed in Mulukutla et al., 2016). In the first step of glycolysis, glucose is trapped in the cell by phosphorylation by a hexokinase forming glucose-6phosphate. An isomerization reaction then takes place forming fructose-6-phosphate. In a key regulatory step of glycolysis, phosphofructokinase-1 then phosphorylates the fructose-6-phosphate forming fructose-1,6bisphosphate. This step is the committed step of glycolysis. Aldolase cleaves fructose-1,6-bisphosphate into glyceraldehyde-3-phosphate and dihydroxyacetone phosphate. Before moving forward through glycolysis, dihydroxyacetone phosphate is isomerized into glyceraldehyde-3-phosphate. Oxidation and subsequent phosphorylation of glyceraldehyde-3-phosphate generate 1,3-bisphosphoglycerate and the electron carrier NADH. Phosphoglycerate kinase catalyzes substratelevel phosphorylation, transferring a phosphate group from 1,3-bisphosphoglycerate to ADP forming ATP and 3-phosphoglycerate. 3-phosphoglycerate is converted to 2-phosphoglycerate and then enolase catalyzes the dehydration of 2-phosphoglycerate to form phosphoenolpyruvate. Pyruvate kinase then catalyzes the transfer of a phosphate group from phosphoenolpyruvate to ADP forming ATP and pyruvate completing the process of glycolysis.
In the presence of oxygen, pyruvate is further oxidized into acetyl-CoA and is carried into the citric acid cycle. The citric acid cycle fully metabolizes the former glucose molecule into CO 2 and H 2 O, while generating NADH and FADH 2 . The NADH and FADH 2 are energy carriers and go through oxidative phosphorylation to generate ATP in the mitochondria (Reece et al., 2014). NADH and FADH 2 act as electron donators, providing electrons that are passed to O 2 through a chain of protein complexes (Reece et al., 2014). This passage of electrons leads to the formation H 2 O and the energy needed to drive ATP synthesis.

Amino acid metabolism
Cells are highly dependent upon amino acids for fuel. Glutamine is the most abundant free amino acid and can be catabolized to generate citric acid cycle interme-diates in a process known as glutaminolysis. The overall reaction converts glutamine to the citric acid cycle intermediate, α-ketoglutarate, which can be used to generate energy or other molecular building blocks (reviewed in Wang et al., 2020). To be converted into other metabolic intermediates, glutamine first needs to undergo a reaction to generate glutamate. This is accomplished by glutaminase catalyzing the deamination of glutamine forming glutamate and ammonia. Then glutamate dehydrogenase forms the citric acid cycle intermediate α-ketoglutarate in the mitochondria. Alternatively, glutamate can undergo transamination producing alanine or aspartate and α-ketoglutarate. As a citric acid cycle intermediate, α-ketoglutarate acts as an energy source for cells by allowing for the cycle to continue producing more of the energy carriers NADH and FADH 2 . Additionally, glutamine serves as a precursor for macromolecules. As key enzymes in the process of glutaminolysis, glutaminase, and glutamate dehydrogenase enzymes are key points of regulation.
Proline has been implicated in a variety of cellular processes including energy utilization, programmed cell death, reactive oxygen species production, stress response and cellular reprogramming and development (reviewed in Phang et al., 2010;Trovato et al., 2019). Proline synthesis begins with Δ-pyrroline-5-carboxylate synthase catalyzing the 2-step reaction in which glutamate in converted into glutamate-γ-semialdehyde (reviewed in Phang et al., 2010;Trovato et al., 2019). This is followed by cyclization forming Δ-pyrroline-5-carboxylate which is subsequently reduced to proline by pyrroline-5-carboxylate reductase 1 (PYCR1). Proline can also be oxidized to form glutamate (reviewed in Phang et al., 2010;Trovato et al., 2019). Since the catabolism of proline is coupled to the electron transport chain and glutamate can be converted to α-ketoglutarate to enter the citric acid cycle, proline breakdown also produces 30 ATP.

Lipid metabolism
Increased lipid metabolism can be altered to allow cells to withstand force. This is often the case in cancer cells that use altered lipid metabolism in order to support tumor formation. It is estimated that 5% of genes play a role in the synthesis of thousands of different cellular lipids in eukaryotic cells (Shimano & Sato, 2017). These lipids serve as energy stores, signaling messengers as well as structural elements of the membrane. Of particular importance are sterols, such as cholesterol, and fatty acids, both of which are major components of cell membranes.
The ER is the main site of lipid biosynthesis, from which sterols are rapidly transported to other organelles such as the Golgi. One way in which cholesterol metabolism is regulated is through the transcriptional regulation of genes. For example, the rate-limiting enzyme of cholesterol biosynthesis, 3hydroxy-3-methylglutaryl CoA reductase can be transcriptionally regulated to maintain lipid homeostasis (Brown et al., 2018;Shimano & Sato, 2017). Additionally, cholesterol levels are regulated through transcriptional regulation of the LDL receptor which increases cholesterol through the endocytosis of LDL's containing cholesterol and fatty acids. Genes related to cholesterol metabolism are regulated by sterol regulatory element (SRE) binding proteins (SREBP) binding to the SRE region of promoters and subsequent stimulation of transcription. SREBP is synthesized as a membrane protein localized in the ER membrane with SREBP cleavageactivating protein (SCAP) (Shimano & Sato, 2017). The SREBP-SCAP complex maintains ER localization in the presence of sterols (Shimano & Sato, 2017). When cholesterol concentration is low the SREBP-SCAP complex is transported to the Golgi where SREBP is processed by proteolytic cleavage (Brown et al., 2018;Shimano & Sato, 2017). In the Golgi, the N-terminal cytoplasmic portion of SREBP is cleaved so it is then able to enter the nucleus, bind to the target SRE sequence, and thus promote transcription (Shimano & Sato, 2017). In addition to regulation through sterol levels, SREBPs can be regulated by ADP ribosylation factor 1 and Lipin 1. The SREBP1a is implicated in regulation of global lipid synthesis while the SREPB1c is implicated in fatty acid synthesis and SREBP2 in cholesterol metabolism. At cytoplasmic membranes, the phosphatase Lipin-1 catalyzes the conversion phosphatidic acid to diacylglycerols as well as inhibits SREBP (Brown et al., 2018;Shimano & Sato, 2017).

Cell-cell junctions and metabolic changes
E-cadherin is the principal cell-cell adhesion receptor that senses and responds to changes in internal and external tension in epithelia. In response to the application of mechanical forces or shear stress, E-cadherin signals for increased Liver kinase B1 (LKB1) recruitment to cell-cell junctions and its activation (Figure 3) (Bays et al., 2017). LKB1, in turn, stimulates the recruitment and phosphorylation of AMP-activate protein kinase (AMPK) in its activation loop, rendering it catalytically active. Activation of AMPK has at least two effects. AMPK stimulates a downstream signal transduction cascade that culminates in the phosphorylation of vinculin at Y822 and the subsequent activation of the small GTPase, RhoA, and myosin II, resulting in increased contractility (Bays et al., 2017). This AMPK-stimulated increase in contractility is required for the growth of cell-cell adhesions and increased actin cytoskeletal reinforcement that allows the cell to resist the force (Bays et al., 2017). AMPK also signals for increased glucose uptake through high-affinity transporters known as glucose transporters or GLUTs . Salvi et al. demonstrated that GLUT1, but not GLUT3 or GLUT4, is the AMPK-activated glucose transporter enriched in cell-cell junctions in response to force. GLUT1 uptakes glucose which is then metabolized to ATP. If force stimulated increases glucose uptake, glycolysis, or ATP production are blocked, cells cannot reinforce their actin cytoskeletons . These studies provide strong evidence that glucosederived ATP fuels the actin cytoskeleton. In addition to AMPK's importance in adhesion junctions, evidence also suggests AMPK is important for the formation and integrity of tight junctions (Olivier et al., 2019;Zhang et al., 2006;Zheng & Cantley, 2007).
How GLUT1 responds to changes in tension and becomes enriched in the plasma membrane in cells exposed to force remains unknown. In muscle, GLUT4 is insulin sensitive and translocates to the plasma membrane in actively contracting muscle (Czech & Buxton, 1993;Satoh et al., 1993). Unlike GLUT4, GLUT1 is thought to be responsible for basal transport and is not thought to translocate in response to insulin or other hormones (Fisher & Frost, 1996). It is possible that GLUT1 itself is force sensitive, but that possibility has yet to be rigorously tested. The more likely possibility is that GLUTs are retained in the plasma membrane in cells experiencing force. In support of this idea, our laboratory discovered that GLUT1 tethered to E-cadherin in response to force . Specifically, force stimulated an association of spectrin with both Ecadherin-bound ankyrin G and GLUT1. When ankyrin G binding to E-cadherin was disrupted, GLUT1 was not enriched in the plasma membrane and cells could not uptake glucose or reinforce their actin cytoskeletons . Thus, GLUT1 is retained at sites of cell-cell adhesion to provide a fuel source for the reinforcing actin cytoskeleton.

Cell-matrix adhesions and metabolism
In response to force on cell-cell and cell-matrix adhesions, cells fortify or reinforce their actin cytoskeletons and the adhesions grow to counter the applied force in a process known as cell stiffening. The reinforcement process is thought to involve the rearrangement of existing actin networks and new polymerization (Balaban et al., 2001;Osborn et al., 2006;Pandit et al., 2020). Studies of platelets and neurons indicate that about 50% of the ATP in a cell is needed to fuel the actin cytoskeleton (Bernstein & Bamburg, 2003;Daniel et al., 1986). Thus, the reinforcement of the actin cytoskeleton that occurs in cells under mechanical tension is thought to be an energetically costly event. In this section, we describe mechanisms that cells employ to adjust their metabolism in response to changes in external tension.

FIGURE 3
Links between metabolism and mechanotransduction at cell-cell junctions. In epithelial cells, E-cadherin mediates the response to force at cell-cell contacts. E-cadherin recruits liver kinase B1 (LKB1), which recruits and activates AMP-activate protein kinase (AMPK). p21-activated protein kinase 2 (PAK2) is then recruited and activated by AMPK. PAK2 stimulates the recruitment and activation of a variety of other adhesion proteins to the E-cadherin complex. These include molecules such as Abelson tyrosine kinase (Abl) and vinculin phosphorylation at Y822 (pVinc). Activation of vinculin leads to a signaling cascade stimulates RhoA and regulates myosin II contractility and cytoskeletal reinforcement. These changes are energetically costly. AMPK signals for increased glucose uptake and its conversion into ATP to provide energy to polymerize new actin filaments needed to reinforce the actin cytoskeleton.

In response to matrix stiffness
During the progression of numerous diseases, such as arthersclerosis, cancer, and fibrosis, there are changes in extracellular matrix composition and deposition that increase the stiffness of the microenvironment. These changes in matrix stiffness are sensed by integrins and are transmitted to the cell interior as described above.
Here we discuss emerging evidence suggesting that some of those signals transmitted to the cell interior alter metabolism (Figure 4).
An adjustment to cell metabolism was observed by Park and colleagues in response to plating human bronchial epithelial on soft versus stiff substrates. On soft substrates, a low level of actomyosin contractility was produced and accompanied by a downregulation of glycolytic and citric acid cycle intermediates in comparison to when plated on stiffer substrates (Park et al., 2020). Notably, there was an accumulation of glucose-6-phosphate, but a reduction in its downstream intermediates. Additionally, a post-transcriptional downregulation of phosphofructokinase 1 expression and activity was observed in the cells plated on soft substrates. A similar decrease in PFK expression was seen when actin bundle formation was disrupted with latrunculin A suggesting a role for actin stress fiber disassembly in observed changes. Proteasomal degradation as responsible for the reduced expression of PFK observed on soft substrates. Moreover, the E3 ubiquitin ligase, TRIM21, was required for proteasomal degradation, and its activity was modulated by binding to stress fibers, thereby suggesting that on stiffer substrates, the formation of F-actin bundles sequesters TRIM21, protecting PFK from degradation, and allowing for increased glycolysis (Park et al., 2020).
Other work indicates the increased glycolysis that occurs in cells plated on stiff extracellular matrices is triggered by activation of the transcriptional coactivators YAP and TAZ (Bertero et al., 2016). Indeed, Bertero et al. discovered that pulmonary vascular endothelial cells plated on stiff matrices had elevated YAP and TAZ activity. This increase in YAP/TAZ elevated a number of metabolic enzymes, including glutaminase, pyruvate carboxylase and lactate dehydrogenase resulting in a coordinately upregulation of glutaminolysis and glycolysis (Bertero et al., 2016). Taken together these studies indicate that integrins respond to changes in matrix stiffness by elevating glycolysis by modulating glycolytic enzyme levels.
Furthermore, cells plated on a stiff matrix had altered mitochondrial dynamics. Tharp and colleagues demonstrated that integrin mechano-signaling tuned flux through mitochondrial respiration by activating a mitochondiral stress response. This stress response is mediated by heat shock factor 1. transcription in order to alter mitochondrial structure and function. This regulation of mitochondrial dynamics lead to a restriction of mitochondrial respiration (Tharp et al., 2021). Integrins alter cell metabolism in response to increased matrix stiffness. (a) On soft matrices TRIM21 targets phosphofructokinase 1(PFK1), the enzyme that catalyzes the committed step of glycolysis, for degradation. Additionally, in cells plated on soft matrices, sterol regulatory element (SRE) binding proteins (SREBP) are cleaved and are then trafficked to the nucleus. In the nucleus, SREBP signals for increased lipid synthesis. (b) On stiff matrices, the integrin-containing adhesions (light purple) and the actin stress fibers (purple cables) are reinforced. TRIM21 is sequestered by actin stress fibers, which protects PFK1 from degradation and promotes glycolysis. The transcriptional co-activators YAP/TAZ are activated by stiff matrices and increase glutaminase (GSL1), thereby promoting glutaminolysis. A stiff matrix also promotes kindlin localization to the mitochondria (purple ovals) where it interacts with (pyrroline-5-carboxylate reductase-1) PYCR1 to promote proline synthesis.
While the evidence supports the role of elevated glycolysis in cells grown on stiff matrices, lipid metabolism appears to prevail in soft cells. Romani and colleagues compared the metabolic profile of MCF-10A cells grown on stiff, tissue culture plastic surfaces or grown on tissue culture plastic and then exposed to ROCK or MLCK inhibitors to decrease actomyosin contractility (Romani et al., 2019). Under conditions of low contractility, there was an upregulation of the synthesis of neutral lipids. When actomyosin contractility was low, lipin-1, a phosphatidate phosphatase, was inhibited, thereby reducing the diacylglycerol at the Golgi. The reduced diacylglycerol content promoted the accumulation of SREBPs. This accumulation of SREBP at the Golgi led to cleavage of SREBP and trafficking to the nucleus where the protein activated transcription factors and increased lipid synthesis (Romani et al., 2019).
Other work suggests integrins modulate proline metabolism in response to force. Kindlins bind to integrin beta subunit cytoplasmic tails and activate them. Guo et al. demonstrated that stiff extracellular matrices induce translocation of kindlin-2 to the mitochondria where it binds and regulates pyrroline-5-carboxylate reductase-1 (PYCR1), an enzyme involved in proline synthesis (Guo et al., 2019). Accordingly, depletion of kindlin-2 reduced PYCR1 and proline synthesis. These results provide a connection between integrins and amino acid metabolism (Guo et al., 2019). Thus, cells plated on stiff extracellular matrices have very different metabolic profiles than cells plated on soft matrices.

Cell spreading and migration
The ability of cells to spread on substrates and migrate requires membrane protrusion. As discussed above, the process of membrane protrusion requires the polymerization of branched actin networks at the protruding edge. This polymerization of actin networks produces a protrusive force against the cell membrane. Increasing evidence suggests that cell metabolism is altered to support these protrusive forces ( Figure 5). Xie et al. demonstrated that cell spreading is accompanied by a drop in the intracellular levels of ATP (Xie et al., 2021). This drop in ATP levels triggers the activation of the AMP-activated protein kinase (AMPK) (Xie et al., 2021). The replenished ATP supply supports cell spreading and reinforcement (Xie et al., 2021). Interestingly, at early time points inhibition of actin dynamics, but not myosin contractility restored cellular ATP levels. This suggests that initially actin polymerization needed generate protrusive force to extend the membrane is the source of ATP expenditure (Xie et al., 2021). Once cells area had reached equilibrium, inhibition of myosin was found to restore ATP levels demonstrating a switch in energy usage to generating contractility and maintaining tension.
Like cell spreading, cell migration requires the ability to form membrane protrusions and adhere those membrane protrusions to the extracellular matrix. Cunniff et al. demonstrated that ATP-generating mitochondria are actively transported to the leading edge of membrane protrusions in migrating cells (Cunniff et al., 2016). The trafficking of mitochondria to the leading edge is mediated by Miro1, a mitochondrial Rho-GTPase 1 (Schuler et al., 2017), and requires AMPK (Cunniff et al., 2016). These findings are supported by the findings of Xie et al. that demonstrate mitochondrial morphology is highly responsive to AMPK activation in response to cell spreading (Xie et al., 2021).
How AMPK is activated in response to cell spreading and migration remains unknown. One simple explanation is that ATP levels decrease and AMP levels in regions of membrane protrusion result in the activation of AMPK. Alternatively, the PI3K pathway is an upstream activator of AMPK and is involved in the regulation of membrane protrusion during cell spreading and migration (reviewed in Xue & Hemmings, 2013). Hu and colleagues demonstrated the glycolytic enzyme aldolase is bound to the actin cytoskeleton and held in an inactive state. In response to PI3K activation, the actin cytoskeleton is rearranged and aldolase is released, resulting in elevated glycolysis . More work is needed to resolve how AMPK is activated in response to elevated cell mechanics at regions rich in integrin-containing adhesions.

Concluding remarks
Here we reviewed the emerging data suggesting that cell mechanics regulate metabolic processes. We focused our attention on the recent data demonstrating links between cell-matrix adhesion and cell-cell adhesion. Both E-cadherin and integrins upregulate glycolysis, suggesting that the processes they modulate are energy-intensive. For E-cadherin, this upregulation of glycolysis fuels the actin rearrangements necessary to reinforce the actin cytoskeleton so that it can withstand the tension. More work is necessary to determine if some integrin-stimulated metabolic changes fuel the actin cytoskeleton or if they fuel some yet undetermined function.
Interestingly, at this time, the mechanisms linking metabolism to mechanotransduction by cadherins and integrins appear to be distinct, despite evidence suggesting that tugging forces can be antagonist, cooperative, and interdependent (Mui et al., 2016). Thus, it will be critical to understand links between mechanics in the context of tissues and the whole organism where both adhesion receptors are under tension.
While this review explores connections between the actin cytoskeleton and metabolism in non-cancerous cells, there are two notable related areas that are emerging. The first is that links between mechanics and metabolism are not limited to the actin cytoskeleton. The observation that increased matrix stiffness reprograms glutamine metabolism to promote the glutamylation and stabilization of microtubules suggests the presence of connections between microtubule dynamics and metabolism (Torrino et al., 2021). Second, it is worth noting that while we largely limited our discussion to normal cells, there are many more studies describing how the mechanics and metabolism are altered in cancer cells (Bertero et al., 2019;Ingber, 2003;Mah et al., 2018;Papalazarou et al., 2020;Sousa et al., 2019). Cancer cells have altered metabolic profiles, aberrant cell-matrix and cell-cell adhesion, and a far more complex microenvironment than the cells we described. Additionally, cancer cells not only migrate but also must invade through that spatially complex matrix to metastasize. Consequently, their metabolic adjustments are proving to be far more complex than what we have described here for untransformed cells. Additionally, while we have focused our attention on how integrins and cadherins stimulate metabolic changes, there is also some data suggesting that the shift to glycolytic metabolism can repress E-cadherin (Krishnamachary et al., 2006). More work is needed to unravel the complex relationship between integrins/cadherins and cell metabolism.