**The Interplay between Cytoskeleton and Calcium Dynamics**

Francisco Javier Martin-Romero, Aida M. Lopez-Guerrero, Carlos Pascual-Caro and Eulalia Pozo-Guisado

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/66862

#### **Abstract**

Cell motility is a complex cellular event that involves reorganization of cytoskeleton. This reorganization encompasses the transient polarization of the cell to facilitate the plasma membrane ruffling, a rearrangement of cortical actin cytoskeleton required for the development of cellular protrusions. It is known that extracellular Ca2+ influx is essential for cell migration and for the positive-feedback cycle that maintains leading-edge structures and ruffling activity. The aim of this review is to summarize our knowledge regarding the Ca2+ dependent signaling pathways, Ca2+ transporters and sensors involved in cell migration. Also, we show here reported evidences that support for a crosstalk between Ca2+ transport and the reorganization of the cytoskeleton required for cell migration. In this regard, we will analyze the role of store-operated Ca2+ entry (SOCE) as a modulator of cytoskeleton and cell migration, but also the modulation of this Ca2+ entry pathway by microtubules and the actin cytoskeleton. As a main conclusion, this review will show that data reported in the last years support a role for SOCE in shaping cytoskeleton, but at the same time, SOCE is strongly dependent on cytoskeletal proteins, in an interesting interplay between cytoskeleton and Ca2+ dynamics.

**Keywords:** calcium, microtubules, actin, STIM1, ORAI1, cell migration, cortical cytoskeleton

### **1. Introduction**

Calcium ions (Ca2+) are essential intracellular transducers for cell signaling because of their role to bind Ca2+-sensitive proteins that mediate key activities in signaling pathways. Upon cell stimulation through a variety of receptors and other types of physicochemical stimulations such as depolarization of plasma membrane, changes in osmolarity, physical distortion

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of cell surface, temperature, etc., a number of intracellular "second messengers" transmit the initial stimulation into the cell to trigger a proper response to these stimuli. The response is attained in some cases by transiently altering ion transport through plasma membrane as well as intracellular membranes, posttranslational modifications of proteins, changes in gene expression, or reshaping the cytoskeleton in a second-messenger-dependent manner.

To properly act as a second messenger the concentration of free cytosolic Ca2+ ([Ca2+] *i* ) is substantially different across plasma membrane. While the physiological extracellular medium contains 1–3 mM Ca2+, the level of the cytosolic milieu is within a narrow range, 80–120 nM [1, 2]. Considering this large chemical gradient of free Ca2+ concentration, cells can trigger a rapid and transient increase of [Ca2+]*<sup>i</sup>* by increasing Ca2+ transport through plasma membrane. This transient Ca2+ increase triggers the action of a wide range of Ca2+-dependent mediators that modify signaling pathways. Because Ca2+ is involved in numerous signaling pathways, differential features of Ca2+ increase are required to respond to diverse initial stimuli. This is achieved by the spatiotemporal control of the Ca2+ increase, i.e., the localization of the cytosolic Ca2+ spike within the cell, as well as the rate of the increase, the extent of the transient increase and the kinetics of the final decrease to basal levels [3].

This Ca2+ transport is therefore tightly regulated by specific channels, which are sensitive to hormones, cytokines, small molecules and other extracellular stimuli. The features of the transporters shape the characteristics of Ca2+ current (Ca2+ influx), modeling the kinetics of Ca2+ entry. Because a specific distribution of Ca2+ channels and transducers is required the cytoskeleton has been involved in the spatial regulation of Ca2+ entry. Interestingly, Ca2+ signaling strongly influences cytoskeleton dynamics, in an interesting interplay that is currently under study. In order to assess the crosstalk between Ca2+ signaling and cytoskeleton we will review here the recent literature regarding this topic, with special focus on the role of store-operated Ca2+ entry (SOCE) in cell migration, focal adhesion turnover and actin filaments reorganization. In addition, we will review the role of the microtubule cytoskeleton in the normal function of SOCE. A major regulator of SOCE is the protein STIM1, an endoplasmic reticulum resident protein that serves as intraluminal Ca2+ sensor and plasma membrane Ca2+ channel modulator. STIM1 is known to be a plus-end microtubule binding protein (+TIP). As a +TIP, STIM1 is transported throughout the cell while bound to microtubules, but it is released upon activation. The molecular basis of this regulation and the role of posttranslational modifications in STIM1 that are known to underlie this regulation, will be also described.

### **2. Ca2+ transporters and Ca2+ influx pathways**

In eukaryotic cells plasma membrane (PM) contains different Ca2+ transport systems to control Ca2+ influx as well as Ca2+ extrusion. Because the temporal control of Ca2+ signaling is also strictly controlled, transporters are highly coordinated to let the Ca2+ spikes/waves last for a precise time, but this time ranges from microseconds (as in exocytosis) to hours, as observed during mammalian oocyte fertilization. To understand this control, we summarize here the most important Ca2+ transporters in eukaryotic cells. Ca2+ entry channels are divided into the following:

of cell surface, temperature, etc., a number of intracellular "second messengers" transmit the initial stimulation into the cell to trigger a proper response to these stimuli. The response is attained in some cases by transiently altering ion transport through plasma membrane as well as intracellular membranes, posttranslational modifications of proteins, changes in gene

stantially different across plasma membrane. While the physiological extracellular medium contains 1–3 mM Ca2+, the level of the cytosolic milieu is within a narrow range, 80–120 nM [1, 2]. Considering this large chemical gradient of free Ca2+ concentration, cells can trigger a

This transient Ca2+ increase triggers the action of a wide range of Ca2+-dependent mediators that modify signaling pathways. Because Ca2+ is involved in numerous signaling pathways, differential features of Ca2+ increase are required to respond to diverse initial stimuli. This is achieved by the spatiotemporal control of the Ca2+ increase, i.e., the localization of the cytosolic Ca2+ spike within the cell, as well as the rate of the increase, the extent of the transient

This Ca2+ transport is therefore tightly regulated by specific channels, which are sensitive to hormones, cytokines, small molecules and other extracellular stimuli. The features of the transporters shape the characteristics of Ca2+ current (Ca2+ influx), modeling the kinetics of Ca2+ entry. Because a specific distribution of Ca2+ channels and transducers is required the cytoskeleton has been involved in the spatial regulation of Ca2+ entry. Interestingly, Ca2+ signaling strongly influences cytoskeleton dynamics, in an interesting interplay that is currently under study. In order to assess the crosstalk between Ca2+ signaling and cytoskeleton we will review here the recent literature regarding this topic, with special focus on the role of store-operated Ca2+ entry (SOCE) in cell migration, focal adhesion turnover and actin filaments reorganization. In addition, we will review the role of the microtubule cytoskeleton in the normal function of SOCE. A major regulator of SOCE is the protein STIM1, an endoplasmic reticulum resident protein that serves as intraluminal Ca2+ sensor and plasma membrane Ca2+ channel modulator. STIM1 is known to be a plus-end microtubule binding protein (+TIP). As a +TIP, STIM1 is transported throughout the cell while bound to microtubules, but it is released upon activation. The molecular basis of this regulation and the role of posttranslational modifica-

tions in STIM1 that are known to underlie this regulation, will be also described.

In eukaryotic cells plasma membrane (PM) contains different Ca2+ transport systems to control Ca2+ influx as well as Ca2+ extrusion. Because the temporal control of Ca2+ signaling is also strictly controlled, transporters are highly coordinated to let the Ca2+ spikes/waves last for a precise time, but this time ranges from microseconds (as in exocytosis) to hours, as observed during mammalian oocyte fertilization. To understand this control, we summarize here the

**2. Ca2+ transporters and Ca2+ influx pathways**

by increasing Ca2+ transport through plasma membrane.

*i* ) is sub-

expression, or reshaping the cytoskeleton in a second-messenger-dependent manner. To properly act as a second messenger the concentration of free cytosolic Ca2+ ([Ca2+]

*i*

increase and the kinetics of the final decrease to basal levels [3].

rapid and transient increase of [Ca2+]

74 Cytoskeleton - Structure, Dynamics, Function and Disease

Voltage-operated channels (VOCs) are regulated by the net electric charge across the plasma membrane in the way that they open upon depolarization. This family consists of three different groups of channels, Ca<sup>v</sup> 1 (L-type channels), Ca<sup>v</sup> 2 (N-, P/Q and R-types) and Ca<sup>v</sup> 3 (T-type channels) [4].

Receptor-operated channels (ROCs), which are gated upon binding to agonists such as ATP, glutamate, or acetylcholine [5]. Within this group we should highlight the transient receptor potential (TRP) ion channel family, a large family of channels involved in sensory perception, smooth muscle contraction-relaxation cycles and cell proliferation [6]. Members of the TRPC (transient receptor potential canonical) family are also involved in the organization of heteromeric Ca2+ channels that respond to intracellular Ca2+ store depletion [7]. This is the case for TRPC1 that has been described to be part of complexes together with members of the ORAI family [8]. Other important members of this family are P2X receptors, which are Ca2+ channels gated by extracellular ATP [9] and glutamate receptors, all of them reviewed in Ref. [10].

Second-messenger-operated channels (SMOCs) are members of an important family of channels gated by intracellular second messengers, such as arachidonic acid-regulated Ca2+ (ARC) channel or TRPC6, a member of the TRPC which is sensitive to diacylglycerol (DAG) [11].

Store-operated channels (SOCs) are channels that are regulated by the filling state of intracellular Ca2+ stores, mainly the sarco(endo)plasmic reticulum (ER). Their activity is actually the result of an initial Ca2+ release from the ER, mainly through the activation of the phosphoinositide pathway. A wide range of stimuli triggers the breakdown of phosphatidylinositol 4,5-bisphosphate (PIP2) into diacylglycerol (DAG) and inositol 1,4,5-trisphosphate (IP3), which activates IP3 receptors (IP3R) at the ER, leading to the transient release of Ca2+. Most of this Ca2+ is taken back to the ER by the sarco(endo)plasmic Ca2+ reticulum ATPase (SERCA), the Ca2+ pump in the ER membrane. However, some of this Ca2+ is extruded out of the cell by plasma membrane Ca2+ ATPases (PMCA), leading to the partial depletion of Ca2+ within the ER when the stimulation is repetitive. Thus, a system is required to replenish this Ca2+ and this action is mediated by SOCs that activate the Ca2+ influx pathway known as the store-operated Ca2+ entry (SOCE), described for the first time in the late 1980s by James W. Putney [12].

As stated above, there are some transporters considered as OFF systems, i.e., they are designed to decrease [Ca2+]*<sup>i</sup>* to basal levels (to approximately 100 nM) and therefore to shut down Ca2+ signaling. These are mainly Ca2+ pumps, ATP-consuming transporters that pump Ca2+ ions against the Ca2+ gradient concentration [13]. PMCAs and SERCAs extrude Ca2+ from the cytosol to extracellular milieu and into the ER vesicles. In addition to these systems, the plasma membrane Na+ /Ca2+ exchanger contributes to Ca2+ extrusion in a Na+ -gradient-dependent manner and it is particularly important in cardiac cells and neuronal tissues [14]. Finally, a mitochondrial Ca2+ uniporter (MCU) and the secretory-pathway Ca2+-ATPase (SPCA) are additional and widespread systems that restore cytosolic Ca2+ levels [15].

In summary, the combination of the activities of channels and pumps, either at the plasma membrane or at subcellular organelles and the particular expression profile of those transporters in different cells and tissues, makes possible for a cell-specific response to a wide range of stimuli.

### **3. Store-operated Ca2+ entry**

As stated above, store-operated Ca2+ entry (SOCE) is a Ca2+ influx transport system that maintains the permanence of the Ca2+-dependent signaling, because it preserves the intraluminal Ca2+ levels in the ER in the high micromolar range. For this extraordinary role, SOCE is a ubiquitous mechanism and is one of the most important pathways for Ca2+ entry in both excitable and nonexcitable cells [16]. However, the molecular nature of the members that control this Ca2+ influx pathway remained elusive until the description of two proteins, STIM1 and ORAI1, 10 years ago [17–22]. The mechanism that links luminal Ca2+ levels with plasma membrane Ca2+ entry is mediated by STIM1, a transmembrane protein located at the ER that acts as a Ca2+ sensor. STIM1 contains a Ca2+-sensitive EF-hand domain at the intraluminal domain and activates plasma membrane Ca2+ channels (SOCs) upon Ca2+ depletion within the intracellular stores. This depletion triggers the oligomerization of STIM1 and the relocalization in ER-PM juxtapositions required for the binding and activation of plasma membrane SOCs.

One of the most important SOCs is ORAI1 (also known as CRACM1), although the ORAI family contains two additional members, ORAI2 and ORAI3. In addition to ORAI1, some of the transient receptor potential canonical (TRPC) channels can function in a STIM1-dependent mode. STIM1 directly activates TRPC1, TRPC4 and TRPC5 channels that can therefore act as SOCs [23]. TRPC1 also binds to ORAI1 and the TRPC1-ORAI1-STIM1 ternary complexes can be therefore considered SOCs [24]. STIM1 also activates TRPC3 and TRPC6, not by direct interaction, but mediating the heteromultimerization of TRPC3 with TRPC1 and TRPC6 with TRPC4 [23].

The mode of STIM1-dependent gating for ORAIs and TRPCs is also different. STIM1 gates ORAI1 by direct binding through the STIM1-ORAI1-activating region, or SOAR, also called CRAC-activating domain or CAD. In contrast, the C-terminal polybasic domain of STIM1 activates TRPC1 by an electrostatic gating mechanism that results in SOC channel activation [25, 26].

STIM1 can bind other types of Ca2+ channels, such as Ca<sup>v</sup> 1.2 channels, but this binding leads to the suppression of the activation of these channels [27, 28]. Interestingly, this inhibitory action is mediated by the domain of STIM1 that activates ORAI1 (SOAR or CAD) and this direct binding to Cav 1.2 causes also the internalization of the channel, which explains the coordinated regulation of Ca2+ entry through VOCCS and SOCs. Thus, the combination of different Ca2+ channels provides a diversity of Ca2+ conductances in response to a wide variety of stimuli. However, the molecular mechanisms underlying the expected differential localization of STIM1 within cells are not clear. In this regard, the cytoskeleton seems to be a requirement for STIM1 function and binding to ORAI1 and therefore for the activation of SOCE. It was early observed that SOCE is sensitive to drugs that modify cytoskeletal components, such as cytochalasin D or nocodazole in NIH 3T3 cells [29], or vascular endothelial cells [30] and later confirmed in RBL-2H3 cells, bone marrow-derived mast cells [31], HEK293 cells [32] and platelets [33–35].

From the information given above, one can conclude that cytoskeleton is not only a critical modulator of SOCE, but also one can hypothesize that cytoskeleton might underlie the differential preference of STIM1 for different Ca2+ transporters.

### **4. Cytoskeletal components involved in the control of SOCE**

In summary, the combination of the activities of channels and pumps, either at the plasma membrane or at subcellular organelles and the particular expression profile of those transporters in different cells and tissues, makes possible for a cell-specific response to a wide

As stated above, store-operated Ca2+ entry (SOCE) is a Ca2+ influx transport system that maintains the permanence of the Ca2+-dependent signaling, because it preserves the intraluminal Ca2+ levels in the ER in the high micromolar range. For this extraordinary role, SOCE is a ubiquitous mechanism and is one of the most important pathways for Ca2+ entry in both excitable and nonexcitable cells [16]. However, the molecular nature of the members that control this Ca2+ influx pathway remained elusive until the description of two proteins, STIM1 and ORAI1, 10 years ago [17–22]. The mechanism that links luminal Ca2+ levels with plasma membrane Ca2+ entry is mediated by STIM1, a transmembrane protein located at the ER that acts as a Ca2+ sensor. STIM1 contains a Ca2+-sensitive EF-hand domain at the intraluminal domain and activates plasma membrane Ca2+ channels (SOCs) upon Ca2+ depletion within the intracellular stores. This depletion triggers the oligomerization of STIM1 and the relocalization in ER-PM juxtapositions required for the binding and activation of plasma membrane SOCs.

One of the most important SOCs is ORAI1 (also known as CRACM1), although the ORAI family contains two additional members, ORAI2 and ORAI3. In addition to ORAI1, some of the transient receptor potential canonical (TRPC) channels can function in a STIM1-dependent mode. STIM1 directly activates TRPC1, TRPC4 and TRPC5 channels that can therefore act as SOCs [23]. TRPC1 also binds to ORAI1 and the TRPC1-ORAI1-STIM1 ternary complexes can be therefore considered SOCs [24]. STIM1 also activates TRPC3 and TRPC6, not by direct interaction, but mediating the heteromultimerization of TRPC3 with TRPC1 and TRPC6 with TRPC4 [23]. The mode of STIM1-dependent gating for ORAIs and TRPCs is also different. STIM1 gates ORAI1 by direct binding through the STIM1-ORAI1-activating region, or SOAR, also called CRAC-activating domain or CAD. In contrast, the C-terminal polybasic domain of STIM1 activates TRPC1 by an electrostatic gating mechanism that results in SOC channel activa-

to the suppression of the activation of these channels [27, 28]. Interestingly, this inhibitory action is mediated by the domain of STIM1 that activates ORAI1 (SOAR or CAD) and this

coordinated regulation of Ca2+ entry through VOCCS and SOCs. Thus, the combination of different Ca2+ channels provides a diversity of Ca2+ conductances in response to a wide variety of stimuli. However, the molecular mechanisms underlying the expected differential localization of STIM1 within cells are not clear. In this regard, the cytoskeleton seems to be a requirement for STIM1 function and binding to ORAI1 and therefore for the activation of SOCE. It was early observed that SOCE is sensitive to drugs that modify cytoskeletal components, such

1.2 causes also the internalization of the channel, which explains the

1.2 channels, but this binding leads

range of stimuli.

tion [25, 26].

direct binding to Cav

STIM1 can bind other types of Ca2+ channels, such as Ca<sup>v</sup>

**3. Store-operated Ca2+ entry**

76 Cytoskeleton - Structure, Dynamics, Function and Disease

After the molecular description of STIM1 and ORAI1 as the two major elements controlling SOCE, it was earlier found that the microtubule cytoskeleton was involved in the regulation of this Ca2+ entry pathway. The group of James W. Putney described that STIM1 tagged with fluorescent proteins colocalized with endogenous tubulin and that the treatment with nocodazole, which induces tubulin depolymerization, triggered the loss of this colocalization in HEK293 cells. Moreover, nocodazole had an inhibitory effect on SOCE that could be reverted by the overexpression of YFP (yellow fluorescent protein)-STIM1 [32], suggesting that the microtubule cytoskeleton has an important role in the activation of SOCE by facilitating the appropriate localization of STIM1 to activate SOCs. Similar findings were reported in COS-7 cells, where the treatment with nocodazole triggered the retraction of tubulin filaments from cell periphery leading to the progressive loss of SOCE, similarly to what is found in mitosis. On the contrary, placlitaxel, which stabilizes microtubules, enhanced SOCE [36], supporting for a role of microtubules in the normal function of SOCE.

It was later demonstrated that STIM1 directly binds to EB1, a microtubule plus-end binding (or tracking) protein (+TIP) and that STIM1 is transported through the surface of the ER network by this microtubule-dependent mechanism [37]. In addition, it was observed that the overexpression of ectopic STIM1 led to the stimulation of the ER extension, an effect that was explained by the direct attachment of the ER to the growing ends of microtubules, which was stimulated by the accumulation of STIM1. Therefore, this observation also revealed that STIM1 concentration could regulate ER extension and remodeling. In addition, Luis Vaca group reported that STIM1 binds to EB1 when traveling through the ER under resting conditions and that there is a dissociation of this STIM1-EB1 complex upon Ca2+ depletion of the ER, an event that facilitates the clustering of STIM1 in the periphery of the cell [38].

STIM1-EB1 binding was studied further and it was found that the sequence Thr-Arg-Ile-Pro (TRIP) in the C-terminus of STIM1 is responsible for the direct binding to EB1 [39]. This sequence belongs to a short polypeptide motif, S/TxIP, found in several +TIPs. For those +TIPs with an S/TxIP motif, a phospho-regulation of the binding with microtubules has been reported. The +TIPs, APC [40, 41], MCAK [42] and CLASP2 [43, 44] are phosphorylated in the vicinity of the S/TxIP sequence, negatively regulating their interaction with microtubules. In these +TIPs we find a high number of proline, serine and basic residues that give a positive charge to the surroundings of the EB1 binding domain. This is why the negative charge of the phosphorylation of residues in the surroundings of the S/TxIP motif explains the blocking of the binding to microtubule ends. In this regard, our group reported that, as it was reported for those +TIPs, the binding of STIM1 to the tip of microtubules growing ends is regulated by phosphorylation. Near the S/TxIP sequence, which in human STIM1 is found in residues 642–645 (Thr-Arg-Ile-Pro), our group have described three phosphorylatable residues, Ser575, Ser608 and Ser621, which are target for ERK1/2 activity *in vitro* and *in vivo* [44, 45]. Using overexpressed STIM1 mutated at ERK1/2 target residues, we reported that dephosphorylated STIM1 (i.e., with Serto-Ala substitution mutations) remained bound to EB1, whereas constitutive phosphorylated STIM1 (mimicked by Ser-to-Glu mutations et ERK1/2 target sites) detached from EB1 [46, 47]. By means of the generation of phospho-specific antibodies directed against the three individual residues, we reported that there is a dynamic phosphorylation of STIM1 during its activation. Thus, activation of SOCE by thapsigargin or 2,5-di-ter-butyl-1,4-benzohydroquinone, two SERCA inhibitors that trigger ER emptying, is accompanied by the increase of phosphorylation at the ERK1/2 target Ser residues (575, 608 and 621). Moreover, the washout of the inhibitor with a Ca2+-containing medium, in order to let the refilling of the ER, carried out in parallel to STIM1 dephosphorylation [46]. In addition, Ser/Ala mutation inhibited SOCE and impaired the binding STIM1-ORAI1. In contrast, Ser/Glu mutations enhanced SOCE, whereas facilitated the clustering of STIM1 in response to store depletion, as well as the binding to ORAI1. Later, we reported that IGF-1 [48] and EGF [49] also trigger Ca2+ release from the ER and phosphorylation of STIM1, an effect that has been proven to be essential for the dissociation from EB1 and finally for the activation of SOCE (see **Figure 1**).

Taken together, those results support a mechanism that explains the reversible interaction of STIM1 and EB1 [46, 50]. This mechanism predicts that those stimuli that induce store depletion and ERK1/2 activation lead to phosphorylation of STIM1 at Ser575, Ser608 and Ser621, an event that promotes the dissociation of STIM1 from EB1. This dissociation enables STIM1 clustering and the binding to SOCs, in order to activate STIM1-dependent Ca2+ entry (SOCE). In addition, the activation of Ca2+ entry and the subsequent refilling of Ca2+ stores induce STIM1 dephosphorylation, promoting the association of STIM1 with EB1 and microtubules.

In addition to EB1, other members of the cytoskeleton have been involved in SOCE regulation. For instance, it has been reported that the microtubule-binding protein adenomatous polyposis coli (APC) is required for STIM1 puncta near ER-PM junctions, because reduced STIM1 was observed at these junctions in APC-depleted cells and this effect correlated well with the inhibition of SOCE [51]. The APC-binding domain was found in the C-terminus of STIM1 (residues 650–685), similarly to what it has been reported for STIM1-EB1. Thus, it can be assumed that upon depletion of the ER, STIM1 dissociates from EB1 and associates to APC to form puncta near ER-PM junctions and to activate ORAI1 and SOCE.

A downstream effector of SOCE is the transcription factor NFAT (nuclear factor of activated T-cells). Once Ca2+ entry becomes activated, the increase of [Ca2+]*<sup>i</sup>* activates the Ca2+-dependent phosphatase calcineurin (or protein phosphatase 2B), which dephosphorylates NFAT, promoting the nuclear translocation of NFAT that can be easily monitored using GFP-tagged NFAT. By means of this well-established feature of SOCE, Sharma et al. [52] designed a genomewide RNA interference screen for NFAT activation in HeLa cells and they identified septin proteins as key regulators of SOCE. Knockdown of SEPT4 gene expression reduced SOCE

microtubule ends. In this regard, our group reported that, as it was reported for those +TIPs, the binding of STIM1 to the tip of microtubules growing ends is regulated by phosphorylation. Near the S/TxIP sequence, which in human STIM1 is found in residues 642–645 (Thr-Arg-Ile-Pro), our group have described three phosphorylatable residues, Ser575, Ser608 and Ser621, which are target for ERK1/2 activity *in vitro* and *in vivo* [44, 45]. Using overexpressed STIM1 mutated at ERK1/2 target residues, we reported that dephosphorylated STIM1 (i.e., with Serto-Ala substitution mutations) remained bound to EB1, whereas constitutive phosphorylated STIM1 (mimicked by Ser-to-Glu mutations et ERK1/2 target sites) detached from EB1 [46, 47]. By means of the generation of phospho-specific antibodies directed against the three individual residues, we reported that there is a dynamic phosphorylation of STIM1 during its activation. Thus, activation of SOCE by thapsigargin or 2,5-di-ter-butyl-1,4-benzohydroquinone, two SERCA inhibitors that trigger ER emptying, is accompanied by the increase of phosphorylation at the ERK1/2 target Ser residues (575, 608 and 621). Moreover, the washout of the inhibitor with a Ca2+-containing medium, in order to let the refilling of the ER, carried out in parallel to STIM1 dephosphorylation [46]. In addition, Ser/Ala mutation inhibited SOCE and impaired the binding STIM1-ORAI1. In contrast, Ser/Glu mutations enhanced SOCE, whereas facilitated the clustering of STIM1 in response to store depletion, as well as the binding to ORAI1. Later, we reported that IGF-1 [48] and EGF [49] also trigger Ca2+ release from the ER and phosphorylation of STIM1, an effect that has been proven to be essential for the dissociation from EB1 and

Taken together, those results support a mechanism that explains the reversible interaction of STIM1 and EB1 [46, 50]. This mechanism predicts that those stimuli that induce store depletion and ERK1/2 activation lead to phosphorylation of STIM1 at Ser575, Ser608 and Ser621, an event that promotes the dissociation of STIM1 from EB1. This dissociation enables STIM1 clustering and the binding to SOCs, in order to activate STIM1-dependent Ca2+ entry (SOCE). In addition, the activation of Ca2+ entry and the subsequent refilling of Ca2+ stores induce STIM1 dephosphorylation, promoting the association of STIM1 with EB1 and microtubules. In addition to EB1, other members of the cytoskeleton have been involved in SOCE regulation. For instance, it has been reported that the microtubule-binding protein adenomatous polyposis coli (APC) is required for STIM1 puncta near ER-PM junctions, because reduced STIM1 was observed at these junctions in APC-depleted cells and this effect correlated well with the inhibition of SOCE [51]. The APC-binding domain was found in the C-terminus of STIM1 (residues 650–685), similarly to what it has been reported for STIM1-EB1. Thus, it can be assumed that upon depletion of the ER, STIM1 dissociates from EB1 and associates to APC

A downstream effector of SOCE is the transcription factor NFAT (nuclear factor of activated

phosphatase calcineurin (or protein phosphatase 2B), which dephosphorylates NFAT, promoting the nuclear translocation of NFAT that can be easily monitored using GFP-tagged NFAT. By means of this well-established feature of SOCE, Sharma et al. [52] designed a genomewide RNA interference screen for NFAT activation in HeLa cells and they identified septin proteins as key regulators of SOCE. Knockdown of SEPT4 gene expression reduced SOCE

activates the Ca2+-dependent

to form puncta near ER-PM junctions and to activate ORAI1 and SOCE.

T-cells). Once Ca2+ entry becomes activated, the increase of [Ca2+]*<sup>i</sup>*

finally for the activation of SOCE (see **Figure 1**).

78 Cytoskeleton - Structure, Dynamics, Function and Disease

**Figure 1.** Involvement of SOCE in cell migration. The binding of ligands to some plasma membrane receptors activates the phosphoinositide pathway, releasing IP3 and activating Ca2+ release from the ER. In parallel, receptor tyrosine kinases trigger the activation of the MAPK pathway, via activation of Ras-Raf-MEK-ERK. Ca2+ mobilization from the ER and phosphorylation of STIM1 are two essential events for the activation of STIM1. Phospho-STIM1 releases from microtubules facilitating STIM1 clustering and translocation to PM-ER junctions to activate ORAI1. This activation increases [Ca2+]*<sup>i</sup>* locally, an event required for the phosphorylation of Src, PYK2 and FAK kinases, as well as MLC2, which are well-known modulators of invadopodia formation, focal adhesion turnover and actomyosin contractility.

without affecting ER-Ca2+ release and without inhibiting SERCA or PMCAs. This knockdown also decreased Ca2+-induced NFAT translocation by >95%. Septins were found to be required for proper organization of ORAI1 in the plasma membrane even before depletion of ER-Ca2+ stores. Septins also promoted the targeting of STIM1 to ER-PM junctions and the formation of stable ORAI1 clusters after store depletion. Because septins are considered scaffold proteins that recruit other proteins, preventing diffusion, septins should be considered promoters of the stable recruitment of STIM1-ORAI1 at ER-PM junctions. More recently it was found that Septin 7 inhibits the activation of Orai channels in Drosophila neurons (dOrai) and that the depletion of Septin 7 levels results in higher dOrai-mediated Ca2+ entry, independently of the filling state of the ER. In fact, overexpression of Septin 7 reduced Ca2+ entry, suggesting that, in Drosophila neurons, Septin 7 is a negative regulator of dOrai channel function [53]. These authors stated that hetero-hexamers septin filaments closely associate with the PM and near the ER in resting neurons and that the reduction of SEPT7 results in breaks in the linear septin filaments present in wild-type cells, leading to the formation of shorter septin filaments. ER-Ca2+ store depletion reorganizes these filaments, moving STIM1 to the peripheral ER, promoting the coupling of STIM/dOrai and the activation of Ca2+ entry through dOrai (SOCE). Shorter septin filaments due to SEPT7 knockdown leads to STIM1 recruitment to the peripheral ER in resting neurons and the activation of dOrai, resulting in a store-independent activation of dOrai [53].

Homer1 is another scaffolding protein that binds to TRPC channels through a PPPF sequence of the Ca2+ channel. However, this sequence is close to the STIM1 binding sequence, thus suggesting that Homer1 and STIM1 could be competitors for binding to TRPC. Yuan et al. [54] proposed that Homer couples TRPC channels to IP3 receptors (IP3R) to keep these channels closed, but dissociation of the TRPC-Homer-IP3R complexes gives STIM1 access to TRPC binding to gate these channels. A similar proposal was later reported for STIM1 and Cav1.2 channels in HEK293 cells [55], where the treatment with thapsigargin induces coimmunoprecipitation of Homer1 with STIM1 and the Cav1.2 α1 subunit. Impairment of Homer1 function with the peptide PPKKFR or by siRNA specific for Homer1 reduced the association of STIM1 to Cav1.2 α1, adding a new scaffolding protein to the list of regulators of Ca2+ entry.

In summary, the increasing number of members of the cytoskeleton involved in the specific interaction of STIM1 with different plasma membrane Ca2+ channels leads to the conclusion that the cytoskeleton strongly modulates SOCE, as well as the activation and inhibition of others Ca2+ transport systems.

### **5. How SOCE modulates cytoskeleton dynamics and cell migration**

Because the localization and function of STIM1 and ORAI1 are strongly dependent on components of the actin and tubulin cytoskeleton, the question of how SOCE and Ca2+ dynamics influence cytoskeleton-dependent events, such as cell adhesion and migration, was rapidly considered by many groups.

Focal adhesions are complexes of macromolecules that serve as mechanical links between the extracellular substrate and the cytoskeleton. These molecular assemblies are highly dynamic. However, the molecular mechanism by which Ca2+ regulates focal adhesion turnover is still unclear. A few proteins that regulate focal adhesion assembly or disassembly are sensitive to changes in the intracellular free Ca2+ concentration. For instance, the Ca2+ dependent protease calpain [56], which could cleave talin at focal adhesion sites, leads to an increase of focal adhesion disassembly rates. Indeed, disassembly of other focal adhesion components, like paxillin, vinculin and zyxin, has been shown to be dependent on the cleavage of talin by calpain, suggesting a role for the Ca2+-dependent talin proteolysis in the regulation of focal adhesion turnover [57]. In this regard, it has been found that the reduction of STIM1 or ORAI1 gene expression, by RNA interference, dramatically affected the rate of focal adhesion turnover, slowing down cell migration *in vitro* and inhibiting metastasis of MDA-MB-231 cancer cells in nude mice [58]. The treatment of cells with SKF96365, a SOC inhibitor, also induced large focal adhesions *in vitro*, due to defective focal adhesion turnover. In addition, this blocking agent also inhibited tumor progression in mice [58].

stable ORAI1 clusters after store depletion. Because septins are considered scaffold proteins that recruit other proteins, preventing diffusion, septins should be considered promoters of the stable recruitment of STIM1-ORAI1 at ER-PM junctions. More recently it was found that Septin 7 inhibits the activation of Orai channels in Drosophila neurons (dOrai) and that the depletion of Septin 7 levels results in higher dOrai-mediated Ca2+ entry, independently of the filling state of the ER. In fact, overexpression of Septin 7 reduced Ca2+ entry, suggesting that, in Drosophila neurons, Septin 7 is a negative regulator of dOrai channel function [53]. These authors stated that hetero-hexamers septin filaments closely associate with the PM and near the ER in resting neurons and that the reduction of SEPT7 results in breaks in the linear septin filaments present in wild-type cells, leading to the formation of shorter septin filaments. ER-Ca2+ store depletion reorganizes these filaments, moving STIM1 to the peripheral ER, promoting the coupling of STIM/dOrai and the activation of Ca2+ entry through dOrai (SOCE). Shorter septin filaments due to SEPT7 knockdown leads to STIM1 recruitment to the peripheral ER in resting neurons and the activation of dOrai, resulting in a store-independent

Homer1 is another scaffolding protein that binds to TRPC channels through a PPPF sequence of the Ca2+ channel. However, this sequence is close to the STIM1 binding sequence, thus suggesting that Homer1 and STIM1 could be competitors for binding to TRPC. Yuan et al. [54] proposed that Homer couples TRPC channels to IP3 receptors (IP3R) to keep these channels closed, but dissociation of the TRPC-Homer-IP3R complexes gives STIM1 access to TRPC binding to gate these channels. A similar proposal was later reported for STIM1 and Cav1.2 channels in HEK293 cells [55], where the treatment with thapsigargin induces coimmunoprecipitation of Homer1 with STIM1 and the Cav1.2 α1 subunit. Impairment of Homer1 function with the peptide PPKKFR or by siRNA specific for Homer1 reduced the association of STIM1 to Cav1.2 α1, adding a new scaffolding protein to the list of regulators

In summary, the increasing number of members of the cytoskeleton involved in the specific interaction of STIM1 with different plasma membrane Ca2+ channels leads to the conclusion that the cytoskeleton strongly modulates SOCE, as well as the activation and inhibition of

Because the localization and function of STIM1 and ORAI1 are strongly dependent on components of the actin and tubulin cytoskeleton, the question of how SOCE and Ca2+ dynamics influence cytoskeleton-dependent events, such as cell adhesion and migration, was rapidly

Focal adhesions are complexes of macromolecules that serve as mechanical links between the extracellular substrate and the cytoskeleton. These molecular assemblies are highly dynamic. However, the molecular mechanism by which Ca2+ regulates focal adhesion turnover is still unclear. A few proteins that regulate focal adhesion assembly or disassembly

**5. How SOCE modulates cytoskeleton dynamics and cell migration**

activation of dOrai [53].

80 Cytoskeleton - Structure, Dynamics, Function and Disease

of Ca2+ entry.

others Ca2+ transport systems.

considered by many groups.

Focal adhesion kinase (FAK) and proline-rich tyrosine kinase 2 beta (PTK2B or PYK2) are two well-known kinases involved in focal adhesion assembly [59]. Because focal adhesion targeting of PYK2 is required for the turnover and the Tyr402 autophosphorylation is required for PYK2 targeting, Chen et al. studied the correlation between EGF stimulation, PYK2 phosphorylation and STIM1 levels in SiHa cells [60]. EGF activates PYK2 phosphorylation at Tyr402, but this phosphorylation has been shown to be inhibited by the silencing of STIM1 [60]. In addition, STIM1 knockdown induced large focal adhesions, independently of the stimulation with EGF, further confirming the role of Ca2+ entry in facilitating the turnover of focal adhesions. Similarly, EGF triggered the phosphorylation of FAK at Tyr397, which is required for focal adhesion turnover and it was demonstrated that this phosphorylation is inhibited by STIM1 knockdown [60].

In this regard, it is known that EGF activates the phosphoinositide pathway, generating IP3 and activating Ca2+ release from the ER. EGF also activates de MAPK (mitogen-activated protein kinases) pathway. As a consequence STIM1 becomes phosphorylated at ERK1/2 target sites (Ser575, Ser608 and Ser621) upon EGF stimulation, leading to the dissociation from microtubules (i.e., EB1) and activating SOCE [49]. Casas-Rua et al. demonstrated that nonphosphorylatable mutants of STIM1, such as STIM1-S575A/S308A/S621A, blocked EGF signaling pathway, inhibiting cell migration in the endometrial adenocarcinoma Ishikawa cell line [49]. The impact of STIM1 was observed also at the genomic response level, because ectopic overexpression of STIM1-S575A/S608A/S621A blocked the epithelial-mesenchymal transition (EMT) of Ishikawa cells treated with EGF. The stimulation of cells with EGF induced a significant switch in E-cadherin localization from subplasma membrane region to a diffuse localization throughout the cytosol, as described for other epithelial cells [61]. EGF also triggered an increase of vimentin expression in well-defined cytoskeletal localization, as for other cells upon EGF stimulation [61, 62]. However, Ishikawa cells overexpressing STIM1- S575A/S608A/S621A-mCherry did not show significant increase in vimentin expression, nor E-cadherin relocalization [49], supporting for a role for phospho-STIM1 in the regulation of cell migration and cell transformation into a mesenchymal phenotype.

Cytosolic Ca2+ levels also regulate actomyosin, the macromolecular complex of actin and myosin that drives the mechanical forces for cell contractility during migration. Nonmuscle cell contractility is controlled by nonmuscle myosin II, through the phosphorylation of its regulatory light chains (MLC2) at Ser19 in a Ca2+/calmodulin-dependent manner [63]. The contractile force that moves the cell body is then transmitted to focal adhesions by phospho-myosin II-based actomyosin contraction. This phosphorylation was abolished by the Ca2+ channel inhibitor SKF96365, or by knocking-down STIM1 expression [64], indicating that STIM1-dependent Ca2+ entry has a significant role in MLC2 activation and in the reorganization of the actin-myosin cytoskeleton in migrating cells.

In addition to focal adhesions, podosomes and invadopodia are also adhesion structures regulated by Ca2+ signaling. Sun et al. reported that melanoma invasion is promoted by STIM1- and ORAI1-mediated Ca2+ oscillations, which promote invadopodia formation and extracellular matrix (ECM) degradation [65]. These authors found that Ca2+ signaling mediated by STIM1 and ORAI1 is essential for invadopodia formation over a collagen matrix and that addition of SKF96365, or chelation of extracellular Ca2+ with EGTA, blocked invadopodia assembly. More interestingly, ectopic expression of STIM1 or STIM1+ORAI1 increased phosphorylation levels of Tyr416 in Src kinase, without affecting phospho-FAK. This effect was revealed for MCF-7 cells (human breast cancer cells), NMuMG (mouse mammary epithelial cell line) and WM793 melanoma cells and the activation of Ca2+ entry with thapsigargin or ionophore A23187 led to the same result, i.e., a rapid increase of pY416 Src levels. These data, together with the fact that STIM1 shRNA, chelation of extracellular Ca2+, or the inhibition of SOCE with 2-APB, reduced pY416 Src levels, strongly suggesting a direct role of STIM1-ORAI1-mediated Ca2+ influx in the preservation of Src activity and invadopodia formation [65].

From the information given above it is now widely accepted that one of the major targets for SOCE is the cytoskeleton and that the dynamics of the focal adhesion assembly as well as the dynamics of actin and tubulin cytoskeleton are strongly influenced by the kinetics of Ca2+ entry through store-operated Ca2+ channels.

### **6. Localization and polarization of SOCE in migrating cells**

From the given information arises the question about the spatial control of STIM1-ORAI1 localization. The polarization of Ca2+ entry pathways has been described in several cell types, especially in exocrine gland cells. In pancreatic acinar cells under stimulation of Ca2+ mobilizing receptors, the ER Ca2+ release is detected in the apical region of the cell and the [Ca2+] *i* increase remains restricted at this region [66], although the signal propagates to basolateral regions at high concentration of agonists. In salivary gland acinar cells, [Ca2+] *i* increase is also detected in the apical region and then propagates to the basal region [67]. Indeed, several studies have demonstrated that agonist-stimulated Ca2+ signaling in exocrine gland cells is highly polarized. TRPC1 is a key factor for SOCE in salivary gland acinar cells and pancreatic acinar cells. In addition, TRPC3 contributes to SOCE and contributes to the receptorstimulated Ca2+ influx in exocrine pancreatic acinar cells [68]. In addition, SOCE is the major contributor to Ca2+ influx in salivary gland and pancreatic acinar cells, so it is expected an asymmetric distribution of STIM1, ORAI1 and TRPC1-3, some of the most widely studied members involved in the control of SOCE.

As expected, a polarized localization has been shown for all Ca2+ signaling proteins in exocrine acinar cells: IP3Rs, SERCAs and PMCA pumps, GPCRs, TRPC channels, ORAI1 channels and STIM1 [66, 69, 70], an asymmetric distribution required for the directed secretion carried out by these cells. However, little is known about the mechanisms involved in the targeting of Ca2+-signaling complexes to these regions.

Because migrating cells are polarized cells, with polarized distribution of receptor tyrosine kinases, the study of Ca2+ entry in migrating cells is a task of particular importance. In this regard, Tsai et al. reported recently that STIM1 is enriched at the leading edge of migrating cells, measured with YFP-STIM1 and CFP-tagged ER marker [71]. The authors concluded that the polarization of STIM1 is microtubule-plus-end dependent, because STIM1-I644N/P645N, a mutant STIM1 that does not bind to EB1, failed to polarize in migrating cells. Because of the enrichment of receptor tyrosine kinases at the front of migrating cells, local Ca2+ pulses were observed with higher frequency at the leading edge, accompanied by a lower level of luminal ER Ca2+ and increased levels of PMCA activity at the front. In this report the authors also described that a DAG gradient is the result of the asymmetric activity of phospholipase C (PLC) at the leading edge of migrating cells, leading to the recruitment and activation of PKCbeta, a kinase involved in migration [72] by phosphorylating myosin [73], or other substrates of the cytoskeleton, such as GAP43, adducin, or fascin [74]. However, the precise mechanism that assembles de STIM1-ORAI1 and/or TRPC1 in a polarized distribution in cells is still unclear and it is expected that additional members of the cytoskeleton will solve this open question.

### **7. Conclusions**

*i*

increase is also

*i*

cell contractility is controlled by nonmuscle myosin II, through the phosphorylation of its regulatory light chains (MLC2) at Ser19 in a Ca2+/calmodulin-dependent manner [63]. The contractile force that moves the cell body is then transmitted to focal adhesions by phospho-myosin II-based actomyosin contraction. This phosphorylation was abolished by the Ca2+ channel inhibitor SKF96365, or by knocking-down STIM1 expression [64], indicating that STIM1-dependent Ca2+ entry has a significant role in MLC2 activation and in the

In addition to focal adhesions, podosomes and invadopodia are also adhesion structures regulated by Ca2+ signaling. Sun et al. reported that melanoma invasion is promoted by STIM1- and ORAI1-mediated Ca2+ oscillations, which promote invadopodia formation and extracellular matrix (ECM) degradation [65]. These authors found that Ca2+ signaling mediated by STIM1 and ORAI1 is essential for invadopodia formation over a collagen matrix and that addition of SKF96365, or chelation of extracellular Ca2+ with EGTA, blocked invadopodia assembly. More interestingly, ectopic expression of STIM1 or STIM1+ORAI1 increased phosphorylation levels of Tyr416 in Src kinase, without affecting phospho-FAK. This effect was revealed for MCF-7 cells (human breast cancer cells), NMuMG (mouse mammary epithelial cell line) and WM793 melanoma cells and the activation of Ca2+ entry with thapsigargin or ionophore A23187 led to the same result, i.e., a rapid increase of pY416 Src levels. These data, together with the fact that STIM1 shRNA, chelation of extracellular Ca2+, or the inhibition of SOCE with 2-APB, reduced pY416 Src levels, strongly suggesting a direct role of STIM1-ORAI1-mediated Ca2+ influx in

From the information given above it is now widely accepted that one of the major targets for SOCE is the cytoskeleton and that the dynamics of the focal adhesion assembly as well as the dynamics of actin and tubulin cytoskeleton are strongly influenced by the kinetics of Ca2+

From the given information arises the question about the spatial control of STIM1-ORAI1 localization. The polarization of Ca2+ entry pathways has been described in several cell types, especially in exocrine gland cells. In pancreatic acinar cells under stimulation of Ca2+ mobilizing receptors, the ER Ca2+ release is detected in the apical region of the cell and the [Ca2+]

increase remains restricted at this region [66], although the signal propagates to basolateral

detected in the apical region and then propagates to the basal region [67]. Indeed, several studies have demonstrated that agonist-stimulated Ca2+ signaling in exocrine gland cells is highly polarized. TRPC1 is a key factor for SOCE in salivary gland acinar cells and pancreatic acinar cells. In addition, TRPC3 contributes to SOCE and contributes to the receptorstimulated Ca2+ influx in exocrine pancreatic acinar cells [68]. In addition, SOCE is the major contributor to Ca2+ influx in salivary gland and pancreatic acinar cells, so it is expected an asymmetric distribution of STIM1, ORAI1 and TRPC1-3, some of the most widely studied

reorganization of the actin-myosin cytoskeleton in migrating cells.

the preservation of Src activity and invadopodia formation [65].

**6. Localization and polarization of SOCE in migrating cells**

regions at high concentration of agonists. In salivary gland acinar cells, [Ca2+]

entry through store-operated Ca2+ channels.

82 Cytoskeleton - Structure, Dynamics, Function and Disease

members involved in the control of SOCE.

In conclusion, cytosolic-free Ca2+ concentration regulates the reorganization of cytoskeleton, focal adhesion turnover, invadopodia formation, actomyosin contractility and it is critical to trigger the development of lamellipodia as the leading structure during migration. But this dependence is reciprocal and Ca2+ influx through store-operated Ca2+ channels at the plasma membrane is fully dependent on the formation of endoplasmic reticulum-plasma membrane juxtapositions that are shaped by the reorganization of the cytoskeleton. In the last few years valuable knowledge has been gained regarding the activation of STIM1 by Ca2+ store depletion and how STIM1 relocalizes and activates ORAI1 at the PM-ER junctions. Here we have shown that both STIM1 and ORAI1 are involved in many aspects of cell migration and that gene silencing and specific inhibitors point out these two proteins in the regulation of Ca2+ influx pathways involved in supporting efficient cell adhesion and migration. However, a major lack of knowledge regarding the polarization of the signaling profile persists. Recent advances in genome editing will be valuable tools to knockout and knockin *STIM* and *ORAI* genes, as well as genes coding for cytoskeletal proteins involved in the reorganization of SOCE. With this coming era we will be able to monitor and study the behavior of tagged proteins at endogenous levels, as well as to study the loss of function of certain genes in any cell type, in an attempt to solve this open question in cell biology and signaling.

### **Author details**

Francisco Javier Martin-Romero¹\*, Aida M. Lopez-Guerrero¹, Carlos Pascual-Caro¹ and Eulalia Pozo-Guisado²

\*Address all correspondence to: fjmartin@unex.es

1 Department of Biochemistry and Molecular Biology, School of Life Sciences, University of Extremadura, Badajoz, Spain

2 Department of Cell Biology, School of Medicine, University of Extremadura, Badajoz, Spain

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**Author details**

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Eulalia Pozo-Guisado²

Extremadura, Badajoz, Spain

\*Address all correspondence to: fjmartin@unex.es

84 Cytoskeleton - Structure, Dynamics, Function and Disease

Francisco Javier Martin-Romero¹\*, Aida M. Lopez-Guerrero¹, Carlos Pascual-Caro¹ and

1 Department of Biochemistry and Molecular Biology, School of Life Sciences, University of

2 Department of Cell Biology, School of Medicine, University of Extremadura, Badajoz, Spain

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#### **Role of Band 3 in the Erythrocyte Membrane Structural Changes Under Isotonic and Hypotonic Conditions Role of Band 3 in the Erythrocyte Membrane Structural Changes Under Isotonic and Hypotonic Conditions**

Ivana Pajic‐Lijakovic and Milan Milivojevic Ivana Pajic‐Lijakovic and Milan Milivojevic

Additional information is available at the end of the chapter Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/64964

#### **Abstract**

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An attempt was made to discuss and connect various modeling approaches which have been proposed in the literature in order to shed further light on the erythrocyte membrane relaxation under isotonic and hypotonic conditions. Roles of the main membrane constituents: (1) the actin‐spectrin cortex, (2) the lipid bilayer, and (3) the transmembrane protein band 3 and its course‐consequence relations were considered to estimate the membrane relaxation phenomena. Cell response to loading conditions includes the successive sub‐bioprocesses: (1) erythrocyte local or global deformation, (2)the cortex‐bilayer coupling, and (3)the rearrangements of band 3. The results indicate that the membrane structural changes include: (1) the spectrin flexibility distribution and (2) the rate of its changes influenced by the number of band 3 molecules attached to spectrin filaments, and phosphorylation of the actin‐spectrin junctions. Band 3 rearrangement also influences: (1) the effective bending modulus and (2) the band 3‐ bilayer interaction energy and on that base the bilayer bending state. The erythrocyte swelling under hypotonic conditions influences the bilayer integrity which leads to the hemolytic hole formation. The hemolytic hole represents the excited cluster of band 3 molecules.

**Keywords:** packing state changes of band 3 clusters, reversible hemolytic hole forma‐ tion, the lipid bilayer bending state changes, the spectrin inter‐ and intrachain interac‐ tions, mathematical modeling

### **1. Introduction**

Erythrocyte mechanics under isotonic and hypotonic conditions has been studied from engineering and biomedical stand points [1–14]. Rheological response of the erythrocyte

© 2016 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

membrane depends on the main membrane constituent rearrangement: (1) the actin cortex, (2) the lipid bilayer, and (3) the transmembrane protein band 3. The membrane fluctuations under isotonic condition induce alternating expansion and compression of the membrane parts in order to ensure surface and volume conservation. The membrane relaxation occurs within (millisecond order) affine regime and (second order) nonaffine regime. The affine regime corresponds to the spectrin interchain interactions while the nonaffine regime corresponds to the spectrin intrachain interactions. However, the membrane fluctuations under hypotonic condition induce volume increase by ensuring surface conservation. The membrane response under hypotonic condition includes the successive sub‐processes: (1) erythrocyte swelling, (2) lifetime of the lipid structural integrity and the rearrangement of transmembrane protein band 3, and (3) the reversible hemolytic hole formation and hemoglobin (Hb) release to surround‐ ing solution. Duration of the membrane relaxation depends on contributions of three sub‐ processes: (1) time for cell swelling *tsw* ∈ [5, 100 *s*], (2) membrane lifetime *τm*∈(0, *tH* ) (where *tH* is the hemolytic time), and (3) time for Hb release from already formed hemolytic hole during successive open‐closed state changes *trelease* ∈ [1,5 *s*] [11, 13].

The band 3 rearrangement significantly influences the state of both, the lipid bilayer and the actin‐spectrin cortex. Space distribution of band 3 molecules and their lateral diffusion influence the bending state of the lipid bilayer and its free energy [15–17]. Local changes of the bilayer bending state enhance anomalous sub‐diffusion and eventually lead to hop‐ diffusion of lipids. These effects induce anomalous nature of energy dissipation during lipids structural ordering and could be quantified by the effective viscosity [18]. The bilayer structural changes have the feedback effects to band 3 protein‐lipids positive hydrophobic mismatch effects [19–21]. De Meyer et al. [19] pointed to the cholesterol role in lipid mediat‐ ed protein‐protein interactions. These effects could lead to the protein tilt angle changes and the protein clustering. This clustering can be modulated by homotypic interactions of the protein transmembrane domains and electrostatic protein‐lipid interactions [21]. Tilt angle changes influence packing state of band 3 clusters and their association‐dissociation to spectrin. Band 3 molecules form various complexes with spectrin and influence its conforma‐ tional changes. In‐homogeneous distribution of band 3 molecules and their ability to cluster‐ ing influence the rheological response of: (1) the bilayer, (2) the cortex, and (3) the nature of the bilayer‐cortex mechanical coupling. Ehrig et al. [22] pointed that the lipid bilayer phase separation can be strongly affected by interaction with the actin cortex, which, depending on the temperature and membrane composition, can either lead to precipitation of highly dynamic membrane domains (rafts), or prevent large‐scale phase separation.

For understanding the influence of band 3 rearrangement on complex nature of the mem‐ brane rheological response, it is necessary to consider three subpopulations of band 3 molecules under isotonic and hypotonic conditions. The first subpopulation (20–40%) as tetramers forms high affinity complexes with ankyrin (quantified by the dissociation con‐ stant ∼5 nM) as reported by Tomishige et al. [23] and Kodippili et al. [24]. Band 3‐ankyrin complexes are located near the center of spectrin tetramers. These complexes could survive the hypotonic conditions. Golan and Veatch [25] reported that 25% of band 3 population remains attached to the cortex underionic strength 26 mM NaPO4 solution at 37°C. The second subpopulation (∼30%) as dimers forms lower affinity complexes with the adducin (the dissociation constant is ∼100 nM) as reported by Franco and Low [26] and Kodippili et al. [24]. This subpopulation is located at the spectrin‐actin junction complexes. The junctions of the network link 4–7 spectrin filaments [27]. The third one is freely diffusing subpopulation (∼30%). The part of freely diffusing subpopulation increases under hypotonic conditions. Golen and Veatch [25] determined that mobile fraction of band 3 molecules under the external solution tonicity 46.0 mM NaPO4 at 21°C was 11 ± 9%. Undertonicity of 5.2 mM NaPO4 at 21°C, the mobile fraction of band 3 molecules was 72 ± 7%. Band 3 molecules are anion at pH = 7. The value of Stokes radius of band 3 dimmer is approximately 7.6 nm while for tetramer is 11  nm at room temperature and pH = 7.2 [28]. The total number of band 3 per single erythro‐ cyte is ∼1 × 10<sup>6</sup> . Thermal fluctuations of the erythrocyte membrane could induce conforma‐ tional changes of cytoplasmic domain of transmembrane protein band 3 [29]. Such conformational changes result in electrostatic interactions between highly anionic N‐termi‐ nal domains of band 3 molecules and on that base intensified their short‐range self‐associa‐ tive tendency [28]. Long‐range self‐associative tendency is induced by positive hydrophobic mismatch effects [21]. The band 3 clustering is pronounced under hypotonic conditions [30]. In this case, ∼25% of band 3 molecules make aggregate of ∼5000 mers [31]. All band 3 subpopulations through these complexes contribute to the spectrin conformational changes by reducing its mobility and influence the cortex stiffening.

membrane depends on the main membrane constituent rearrangement: (1) the actin cortex, (2) the lipid bilayer, and (3) the transmembrane protein band 3. The membrane fluctuations under isotonic condition induce alternating expansion and compression of the membrane parts in order to ensure surface and volume conservation. The membrane relaxation occurs within (millisecond order) affine regime and (second order) nonaffine regime. The affine regime corresponds to the spectrin interchain interactions while the nonaffine regime corresponds to the spectrin intrachain interactions. However, the membrane fluctuations under hypotonic condition induce volume increase by ensuring surface conservation. The membrane response under hypotonic condition includes the successive sub‐processes: (1) erythrocyte swelling, (2) lifetime of the lipid structural integrity and the rearrangement of transmembrane protein band 3, and (3) the reversible hemolytic hole formation and hemoglobin (Hb) release to surround‐ ing solution. Duration of the membrane relaxation depends on contributions of three sub‐ processes: (1) time for cell swelling *tsw* ∈ [5, 100 *s*], (2) membrane lifetime *τm*∈(0, *tH* ) (where *tH* is the hemolytic time), and (3) time for Hb release from already formed hemolytic hole during

The band 3 rearrangement significantly influences the state of both, the lipid bilayer and the actin‐spectrin cortex. Space distribution of band 3 molecules and their lateral diffusion influence the bending state of the lipid bilayer and its free energy [15–17]. Local changes of the bilayer bending state enhance anomalous sub‐diffusion and eventually lead to hop‐ diffusion of lipids. These effects induce anomalous nature of energy dissipation during lipids structural ordering and could be quantified by the effective viscosity [18]. The bilayer structural changes have the feedback effects to band 3 protein‐lipids positive hydrophobic mismatch effects [19–21]. De Meyer et al. [19] pointed to the cholesterol role in lipid mediat‐ ed protein‐protein interactions. These effects could lead to the protein tilt angle changes and the protein clustering. This clustering can be modulated by homotypic interactions of the protein transmembrane domains and electrostatic protein‐lipid interactions [21]. Tilt angle changes influence packing state of band 3 clusters and their association‐dissociation to spectrin. Band 3 molecules form various complexes with spectrin and influence its conforma‐ tional changes. In‐homogeneous distribution of band 3 molecules and their ability to cluster‐ ing influence the rheological response of: (1) the bilayer, (2) the cortex, and (3) the nature of the bilayer‐cortex mechanical coupling. Ehrig et al. [22] pointed that the lipid bilayer phase separation can be strongly affected by interaction with the actin cortex, which, depending on the temperature and membrane composition, can either lead to precipitation of highly

dynamic membrane domains (rafts), or prevent large‐scale phase separation.

For understanding the influence of band 3 rearrangement on complex nature of the mem‐ brane rheological response, it is necessary to consider three subpopulations of band 3 molecules under isotonic and hypotonic conditions. The first subpopulation (20–40%) as tetramers forms high affinity complexes with ankyrin (quantified by the dissociation con‐ stant ∼5 nM) as reported by Tomishige et al. [23] and Kodippili et al. [24]. Band 3‐ankyrin complexes are located near the center of spectrin tetramers. These complexes could survive the hypotonic conditions. Golan and Veatch [25] reported that 25% of band 3 population

successive open‐closed state changes *trelease* ∈ [1,5 *s*] [11, 13].

902 Cytoskeleton - Structure, Dynamics, Function and Disease Cytoskeleton

The band 3 molecules within the freely diffusing subpopulation could form low affinity complexes to spectrin (the dissociation constant is ∼1–10 μM) [25]. On that base, they influence spectrin conformations [32]. Gov [32] reported that the parts of the spectrin filament be‐ tween two mid‐point attachments behave as independent blobs. The main factor which influences the spectrin flexibility and conformations is *<sup>l</sup> <sup>L</sup> <sup>p</sup>* (where *<sup>l</sup>* is the length of the filament part between two mid-point attachments and *L <sup>p</sup>* is the spectrin persistence length *<sup>L</sup> <sup>p</sup>* =15−<sup>25</sup> *nm* [33]). The spectrin filament parts are as follows: (1) flexible if *<sup>l</sup> <sup>L</sup> <sup>p</sup>* <sup>≻</sup> ≻1, (2) semiflexible if *<sup>l</sup> <sup>L</sup> <sup>p</sup>* ≈1, and (3) rod‐like if *<sup>l</sup> <sup>L</sup> <sup>p</sup>* <sup>≺</sup> ≺1. The average length of spectrin parts is equal to *<sup>l</sup>* <sup>=</sup> *<sup>L</sup> <sup>c</sup> NB* (where *L <sup>c</sup>* ≈200nm is the spectrin contour length [27] and *NB* ≈4−10 is the average number of attached band 3 molecules per single spectrin filament [34]). The length of the spectrin parts depends on the rearrangement of band 3 molecules which include their lateral diffusion and self‐associative tendency [28]. Consequently, rheological behavior of the cortex is related to the spectrin flexibility distribution and the rate of its changes [35, 36]. Deeper insight into coarse‐consequence relations between the band 3 rearrangement and the spec‐ trin inter‐ and intrachain interactions as well as the bilayer bending offers a possibility for understanding the complex nature of the membrane relaxation phenomena.

### **2. Rearrangement of band 3 molecules‐cluster packing state changes**

Packing state of band 3 clusters and its changes during short‐time lateral motion under isotonic and hypotonic conditions could be estimated by applying Edwards' statistics [30, 35, 37, 38]. Short‐time motion of band 3 molecules includes: (1) Brownian diffusion within the mesh compartment of the spectrin‐actin cortex and (2) hop diffusion between two compartments. Hop is observed at every 350 ms [23]. Band 3 molecules form clusters caused by positive hydrophobic mismatch effects during their lateral diffusion. This statistical approach is suitable for describing the cluster packing state changes under vibration field. Edwards introduced a new and very significant parameter for description of particle clusters named compactivity of the cluster part *X* representing the change of cluster part volume *Vc <sup>r</sup>* with entropy *Sc <sup>r</sup>* for given number of molecules *Np* (i.e., considering it as a canonical ensemble):

$$X = \left(\frac{\partial V\_{cr}}{\partial S\_{cr}}\right)\_{N\_P} \tag{1}$$

The two limits of the compactivity have been introduced: (1) *X* →0 corresponds to the most compact particle rearrangement and (2) *X* →*∞* corresponds to the least compact particle rearrangement. When *X* →0, the system picks out one particular configuration as being most likely, and when *X* →*∞*, the system picks out all configurations as being equally likely. Probability distribution of band 3 cluster states could be expressed as:

$$P = e^{\left(Y\_r - W\_r\right) / \mathcal{A} / X} \tag{2}$$

where *Yr* is effective volume of the cluster part (corresponding to the free energy *F* in classical statistical mechanics) and *Wr* is the "volume function" corresponding to the Hamiltonian and *λ* is constant adjusting dimensions. Then, we can write:

$$Y\_r = V\_{cr} + X \frac{\partial Y\_r}{\partial X} = V\_{cr} - XS\_{cr} \tag{3}$$

Cluster is considered as canonical ensemble during short‐time rearrangement. The partition function relating *Yr* with the volume function of a cluster part *Wr* (*q*) can be written in the form:

$$Z\_p = e^{-\frac{Y\_r}{\lambda\_r X}} = \iint \dots \int \Omega\_n e^{-\frac{W\_r(q\_1, q\_2, \dots, q\_n)}{\lambda\_r X}} dq\_1 dq\_2 \dots dq\_n \tag{4}$$

where integration goes over all geometrical degrees of freedom (DOFs) *qi* of all molecules in the cluster, *Zp* =*Zp*(*r*, *teq*) is the partition function. The key step is the identification of an exact volume function which makes it possible to pinpoint the configuration phase space and evaluate its dimensionality. We consider weakly interacting systems. To be precise, the interactions must be sufficient to lead to thermodynamical equilibrium, but weak enough that these interactions have negligible effects on the effective volume of individual molecules.

$$\boldsymbol{W}\_r(\boldsymbol{n}\_q) = \boldsymbol{N}\_{pr}\boldsymbol{w}\_p(\boldsymbol{q}\_1, \boldsymbol{q}\_2, \dots, \boldsymbol{q}\_n) \tag{5}$$

where is the volume function of single molecule. Rearrangement of rigid molecules such as band 3 within the cluster could be described using low degrees of freedom (DOFs). For this case, the DOFs describe molecule's orientation in the cluster part located at *r* and its coordination number.

**The band 3 cluster excitation under isotonic condition** induces the molecule orientation changes presented in **Figure 1**. The excitation occurs during alternating expansion and compression of the membrane parts in order to ensure surface and volume conservation.

The corresponding volume function could be expressed as [30, 35]:

**2. Rearrangement of band 3 molecules‐cluster packing state changes**

924 Cytoskeleton - Structure, Dynamics, Function and Disease Cytoskeleton

Packing state of band 3 clusters and its changes during short‐time lateral motion under isotonic and hypotonic conditions could be estimated by applying Edwards' statistics [30, 35, 37, 38]. Short‐time motion of band 3 molecules includes: (1) Brownian diffusion within the mesh compartment of the spectrin‐actin cortex and (2) hop diffusion between two compartments. Hop is observed at every 350 ms [23]. Band 3 molecules form clusters caused by positive hydrophobic mismatch effects during their lateral diffusion. This statistical approach is suitable for describing the cluster packing state changes under vibration field. Edwards introduced a new and very significant parameter for description of particle clusters named compactivity of the cluster part *X* representing the change of cluster part volume *Vc <sup>r</sup>* with entropy *Sc <sup>r</sup>* for given number of molecules *Np* (i.e., considering it as a canonical ensemble):

*P*

The two limits of the compactivity have been introduced: (1) *X* →0 corresponds to the most compact particle rearrangement and (2) *X* →*∞* corresponds to the least compact particle rearrangement. When *X* →0, the system picks out one particular configuration as being most likely, and when *X* →*∞*, the system picks out all configurations as being equally likely.

( )/ *YW X r r P e* -

¶ =+ =- ¶ *<sup>r</sup> r cr cr cr <sup>Y</sup> Y V X V XS*

*Yr W qq q r n X X Z e pw n <sup>e</sup> dq dq dq*


l

where *Yr* is effective volume of the cluster part (corresponding to the free energy *F* in classical statistical mechanics) and *Wr* is the "volume function" corresponding to the Hamiltonian and

Cluster is considered as canonical ensemble during short‐time rearrangement. The partition

( 1 2 , ,... ) 1 2 ... ...

= = W òò ò (4)

 l

¶ è ø (1)

= (2)

*<sup>X</sup>* (3)

(*q*) can be written in the form:

*cr cr N*

*<sup>V</sup> <sup>X</sup> S* æ ö ¶ <sup>=</sup> ç ÷

Probability distribution of band 3 cluster states could be expressed as:

*λ* is constant adjusting dimensions. Then, we can write:

function relating *Yr* with the volume function of a cluster part *Wr*

l

$$
\Delta \mathbf{v}\_p \left( q\_o, q\_c \right) = \mathbf{v}\_0 + \Delta \mathbf{v}\_o q\_o^2 \tag{6}
$$

**Figure 1.** Schematic representation of the packing state changes for excited band 3 cluster under isotonic and hypoton‐ ic conditions.

where *v*<sup>0</sup> is the minimum specific volume of single molecule of band 3 *rs* min is the minimum of the molecule Stokes radius, is the volume increment caused by the molecule orientation equal to is the maximum specific volume of single molecule of band 3 *v*max <sup>=</sup> <sup>4</sup> <sup>3</sup> *rs* max3*π*, *rs* max is the maximum of the molecule Stokes radius, *qo* accounts the molecule's orientation equal to *qo* <sup>2</sup> <sup>=</sup> <sup>1</sup> <sup>2</sup> ∑ *i qo i* 2 while *qo <sup>i</sup>* accounts various directions, *Δvo* is the volume increment contributions caused by the DOFs *qo* changes. For molecule loose packing, we can take and a molecule has high values of the degrees of freedom in such system so *qi* values are close to 1.

**The band 3 cluster excitation under hypotonic conditions** is more intensive than that obtained under isotonic condition due to changes the bilayer bending state during erythrocyte swelling. The excitation could induce changes the packing state from close packing to ring‐like structure which represents the reversible hemolytic hole. These changes are influenced by the positive hydrophobic mismatch effects which could induce protein tilting [20]. Zade‐Oppen [13] reported that the average hole opening time period is 270 ms, while the average hole closing time period is 260 ms. Seeman et al. [8] experimentally determined the diameter of the reversible osmotic holes in the range between 10 and 100 nm for human erythrocyte under hypotonic condition at pH = 7. Accordingly, the smaller hemolytic hole corresponds to ∼4 band 3 molecules while the larger one corresponds to ∼40 molecules [18, 30]. The result points out that cluster size is not the main factor for the hole formation. The main factor could be the hydrophobic mismatch effects between band 3 and the surrounding lipid bilayer as was shown in **Figure 1**. The corresponding volume function is as follows:

$$
\Delta \mathbf{w}\_p \left( q\_o, q\_c \right) = \mathbf{v}\_0 + \Delta \mathbf{v}\_o q\_o^2 + \Delta \mathbf{v}\_c q\_c^2 \tag{7}
$$

where *qc* accounts the coordination number, is the volume increment caused by changes

the molecule coordination number equal to *tR* is the relaxation time, *h <sup>m</sup>* is the thickness of the lipid bilayer for already swollen erythrocyte, *RH* (*tR*) is the radius of hemolytic hole, *Nr*(*tR*) is the number of molecules per cluster located at *r*. DOFs can have values in the range 0 to 1. When *qc* =0 and *qoi* =0 at *tR* →*tR eq*, the volume function is *wp* =*v*0. When *qc* =1 and *qoi* =1 at *tR* =0, the volume function is equal to . This excited cluster state represents the hemolytic hole.

Band 3 molecules change their states during migration. Consequently, for estimating the DOFs temporal changes it is necessary to consider the temporal changes of single molecule velocity. Temporal changes of DOFs under vibration field could be described in the form of Langevin‐ type equations [30, 37]:

where *v*<sup>0</sup> is the minimum specific volume of single molecule of band 3 *rs* min is the minimum of the molecule Stokes radius, is the volume increment caused by the molecule orientation equal to is the maximum specific volume of single

> <sup>2</sup> <sup>=</sup> <sup>1</sup> <sup>2</sup> ∑ *i qo i*

*Δvo* is the volume increment contributions caused by the DOFs *qo* changes. For molecule loose packing, we can take and a molecule has high values of the degrees of freedom

**The band 3 cluster excitation under hypotonic conditions** is more intensive than that obtained under isotonic condition due to changes the bilayer bending state during erythrocyte swelling. The excitation could induce changes the packing state from close packing to ring‐like structure which represents the reversible hemolytic hole. These changes are influenced by the positive hydrophobic mismatch effects which could induce protein tilting [20]. Zade‐Oppen [13] reported that the average hole opening time period is 270 ms, while the average hole closing time period is 260 ms. Seeman et al. [8] experimentally determined the diameter of the reversible osmotic holes in the range between 10 and 100 nm for human erythrocyte under hypotonic condition at pH = 7. Accordingly, the smaller hemolytic hole corresponds to ∼4 band 3 molecules while the larger one corresponds to ∼40 molecules [18, 30]. The result points out that cluster size is not the main factor for the hole formation. The main factor could be the hydrophobic mismatch effects between band 3 and the surrounding lipid bilayer as was shown

( ) 2 2

where *qc* accounts the coordination number, is the volume increment caused by changes

the molecule coordination number equal to *tR* is the relaxation time, *h <sup>m</sup>* is the thickness of the lipid bilayer for already swollen erythrocyte, *RH* (*tR*) is the radius of hemolytic hole, *Nr*(*tR*) is the number of molecules per cluster located at *r*. DOFs can have values in the range 0 to 1. When *qc* =0 and *qoi* =0 at *tR* →*tR eq*, the volume function is *wp* =*v*0. When *qc* =1 and *qoi* =1 at *tR* =0, the volume function is equal to . This excited cluster state

Band 3 molecules change their states during migration. Consequently, for estimating the DOFs temporal changes it is necessary to consider the temporal changes of single molecule velocity.

<sup>3</sup> *rs* max3*π*, *rs* max is the maximum of the molecule Stokes radius, *qo*

2 while *qo <sup>i</sup>*

<sup>0</sup> , *w q q v vq vq p o c oo cc* = +D +D (7)

accounts various directions,

molecule of band 3 *v*max <sup>=</sup> <sup>4</sup>

in such system so *qi*

represents the hemolytic hole.

accounts the molecule's orientation equal to *qo*

946 Cytoskeleton - Structure, Dynamics, Function and Disease Cytoskeleton

values are close to 1.

in **Figure 1**. The corresponding volume function is as follows:

$$\frac{dq\_l\left(t\_R\right)}{dt\_R} = -\frac{1}{\gamma\_q} \frac{\partial \mathbf{w}\_p\left(q\_l\left(t\_R\right)\right)}{\partial q\_l\left(t\_R\right)} + \phi\_q\left(t\_R\right) \tag{8}$$

where the stochastic random force *ϕq*(*tR*) is formulated as white noise with correlation function *ϕq*(*tR*)*ϕq*(*tR* ') =2*λX γqδij δ*(*tR* −*tR* '), and *γq* is the analog of frictional resistance. System structural changes could induce anomalous nature of energy dissipation derivative during molecules migration within the cluster. If the molecule migration causes damping effects as described by Tomishige et al. [23] (subdiffusion phenomenon), the derivative *d qi* (*tR*) *<sup>d</sup> tR* could be replaced by the fractional derivative *Dt <sup>γ</sup>* (where *Dt <sup>γ</sup>* is the Caputo's fractional derivative, *γ* is the order of the fractional derivative such that *γ* ≺1). Caputo's definition of the fractional derivative of some function *f* (*t*) is given as follows [39]: *Dt <sup>γ</sup>*( *<sup>f</sup>* (*t*))= <sup>1</sup> *Γ*(1 − *γ*) *d dt ∫* 0 *t f* (*t* ') (1) (*<sup>t</sup>* <sup>−</sup>*<sup>t</sup>* ')*<sup>γ</sup> dt* ' (where *Γ*(1−*γ*) is the gamma

function). Average equilibrium volume of single molecule is obtained as:

$$\nu\_{p\text{ }eq} = \frac{\iint \dots \int \nu\_{pr}(\bullet) \, e^{-\frac{\mathcal{W}\_{pr}(\bullet)}{\hat{\lambda} \, X}} d[\bullet]}{Z\_p} \tag{9}$$

where *Zp*(*r*, *t*Re*<sup>q</sup>*) is the partition function, and is the cluster volume.

### **3. Band 3 rearrangement influences the spectrin inter‐ and intrachain interactions and the bilayer bending**

The spectrin interchain interactions depend on the number of band 3 molecules attached per single spectrin filament as was shown in **Figure 2**.

**Figure 2.** Spectrin conformational changes influenced by the number of attached band 3 molecules.

The spectrin filaments have been treated as flexible ( *<sup>L</sup> <sup>c</sup> <sup>L</sup> <sup>p</sup>* <sup>≻</sup> <sup>≻</sup>1) [27] and semiflexible ( *<sup>L</sup> <sup>c</sup> <sup>L</sup> <sup>p</sup>* <sup>≈</sup>1) [40] (where *<sup>L</sup> <sup>c</sup>* is the spectrin contour length equal to *<sup>L</sup> <sup>c</sup>* <sup>=</sup> ∑ *i*=1 *NB*−1 *l i* , *l <sup>i</sup>* is the length of i‐th filament part between two mid-point attachments of band 3). The flexibility depends on the number of band 3 molecules attached per single spectrin filament. Spectrin filament without band 3‐ spectrin complexes behaves as flexible. Its conformations have been described as [27]:

$$F\_s \approx N \mu\_s \left( R - \left\langle r\_{\mathcal{S}}^2 \right\rangle^{1/2} \right) \tag{10}$$

Where *N* ≈3 is the number of spectrin filaments per network units, *μs* is the surface shear modulus of the cortex equal to *μs* <sup>=</sup> *kBT rg* <sup>2</sup> , *kB* is Boltzmann constant and *T* is temperature, *R* is the end‐to‐end distance of spectrin filaments in the cortex, *rg* 2 1/2 is the average filaments radius of gyration. If the band 3‐spectrin low affinity complexes exist, the parts of the spectrin filament between the complexes behave as independent blobs [32]. The conformation changes within the blobs are the milliseconds order [32]. When *<sup>l</sup> <sup>L</sup> <sup>p</sup>* <sup>≻</sup> <sup>≻</sup>1 the filament parts are flexible, but if *l <sup>L</sup> <sup>p</sup>* <sup>≈</sup><sup>1</sup> they become semiflexible, while if *<sup>l</sup> <sup>L</sup> <sup>p</sup>* <sup>≺</sup> <sup>≺</sup><sup>1</sup> they behave as rod‐like polymers (where *<sup>l</sup>*

is the average length of the filament part) [41]. Li et al. [40] treated the whole spectrin filaments as a semiflexible and proposed worm‐like force for modeling of the spectrin conformations:

$$F\_{wlc} = \frac{k\_B T}{L\_p} \left\{ \frac{1}{4\left(1 - \chi\_r\right)^2} - \frac{1}{4} + \chi\_r \right\} \tag{11}$$

where *<sup>x</sup>* <sup>=</sup> *<sup>R</sup>* <sup>−</sup> *rg* 2 1/2 *<sup>L</sup> <sup>c</sup>* is the stretch ratio. The worm‐like force corresponds to the condition *L <sup>c</sup> <sup>L</sup> <sup>p</sup>* ≈1 [41]. It is in accordance with the fact that the spectrin‐band 3 complexes lead to decrease in the spectrin flexibility. It could be quantified by apparent increase in the spectrin persistence length *L <sup>p</sup>* → *L <sup>p</sup> eff* (where *L <sup>p</sup> eff* is the effective spectrin persistence length). The concept of effective persistence length has been introduced for describing the nature of interchain structural changes for worm‐like chains such as proteins under stretching [42, 43]. On that base, the effective persistence length in our case could be expressed as [36]:

$$L\_{p\text{ eff}}\left(T, N\_B\right) = L\_p\left(T\right) + \Delta L\_p\left(N\_B\right) \tag{12}$$

Where *L <sup>p</sup> eff* (*T* , *NB*) is the effective persistence length of spectrin filament, *L <sup>p</sup>*(*T* ) is the spectrin persistence length for the filaments without mid‐point attachments at the same temperature conditions and *ΔL <sup>p</sup>*(*NB*) is the contribution to the persistence length caused by the band 3 midpoint attachments. The collective phenomena among variously flexible spectrin filaments induce generation of the cortex in‐homogeneities [36]. The in‐homogeneities in the context of the cortex micro domains influence the cortex relaxation. The cortex relaxation modulus *GC*(*tR*) could be expressed as [18, 36]:

**Figure 2.** Spectrin conformational changes influenced by the number of attached band 3 molecules.

part between two mid-point attachments of band 3). The flexibility depends on the number of band 3 molecules attached per single spectrin filament. Spectrin filament without band 3‐

spectrin complexes behaves as flexible. Its conformations have been described as [27]:

1/2 <sup>2</sup> *FN Rr ss g* m

*rg*

the end‐to‐end distance of spectrin filaments in the cortex, *rg*

the blobs are the milliseconds order [32]. When *<sup>l</sup>*

*<sup>L</sup> <sup>p</sup>* <sup>≈</sup><sup>1</sup> they become semiflexible, while if *<sup>l</sup>*

æ ö » - ç ÷

Where *N* ≈3 is the number of spectrin filaments per network units, *μs* is the surface shear

of gyration. If the band 3‐spectrin low affinity complexes exist, the parts of the spectrin filament between the complexes behave as independent blobs [32]. The conformation changes within

*<sup>L</sup> <sup>p</sup>* <sup>≻</sup> <sup>≻</sup>1) [27] and semiflexible ( *<sup>L</sup> <sup>c</sup>*

è ø (10)

<sup>2</sup> , *kB* is Boltzmann constant and *T* is temperature, *R* is

*<sup>L</sup> <sup>p</sup>* <sup>≻</sup> <sup>≻</sup>1 the filament parts are flexible, but if

*<sup>L</sup> <sup>p</sup>* <sup>≺</sup> <sup>≺</sup><sup>1</sup> they behave as rod‐like polymers (where *<sup>l</sup>*

2 1/2

*<sup>i</sup>* is the length of i‐th filament

is the average filaments radius

*i*=1 *NB*−1 *l i* , *l* *<sup>L</sup> <sup>p</sup>* <sup>≈</sup>1)

The spectrin filaments have been treated as flexible ( *<sup>L</sup> <sup>c</sup>*

968 Cytoskeleton - Structure, Dynamics, Function and Disease

Cytoskeleton

modulus of the cortex equal to *μs* <sup>=</sup> *kBT*

*l*

[40] (where *<sup>L</sup> <sup>c</sup>* is the spectrin contour length equal to *<sup>L</sup> <sup>c</sup>* <sup>=</sup> ∑

$$\mathcal{G}\_C\left(t\_R\right) = \int\_{L\_{p\text{ min}}}^{L\_{p\text{ max}}} \rho\_C\left(L\_{p\text{ eff}}, t\_R\right) \, G\_C\left(L\_{p\text{ eff}}, t\_R\right) \, dL\_{p\text{ eff}} \tag{13}$$

where *GC*(*L <sup>p</sup> eff* , *tR*) is the cortex relaxation modulus within the domain and *ρC*(*L <sup>p</sup> eff* , *tR*) is the spectrin flexibility distribution caused by the band 3 rearrangement.

The presence of the cortex micro domains is related to in‐homogeneous distribution of: (1) band 3 molecules, (2) spectrin flexibility, and (3) the presence of the cortex defects as was shown in **Figure 3**. Cumulative effects as: (1) the spectrin intrachain interactions which lead to formation of the cortex micro domains, (2) longtime diffusion of band 3 molecules, and (3) longtime bending relaxation of the lipid bilayer are at the order of seconds [18].

**Figure 3.** The cortex micro domains—schematic representation.

These local in‐homogeneities of the cortex are caused by alternating expansion and compres‐ sion of the membrane and the cortex‐bilayer coupling. The bilayer bending is influenced by conformational changes of the two types of spectrin filaments [44]: (1) type 1—corresponds to the filaments grafted at one end or at both ends but not connected to the stretched cortex and (2) type 2—corresponds to the filaments grafted at both ends and on that base represents a part of the connected stretched cortex. Filaments within the type 1 induce a concave curvature of radius *RL* <sup>1</sup>, while the type 2 induce a concave curvature of radius *RL* <sup>2</sup> such that *RL* <sup>1</sup> = −*RL* <sup>2</sup>. The bilayer‐cortex coupling has been expressed by Helfrich‐type bending‐free energy functional [44]:

$$E\left(n\_{\rm l}, n\_{\rm 2}\right) = \frac{1}{2}\kappa\nu\left[\left(H - \overline{H}\_{\rm l}n\_{\rm l}\left(r, s, t\_{R}\right) - \overline{H}\_{\rm 2}n\_{\rm 2}\left(r, s, t\_{R}\right)\right)^{2}d^{2}s\right.\tag{14}$$

Where *s* is the coordinate along the contour, *κ* is the bending modulus of the bilayer, *w* is the bilayer width, *n*1(*r*, *s*, *tR*) and *n*2(*r*, *s*, *tR*) are the relative densities of the types 1 and 2 of spectrin filaments, and the corresponding local mean curvature are *H*¯ <sup>1</sup> <sup>=</sup> <sup>1</sup> *RL* <sup>1</sup> and *H*¯ <sup>2</sup> <sup>=</sup> <sup>1</sup> *RL* <sup>2</sup> . The overall curvature by spectrin filaments is expressed as *H*¯ <sup>1</sup>*n*<sup>1</sup> <sup>+</sup> *<sup>H</sup>*¯ <sup>2</sup>*n*2. Band 3 molecules influence the lipid bilayer bending modulus. Shlomovitz and Gov [45] formulated the apparent bending modulus equal to:

Role of Band 3 in the Erythrocyte Membrane Structural Changes Under Isotonic and Hypotonic Conditions Role of Band 3 in the Erythrocyte Membrane Structural Changes Under Isotonic and Hypotonic Conditions 11 http://dx.doi.org/10.5772/64964 99

$$
\kappa\_{app}\left(\varphi\right) = \kappa\left(1 - \varphi\left(r, t\_R\right)\right) + \kappa'\varphi\left(r, t\_R\right) \tag{15}
$$

where *κapp*(*φ*) is the apparent bending modulus, *κ* is the bending modulus of the bilayer without band 3 molecules, *κ* ' is the contribution of band 3 molecules to the bending modulus, *φ*(*r*, *t*) is the local surface fraction of the band 3 molecules. Shlomovitz and Gov [45] expressed the influence of inclusions (band 3 molecules) on the lipid bilayer bending by formulating the free energy functional as:

$$E\left(\phi\right) = \int \frac{1}{2} \kappa\_{app}\left(\phi\right) \left(H - \phi \overline{H}\right)^2 d^2r\tag{16}$$

Consequently, the model Eqs. (15) and (16) could be combined to describe the influence of: (1) spectrin conformationals and (2) band 3 molecules migration on the lipid bilayer bending state

expressed in the form: . The collective phenomena related to the spectrin filaments migration is expressed by spatial‐ temporal changes of the conservative variable *n* =*n*(*r*, *s*, *tR*) [46] as:

**Figure 3.** The cortex micro domains—schematic representation.

9810 Cytoskeleton - Structure, Dynamics, Function and Disease

Cytoskeleton

energy functional [44]:

modulus equal to:

These local in‐homogeneities of the cortex are caused by alternating expansion and compres‐ sion of the membrane and the cortex‐bilayer coupling. The bilayer bending is influenced by conformational changes of the two types of spectrin filaments [44]: (1) type 1—corresponds to the filaments grafted at one end or at both ends but not connected to the stretched cortex and (2) type 2—corresponds to the filaments grafted at both ends and on that base represents a part of the connected stretched cortex. Filaments within the type 1 induce a concave curvature of radius *RL* <sup>1</sup>, while the type 2 induce a concave curvature of radius *RL* <sup>2</sup> such that *RL* <sup>1</sup> = −*RL* <sup>2</sup>. The bilayer‐cortex coupling has been expressed by Helfrich‐type bending‐free

( ) ( ( ) ( ))

Where *s* is the coordinate along the contour, *κ* is the bending modulus of the bilayer, *w* is the bilayer width, *n*1(*r*, *s*, *tR*) and *n*2(*r*, *s*, *tR*) are the relative densities of the types 1 and 2 of spectrin

lipid bilayer bending modulus. Shlomovitz and Gov [45] formulated the apparent bending

<sup>1</sup>*n*<sup>1</sup> <sup>+</sup> *<sup>H</sup>*¯

1 2 1 1 2 2 <sup>1</sup> , ,, ,, <sup>2</sup> *E n n w H Hn rst H n rst d s* =- -

k

filaments, and the corresponding local mean curvature are *H*¯

curvature by spectrin filaments is expressed as *H*¯

<sup>2</sup> <sup>2</sup>

and *H*¯

<sup>2</sup> <sup>=</sup> <sup>1</sup> *RL* <sup>2</sup>

<sup>2</sup>*n*2. Band 3 molecules influence the

. The overall

*R R* ò (14)

<sup>1</sup> <sup>=</sup> <sup>1</sup> *RL* <sup>1</sup>

$$\frac{1}{\dot{s}} \frac{\partial \left(\dot{s} \ n\right)}{\partial t\_R} = \frac{D}{\dot{s}} \nabla\_s^2 \left(\dot{s} \ n\right) + \frac{1}{\dot{s}} \frac{\Lambda}{n\_{sat}} \nabla\_s \left(\dot{s} \ n \nabla\_s \left(\frac{1}{\dot{s}} \frac{\delta E}{\delta \ n}\right)\right) \tag{17}$$

where *<sup>D</sup>* <sup>=</sup> *kBT <sup>Λ</sup>* is the spectrin collective diffusion coefficient which accounted for the intrachain interactions, *Λ* is the filaments mobility parameter, *nsat* is the maximal packing density of the filaments, and ∇*<sup>s</sup>* is the derivative along the contour *s*. Spectrin filament mobility depends on the number of attached band 3 molecules. Consequently, the effective diffusivity could be formulated as *Deff* =*Deff* (*φ*)and introduced in eq. 17. Shlomovitz and Gov [45] modeled the collective migration of band 3 molecules as:

$$\frac{\partial \mathcal{O}}{\partial t\_R} = D\_B \nabla^2 \mathcal{O} + \Lambda\_B \nabla \left( \mathcal{g} \nabla \left( \frac{\delta E}{\delta \mathcal{J} \mathcal{g}} \right) \right) \tag{18}$$

where ∇ is the derivative along the space, *Λ<sup>B</sup>* is the band 3 lateral mobility and *DB* is the band 3 diffusion coefficient equal to *DB* <sup>=</sup> *kBT <sup>Λ</sup><sup>B</sup>* . Lateral motion of the band 3 molecules induces the anomalous nature of energy dissipation which includes damping effects [23]. These damping effects are induced from band 3 association‐dissociation to spectrin filaments. Pajic‐Lijakovic [35, 36] proposed fractional Langevin equation for describing the lateral diffusion by applying the fractional derivatives [39]. Consequently, the time derivatives from Eqs. (17) to (18) could be replaced by the fractional derivative *Dt <sup>α</sup>*(•).

### **4. Conclusion**

Rheological behavior of the cortex depends on the spectrin flexibility distribution and the rate of its changes [35, 36]. The spectrin flexibility primarily depends on the number of band 3 molecules attached per single spectrin filaments. Rearrangement of the band 3 molecules and their lateral diffusion also influence the bending modulus of the lipid bilayer and the band 3‐ bilayer interaction energy. Consequently, the band 3 rearrangement influences the cortex‐ bilayer coupling and on that base influences the membrane rheological behavior as a whole. The membrane structural changes induce anomalous nature of energy dissipation caused by these complex multi scale molecular dynamics.

### **Acknowledgements**

This research was funded by grant III46001 from the Ministry of Science and Environmental Protection, Republic of Serbia.

### **Author details**

Ivana Pajic‐Lijakovic\* and Milan Milivojevic

\*Address all correspondence to: iva@tmf.bg.ac.rs

Faculty of Technology and Metallurgy, Belgrade University, Belgrade, Serbia

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the fractional derivatives [39]. Consequently, the time derivatives from Eqs. (17) to (18) could

Rheological behavior of the cortex depends on the spectrin flexibility distribution and the rate of its changes [35, 36]. The spectrin flexibility primarily depends on the number of band 3 molecules attached per single spectrin filaments. Rearrangement of the band 3 molecules and their lateral diffusion also influence the bending modulus of the lipid bilayer and the band 3‐ bilayer interaction energy. Consequently, the band 3 rearrangement influences the cortex‐ bilayer coupling and on that base influences the membrane rheological behavior as a whole. The membrane structural changes induce anomalous nature of energy dissipation caused by

This research was funded by grant III46001 from the Ministry of Science and Environmental

[1] Amin MS, Park YK, Lue N, Dasari RR, Badizadegan K, Feld MS, Popescu G. Micro‐ rheology of red blood cell membrane using dynamics scattering microscopy. Opt

[2] Popescu G, Park YK, Dasari RR, Badizadegan K, Feld MS. Coherence Properties of red

[3] Puig‐de‐Morales‐Marinkovic M, Turner KT, Butler JP, Fredberg JJ, Suresh S. Viscoe‐ lasticity of the human red blood cell. Am J Physiol Cell Physiol. 2007; 293:C597–C605.

*<sup>α</sup>*(•).

be replaced by the fractional derivative *Dt*

10012 Cytoskeleton - Structure, Dynamics, Function and Disease Cytoskeleton

these complex multi scale molecular dynamics.

and Milan Milivojevic

Faculty of Technology and Metallurgy, Belgrade University, Belgrade, Serbia

blood cell membrane motion. Phys Rev E. 2007; 76:031902 1–5.

\*Address all correspondence to: iva@tmf.bg.ac.rs

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**4. Conclusion**

**Acknowledgements**

**Author details**

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Ivana Pajic‐Lijakovic\*

Protection, Republic of Serbia.


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## **Dystrophin–Glycoprotein Complex in Blood Cells**

### Doris Cerecedo

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/66857

#### **Abstract**

The Dystrophin-Associated Protein Complex (DAPC), known as the Dystrophin– Glycoprotein Complex (DGC), comprises an array of glycoproteins that are essential for the normal function of striated muscle, in which they were first described, and for many other tissues, including blood. Understanding the role that these molecules play in muscle function has increased over the last decade, and some of the knowledge derived can be applied to other biological systems. However, there is no doubt that to date, some progress has been achieved in blood cells.

Multiple interactions have been described among the proteins comprising the DGC, it is now well established that the DGC possesses a crucial role for numerous signaling pathways, recruiting and regulating various signaling proteins into a macromolecular complex. The aim of this chapter is to summarize the current state of knowledge regarding DGC processing and assembly, mainly in muscle tissue and in blood cells, with a primary focus on the dystroglycan heterodimer and associated proteins, including ion channels and membrane lipids. In addition, and due to increasing evidence involving dystroglycan proteins in the pathophysiology of solid tissue cancer, Duchenne muscular dystrophy, and leukemia, current information on these topics will be included.

**Keywords:** DGC, dystroglycan, intermediate filaments, leukemia cells, adhered platelets

### **1. Introduction**

Dystrophin-associated glycoprotein complex, known as the DGC, is a multimeric and multifaceted protein complex located in the plasma membrane and mediates interactions among the cytoskeleton, cell membrane, and extracellular matrix (ECM) of the muscle and nonmuscle tissues. Therefore, the DGC is involved in signaling pathways that regulate the structural organization of specialized membrane-contact zones, and on the basis of its different biochemical characteristics and localization, the DGC can be divided into the following three

© 2017 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

subcomplexes: the dystroglycan (Dg), the sarcoglycan, and the cytoplasmic, dystrophin-containing complex.

The dystroglycan subcomplex comprises α- and β-dystroglycan. α-Dystroglycan is the extracellular component that binds to α-laminin and to other components of the basal lamina (ECM), while β-dystroglycan is the transmembrane component. Both attach the intracellular cytoskeleton to the ECM, a task that is widespread in all human tissues and cells [1].

The sarcoglycan subcomplex is a multimember complex that, in association with dystroglycan, stabilizes interactions with the extracellular and transmembrane components of the DGC, as well as with dystrophin and its associated proteins. To date, six sarcoglycan genes have been identified that give rise to their respective proteins *α-, β-, γ-, δ-, ε-*, and *ζ-*sarcoglycan, which are organized in a tetrameric arrangement; however, it has been hypothesized that the six sarcoglycans can be arranged in an exameric structure [2].

The dystrophin subcomplex can form a mechanically strong bond with any costameric protein, forming a mechanically strong link between the sarcolemma and the costameric cytoskeleton through interaction with *γ*-actin filaments. Additionally, based on its structure, protein interactions, and the membrane defects associated with its absence or abnormality in dystrophic muscle, the dystrophin complex provides mechanical stabilization of the sarcolemmal membrane against the stresses imposed upon it during muscle contraction or stretch [3].

Therefore, the DGC appears to play both mechanical and nonmechanical roles in skeletal muscle and in nonmuscle cells, although neither the DGC structure nor the functions are completely understood at present.

This chapter focuses on recent insights into the specific roles of the DGC in different tissue cells, including blood cells, with special focus on dystroglycan biology and its feasible pathophysiologic implications in human leukemia cells and dystrophies.

### **2. Dystrophin–glycoprotein complex**

Dystrophin is the protein that plays a central role in trans-sarcolemmal linkage between the basement membrane and the intracellular actin cytoskeleton, and is the product of the largest identified gene in the human genome [4].

The complexity of Duchenne muscular dystrophy (*DMD*) gene expression, which results in multiple transcripts and protein isoforms, has hampered understanding of the functions of individual dystrophin protein isoforms. The transcription of human DMD is controlled by the following three independent promoters, brain (B), muscle (M), and Purkinje (P) promoters, which indicate the tissue distribution of dystrophin expression, as well as four internal promoters (R for retinal, B for brain, S for Schwann cells, and G for general), which give rise to shorter transcripts encoding for the truncated COOH-terminal isoforms formed from the alternative splicing that generates dystrophin isoforms of 260 kDa (Dp260), 140 kDa (Dp140) [5], 116 kDa (Dp116) [6], and 71 kDa (Dp71) [7, 8]. When these COOH-terminal dystrophin submembrane cytoskeletal proteins interact with a large macromolecular protein complex, they constitute the dystrophin-associated protein complex (DAPC). The crucial structural role of this complex is based on its strategic localization, spanning the plasma membrane and linking with the ECM and the actin cytoskeleton. Since the original discovery of the dystrophin-glycoprotein complex (DGC) [9], a large number of studies have characterized the various components involved in dystrophin [10]. Dystrophin-associated proteins can be divided into sarcolemmal proteins (β-dystroglycan, α-sarcoglycan, β-sarcoglycan, γ-sarcoglycan, and δ-sarcoglycan, sarcospan), cytosolic proteins (dystrobrevins, syntrophins, neuronal nitric-oxide synthase [nNOS]), and extracellular proteins (α-dystroglycan and laminin) [11]. Several DGC components are also found in two or more isoforms, which are either generated by alternative splicing of a single gene or originate from distinct genes [12, 13].

The large, multi-subunit DGC is found in the sarcolemma of striated muscle fibers, and this is essential for maintaining the structural integrity of these fibers during contraction; therefore, the generally accepted role for the DGC is its acting as a molecular shock absorber and stabilizing the plasma membrane during muscle contraction. However, its role goes beyond that solely of a passive scaffold among the elements of the complex, anchoring these near sitesof-action or important partners, since genetic disruption of any of the DGC elements causes mislocalization, destabilization, and the loss-of-function of the cell [14].

As evidence of DGC signaling capacity, it has been reported that nNOS is associated with the DGC via α-dystrobrevin, and that there is a loss of nNOS from the sarcolemma in Duchenne muscular dystrophy (DMD) [15]. Additionally, the DGC promotes the mechanical activation of cardiac nNOS by acting as a mechanosensor in the regulation of AMP-activated protein kinase AMPK activity [16].

The complex also constitutes a scaffold for signaling molecules based on its association with several signaling proteins, including Grb2-Sos1 [17], MEK and ERK [18], heterotrimeric G protein subunits [19], archvillin [20], and nNOS [21].

### **3. Dystrophin-related proteins**

subcomplexes: the dystroglycan (Dg), the sarcoglycan, and the cytoplasmic, dystrophin-con-

The dystroglycan subcomplex comprises α- and β-dystroglycan. α-Dystroglycan is the extracellular component that binds to α-laminin and to other components of the basal lamina (ECM), while β-dystroglycan is the transmembrane component. Both attach the intracellular

The sarcoglycan subcomplex is a multimember complex that, in association with dystroglycan, stabilizes interactions with the extracellular and transmembrane components of the DGC, as well as with dystrophin and its associated proteins. To date, six sarcoglycan genes have been identified that give rise to their respective proteins *α-, β-, γ-, δ-, ε-*, and *ζ-*sarcoglycan, which are organized in a tetrameric arrangement; however, it has been hypothesized that the

The dystrophin subcomplex can form a mechanically strong bond with any costameric protein, forming a mechanically strong link between the sarcolemma and the costameric cytoskeleton through interaction with *γ*-actin filaments. Additionally, based on its structure, protein interactions, and the membrane defects associated with its absence or abnormality in dystrophic muscle, the dystrophin complex provides mechanical stabilization of the sarcolemmal membrane against the stresses imposed upon it during muscle contraction or stretch [3].

Therefore, the DGC appears to play both mechanical and nonmechanical roles in skeletal muscle and in nonmuscle cells, although neither the DGC structure nor the functions are

This chapter focuses on recent insights into the specific roles of the DGC in different tissue cells, including blood cells, with special focus on dystroglycan biology and its feasible patho-

Dystrophin is the protein that plays a central role in trans-sarcolemmal linkage between the basement membrane and the intracellular actin cytoskeleton, and is the product of the largest

The complexity of Duchenne muscular dystrophy (*DMD*) gene expression, which results in multiple transcripts and protein isoforms, has hampered understanding of the functions of individual dystrophin protein isoforms. The transcription of human DMD is controlled by the following three independent promoters, brain (B), muscle (M), and Purkinje (P) promoters, which indicate the tissue distribution of dystrophin expression, as well as four internal promoters (R for retinal, B for brain, S for Schwann cells, and G for general), which give rise to shorter transcripts encoding for the truncated COOH-terminal isoforms formed from the alternative splicing that generates dystrophin isoforms of 260 kDa (Dp260), 140 kDa (Dp140) [5], 116 kDa (Dp116) [6], and 71 kDa (Dp71) [7, 8]. When

cytoskeleton to the ECM, a task that is widespread in all human tissues and cells [1].

six sarcoglycans can be arranged in an exameric structure [2].

physiologic implications in human leukemia cells and dystrophies.

completely understood at present.

**2. Dystrophin–glycoprotein complex**

identified gene in the human genome [4].

taining complex.

106 Cytoskeleton - Structure, Dynamics, Function and Disease

Dystrophins share structural homology with a range of paralog proteins denominated the dystrophin-related proteins (DRP), such as utrophin, DRP2, dystrobrevin, and dystrotelin [22].

The utrophin gene possesses internal promoters and shorter protein products and is also modulated by alternative splicing [23]. Transcription of full-length utrophin (Up395) is driven by two independent promoters: Utrn-A and Utrn-B. The Utrn-A protein is the main isoform in adult skeletal muscles, in contrast with Utrn-B, which is found in the vascular muscle endothelium [24].

G-utrophin, or Up113, was the first short product identified as a structural homolog of Dp116, while Up140 and Up71 are homologous to the short dystrophins Dp140 and Dp71, respectively; these short utrophins do not possess actin-binding sites in the N-terminal domain of the molecule [25]. Up71 is detected in nonmuscle tissues such as lung, kidney, thymus, liver, and brain, while Up140 is found in lung, muscle, kidney, thymus, liver, testes, and brain. Full-length utrophins are also detected in nonmuscle tissues, such ass those of the central nervous system (CNS), peripheral nerves, testes, kidney, spleen, liver, and lung, and in small arteries and veins [26, 27]. In 1995, utrophin was described as a component of the platelet cytoskeleton, participating in its reorganization [28], while in hematopoietic stem/progenitor cells, Up400 and Up140 comprised the main gene products [29]. In addition, Up71 has been described in platelets [30], as well as in neutrophils [31].

It has long been considered that utrophin and dystrophin share comparable functions during fetal development and adulthood, maintaining utrophin expression in adult dystrophic tissues, compensating for dystrophin loss, as has been observed in mdx skeletal and cardiac muscles [24, 32]. However, spontaneous upregulations also occur in nonmuscle tissues, such as in Dp71-deficient platelets [33] and, most importantly, in the brains of DMD mouse models [34]. However, their expression in distinct structures, as compared with dystrophin, may not reflect functional compensation [24].

### **4. Dual role of the Dp71 isoform**

Dp71 (70–75 kDa) is the first product of the *DMD* gene detectable in pluripotent embryonic stem cells (ESC) during development. It decreases in differentiated ESC cultures and tumors [35] and is the major dystrophin expressed in nonmuscle cells, such as neural tissue [36], glia [37], spermatozoa [38], and astrocytoma cells [39]; in platelets, its participation has been suggested in cytoskeletal reorganization and/or signaling, and in thrombin-mediated platelet adhesion [28].

The variation in the molecular mass of Dp71 transcripts is consistent with the expression of Dp71 isoforms derived from transcripts alternatively spliced for exons 71 and/or 78 [40]. The splicing product of exon 78 produces the isoform known as Dp71d, which preserves the C-terminal, while Dp71f is the product of the absence of exon 78. Two other gene products resulting from an alternative splicing at exons 71–74 and/or 78 transcripts, Dp71Δ110<sup>a</sup> and Dp71Δ110<sup>m</sup>, respectively, with a relative mass of 55 kDa, have been recently characterized [40, 41].

In 2005, Dp71d/Dp71Δ110<sup>m</sup>~DGC and Up400/Up71~DGC were described as participating with structural roles associated with the actin cytoskeleton in the formation of membrane scaffolds. They were probably involved in defining platelet shape, substrate adhesion, and granule migration, as well as possessing a signaling role, participating in signaling triggered by adhesion to glass and by interaction with agonists such as thrombin [30].

The presence of Dp71 and some DGC elements that form a nuclear complex at the plasma membrane and in the nucleus of muscle cells suggested their participation in nuclear structure and in the modulation of nuclear processes [42].

these short utrophins do not possess actin-binding sites in the N-terminal domain of the molecule [25]. Up71 is detected in nonmuscle tissues such as lung, kidney, thymus, liver, and brain, while Up140 is found in lung, muscle, kidney, thymus, liver, testes, and brain. Full-length utrophins are also detected in nonmuscle tissues, such ass those of the central nervous system (CNS), peripheral nerves, testes, kidney, spleen, liver, and lung, and in small arteries and veins [26, 27]. In 1995, utrophin was described as a component of the platelet cytoskeleton, participating in its reorganization [28], while in hematopoietic stem/progenitor cells, Up400 and Up140 comprised the main gene products [29]. In addition, Up71 has been described in platelets [30],

It has long been considered that utrophin and dystrophin share comparable functions during fetal development and adulthood, maintaining utrophin expression in adult dystrophic tissues, compensating for dystrophin loss, as has been observed in mdx skeletal and cardiac muscles [24, 32]. However, spontaneous upregulations also occur in nonmuscle tissues, such as in Dp71-deficient platelets [33] and, most importantly, in the brains of DMD mouse models [34]. However, their expression in distinct structures, as compared with dystrophin, may not

Dp71 (70–75 kDa) is the first product of the *DMD* gene detectable in pluripotent embryonic stem cells (ESC) during development. It decreases in differentiated ESC cultures and tumors [35] and is the major dystrophin expressed in nonmuscle cells, such as neural tissue [36], glia [37], spermatozoa [38], and astrocytoma cells [39]; in platelets, its participation has been suggested in cytoskeletal reorganization and/or signaling, and in thrombin-mediated platelet

The variation in the molecular mass of Dp71 transcripts is consistent with the expression of Dp71 isoforms derived from transcripts alternatively spliced for exons 71 and/or 78 [40]. The splicing product of exon 78 produces the isoform known as Dp71d, which preserves the C-terminal, while Dp71f is the product of the absence of exon 78. Two other gene products resulting from an alternative splicing at exons 71–74 and/or 78 transcripts, Dp71Δ110<sup>a</sup>

Dp71Δ110<sup>m</sup>, respectively, with a relative mass of 55 kDa, have been recently characterized

In 2005, Dp71d/Dp71Δ110<sup>m</sup>~DGC and Up400/Up71~DGC were described as participating with structural roles associated with the actin cytoskeleton in the formation of membrane scaffolds. They were probably involved in defining platelet shape, substrate adhesion, and granule migration, as well as possessing a signaling role, participating in signaling triggered

The presence of Dp71 and some DGC elements that form a nuclear complex at the plasma membrane and in the nucleus of muscle cells suggested their participation in nuclear struc-

by adhesion to glass and by interaction with agonists such as thrombin [30].

ture and in the modulation of nuclear processes [42].

and

as well as in neutrophils [31].

108 Cytoskeleton - Structure, Dynamics, Function and Disease

reflect functional compensation [24].

adhesion [28].

[40, 41].

**4. Dual role of the Dp71 isoform**

**Figure 1.** Schematic diagram of the dystrophin–glycoprotein complex (DGC) composed of Dp71 (left) and utrophin (right) in adhered platelets. Dystrophin is a linker between the cytoskeleton and the extracellular matrix (ECM). Dp71 and utrophin are associated with the dystroglycan complex and the dystrobrevin/syntrophin complex (α-Db/α-Syn). α-Dystroglycan (α-Dg) binds to ECM proteins and β-dystroglycan (β-Dg); β-Dg binds to the dystrophin, completing the link between the actin cytoskeleton and the ECM. Focal adhesion (magnified at the bottom of the figure) clusters the α- and β-integrin receptors and induces recruitment of focal adhesion proteins vinculin (Vin), talin (Tal), and α-actinin (α-Act), which connect directly with microfilaments and short dystrophins (Dp71) and indirectly with microtubules and intermediate filaments. The adhesion complex activates integrin-associated signaling cascades, including focal adhesion kinase (FAK). Dystroglycan plays a scaffold role, modulating the cytoplasmic protein kinases, and is in close association with integrin β1.

The neuronal cell line PC12 expresses at least two different Dp71 protein isoforms generated by the alternative splicing of exon 78 [43, 44]. The splicing isoform of Dp71 (Dp71d) contains 13C-terminal amino acids encoded by exon 78, which are replaced by 31 new amino acids encoded by exon 79 in the Dp71f isoform upon removal of exon 78 [40]. Depletion of total Dp71 protein levels gives rise to impairment in nerve growth factor (NGF)-induced neurite outgrowth [45] and in the cell adhesion activity of PC12 cells [46], indicating that Dp71 is required for these neuronal functions. Dp71f assembles an adhesion complex comprising talin, α-actinin, paxillin, focal adhesion kinase (FAK), and actin, but not vinculin, contributing to cell stability [47].

During the platelet adhesion process, short dystrophins (Dp71d/Dp71Δ110m) and utrophins (Up400/Up71) have demonstrated potential association with the integrin β-1 fraction and with focal adhesion system that includes α-actinin, vinculin, and talin. Apparently, in order to fulfill this hemosatic role, the coexistence of the DGC composed of short dystrophins or utrophins plays both a structural role in participation in stress-fiber assembly and in the centralization of cytoplasmic granules, and a regulatory role, incorporating FAK into the complex. The coexistence of dystrophin and utrophin complexes indicates structural and signaling mechanisms that are complementary to the actin network during the adhesion process [48] (**Figure 1**).

### **5. DGC components**

The findings described in systematic proteomic studies indicate that dystrophin interacts closely with core members of the dystrophin-associated glycoprotein complex, such as dystroglycans, sarcoglycans, syntrophins, dystrobrevins, and sarcospan, but that it also forms indirect linkages with a large variety of other protein species, including tubulin, vimentin, desmin, annexin, and collagens [49].

#### **5.1. Dystrobrevins**

Dystrobrevins are proteins among dystrophin-related proteins that are encoded by two different genes, *α* and *β* and that possess significant homology to dystrophin. *α-Dystrobrevin* is expressed predominantly in muscle and brain, whereas *β-dystrobrevin* is expressed in nonmuscle tissues, which is abundant in brain, kidney, lung, and liver. Dystrobrevins have also been involved in intracellular signaling in muscle and nonmuscle tissues, either directly or through interaction with syntrophin, another element of the DGC. In humans, Sadoulet-Puccio et al. [50] found six isoforms of dystrobrevin (designated α-, β-, γ-, δ-, ε-, and ζ-dystrobrevin), which ranged in size from 22 to 80 kDa.

Human α-dystrobrevin and its few isoforms are expressed in the cytosol and the nucleus of the promyelocytic HL-60 cell line. A distinct distribution pattern of α-dystrobrevin, including colocalization with actin, was described in HL-60 promyelocytes, differentiated mature granulocytes, and in human neutrophils, supporting a signaling role [51]. In adhered platelets, it was suggested that actin filaments and microtubules contribute to α-granule and dense granule mobilization in adhered platelets, identifying α-dystrobrevins as part of the platelet transport machinery that is closely associated with the ubiquitous kinesin heavy chain (UKHC), this system is depicted in **Figure 2** [52].

#### **5.2. Sarcoglycans**

The sarcoglycan complex (SGC) is composed of α-, β-, γ-, and δ-sarcoglycan isoforms encoded by separate genes, and of sarcospan. Sarcoglycans are single transmembrane glycoproteins with the N-terminus oriented extracellularly for α-sarcoglycan and intracellularly for β-,

**Figure 2.** Platelet distribution of cytoskeleton elements. Schematic diagram of actin filaments, microtubules, and intermediate filaments in adhered platelets. Plectin is the protein that acts as a link among the three main components of the cytoskeleton.

γ-, and δ-sarcoglycans [53]. Contrariwise, sarcospan is composed of four transmembranespanning segments that are homologous to the tetraspanin family. The function of the SGC is not fully understood, but it appears to strengthen the interaction of β-dystroglycan with α-dystroglycan and dystrophin, as well as to play a role in intracellular signal transduction for sarcoglycan [54]. Sarcospan, a 25-kDa transmembrane protein, improves the cell-surface expression of the three major laminin-binding complexes, i.e., the dystrophin– and utrophin– glycoprotein complexes, as well as of an α7β1 integrin [55].

#### **5.3. Syntrophins**

During the platelet adhesion process, short dystrophins (Dp71d/Dp71Δ110m) and utrophins (Up400/Up71) have demonstrated potential association with the integrin β-1 fraction and with focal adhesion system that includes α-actinin, vinculin, and talin. Apparently, in order to fulfill this hemosatic role, the coexistence of the DGC composed of short dystrophins or utrophins plays both a structural role in participation in stress-fiber assembly and in the centralization of cytoplasmic granules, and a regulatory role, incorporating FAK into the complex. The coexistence of dystrophin and utrophin complexes indicates structural and signaling mechanisms that are complementary to the actin network during the adhesion

The findings described in systematic proteomic studies indicate that dystrophin interacts closely with core members of the dystrophin-associated glycoprotein complex, such as dystroglycans, sarcoglycans, syntrophins, dystrobrevins, and sarcospan, but that it also forms indirect linkages with a large variety of other protein species, including tubulin, vimentin,

Dystrobrevins are proteins among dystrophin-related proteins that are encoded by two different genes, *α* and *β* and that possess significant homology to dystrophin. *α-Dystrobrevin* is expressed predominantly in muscle and brain, whereas *β-dystrobrevin* is expressed in nonmuscle tissues, which is abundant in brain, kidney, lung, and liver. Dystrobrevins have also been involved in intracellular signaling in muscle and nonmuscle tissues, either directly or through interaction with syntrophin, another element of the DGC. In humans, Sadoulet-Puccio et al. [50] found six isoforms of dystrobrevin (designated α-, β-, γ-, δ-, ε-, and ζ-dystrobrevin),

Human α-dystrobrevin and its few isoforms are expressed in the cytosol and the nucleus of the promyelocytic HL-60 cell line. A distinct distribution pattern of α-dystrobrevin, including colocalization with actin, was described in HL-60 promyelocytes, differentiated mature granulocytes, and in human neutrophils, supporting a signaling role [51]. In adhered platelets, it was suggested that actin filaments and microtubules contribute to α-granule and dense granule mobilization in adhered platelets, identifying α-dystrobrevins as part of the platelet transport machinery that is closely associated with the ubiquitous kinesin heavy chain

The sarcoglycan complex (SGC) is composed of α-, β-, γ-, and δ-sarcoglycan isoforms encoded by separate genes, and of sarcospan. Sarcoglycans are single transmembrane glycoproteins with the N-terminus oriented extracellularly for α-sarcoglycan and intracellularly for β-,

process [48] (**Figure 1**).

**5. DGC components**

**5.1. Dystrobrevins**

**5.2. Sarcoglycans**

desmin, annexin, and collagens [49].

110 Cytoskeleton - Structure, Dynamics, Function and Disease

which ranged in size from 22 to 80 kDa.

(UKHC), this system is depicted in **Figure 2** [52].

Syntrophins are a multigene family of intracellular membrane-associated adaptor proteins and consist of five homologous isoforms: α1-syntrophin, β1-syntrophin, β2-syntrophin, γ1-syntrophin, and γ2-syntrophin; they possess a different cellular and subcellular localization, suggesting a distinct functional role [56]. In human platelets, a 54-kDa band corresponding to α-syntrophin is well expressed [30].

The pleckstrin homology (PH) and PDZ domains of syntrophins were shown to bind various proteins, including nitric oxide synthase (NOS), and have been implicated in the regulation of various plasma membrane ion channels, such as voltage-operated sodium channels and other nonvoltage gated channels, such as mechanosensitive Na<sup>+</sup> channels [57].

#### **5.4. Dystrolglycans**

The single dystroglycan gene encodes for a precursor protein that undergoes posttranslational proteolytic cleavage, which in turn produces two noncovalant DGC subunits: α- and β-dystroglycan. α-Dystroglycan is a dumbbell-shaped protein that binds to the laminin G domain in ECM components such as laminins, agrin, and perlecan. β-Dystroglycan (β-Dg) possesses a single transmembrane domain spanning the plasma membrane and an extracellular amino-terminal extracellular domain binding to the carboxy-terminal globular domain of α-Dg [58].

#### *5.4.1. Dg involved in the signaling process*

The β-Dg dual role (structural and signaling) has been demonstrated in various cell types and tissues. Examples of the former role are represented by the participation of β-Dg in cytoskeleton remodeling, where it is associated with actin [59, 60], while its signaling role is represented by its association with the extracellular signal-related kinase-mitogen-activated protein (ERK-MAP) kinase cascade [18], or with integrins modulates myoblast anchorage and migration [61]; this latter process is critically regulated by Src-mediated phosphorylation of β-Dg at tyrosine 890 [62].

Grb2–β-Dg interaction could facilitate the transduction of signals between the DGC and extracellular proteins and other signaling pathways [62]. However, when Dg is localized at the tips of dynamic filopodia, it directs local Cdc42 activation and recruits the guanine nucleotide exchange factor (GEF) Dbl to generate actin protrusions [60].

Dystroglycan is also a multifunctional adaptor or scaffold capable of interacting with components of the ERK-MAP kinase cascade, including MEK and ERK [18]. However, it has been established that integrin α6Aβ1 and dystroglycan play antagonistic roles in signaling to the Ras-Raf-MEK-ERK pathway in response to laminin [63].

#### *5.4.2. Dg promoter of the adhesion process*

Since 1995, dystroglycan-associated proteins, such as utrophin, have been considered residents of focal adhesions in nonmuscle cells [59, 64, 65] and, after direct interaction of the cytoplasmic tail of β-Dg with F-actin was described [66], Dg has been implicated in cell adhesion and spreading.

Dg was identified in podosomes at the early stages of myoblast spreading; these structures contain a regulatory complex comprising dystroglycan, Tks5, and Src [67]. Myoblast spreading occurred in relation to dystroglycan expression levels, which in turn altered the size and number of focal contacts, focal adhesions, and fibrillar adhesions. Dystroglycan-mediated cell adhesion and spreading took place through indirect interaction with vinculin by binding to the vinculin-binding protein vinexin [61], while an adhesome was made up of by laminin-Dgmyosin IIA, crucial for maintaining the shape of notochordal cells [65].

In addition to a specific role in the maintenance of muscle integrity, Dg plays a more ubiquitous role in cell adhesion, signaling, and polarity. During embryogenesis, the follicle-cell epithelium (FCE) maintains the cell polarity promoted by the association between perlecan and Dg [68], while in astrocytes, end-feet in brain laminin induced a dramatic, polarized redistribution of cell-surface clusters or macrodomains, which colocalized extensively with β-Dg and AQP4 [69].

**5.4. Dystrolglycans**

of α-Dg [58].

*5.4.1. Dg involved in the signaling process*

112 Cytoskeleton - Structure, Dynamics, Function and Disease

β-Dg at tyrosine 890 [62].

The single dystroglycan gene encodes for a precursor protein that undergoes posttranslational proteolytic cleavage, which in turn produces two noncovalant DGC subunits: α- and β-dystroglycan. α-Dystroglycan is a dumbbell-shaped protein that binds to the laminin G domain in ECM components such as laminins, agrin, and perlecan. β-Dystroglycan (β-Dg) possesses a single transmembrane domain spanning the plasma membrane and an extracellular amino-terminal extracellular domain binding to the carboxy-terminal globular domain

The β-Dg dual role (structural and signaling) has been demonstrated in various cell types and tissues. Examples of the former role are represented by the participation of β-Dg in cytoskeleton remodeling, where it is associated with actin [59, 60], while its signaling role is represented by its association with the extracellular signal-related kinase-mitogen-activated protein (ERK-MAP) kinase cascade [18], or with integrins modulates myoblast anchorage and migration [61]; this latter process is critically regulated by Src-mediated phosphorylation of

Grb2–β-Dg interaction could facilitate the transduction of signals between the DGC and extracellular proteins and other signaling pathways [62]. However, when Dg is localized at the tips of dynamic filopodia, it directs local Cdc42 activation and recruits the guanine nucleotide

Dystroglycan is also a multifunctional adaptor or scaffold capable of interacting with components of the ERK-MAP kinase cascade, including MEK and ERK [18]. However, it has been established that integrin α6Aβ1 and dystroglycan play antagonistic roles in signaling to the

Since 1995, dystroglycan-associated proteins, such as utrophin, have been considered residents of focal adhesions in nonmuscle cells [59, 64, 65] and, after direct interaction of the cytoplasmic tail of β-Dg with F-actin was described [66], Dg has been implicated in cell adhesion

Dg was identified in podosomes at the early stages of myoblast spreading; these structures contain a regulatory complex comprising dystroglycan, Tks5, and Src [67]. Myoblast spreading occurred in relation to dystroglycan expression levels, which in turn altered the size and number of focal contacts, focal adhesions, and fibrillar adhesions. Dystroglycan-mediated cell adhesion and spreading took place through indirect interaction with vinculin by binding to the vinculin-binding protein vinexin [61], while an adhesome was made up of by laminin-Dg-

In addition to a specific role in the maintenance of muscle integrity, Dg plays a more ubiquitous role in cell adhesion, signaling, and polarity. During embryogenesis, the follicle-cell

myosin IIA, crucial for maintaining the shape of notochordal cells [65].

exchange factor (GEF) Dbl to generate actin protrusions [60].

Ras-Raf-MEK-ERK pathway in response to laminin [63].

*5.4.2. Dg promoter of the adhesion process*

and spreading.

The cytoskeletal polymers—actin, microtubules, and intermediate filaments—are interlinked by coordinated protein interactions to form a complex three-dimensional (3D) cytoskeletal network; these components are depicted in **Figure 3**. Although these systems are composed of distinctly different proteins, they are in constant and intimate communication with each another and with intermediate filaments, and their associated proteins are important components in mediating this crosstalk [70].

In platelets, two members of type-III intermediate filament (IF) proteins, desmin and vimentin, maintain a close relationship with DGC components, such as β-dystroglycan [β-dg], α-syntrophin [α-syn], and α-dystrobrevin [α-db], and are codistributed at the granulomere zone, participating in α-granule distribution [71].

The epithelial sodium channel (ENaC) is associated with IF and with dystrophin-associated proteins (DAP) via α-syntrophin and β-dystroglycan. ENaC is apparently dispensable for

**Figure 3.** Schematic diagram of microtubules and actin filaments participating in the transport of alpha and dense granules in the platelet adhesion process, during which α-dystrobrevins are the regulatory and adaptor proteins for governing trafficking events.

migration and alpha- and dense-granule secretion, whereas Na<sup>+</sup> influx through this channel is fundamental for platelet collagen activation [72]. This channel is overexpressed in platelets from hypertensive subjects in relation with control subjects, and β-Dg is a scaffold for the organization of ENaC and associated proteins [73].

#### *5.4.3. Dg and its posttranscriptional modifications*

Posttranscriptional modifications in the Dg protein possess important implications in cellular functions. The transmembrane β-subunit, which interacts with α-Dg extracellularly and which also connects with several different cytolinker proteins intracellularly, is additionally subject to altered N-linked glycosylation [74]. Additional modifications to β-Dg, however, include phosphorylation on tyrosine [75, 76] and specific proteolytic cleavage events. Tyrosine phosphorylation of β-Dg serves as a molecular switch to regulate the binding of different cellular-binding partners [77], but it is also a signal of the internalization of Dg from the plasma membrane [78, 79] and may mediate some proteolytic events and nuclear translocation [80, 81].

β-Dg is subject to proteolysis at several key sites: matrix metalloproteinase (MMP)-mediated cleavage liberates the extracellular portion of β-Dg, MMP-9-mediated proteolytic cleavage of the β-Dg, and it has been implicated in dendritic outgrowth and arborization in primary hippocampal neurons [82]. The remaining 31 kDa transmembrane stub and cytoplasmic domain can be detected with antibodies at the carboxy terminus of the cytoplasmic domain. As yet unknown proteases generate smaller fragments corresponding to the cytoplasmic region of β-Dg [83, 84], most typically observed as a 26-kDa fragment, but occasionally as a 17-kDa fragment.

In hematopoietic stem/progenitor cells, a 50-kDa β-Dg is the main product, while in differentiated cells, such as neutrophils and platelets, the characteristic glycosylated 43 kDa band is present [29, 31]. A 65-kDa band was also observed in neutrophils; perhaps this molecular weight (MW) is due to a posttranscriptional modification such as SUMOylation.

Ezrin is able to interact with dystroglycan through a cluster of basic residues in the juxtamembrane region, and appears to be responsible for dystroglycan-mediated formation of filopodia [18]. Colocalization of endogenous dystroglycan with ezrin at the cleavage furrow and midbody during cytokinesis not only affords dystroglycan a role in organizing the contractile ring through direct or indirect associations with actin, but also can modulate the cell cycle by affecting extracellular signal-regulated kinase levels [85]. Recent experiments have demonstrated β-Dg trafficking from the cytoplasm to the nucleus by ezrin-mediated cytoskeleton reorganization, the latter dependent on IMPα2/β1 [86].

Due to the presence of a conventional nuclear localization sequence (NLS)-/Imp-dependent nuclear import pathway in the cytoplasmic juxtamembrane region of β-Dg [87], β-Dg and proteolytic fragments containing the nuclear localization signal can be targeted to the nucleus via an importin-dependent pathway [88], where it can exert effects on nuclear architecture [89].

#### *5.4.4. Dg in the differentiation process*

migration and alpha- and dense-granule secretion, whereas Na<sup>+</sup>

organization of ENaC and associated proteins [73].

*5.4.3. Dg and its posttranscriptional modifications*

114 Cytoskeleton - Structure, Dynamics, Function and Disease

location [80, 81].

fragment.

tecture [89].

is fundamental for platelet collagen activation [72]. This channel is overexpressed in platelets from hypertensive subjects in relation with control subjects, and β-Dg is a scaffold for the

Posttranscriptional modifications in the Dg protein possess important implications in cellular functions. The transmembrane β-subunit, which interacts with α-Dg extracellularly and which also connects with several different cytolinker proteins intracellularly, is additionally subject to altered N-linked glycosylation [74]. Additional modifications to β-Dg, however, include phosphorylation on tyrosine [75, 76] and specific proteolytic cleavage events. Tyrosine phosphorylation of β-Dg serves as a molecular switch to regulate the binding of different cellular-binding partners [77], but it is also a signal of the internalization of Dg from the plasma membrane [78, 79] and may mediate some proteolytic events and nuclear trans-

β-Dg is subject to proteolysis at several key sites: matrix metalloproteinase (MMP)-mediated cleavage liberates the extracellular portion of β-Dg, MMP-9-mediated proteolytic cleavage of the β-Dg, and it has been implicated in dendritic outgrowth and arborization in primary hippocampal neurons [82]. The remaining 31 kDa transmembrane stub and cytoplasmic domain can be detected with antibodies at the carboxy terminus of the cytoplasmic domain. As yet unknown proteases generate smaller fragments corresponding to the cytoplasmic region of β-Dg [83, 84], most typically observed as a 26-kDa fragment, but occasionally as a 17-kDa

In hematopoietic stem/progenitor cells, a 50-kDa β-Dg is the main product, while in differentiated cells, such as neutrophils and platelets, the characteristic glycosylated 43 kDa band is present [29, 31]. A 65-kDa band was also observed in neutrophils; perhaps this molecular

Ezrin is able to interact with dystroglycan through a cluster of basic residues in the juxtamembrane region, and appears to be responsible for dystroglycan-mediated formation of filopodia [18]. Colocalization of endogenous dystroglycan with ezrin at the cleavage furrow and midbody during cytokinesis not only affords dystroglycan a role in organizing the contractile ring through direct or indirect associations with actin, but also can modulate the cell cycle by affecting extracellular signal-regulated kinase levels [85]. Recent experiments have demonstrated β-Dg trafficking from the cytoplasm to the nucleus by ezrin-mediated cytoskeleton

Due to the presence of a conventional nuclear localization sequence (NLS)-/Imp-dependent nuclear import pathway in the cytoplasmic juxtamembrane region of β-Dg [87], β-Dg and proteolytic fragments containing the nuclear localization signal can be targeted to the nucleus via an importin-dependent pathway [88], where it can exert effects on nuclear archi-

weight (MW) is due to a posttranscriptional modification such as SUMOylation.

reorganization, the latter dependent on IMPα2/β1 [86].

influx through this channel

The expression has been described as the major components of DAPC visceral and subcutaneous rat adipose depots that are regulated during adipogenesis and by ECM components, suggesting an important role in adipocyte differentiation [90].

The human myeloid leukemia cell line HL-60 achieves increasing cessation after its exposure to all-trans-retinoic acid (ATRA) and dimethyl sulfoxide (DMSO) and becomes differentiated into granulocytes, evoking the biology of the disease *in vitro* [91, 92]. Recently, it was demonstrated that dystroglycans actively participate in the differentiation process, in that the expression levels of α-Dg (160 kDa), β-Dg (42kDa), and β-DgpY892 (42 kDa) were increased in differentiated compared with nondifferentiated cells. Additionally, low levels of β-Dg in differentiated HL-60 cells are accompanied by reducing actin-based protrusions, such as in filopodia and lamellipodia extrusion, avoiding motility or phagocytic capabilities, respectively [93]. Similar changes were also observed when HL-60 cells were transfected with a shRNA directed to dystroglycan; therefore, a direct consequence of the reduction in dystroglycan exerted a direct effect on actin cytoskeletal dynamics, either on its direct or indirect interaction with actin, but also interfering with actin regulatory pathways [66].

The Kasumi-1 cell line is a model system of acute myeloid leukemia (AML) with *t*(8;21) translocation and the corresponding functional consequences of the AML1–ETO fusion oncogene on myeloid differentiation [94]. *In vitro*, macrophages differentiated from myelomonocytic cell lines exhibited downregulation of adhesion molecules after tissue plasminogen activator (TPA) treatment [95]. The biochemical analysis of cytoplasmic or nuclear Kasumi-1 cell extracts revealed bands of 50, 38, and 30 kDa present in the nucleus of the cells, while the majority of 43 kDa β-Dg was found mainly in the cytoplasmic compartment, with the 38-kDa band also abundant in the cytoplasm of nondifferentiated Kasumi-1 and differentiated Kasumi-1 cells. The phosphorylated 31-kDa fragment of dystroglycan is the species that is most translocated to the nucleus of nondifferentiated cells, while the 50-kDa fragment comprised the most abundant species at the nucleus of differentiated cells. The diminished expression levels of Dg in differentiated Kasumi-1 cells compared with nondifferentiated cells could facilitate cell recruitment in solid tissues; apparently, the phosphorylated species may be ubiquitinated and processed by the proteasome. However, a direct consequence of a reduction in dystroglycan exerts an effect on actin cytoskeletal dynamics, but does not impair the differentiation process [96].

#### *5.4.5. Dg in cell membrane organization*

Several structures of the cell membrane play major roles in physiological functions through signaling and adhesion to neighbor cells and to ECM. Generic features, such as the cytoskeleton meshwork, rafts, and protein complexes, which are subjected to thermal motion, contribute to building membrane structures such as focal adhesions (FA) [97] and immune [98] and neuronal [99] synapses. The rapid and transient association of the partners of a given signaling pathway, localized in close proximity within narrow structures/domains, is a requirement for rapid and reliable signal transmission [100].

The existence of "rafts" supposes that membrane lipids and proteins associate with each other according to their affinities, due to their hydrophobicity and geometry [101]. Rafts were initially proposed as contributing to protein sorting along the synthesis pathway, and have also been associated with several membrane features, including signaling platforms and adhesion structures. Caveolae are cholesterol- and sphingolipid-enriched membrane invaginations [102], and caveolin-1 is the primary caveolae structural protein in several cells [103]. Therefore, caveolae and caveolin-1 play a key role in orchestrating the activation of pathways that underpin cell proliferation, migration, and contraction [104]. For example, direct interaction between caveolin-1 and β-Dg was demonstrated in contractile smooth muscle, where the distribution of caveolae is determined by their tethering to the actin cytoskeleton via caveolin-1 and the DGC [105].

In this regard, cholesterol demonstrated to be essential to modulate platelet cytoskeleton reorganization, while the association of caveolin-1 PY14 with intermediate filaments, as well as with focal adhesion proteins via vinculin, was a determinant in adhered platelets, where β-Dg participation was a key scaffold component for caveolin-1 and FAK [106].

In general, diseases of the DGC are incurable, in part because the majority of these give rise to great damage resulting from the loss of these proteins. However, there is increasing evidence that proteins in the DGC may play a significant role in the pathophysiology of more common diseases such as cancer, in which the DGC has been implicated.

Throughout these years of basic research, it has been observed that dystroglycan functional changes, either for posttranscriptional modifications or for deregulation of the protein, simultaneously affect both scaffolding and signaling roles. These changes modify cell adhesion and motility, MAPK signaling, or its translocation to the nuclei that, in the prostate, is associated with the ETV1 transcription factor, acting directly on cancer progression and the pathophysiology of the disease [81]. Therefore, a complete understanding of the role of DGC elements in the pathophysiology of a disease would allow the identification of strategies for the development of specific therapeutics.

Previous studies demonstrated that preventing tyrosine phosphorylation of β-Dg in mdx mouse alleviated the dystrophic phenotype in a genetic mouse model, ameliorating many of the main pathological symptoms associated with dystrophin deficiency [78]. The use of dasatinib was found to decrease β-Dg phosphorylation levels in tyrosine and to increase the relative levels of nonphosphorylated β-Dg in the sapje zebrafish, improving its physical condition [79].

### **6. Conclusion**

Since 1980, the dystrophin–glycoprotein complex has been considered only as a group of multiproteins working together to ensure the function of muscle tissue; however, along these years and according to basic research, dystrophin has acquired prime status and has become the central component of a scaffold of proteins expressed in a variety of tissues including blood. Within the complex elements, dystroglycan has received the majority of our attention and has been identified as participating in the clustering of membrane receptors, integrins, and ion channels, modulating cellular signal integration, such as in the differentiation process.

Despite all of these advances, it remains difficult to dissect the specific function of a particular protein and, given the close association and interdependence of the different elements of the complex, it should be difficult to define the specific contribution of each of the complex's protein elements. However, the improvement and development of biochemical and molecular tolls will undoubtedly aid in elucidating novel therapies to counteract common diseases such as cancer.

### **Author details**

The existence of "rafts" supposes that membrane lipids and proteins associate with each other according to their affinities, due to their hydrophobicity and geometry [101]. Rafts were initially proposed as contributing to protein sorting along the synthesis pathway, and have also been associated with several membrane features, including signaling platforms and adhesion structures. Caveolae are cholesterol- and sphingolipid-enriched membrane invaginations [102], and caveolin-1 is the primary caveolae structural protein in several cells [103]. Therefore, caveolae and caveolin-1 play a key role in orchestrating the activation of pathways that underpin cell proliferation, migration, and contraction [104]. For example, direct interaction between caveolin-1 and β-Dg was demonstrated in contractile smooth muscle, where the distribution of caveolae is determined by their tethering to the actin cytoskeleton via caveo-

In this regard, cholesterol demonstrated to be essential to modulate platelet cytoskeleton reorganization, while the association of caveolin-1 PY14 with intermediate filaments, as well as with focal adhesion proteins via vinculin, was a determinant in adhered platelets, where β-Dg

In general, diseases of the DGC are incurable, in part because the majority of these give rise to great damage resulting from the loss of these proteins. However, there is increasing evidence that proteins in the DGC may play a significant role in the pathophysiology of more common

Throughout these years of basic research, it has been observed that dystroglycan functional changes, either for posttranscriptional modifications or for deregulation of the protein, simultaneously affect both scaffolding and signaling roles. These changes modify cell adhesion and motility, MAPK signaling, or its translocation to the nuclei that, in the prostate, is associated with the ETV1 transcription factor, acting directly on cancer progression and the pathophysiology of the disease [81]. Therefore, a complete understanding of the role of DGC elements in the pathophysiology of a disease would allow the identification of strategies for the develop-

Previous studies demonstrated that preventing tyrosine phosphorylation of β-Dg in mdx mouse alleviated the dystrophic phenotype in a genetic mouse model, ameliorating many of the main pathological symptoms associated with dystrophin deficiency [78]. The use of dasatinib was found to decrease β-Dg phosphorylation levels in tyrosine and to increase the relative levels of nonphosphorylated β-Dg in the sapje zebrafish, improving its physical

Since 1980, the dystrophin–glycoprotein complex has been considered only as a group of multiproteins working together to ensure the function of muscle tissue; however, along these years and according to basic research, dystrophin has acquired prime status and has become the central component of a scaffold of proteins expressed in a variety of tissues including blood. Within the complex elements, dystroglycan has received the majority of our attention

participation was a key scaffold component for caveolin-1 and FAK [106].

diseases such as cancer, in which the DGC has been implicated.

lin-1 and the DGC [105].

116 Cytoskeleton - Structure, Dynamics, Function and Disease

ment of specific therapeutics.

condition [79].

**6. Conclusion**

Doris Cerecedo

Address all correspondence to: dcereced@prodigy.net.mx

Laboratorio de Hematobiología, Escuela Nacional de Medicina y Homeopatía (ENMH), Instituto Politécnico Nacional (IPN), Mexico City, Mexico

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