**Table 3.** *Std Z*½ � *d*ð Þ*t for* λ*1* ¼ *1*, λ*2* ¼ *10*, δ ¼ *3*%*.*

**Table 4.** *Std Z*½ � *d*ð Þ*t for* λ*1* ¼ *5*, λ*2* ¼ *10*, δ ¼ *3*%*.*

#### **4.2 Premium calculation**

From the results in Section 4.1, we can compute the premium related to the risk of an insurance portfolio represented by *G t*ð Þ, depending on the premium calculation principles adopted by the insurance company. The loaded premium *Zd*ð Þ*t* consists in the sum of the pure premium *E Z*½ � *d*ð Þ*t* , the expected value of the costs related to the portfolio, and a loading for the risk *M t*ð Þ as

$$\mathbf{G}(t) = \mathbf{E}[\mathbf{Z}\_d(t)] + \mathbf{M}(t) \tag{73}$$

The loading for the risk differs according to the premium calculation principles.

#### *4.2.1 The expected value principle*

Denote by θ . 0 the safety loading. The expected value principle defines the loaded premium as:

$$\mathbf{G}(\mathbf{t}) = \mathbf{E}[\mathbf{Z}\_d(\mathbf{t})] + \theta \mathbf{E}[\mathbf{Z}\_d(\mathbf{t})],\tag{74}$$

where *M t*ðÞ¼ *θE Z*½ � *d*ð Þ*t* .

#### *4.2.2 The variance principle*

Denote by θ . 0 the safety loading. The variance principle defines the loaded premium as:

*Moments of the Discounted Aggregate Claims with Delay Inter-Occurrence Distribution… DOI: http://dx.doi.org/10.5772/intechopen.88699*

$$\mathbf{G}(\mathbf{t}) = \mathbf{E}[\mathbf{Z}\_d(\mathbf{t})] + \theta \mathbf{Var}[\mathbf{Z}\_d(\mathbf{t})],\tag{75}$$

where *M t*ðÞ¼ *θVar Z*½ � *d*ð Þ*t* .

#### *4.2.3 The standard deviation principle*

Denote by θ . 0 the safety loading. The standard deviation principle defines the loaded premium as:

$$\mathbf{G}(\mathbf{t}) = \mathbf{E}[\mathbf{Z}\_d(\mathbf{t})] + \Theta \sqrt{\mathbf{Var}[\mathbf{Z}\_d(\mathbf{t})]},\tag{76}$$

where *M t*ðÞ¼ θ ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi *Var Z*½ � *d*ð Þ*t* p .

#### *4.2.4 The quantile principle*

The standard deviation principle defines the loaded premium as:

$$G(t) = F\_{Z\_d(t)}^{-1}(1 - \varepsilon),\tag{77}$$

where *ε* is smallest (for example: *ε* ¼ 0*:*5%*,* 1%*,* 2*:*5%*,* 5%Þ. In this case, the safety loading *M t*ð Þ is given by

$$\mathbf{M}(t) = F\_{Z\_d(t)}^{-1}(1 - \varepsilon) - E[Z\_d(t)] \tag{78}$$

The principles of standard deviation and variance only require partial information on the distribution of the random variable, *Zd*ð Þ*t* , i.e., its expectation and its variance.

Often, the actuary only has this information for different reasons (time constraints, information …).

If the actuary has more information about the random variable, *Zd*ð Þ*t* i.e., he knows the form of *FZd*ð Þ*t* , then he can apply the quantile principle.

But he does not know much about *FZd*ð Þ*t* , then he can approximate the distribution of *Zd*ð Þ*t* using the matching moments technique.


**Novel Insights into the Role of the Cytoskeleton in Cancer**

Xuan Zhang, Zenglin Pei, Chunxia Ji, Xiaoyan Zhang, Jianqing Xu and Jin Wang

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/66860

#### **Abstract**

The cytoskeleton is a complex network of highly ordered intracellular filaments that plays a central role in controlling cell shape, division, functions, and interactions in human organs and tissues, but dysregulation of this network can contribute to numerous human diseases, including cancer. To clarify the various functions of the cytoskeleton and its role in cancer progression, in this chapter, we will discuss the microfilament (actin cytoskeleton), microtubule (β‐tubulin), and intermediate filament (keratins, cytokera‐ tins, vimentin, and lamins) cytoskeletal structures; analyze the physiological functions of the cytoskeleton and its regulation of cell differentiation; and investigate the roles of the cytoskeleton in cancer progression, the epithelial‐mesenchymal transition process (EMT), and the mechanisms of multidrug resistance (MDR) in relation to the cytoskel‐ eton. Importantly, the cytoskeleton, as a key regulator of the transcription and expres‐ sion of many genes, is known to be involved in various physiological and pathological processes, which makes the cytoskeleton a novel and highly effective target for assessing the response to antitumor therapy in cancer.

**Keywords:** Cytoskeleton, Regulator, cell differentiation, drug resistance, EMT

### **1. Introduction**

The cytoskeleton is a structure similar to a bird's nest; it can be tightly packed or sparse and is found in both prokaryotes and eukaryotes [1]. The main component of the cytoskeleton is protein, and the specific differences in structure never affect the type of proteins incorporated [2]. The cytoskeletons of prokaryotes display apparent plasticity in composition, without con‐ servation of the core filament‐forming proteins. However, the eukaryotic cytoskeleton has evolved in a variety of functions through the addition of accessory proteins and extensive

© 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.

gene duplication. The distribution of cytoskeletal filaments puts constraints on the likely pro‐ karyotic line that made the leap into eukaryogenesis [3].

There are three main cytoskeletal structures in eukaryotes, microfilaments (MFs, ≈7 nm diame‐ ter), microtubules (MTs, ≈25 nm diameter), and intermediate filaments (IFs, ≈10 nm diameter) [1]. MFs are responsible for cell contraction and reinforcement of the cell surface and allow changes in cell morphology. Actin and tubulin are the main globular proteins that form MFs and MTs, respectively. Actin is a ubiquitous eukaryotic filament‐forming protein. Actin fila‐ ments (also called microfilaments or F‐actin) consist of two proto filament polymers wound together in a right‐handed helix [3]. Eukaryotic actin is a member of a large and diverse super‐ family of ATPases that includes Hsp70 chaperones and several classes of sugar/sugar alcohol kinases [4, 5] as well as eukaryotic actin‐related proteins (ARPs) [6, 7]. The actin cytoskeleton is involved in actin‐based cytoskeletal structures, including various functionally distinct struc‐ tures of actin organization, and can be regulated by actin regulatory proteins. It is well known that the actin cortex consists of a dense mesh‐like array of F‐actin anchored to the cell mem‐ branes [8, 9]. This structure provides the core "skeleton" of the cell, functioning to define cell shape and provide resistance to mechanical stress. Reorganization of the actin cytoskeleton describes a process where cells actively alter the architecture of actin filaments to adjust cell shape in response to environmental requirements. Globular‐ (G‐) actin is a highly conserved, polar protein with a molecular weight of 42 kDa that forms dimers and trimers in a process called actin nucleation; these structures then assemble into a double‐stranded helical filament (F‐actin) with a diameter of 7–9 nm (actin polymerization) [10–15]. Filopodia are thin, hair‐like cellular protrusions that consist of parallel actin bundles cross‐linked by interacting protein partners such as fascin, α‐actinin, fimbrin, and formins [16]. Filopodia sense changes in the cellular microenvironment, such as growth factor concentrations, to guide cellular movement through the surrounding matrix [10–12]. Fascin is a highly conserved actin‐bundling pro‐ tein with three isoforms. While Fascin‐1 is ubiquitously expressed during embryogenesis, its expression is later restricted to the endothelium, neuronal tissue, and testis [16]. Fascin‐2 and Fascin‐3 are expressed in the retinal epithelium and testis only [17]. Fascin is phosphorylated by protein kinase C (PKC), which regulates its actin bundling activity in accordance with the current microenvironmental conditions, which are communicated via surface integrins [16].

Microtubules are responsible for structural strength and cell shape. They allow organelles to move within cells. These structures act like rails on which kinesin and dynein can pull organ‐ elles. Most microtubules consist of 13 protofilaments that interact laterally to form a hollow tube and arise from the polymerization of heterodimers of a‐ and ß‐tubulin, which are added to the plus‐end of microtubules containing GTP in both subunits [3, 18].

As the major components of the cytoskeleton, intermediate filaments (IFs) are ubiquitous cyto‐ skeletal elements that are encoded by about 70 genes in the human genome [19–21] and are divided into six groups based on their structure. These groups include the keratins, cytokera‐ tins, mesoderm‐specific intermediate filaments, neurofilaments and related proteins, lamins, and beaded filament proteins of the eye lens [21–24]. Although these IF proteins have very different amino acid sequences, the organization of the structural domain is similar [24]. The keratin group is defined as the group of intermediate filaments within epithelial cells, forming particles from 44 kDa to approximately 66 kDa that are characterized by high stability and chemical resistance [25]. As the major structural proteins of the nuclear lamina, the lamins are subdivided into A‐ and B‐types, both of which belong to the type V intermediate filament protein family [26].

gene duplication. The distribution of cytoskeletal filaments puts constraints on the likely pro‐

There are three main cytoskeletal structures in eukaryotes, microfilaments (MFs, ≈7 nm diame‐ ter), microtubules (MTs, ≈25 nm diameter), and intermediate filaments (IFs, ≈10 nm diameter) [1]. MFs are responsible for cell contraction and reinforcement of the cell surface and allow changes in cell morphology. Actin and tubulin are the main globular proteins that form MFs and MTs, respectively. Actin is a ubiquitous eukaryotic filament‐forming protein. Actin fila‐ ments (also called microfilaments or F‐actin) consist of two proto filament polymers wound together in a right‐handed helix [3]. Eukaryotic actin is a member of a large and diverse super‐ family of ATPases that includes Hsp70 chaperones and several classes of sugar/sugar alcohol kinases [4, 5] as well as eukaryotic actin‐related proteins (ARPs) [6, 7]. The actin cytoskeleton is involved in actin‐based cytoskeletal structures, including various functionally distinct struc‐ tures of actin organization, and can be regulated by actin regulatory proteins. It is well known that the actin cortex consists of a dense mesh‐like array of F‐actin anchored to the cell mem‐ branes [8, 9]. This structure provides the core "skeleton" of the cell, functioning to define cell shape and provide resistance to mechanical stress. Reorganization of the actin cytoskeleton describes a process where cells actively alter the architecture of actin filaments to adjust cell shape in response to environmental requirements. Globular‐ (G‐) actin is a highly conserved, polar protein with a molecular weight of 42 kDa that forms dimers and trimers in a process called actin nucleation; these structures then assemble into a double‐stranded helical filament (F‐actin) with a diameter of 7–9 nm (actin polymerization) [10–15]. Filopodia are thin, hair‐like cellular protrusions that consist of parallel actin bundles cross‐linked by interacting protein partners such as fascin, α‐actinin, fimbrin, and formins [16]. Filopodia sense changes in the cellular microenvironment, such as growth factor concentrations, to guide cellular movement through the surrounding matrix [10–12]. Fascin is a highly conserved actin‐bundling pro‐ tein with three isoforms. While Fascin‐1 is ubiquitously expressed during embryogenesis, its expression is later restricted to the endothelium, neuronal tissue, and testis [16]. Fascin‐2 and Fascin‐3 are expressed in the retinal epithelium and testis only [17]. Fascin is phosphorylated by protein kinase C (PKC), which regulates its actin bundling activity in accordance with the current microenvironmental conditions, which are communicated via surface integrins [16].

Microtubules are responsible for structural strength and cell shape. They allow organelles to move within cells. These structures act like rails on which kinesin and dynein can pull organ‐ elles. Most microtubules consist of 13 protofilaments that interact laterally to form a hollow tube and arise from the polymerization of heterodimers of a‐ and ß‐tubulin, which are added

As the major components of the cytoskeleton, intermediate filaments (IFs) are ubiquitous cyto‐ skeletal elements that are encoded by about 70 genes in the human genome [19–21] and are divided into six groups based on their structure. These groups include the keratins, cytokera‐ tins, mesoderm‐specific intermediate filaments, neurofilaments and related proteins, lamins, and beaded filament proteins of the eye lens [21–24]. Although these IF proteins have very different amino acid sequences, the organization of the structural domain is similar [24]. The keratin group is defined as the group of intermediate filaments within epithelial cells, forming particles from 44 kDa to approximately 66 kDa that are characterized by high stability and

to the plus‐end of microtubules containing GTP in both subunits [3, 18].

karyotic line that made the leap into eukaryogenesis [3].

300 Cytoskeleton - Structure, Dynamics, Function and Disease

On the other hand, prokaryotes also have homologs of the eukaryotic microfilaments (actin), microtubules (tubulin), and intermediate filament proteins [27]. FtsZ was first found in the prokaryote as the cytoskeleton and forms filaments but not tubules [28]. The MreB and ParM proteins in prokaryotic cells function like actin in eukaryotic cells [29]. The third type of cyto‐ skeleton in prokaryotes is crescentin, which is responsible for the shape of cells [30].

### **2. The physiological functions of the cytoskeleton and the regulation of cell differentiation**

The cytoskeleton is the frame around or within the cell, and a system of intracellular filaments is crucial for cell shape, division, and function in all three domains of life [3, 4]. The classi‐ cal functions of the cytoskeleton have been summarized as morphology determination, cell polarity formation, structural support, membrane constitution, cell motility, and receptor or channel localization in the plasma membrane [31–33].

The activity of actin, the main type of microfilament, is regulated by GTPases, which control the formation of actin filaments [34–36]. In patients with Alzheimer's disease (AD), actin fila‐ ments play a central role in maintaining and modifying synaptic connections [37]. As the key mediator between receptor activation during learning and a protein involved in regulating spine morphology [38, 39], actin not only plays a role in the nervous system but also has func‐ tions in the immune system [40]. For example, F‐actin can mediate signaling in B cells and T cells [41, 42]. The dynamics of the actin cytoskeleton regulate adhesion and signal transduc‐ tion in T cells/APCs [40]. SWAP‐70 and HS1 are important downstream components of the TCR signaling pathway and are regulated by actin [43, 44].

The key function of intermediate filaments is to support the cell membrane, serving as a structural scaffold to maintain cell shape. Cell motility is significantly enhanced as a result of changes in intermediate filaments. Intermediate filaments are fixed to the membrane through transmembrane proteins such as cadherins, which are involved in the formation of cell‐cell tight junctions and the distribution of traction forces that arise in the interspace between cells. Under stress stimulation, intermediate filaments are significantly upregulated to induce the rearrangement of the cytoskeleton [45, 46]. Keratins are proteins that form intermediate fila‐ ments of epithelial cell cytoskeleton and have an antiapoptotic function, regulate protein syn‐ thesis, and play a role in wound healing [25]. Epithelial cells obtain a specific pattern of keratin expression during differentiation and maturation; this pattern reflects the specificity of the tissue and the degree of maturation [25]. The different components of the cytoskeleton do not work alone, and microtubules, microfilaments, and intermediate filaments often interact with each other to complete cellular processes. They always participate in protein localization and cell signaling. A characteristic of differentiation is a change in cell shape that is dependent on the cytoskeleton. During mesenchymal stem cell differentiation, the actin cytoskeleton of mesenchymal stem cells changes during osteogenic and chondrogenic differentiation [47]. Previous studies have shown that adipocyte differentiation is associated with actin structure [48, 49]. Disruption of the actin cytoskeleton can regulate MKL1 and result in adipocyte dif‐ ferentiation [50]. Rearrangements in and the formation of processes by the cytoskeleton are associated with the synthesis of synaptopodin, which is a marker of differentiated podocytes [51]. Moreover, cell differentiation is regulated by activating or repressing some transcription factors and is linked to the cytoskeleton [52–57]. For example, Zoubiane demonstrated that microtubules were required for β‐casein expression, which resulted in epithelial cell differen‐ tiation [55]; Ahmad discovered that the pattern of microtubules in HL‐60 cells changed follow‐ ing differentiation, with α‐tubulin appearing more regularly organized in the differentiated HL‐60 cells [56]; Takiqawa also confirmed that the cytoskeleton, including microtubules and microfilaments, regulates the expression of a differentiated phenotype in chondrocytes [57].

### **3. The cytoskeleton and its role in cancer progression**

The cytoskeleton is known to contribute to cancer. The cytoskeleton may induce cell prolifera‐ tion and activate oncogenes, resulting in tumorigenesis [58]. In mammary carcinoma cells, the upregulation of WNT4 increased mesenchymal and cytoskeleton remodeling markers [59].  DKK1 is considered a factor that can modulate the β‐catenin pathway and stimulate cancer cells or noncancerous proliferation [61, 62]. Zyxin localizes to focal adhesion sites and stress fibers in response to mechanical cues and has been shown to control the assembly of the cytoskeleton, the generation of traction force, cell movement, and signal transduction. If zyxin is nonfunc‐ tional, the cytoskeleton may be disturbed, leading to cancer [59].

Many actin‐bundling proteins are also linked to cancer progression and tumor chemoresis‐ tance [63]. Fascin proteins organize F‐actin into parallel bundles and are required for the formation of actin‐based cellular protrusions. The inhibition of fascin can interfere with the formation of filopodia and suppress the migration and invasion of tumors [64], making it pos‐ sible to use fascin as a small molecular target to inhibit cancer metastasis.

Intermediate filaments interact with arcs and can inhibit the activity of arcs, which can trans‐ port intermediate filaments to cell nucleus. However, fewer intermediate filaments may alter the cell shape and lead to diseases such as tumors [65]. For example, the changes in nuclear architecture that are the pathological hallmarks of cancer cells are related to alterations in the lamins, with alterations in lamins A/C being verified in the colon [66], gut [67], lung [68], and prostate cancer [69].

### **4. The role of the cellular cytoskeleton in epithelial‐mesenchymal transition (EMT)**

Epithelial‐mesenchymal transition (EMT) is a biological process resulting in the loss of polar‐ ity and cell junctions, and disturbances in the cytoskeleton [70]. The reorganization of the actin cytoskeleton is important for metastasis and the differentiation of epithelial cells to mesenchymal cells [71]. When cells undergo EMT, the number of β‐actin fibers is reduced and the distribution of β‐actin becomes diffused. RhoA induces action fiber formation and regulates the cytoskeleton. The Rho family also plays a role in tumor migration and EMT [72]. Cellular transformation was closely associated with changes in the distribution and amount of cytoplasmic actin isoforms [73]. Actin filaments are formed by the polymerization of G‐actin, which induces the formation of a leading edge in cancer cells undergoing EMT through inter‐ actions with binding proteins and contractile proteins such as myosin II, which accelerates the movement of actin fibers on the substrate to the leading edge [74].

Previous studies have shown that adipocyte differentiation is associated with actin structure [48, 49]. Disruption of the actin cytoskeleton can regulate MKL1 and result in adipocyte dif‐ ferentiation [50]. Rearrangements in and the formation of processes by the cytoskeleton are associated with the synthesis of synaptopodin, which is a marker of differentiated podocytes [51]. Moreover, cell differentiation is regulated by activating or repressing some transcription factors and is linked to the cytoskeleton [52–57]. For example, Zoubiane demonstrated that microtubules were required for β‐casein expression, which resulted in epithelial cell differen‐ tiation [55]; Ahmad discovered that the pattern of microtubules in HL‐60 cells changed follow‐ ing differentiation, with α‐tubulin appearing more regularly organized in the differentiated HL‐60 cells [56]; Takiqawa also confirmed that the cytoskeleton, including microtubules and microfilaments, regulates the expression of a differentiated phenotype in chondrocytes [57].

The cytoskeleton is known to contribute to cancer. The cytoskeleton may induce cell prolifera‐ tion and activate oncogenes, resulting in tumorigenesis [58]. In mammary carcinoma cells, the upregulation of WNT4 increased mesenchymal and cytoskeleton remodeling markers [59].  DKK1 is considered a factor that can modulate the β‐catenin pathway and stimulate cancer cells or noncancerous proliferation [61, 62]. Zyxin localizes to focal adhesion sites and stress fibers in response to mechanical cues and has been shown to control the assembly of the cytoskeleton, the generation of traction force, cell movement, and signal transduction. If zyxin is nonfunc‐

Many actin‐bundling proteins are also linked to cancer progression and tumor chemoresis‐ tance [63]. Fascin proteins organize F‐actin into parallel bundles and are required for the formation of actin‐based cellular protrusions. The inhibition of fascin can interfere with the formation of filopodia and suppress the migration and invasion of tumors [64], making it pos‐

Intermediate filaments interact with arcs and can inhibit the activity of arcs, which can trans‐ port intermediate filaments to cell nucleus. However, fewer intermediate filaments may alter the cell shape and lead to diseases such as tumors [65]. For example, the changes in nuclear architecture that are the pathological hallmarks of cancer cells are related to alterations in the lamins, with alterations in lamins A/C being verified in the colon [66], gut [67], lung [68], and

**4. The role of the cellular cytoskeleton in epithelial‐mesenchymal**

Epithelial‐mesenchymal transition (EMT) is a biological process resulting in the loss of polar‐ ity and cell junctions, and disturbances in the cytoskeleton [70]. The reorganization of the actin cytoskeleton is important for metastasis and the differentiation of epithelial cells to

**3. The cytoskeleton and its role in cancer progression**

302 Cytoskeleton - Structure, Dynamics, Function and Disease

tional, the cytoskeleton may be disturbed, leading to cancer [59].

prostate cancer [69].

**transition (EMT)**

sible to use fascin as a small molecular target to inhibit cancer metastasis.

Tubulin plays an important role in EMT induction and contributes to TGF‐β‐induced membrane extensions or protrusions of human carcinoma cells undergoing EMT in three‐dimensional col‐ lagen gels [75]. Acetylated α‐tubulin can serve as a new marker of EMT and is expressed at a high level on normal epithelial cells, while the expression of acetylated α‐tubulin decreases during TGF‐β‐induced EMT [76]. β‐III tubulin can modulate snail expression during EMT in HT‐29 and LS180 colon cancer cells [77]. When human mammary epithelial cells undergo EMT, the expression of Twist or Snail downregulates the tubulin tyrosine ligase enzyme, leading to the detyrosination of α‐tubulin. The accumulation of detyrosinated Glu‐tubulin is vital for the formation of microtentacles. These results provide new insight into tumor progression, as increasing α‐tubulin detyrosination could promote EMT [78]. Because of their effect on tumor migration during EMT, the inhibition of microtubules can be a useful target for antitumor drugs [79, 80].

During the EMT process, intermediate filaments are significantly rearranged, typically switching from cytokeratin‐rich to vimentin‐rich networks [81]. Intermediate filaments can be expressed in different types of cells. For example, keratins are specifically expressed in epithelial cells; type III (mostly mesodermal) proteins are expressed in mesenchymal cells [82–84]. Epithelial cells express different keratins that are considered almost specific markers, whereas mesenchymal cells, endothelial cells, and hematopoietic cells express vimentin [82, 85, 86]. Vimentin is a type III intermediate filament that is significantly upregulated during EMT in epithelial cells; thus, vimentin can be used as a marker of EMT [87]. E‐cadherin is one type of cell adhesion molecule that regulates EMT [88]. Reduced CK8 expression is regarded as an indicator of EMT, leading to more malignant forms of cancer [89]. Breast milk exosomes containing high levels of TGF‐β2 can induce changes in both benign and malignant breast epithelial cells, consistent with the development and progression of breast cancer, which occurs due to alterations in cellular shape and the actin cytoskeleton and the loss of cell‐cell junctions [90]. TGF‐β‐induced EMT was found to restrain cell invasion, which may be allevi‐ ated by the overexpression of hyperactivated Ras [91]. Endocytosis has emerged as a highly interconnected infrastructure of various cellular circuitries that is essential for the execution of different cellular programs, including those promoting a canonical EMT program and rely‐ ing on the activation of Wnt, Notch, or TGF‐β signaling [92]. On the other hand, miR‐200 can inhibit EMT and the migration of cervical cancer cells through RhoE, which is an actin‐bind‐ ing protein [93]. Therefore, the cellular cytoskeleton plays a role in EMT by activating Wnt, Notch, or TGF‐β signaling pathways, triggering the reprogramming of gene expression pat‐ terns via transcriptional changes and the altered expression of mRNA, including epithelial (E‐cadherin, claudins, occludins, desmoplakin, mucin‐1, and cytokeratin‐8, ‐9, ‐18) and mes‐ enchymal markers (fibronectin, FSP1, vitronectin, vimentin, smooth‐muscle actin, and FGFR2 IIIb and IIIc species variants) (**Figure 1**). Further in‐depth study is required to determine the features of the dynamic expression and arrangement of intracellular filaments during cancer invasion and migration.

**Figure 1.** Cellular cytoskeletons in epithelial‐mesenchymal transition process (EMT).

### **5. Mechanism of multidrug resistance (MDR) in relation to the cytoskeleton**

Many patients develop drug resistance to anticancer agents, with the mechanisms includ‐ ing alterations in the ATP‐binding cassette [94], microtubule dynamics, drug transport, and cell death [89], all of which involve tubulin and microtubules [95–97]. Microtubules have been considered a highly significant molecular target for anticancer agents, including micro‐ tubule‐stabilizing agents. For example, paclitaxel binds to the β‐tubulin subunit, accelerates the polymerization of tubulin, and stabilizes the resultant microtubules [98, 99]. Moreover, the paclitaxel‐induced resistance of vimentin intermediate filaments to okadaic acid may occur through a microtubule‐independent mechanism [100]. Townson also demonstrated that K8/18 filaments provided resistance to apoptosis in granulosa cell tumors by impairing FAS expression [101]. The organization of actin filaments associated with cellular differentia‐ tion may also influence the expression of P‐glycoprotein (P‐gp) through ezrin, radixin, and moesin in MDR osteosarcoma cells [92–104], which exhibit a significant increase in well‐orga‐ nized actin stress fibers [103], while inhibiting actin remodeling can suppress drug resistance in cancer [105].

The balance between polymerized and nonpolymerized tubulin will be a key determinant of the response to antimitotic‐based chemotherapy. Cancer cells obtain mitotic drug resistance properties through β I‐tubulin mutations [106], which is important for maintaining micro‐ tubule structure and sensitivity to microtubule‐targeting agents. β‐tubulin mutations confer resistance to epothilone and taxanes in ovarian cancer cells. Moreover, mutations in both a‐ and β‐tubulin have been found to confer resistance to colchicine and vinblastine in Chinese hamster ovary (CHO) cells [107, 108]. The upregulation of β III‐tubulin was further associated with resistance to paclitaxel and docetaxel [109–113]. On the other hand, as a negative regula‐ tor of β III‐tubulin, HDAC3 can also mediate drug resistance [113].

As a major intermediate filament in the cells of epithelial‐derived tumors, cytokeratin K8/18 expression is involved in cytokeratin‐dependent drug resistance [114]. Hepatocyte cytokera‐ tin plays a role in bile formation and resistance to bile acid challenge; however, the loss of K8 significantly increased liver injury in response to toxic stress in mice [115]. Caulin also dem‐ onstrated that normal and malignant epithelial cells deficient in K8/18 were approximately 100 times more sensitive to TNF‐induced death [116], indicating that interaction between the damaging agent and cytokeratin might trigger signaling responses for cell survival.

In our previous study, we found that tissue remodeling proteins such as KRTHB3, KRT7, KRT8, KRT17, TPM4, CRYAB, SEPW1, LGALS3BP, and VATI were overexpressed in resis‐ tant KB‐v1 cells, implying that the intracellular vesicular transport of many drugs is partly controlled by cytoskeletal filaments [117]. Research into the cytoskeleton is experiencing increased interest and rapid advancement, which will provide a greater mechanistic under‐ standing of the molecular pathways and mechanisms contributing to drug resistance and will enable the development of more patient‐tailored therapies.

### **Acknowledgements**

IIIb and IIIc species variants) (**Figure 1**). Further in‐depth study is required to determine the features of the dynamic expression and arrangement of intracellular filaments during cancer

**5. Mechanism of multidrug resistance (MDR) in relation to the**

**Figure 1.** Cellular cytoskeletons in epithelial‐mesenchymal transition process (EMT).

Many patients develop drug resistance to anticancer agents, with the mechanisms includ‐ ing alterations in the ATP‐binding cassette [94], microtubule dynamics, drug transport, and cell death [89], all of which involve tubulin and microtubules [95–97]. Microtubules have been considered a highly significant molecular target for anticancer agents, including micro‐ tubule‐stabilizing agents. For example, paclitaxel binds to the β‐tubulin subunit, accelerates the polymerization of tubulin, and stabilizes the resultant microtubules [98, 99]. Moreover, the paclitaxel‐induced resistance of vimentin intermediate filaments to okadaic acid may occur through a microtubule‐independent mechanism [100]. Townson also demonstrated that K8/18 filaments provided resistance to apoptosis in granulosa cell tumors by impairing FAS expression [101]. The organization of actin filaments associated with cellular differentia‐ tion may also influence the expression of P‐glycoprotein (P‐gp) through ezrin, radixin, and moesin in MDR osteosarcoma cells [92–104], which exhibit a significant increase in well‐orga‐ nized actin stress fibers [103], while inhibiting actin remodeling can suppress drug resistance

The balance between polymerized and nonpolymerized tubulin will be a key determinant of the response to antimitotic‐based chemotherapy. Cancer cells obtain mitotic drug resistance

invasion and migration.

304 Cytoskeleton - Structure, Dynamics, Function and Disease

**cytoskeleton**

in cancer [105].

This work was sponsored by a grant (16PJ1408800) to Dr. Jin Wang from the Shanghai Pujiang Program, a grant (15DZ2290200) from the Shanghai Science and Technology Commission, and a grant (SHDC12014104) from the Shanghai ShenKang Hospital Development Center, Shanghai, China.

### **Author details**

Xuan Zhang, Zenglin Pei, Chunxia Ji, Xiaoyan Zhang, Jianqing Xu and Jin Wang\*

\*Address all correspondence to<EMAIL_ADDRESS>
Scientific Research Center, Shanghai Public Health Clinical Center, Shanghai, P.R. China

### **References**

[1] Hardin J, Bertoni G, Kleinsmith LJ. Becker's World of the Cell (8th ed). New York: Pearson. 2015, 422–446.

[21] Evans RM. Vimentin: the conundrum of the intermediate filament gene family. Bioessays. 1998, 20(1): 79–86.

[2] McKinley M, Valerie DO, Pennefather‐O'Brien E, Harris R. Human Anatomy (4th Ed).

[3] Wickstead B, Gull K. The evolution of the cytoskeleton. J Cell Biol. 2011, 194 (4): 513–525. [4] Flaherty KM, McKay DB, Kabsch W, Holmes KC. Similarity of the three‐dimensional structures of actin and the ATPase fragment of a70‐kDa heat shock cognate protein.

[5] Bork P, Sander C, Valencia A. An ATPase domain common to prokaryotic cell cycle proteins, sugar kinases, actin, and hsp70 heat shock proteins. Proc Natl Acad Sci USA.

[6] Frankel S, Mooseker MS. The actin‐related proteins. Curr Opin Cell Biol. 1996, 8: 30–37. [7] Schafer DA, Schroer TA. Actin‐related proteins. Annu Rev Cell Dev Biol. 1999, 15: 341–363.

[9] Charras GT, Hu CK, Coughlin M, Mitchison TJ. Reassembly of contractile actin cortex

[10] Gerhardt H, Golding M, Fruttiger M, Ruhrberg C, Lundkvist A, Abramsson A, et al. VEGF guides angiogenic sprouting utilizing endothelial tip cell filopodia. J Cell Biol.

[11] Zheng JQ, Wan JJ, Poo MM. Essential role of filopodia in chemotropic turning of nerve growth cone induced by a glutamate gradient. J Neuro. 1996, 16(3): 1140–1149.

[12] Kalil K, Dent EW. Touch and go: guidance cues signal to the growth cone cytoskeleton.

[13] Blanchoin L, Boujemaa‐Paterski R, Sykes C, Plastino J. Actin dynamics, architecture,

[14] Higgs HN. Discussing the morphology of actin filaments in lamellipodia. Trends Cell

[15] Holmes KC, Popp D, Gebhard W, Kabsch W. Atomic model of the actin filament.

[16] Machesky LM, Li A. Invasive filopodia promotingmetastasis. Commun Integr Biol.

[17] Adams JC. Roles of fascin in cell adhesion and motility. Curr Opin Cell Biol. 2004, 16(5):

[18] Howard J, Hyman AA. Dynamics and mechanics of the microtubule plus end. Nature.

[19] Nicholl ID, Quinlan RA. Chaperone activity of alpha‐crystallins modulates intermedi‐

[20] Helfand BT, Chang L, Goldman RD. Intermediate filaments are dynamic and motile

elements of cellular architecture. J Cell Sci. 2004, 117(Pt2): 133–141.

and mechanics in cell motility. Physiol Rev. 2014, 94(1): 235–263.

[8] Bray D, White JG. Cortical flow in animal cells. Science. 1988, 239: 883–888.

New York: McGraw Hill Education. 2015, 29.

306 Cytoskeleton - Structure, Dynamics, Function and Disease

Proc Natl Acad Sci USA. 1991, 88: 5041–5045.

in cell blebs. J Cell Biol. 2006, 175: 477–490.

Curr Opin Neurobiol. 2005, 15(5): 521–526.

2003, 422(6933): 753–758. doi:10.1038/nature01600.

ate filament assembly. EMBO J. 1994, 13(4): 945–953.

1992, 89: 7290–7294.

2003, 161(6): 1163–1177.

Biol. 2011, 21(1): 2–4.

2010, 3(3): 263–270.

590–596.

Nature. 1990, 347(6288): 44–49.

[52] Braun T, Gautel M. Transcriptional mechanisms regulating skeletal muscle differentia‐ tion, growth and homeostasis. Nat Rev Mol Cell Biol. 2011, 12: 349–361.

[37] Stefen H, Chaichim C, Power J, Fath T. Regulation of the postsynaptic compartment of excitatory synapses by the actin cytoskeleton in health and its disruption in disease.

[38] Hotulainen P, Hoogenraad CC. Actin in dendritic spines: connecting dynamics to func‐

[39] Lamprecht R. The role of actin cytoskeleton in memory formation in amygdala. Front

[40] Comrie WA, Burkhardt JK. Action and traction: cytoskeletal control of receptor trig‐ gering at the immunological synapse. Front Immunol. 2016, 7: 68. doi: 10.3389/

[41] Treanor B, Depoil D, Gonzalez‐Granja A, Barral P, Weber M, Dushek O, et al. The mem‐ brane skeleton controls diffusion dynamics and signaling through the B cell receptor.

[42] Campi G, Varma R, Dustin ML. T cell receptor microclusters as scaffolds for signaling.

[43] Gomez TS, McCarney SD, Carrizosa E, Labno CM, Comiskey EO, Nolz JC, et al. HS1 functions as an essential actin‐regulatory adaptor protein at the immune synapse.

[44] Becart S, Altman A. SWAP‐70‐like adapter of T cells: a novel Lck‐regulated guanine nucleotide exchange factor coordinating actin cytoskeleton reorganization and Ca2+

[45] Toivola DM, Strnad P, Habtezion A, Omary MB. Intermediate filaments take the heat

[46] Herrmann H, Strelkov SV, Burkhard P, Aebi U. Intermediate filaments: Primary deter‐ minants of cell architecture and plasticity. J Clin Invest. 2009, 119: 1772–1783

[47] Yourek G, Hussain MA, Mao JJ. Cytoskeletal changes of mesenchymal stem cells dur‐

[48] Kanzaki M, Pessin JE. Insulin‐stimulated GLUT4 translocation in adipocytes is depen‐

[49] Noguchi M, Hosoda K, Fujikura J, Fujimoto M, Iwakura H, Tomita T, et al. Genetic and pharmacological inhibition of Rho‐associated kinase II enhances adipogenesis. J Biol

[50] Nobusue H, Onishi N, Shimizu T, Sugihara E, Oki Y, Sumikawa Y, et al. Regulation of MKL1 via actin cytoskeleton dynamics drives adipocyte differentiation. Nat Commun.

[51] Mundel P, Reiser J, Zúñiga Mejía Borja A, Pavenstädt H, Davidson GR, Kriz W, et al. Rearrangements of the cytoskeleton and cell contacts induce process formation during differentiation of conditionally immortalized mouse podocyte cell lines. Exp Cell Res.

dent upon cortical actin remodeling. J Biol Chem. 2001, 276: 42436–42444.

Neural Plast. 2016, 2016: 2371970. doi: 10.1155/2016/2371970.

Immunity. 2006, 24(6): 741–752. 10.1016/j.immuni.2006.03.022

signaling in T cells. Immunol Rev. 2009, 232(1): 319–333.

as stress proteins. Trends Cell Biol. 2010, 20: 79–91.

ing differentiation. ASAIO J. 2007, 53(2): 219–228.

Chem. 2007, 282(40): 29574–29583.

1997, 236(1): 248–258.

2014, 5: 3368. doi: 10.1038/ncomms4368.

tion. J Cell Biol. 2010, 189(4): 619–629.

fimmu.2016.00068. eCollection 2016.

Immunity. 2010, 32(2): 187–199.

J Exp Med. 2005, 202(8): 1031–1036.

Mol Neurosci. 2016; 9: 23.

308 Cytoskeleton - Structure, Dynamics, Function and Disease

[81] Yamasaki T, Seki N, Yamada Y, Yoshino H, Hidaka H, Chiyomaru T, et al. Tumor sup‐ pressive microRNA‐138 contributes to cell migration and invasion through its target‐ ing of vimentin in renal cell carcinoma. Int J Oncol. 2012, 41: 805–817.

[67] Machiels BM, et al. Abnormal A‐type lamin organization in a human lung carcinoma

[68] Wu Z, et al. Reduced expression of lamin A/C correlates with poor histological differen‐ tiation and prognosis in primary gastric carcinoma. J Exp Clin Cancer Res. 2009, 28: 8.

[69] Helfand BT, et al. Chromosomal regions associated with prostate cancer risk localize to lamin B deficient microdomains and exhibit reduced gene transcription. J Pathol. 2011,

[70] Kalluri R, Neilson EG. Epithelial‐mesenchymal transition and its implications for fibro‐

[71] Thiery JP, Acloque H, Huang RY, Nieto MA. Epithelial‐mesenchymal transitions in

[72] Vega FM, Fruhwirth G, Ng T, Ridley AJ. RhoA and RhoC have distinct roles in migra‐ tion and invasion by acting through different targets. J Cell Biol. 2011, 193(4): 655–665.

[73] Nakashima J, Liao F, Sparks JA, Tang Y, Blancaflor EB. The actin cytoskeleton is a sup‐ pressor of the endogenous skewing behaviour of Arabidopsis primary roots in micro‐

[74] Anderson TW, Vaughan AN, Cramer LP. Retrograde flow and myosin II activity within the leading cell edge deliver F‐actin to the lamella to seed the formation of graded polar‐ ity actomyosin II filament bundles in migrating fibroblasts. Mol Biol Cell. 2008, 19:

[75] Oyanagi J, Ogawa T, Sato H, Higashi S, Miyazaki K. Epithelial‐mesenchymal transition stimulates human cancer cells to extend microtubule‐based invasive protrusions and

[76] Gu S, Liu Y, Zhu B, Ding K, Yao TP, Chen F, et al. Loss of α‐tubulin acetylation is associated with TGF‐β‐induced epithelial‐mesenchymal transition. J Biol Chem. 2016,

[77] Sobierajska K, Wieczorek K, Ciszewski WM, Sacewicz‐Hofman I, Wawro ME, Wiktorska M, et al. β‐III tubulin modulates the behavior of Snail overexpressed during the epithelial‐to‐mesenchymal transition in colon cancer cells. Biochim Biophys Acta.

[78] Whipple RA, Matrone MA, Cho EH, Balzer EM, Vitolo MI, Yoon JR, et al. Epithelial‐ to‐mesenchymal transition promotes tubulin detyrosination and microtentacles that

[79] Li WT, Yeh TK, Song JS, Yang YN, Chen TW, Lin CH, et al. BPR0C305, an orally active microtubule‐disrupting anticancer agent. Anticancer Drugs. 2013, 24: 1047–1057. [80] Shin SY, Kim JH, Yoon H, Choi YK, Koh D, Lim Y, et al. Novel antimitotic activity of 2‐hydroxy‐4‐methoxy‐2′,3′‐benzochalcone (HymnPro) through the inhibition of tubu‐

suppresses cell growth in collagen gel. PLoS One. 2012, 7: e53209.

enhance endothelial engagement. Cancer Res. 2010, 70: 8127–8137.

lin polymerization. J Agric Food Chem. 2013, 61: 12588–12597.

cell line. Eur J Cell Biol. 1995, 67: 328–335.

310 Cytoskeleton - Structure, Dynamics, Function and Disease

sis. J Clin Invest. 2003, 112: 1776–1784.

development and disease. Cell. 2009, 139: 871–890.

gravity. Plant Biol Stuttg. 2014, 16(Suppl 1): 142–150.

226(5): 735–45.

5006–5018.

291(10): 5396–5405.

2016, 1863(9): 2221–2233.

[108] Hari M, Wang Y, Veeraraghavan S, Cabral F. Mutations in alpha‐ and beta‐tubulin that stabilize microtubules and confer resistance to colcemid and vinblastine. Mol Cancer Ther. 2003, 2: 597–605.

[96] Haber M, Burkhart CA, Regl DL, Madafiglio J, Norris MD, Horwitz SB. Altered expres‐ sion of M beta 2, the class II beta‐tubulin isotype, in a murine J774.2 cell line with a high

[97] Kavallaris M, Kuo DY, Burkhart CA, Regl DL, Norris MD, Haber M, et al. Taxol‐resis‐ tant epithelial ovarian tumors are associated with altered expression of specific beta‐

[98] Schiff PB, Fant J, Horwitz SB. Promotion of microtubule assembly in vitro by taxol.

[99] Schiff PB, Horwitz SB. Taxol stabilizes microtubules in mouse fibroblast cells. Proc Natl

[100] Vilalta PM, Zhang L, Hamm‐Alvarez SF. A novel taxol‐induced vimentin phosphoryla‐ tion and stabilization revealed by studies on stable microtubules and vimentin interme‐

[101] Trisdale SK, Schwab NM, Hou X, Davis JS, Townson DH. Molecular manipulation of keratin 8/18 intermediate filaments: modulators of FAS‐mediated death signaling in

[102] Takeshita H, Kusuzaki K, Ashihara T, Gebhardt MC, Mankin HJ, Hirasawa Y. Actin organization associated with the expression of multidrug resistant phenotype in osteo‐ sarcoma cells and the effect of actin depolymerization on drug resistance. Cancer Lett.

[103] Luciani F, Molinari A, Lozupone F, Calcabrini A, Lugini L, Stringaro A, et al. P‐glyco‐ proteinactin association through ERM family proteins: a role in P glycoprotein function

[104] Wang J, Chan JY, Fong CC, Tzang CH, Fung KP, Yang M. Transcriptional analysis of doxorubicin‐induced cytotoxicity and resistance in human hepatocellular carcinoma

[105] Kim MH, Kim J, Hong H, Lee SH, Lee JK, Jung E, et al. Actin remodeling confers BRAF inhibitor resistance to melanoma cells through YAP/TAZ activation. EMBO J. 2016,

[106] Cheung CH, Wu SY, Lee TR, Chang CY, Wu JS, Hsieh HP, Chang JY. Cancer cells acquire mitotic drug resistance properties through beta I‐tubulin mutations and alter‐ ations in the expression of beta‐tubulin isotypes. PLoS One. 2010, 5(9): e12564. doi:

[107] Giannakakou P, Gussio R, Nogales E, Downing KH, Zaharevitz D, Bollbuck B, Poy G, Sackett D, Nicolaou KC, Fojo T. A common pharmacophore for epothilone and taxanes: molecular basis for drug resistance conferred by tubulin mutations in human cancer

level of taxol resistance. J Biol Chem. 1995, 270(52): 31269–31275.

tubulin isotypes. J Clin Invest. 1997, 100(5): 1282–1293.

diate filaments. J Cell Sci. 1998, 111(Pt 13): 1841–1852.

human ovarian granulosa tumor cells. J Ovarian Res. 2016, 9: 8.

in human cells of lymphoid origin. Blood. 2002, 99(2): 641–648.

cell lines. Liver Int. 2009, 29(9): 1338–1347.

cells. Proc Natl Acad Sci USA. 2000, 97: 2904–2909.

Nature. 1979, 277(5698): 665–667.

312 Cytoskeleton - Structure, Dynamics, Function and Disease

1998, 126(1): 75–81.

35(5): 462–478.

10.1371/journal.pone.0012564.

Acad Sci USA. 1980, 77(3): 1561–1565.


**5.5. Qualification of active components**

112 Nuclear Power – Practical Aspects

springs and dampers.

integrity (Katona et al, 1999).

has been applied;

required. The soil-structure interaction was be modelled by frequency independent soil

Unique blast tests have been performed for empirical modal analysis of the dynamic behaviour of the main building structures and for the verification of the models developed (Katona et al, 1992, 1993a; Halbritter at al, 1993a). These tests provide good information

For optimal modelling of primary system responses, a coupled mechanical and structural

The selection of upgrading concept for buildings has been made iteratively. For all options of upgrading, the response and resistance of modified structure has been made and the optimal solution selected via comparison of response and strength achieved. After selection of final upgrading solution the dynamic calculations have been repeated for the modified configuration for justification of the adequacy of the upgrades and development of the floor response spectra. Latter has little importance for the reinforced concrete containment part of the main building complex, but it was essential, e.g. in the gallery buildings. The same iterative procedure has been applied in case of Reactor Coolant System upgrade, and the fixed configuration has been re-calculated for the justification of code compliance of the

The methods for evaluation of as-built capacity of structures and components (passive SCs)

The floor response spectra used for the component capacity evaluation was defined according to the design codes (see e.g. ASCE-4-86). However, in case of Class 3 SCs that failed when conservative floor response was used, the calculation was performed for the best estimate floor response spectra (FRS). The best estimate FRS has been obtained either via probabilistic method, or taking into account the inelastic energy absorption, or

In those cases when the existing supports of pipelines are modified in order to provide adequate seismic capacity, e.g. when the number or type of the supports is changed, it shall be demonstrated on the basis of the relevant nuclear standards (ASME BPVC Section III (ANSI ANS N690) or KTA 3201, 3211, 3205) that the upgraded high energy pipelines and

have been selected in accordance of safety and seismic class, as follows:

accounting the equipment-structure mass ratio.

their supports comply with the following criteria:

model has been developed (Halbritter et al, 1993b; Katona et al, 1993b, 1994, 1999).

regarding soil-structure interaction under small-strain excitation.

The qualification of active components has been made by several methods:

For example, the relays have been qualified by replacing the not to be qualified by new one, shaking table testing of samples for in-rack response spectra (Katona et al, 1995b), experience based method, where it was applicable.

Since the GIP database does not specifically include all the equipment of Paks NPP (manufactured in the Soviet Union or Eastern European countries), it was necessary to apply GIP-VVER (Masopust, 2003) incorporating the knowledge and experience gained during the evaluation of VVER type power plants.

The comparison of 1.5 times bounding spectra (BS) to the floor response spectra is always recommended instead of the comparison of bounding spectra to the ground motion response spectra even below the 12 m level of the building.


*Edited by Aise Seda Artis*

Memory is mainly the outcome of learning. And forgetting is sometimes a blessed physiological event, and sometimes part of a serious pathology. Recent findings suggest that alterations in the gut microbiome may play a pathophysiological role in human brain diseases. It has been a challenge for neuroscientists to understand the basic processes of memory storage in both health and disease conditions. Our ability to store and process what is going on and use the classified information basically relies on memory being a constructive, fallible process. Here you will find a tiny reflection of the accumulated knowledge for scientific practice. In your daily life do not forget to eat to live, live to learn, and remember to value everything just as much as you deserve.

ISBN 978-1-78985-165-6

Eat, Learn, Remember

Published in London, UK © 2019 IntechOpen © zhaojiankang / iStock


**1. Introduction**

B cell lymphoma 2 (Bcl2)-associated athanogene (BAG) family proteins are conserved across a wide range of eukaryotes including animals, yeast, and plants. BAG1, the first BAG protein to be discovered, was identified by screening for human Bcl-2-interacting proteins in a mouse embryo cDNA library [1–3]. To date, six BAG proteins (BAG1–BAG6), harboring a conserved BAG domain (BD) at the C-terminus have been reported in humans [4]. The three-dimensional structure of the BD, which contains approximately 110 amino acids (aa), comprises three α-helices, each containing 30–40 aa [5,6]. Through the BD, BAG proteins modulate the refolding activity of the heat shock-inducible heat shock protein 70 (HSP70) and the constitutively expressing heat shock cognate protein 70 (HSC70) by interacting with their ATPase domain [5]. In mammals, BAG proteins interact with members of diverse protein families to regulate various cellular phenomena such as protein degradation and cell apoptosis, migration, and proliferation [7,8].

In plants, *BAG* genes are associated with development and stress responses [9]. An advanced bioinformatics analysis revealed that the model plant *Arabidopsis thaliana* contains seven BAG proteins (AtBAG1–AtBAG7) [10,11]. Among these, AtBAG1–AtBAG4 carry

**Citation:** Kang, C.H.; Lee, J.H.; Kim, Y.-J.; Kim, C.Y.; Lee, S.I.; Hong, J.C.; Lim, C.O. Characterization of AtBAG2 as a Novel Molecular Chaperone. *Life* **2023**, *13*, 687. https://doi.org/10.3390/ life13030687

Academic Editors: Hakim Manghwar and Wajid Zaman

Received: 22 November 2022 Revised: 31 December 2022 Accepted: 31 December 2022 Published: 3 March 2023

**Copyright:** © 2023 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

an N-terminal ubiquitin-like (UBL) domain, whereas AtBAG5–AtBAG7 are plant-specific proteins harboring a calmodulin (CaM)-binding motif near the BD [10,11]. AtBAG proteins are reportedly involved in various processes related to plant development and environmental stress response. For example, AtBAG1 plays a critical role in the Hsc70-mediated proteasomal degradation of misfolded and unimported plastid proteins in the cytosol, and an optimal level of this protein is important for normal plant growth. AtBAG6, a CaM-binding protein, induces programmed cell death (PCD) in Arabidopsis and yeast, and its 134 aa stretch, which encompasses both the CaM-binding IQ motif and BD, is sufficient to induce cell death [11]. Additionally, AtBAG4, AtBAG6, and AtBAG7 are involved in the plant response to pathogen attack as well as the tolerance to cold, heat, ultraviolet (UV), and endoplasmic reticulum (ER) stresses [9,10,12]. Recently, *AtBAG2* and *AtBAG6* were reported to be involved in the tolerance to multiple abiotic stresses in Arabidopsis [13]. However, information on the biochemical and molecular functions of BAG proteins that affect plant physiology is extremely limited. In this study, we explored the intrinsic biochemical and molecular properties of AtBAG2 and characterized this protein as a novel molecular chaperone.

#### **2. Materials and Methods**

#### *2.1. Plant Material and Growth Conditions*

*Arabidopsis thaliana* ecotype Columbia (Col-0) was used as the wild type (WT) in this study. The homozygous T-DNA mutant *atbag2* (SALK\_030295; Col-0 background) was obtained from the Arabidopsis Biological Resource Center (ABRC; Ohio State University, Columbus, OH, USA). Seeds were sown in Petri dishes containing full-strength Murashige and Skoog (MS) medium (Duchefa Biochemie B.V., Haarlem, The Netherlands) supplemented with 2% (*w*/*v*) sucrose and 1% (*w*/*v*) agar, and cold-stratified in the dark at 4 ◦C for 3 days. The plates were then transferred to an environmentally controlled growth chamber maintained at 22 ◦C temperature, a 16 h light/8 h dark photoperiod, and 100 μmol m−2 s−1 light intensity. The chemicals were purchased from Sigma-Aldrich (Sigma-Aldrich Chemical, Saint Louis, MO, USA).

#### *2.2. Heat Shock Treatment and Phenotypic Analysis of Plants*

To examine the thermotolerance of the WT and *atbag2* mutant, seedlings were grown on solid nutrient medium containing 2% (*w*/*v*) sucrose, and incubated under the abovementioned conditions [14]. Plates containing 7-day-old seedlings were sealed with plastic electrical tape and submerged for 30 min in a water bath maintained at 44 ◦C. The plates were then transferred to normal growth conditions, and the thermotolerance of seedlings was determined by measuring their survival rate. To measure electrolyte leakage, the seedlings were submerged in 5 mL of deionized water and placed on a shaker at 22 ◦C. The initial conductivity of the solution was measured using an Orion 3-Star Benchtop Conductivity Meter (Thermo Electron Cooperation, Rosemount, MN, USA). The seedlings were then autoclaved for 15 min, and conductivity was measured again (final conductivity) to determine the total amount of ions in solution. Percent ion leakage was estimated as the ratio of initial conductivity to final conductivity.

#### *2.3. Yeast Strain, Survivability Assay, and TB Exclusion Assay*

The yeast (*Saccharomyces cerevisiae*) strain W303 (SVL82; *MaTα*, *ade2*, *his3*, *leu2*, *trp1*, *ura3*, *can1*) was grown in Yeast Extract-Peptone-Dextrose (YPD) medium at 27 ◦C [15]. Cells transformed with the glutathione S-transferase (GST)-only vector (control) and various GST-AtBAG2 fusions (G-AtBAG2, G-AtBAG2-Nt, G-AtBAG2-M, and G-AtBAG2-Ct) were grown in the YPD medium overnight. The cells were then transferred to fresh YPD medium and incubated at a concentration of 5 × 107 cells/mL at 27 ◦C or 55 ◦C. Cells were sampled at 0, 10, 20, 30, 40, 50, and 60 min after incubation, and suitable dilutions were plated on YPD agar plates. The number of viable cells or colony forming units (CFU) was counted after 2–3 days of incubation at 27 ◦C.

Samples of vector, *G-AtBAG2*, *G-AtBAG2-Nt*, *G-AtBAG2-M*, and *G-AtBAG2-Ct* transformants collected at the 60 min time point were subjected to trypan blue (TB) staining and spot assays. The TB staining assay was performed as described previously [16]. To conduct spot assays, the concentration of each culture was adjusted to an optical density (OD600) of 1.0. Next, 6 μL aliquots of 10-fold serial dilutions were spotted onto YPD agar plates, incubated at 27 ◦C, and examined after 2–3 days.

#### *2.4. Construction of Expression Plasmids*

The *AtBAG2* coding sequence was PCR-amplified using the following primer pair: At-BAG2(*Bam*HI)-F (5- -GGATCCTAATGATGAAAATGAGTATCGGA-3- ) and AtBAG2(*Xho*I)- R (5- -CTCGAGTTAATTGAATAATTCCCATTTA-3- ). The PCR products were cloned into the pGEM-T Easy vector (Promega, Madison, WI, USA), and the *Bam*HI/*Xho*I fragments were subcloned into the pGEX-2T vector (GE Healthcare, Piscataway, NJ, USA). To identify the amino acid sequence required for the molecular chaperone activity of AtBAG2 and the thermotolerance of yeast cells, deletion fragments of *AtBAG2* were generated by PCR using the following primers: AtBAG2(*Bam*HI)-F and AtBAG2-Nt(*Xho*I)-R (5- - CTCGAGTTAGATTGCTTTAGATGATTT-3- ) for *AtBAG2-Nt* (1–133 aa); AtBAG2-M(*Bam*HI)- F (5- -GGATCCTAATGAAAGAGAAATCATCTAAA-3- ) and AtBAG2-M(*Xho*I)-R (5- -CTCGAGTTACTGCATCTTCTTCTTCAA-3- ) for *AtBAG2-M* (131–195 aa); and AtBAG2- Ct(*Bam*HI)-F (5- -GGATCCTAATGAAGAAGAAGATGCAGAAT-3- ) and At-BAG2(*Xho*I)-R for *AtBAG2-Ct* (191–296 aa). The amplified products were cloned into the pGEM-T Easy vector, and subcloned into the pGEX-2T expression vector digested with *Bam*HI and *Xho*I (for expression in *E. coli*) or into the pYES2-GST fusion vector digested with *Bam*HI and *Xho*I (for expression in yeast).

### *2.5. Recombinant Protein Production and Purification*

The pGEX-AtBAG2, pGEX-AtBAG2-Nt, pGEX-AtBAG2-M, and pGEX-AtBAG2-Ct vectors were transformed into *Escherichia coli* BL21(DE3) pLysS cells. Recombinant protein production and purification were performed according to the procedure described previously [17]. The AtBAG2 deletion variants were further purified using the TSK heparin-5PW HPLC column (7.5 mm × 75 mm), as described previously [17,18]. DnaK, a possible co-purifying contaminant on GSH columns, was removed using the ATP-agarose column (Amersham Pharmacia Biotech, Piscataway, NJ, USA), according to the manufacturer's instructions. The purified AtBAG2 proteins were dialyzed against 20 mM HEPES (pH 8.0) before use.

#### *2.6. Protein Quantification and Chaperone Activity Assay*

Mixtures containing a given recombinant AtBAG2 protein (30 μg/mL protein in 20 mM HEPES [pH 8.0]) and 10 μM 4,4- -bis (1-anilinonaphthalene 8-sulfonate) (bis-ANS; Sigma-Aldrich) were incubated at various temperatures for 30 min. The fluorescence of each mixture was then measured using a SFM25 spectrofluorometer (Kontron, Zurich, Switzerland) at an excitation wavelength of 380 nm and emission wavelengths ranging from 400–600 nm.

The chaperone activity of recombinant proteins was measured using citrate synthase (CS), malate dehydrogenase (MDH), and insulin (Sigma-Aldrich) as substrates, as described previously [19,20]. Turbidity caused by substrate aggregation was monitored using the DU800 spectrophotometer (Beckman Coulter, Indianapolis, IN, USA) equipped with a thermostatic cell holder.

#### **3. Results**

#### *3.1. AtBAG2 Enhances Heat Shock Tolerance in Arabidopsis*

To confirm the physiological significance of *AtBAG2* in plant stress response, we selected homozygous lines of the *atbag2* T-DNA insertion mutant originally obtained from ABRC (OH, USA) (Figures 1A,B, S1 and S2). Transcript levels of *AtBAG2* were compared

between WT (Col-0) and *atbag2* mutant plants by semi-quantitative reverse transcription PCR (sqRT-PCR) (Figures 1C, S3 and S4). Although the *AtBAG2* signal was clearly detected in WT plants, it was barely detectable in the *atbag2* mutant, as expected (Figures 1C and S3). By contrast, the signal for the *Actin-2* (*ACT2*) gene (*AT3G18780*), which was used as a control, was nearly identical in both WT and *atbag2* samples (Figures 1C and S4). Next, we investigated the physiological function of *AtBAG2* in 7-day-old WT and *atbag2* seedlings under heat shock conditions (Figure 1D–F). Under normal conditions (22 ◦C), the two genotypes exhibited no difference in seed germination and seedling survival rates (Figure 1D,E). However, when incubated at 44 ◦C for 30 min, the 7-day-old *atbag2* seedlings exhibited outstanding thermosensitivity (Figure 1D,E), as indicated by their low survival rate (31.4 ± 5.6%) compared with the WT (56.6 ± 5.2%) (Figure 1E). Because heat shock causes considerable damage to the channel and transporter proteins, fluidity, and other properties of the cell membrane, resulting in electrolyte leakage [21,22], we compared ion leakage between the WT and *atbag2* plants. Following incubation at 44 ◦C for 30 min, the seedlings of both genotypes exhibited an increase in ion leakage; however, the level of ion leakage was higher in *atbag2* seedlings than in the WT (Figure 1F). The WT and *atbag2* seedlings took 53 and 37 min, respectively, to reach 50% ion leakage. Overall, these results indicate that *AtBAG2* plays a major role in plant survival under heat stress.

**Figure 1.** *AtBAG2* regulates heat stress tolerance in Arabidopsis. (**A**) Schematic representation of the predicted gene structure of *AtBAG2*. Black elbow arrows represent the start codon (ATG). Filled boxes and horizontal black lines indicate exons and introns, respectively. Empty boxes at either end of the gene represent 5 and 3 untranslated regions (UTRs). A black arrowhead indicates the T-DNA position in *atbag2*. Black arrows represent the binding sites of primers used for genotyping. (**B**,**C**) Confirmation of homozygous *atbag2* mutant lines by genomic DNA-based PCR (**B**) and sqRT-PCR (**C**). (**D**) Photographs of WT (Col-0) and *atbag2* seedlings grown at optimal temperature (22 ◦C) for 14 days (control) or subjected to heat stress conditions (7-day-old seedlings treated with heat shock [44 ◦C] for 30 min, and then grown at 22 ◦C for 7 days). (**E**,**F**) Comparison of survival rate (**E**) and ion leakage (**F**) between 7-day-old WT (Col-0) and *atbag2* seedlings subjected to the heat shock treatment, as described in (**D**). In (**E**), the survival rates of heat-stressed Col-0 and *atbag2* seedlings

(heat shock) were compared with those of unstressed plants (control). Data represent the mean ± standard deviation (SD) of at least three independent experiments. To measure the ion leakage of Col-0 (-•-) and *atbag2* (---) seedlings shown in (**F**), samples were collected at the indicated times and submerged in deionized water for 1 day. The conductivity of at least 10 seedlings was measured before autoclaving (initial conductivity) and after autoclaving (final conductivity). Data represent the mean ± SD of at least three independent experiments.

#### *3.2. AtBAG2 Overexpression Enhances Thermotolerance in Yeast*

Because *atbag2* plants were more sensitive to heat shock than WT plants, we investigated whether *AtBAG2* imparts heat stress tolerance at the cellular level by monitoring the effects of *AtBAG2* overexpression in yeast cells. To conduct this experiment, we cloned the *GST*-*AtBAG2* gene fusion under the control of the *GAL1* promoter to generate the *pYES2-G-AtBAG2* construct. We then transformed *pYES2-G-AtBAG2* and the *GST (G)*-only construct (*pYES2-G*) separately into yeast W303 cells for the conditional overexpression of recombinant proteins. To confirm the physiological significance of *AtBAG2* overexpression in yeast, we compared the viability of heat-shocked *G* and *G-AtBAG2* transformants. Cultures of the two transformants were adjusted to equal cell densities at the mid-exponential growth stage, and aliquots were incubated at 27 ◦C or 55 ◦C. Viable cell counts were measured at regular intervals (Figure 2A). The *G*- and *G-AtBAG2*-transformed cells showed no significant difference in percent survival upon incubation at 27 ◦C for up to 60 min. However, at 55 ◦C, the survival rate of *G* transformants was dramatically lower than that of *G-AtBAG2* transformants (57.6 ± 4.73% and 80.9 ± 3.97%, respectively, at 30 min; 43.3 ± 4.22% and 73.6 ± 5.37%, respectively, at 60 min). To confirm these results, we performed the TB exclusion assay [23]; in this assay, only dead cells are expected to stain blue as they cannot exclude the dye. No TB-positive *G*- and *G-AtBAG2*-transformed cells were observed at 27 ◦C after 60 min (Figure 2B). However, after a 60 min incubation at 55 ◦C, more than 50% of the *G*-transformed cells appeared to be dead (intense blue staining), while the *G-AtBAG2* transformants were alive (mild blue staining) (Figure 2B). Taken together, our results consistently demonstrated that *G-AtBAG2* overexpression resulted in a remarkable increase in the thermotolerance of yeast cells.

**Figure 2.** Ectopic expression of *AtBAG2* enhances the thermotolerance of yeast cells. (**A**) Effect of heat stress on yeast cell viability. Yeast cells transformed with pYES2-G-AtBAG2 (*G-AtBAG2*) or pYES2-G (*G*) were grown in YPD media supplemented with 2% galactose (Gal). The transformed cells (5 × 107 cells/mL) were incubated at 27 ◦C or 55 ◦C and sampled at the indicated time points to count the number of viable cells. The cell survival rate (%) of each transformant was calculated as the ratio of the viable cell count at a given time point to the viable cell count at the 0 min time point. Data represent the mean ± SD of at least three independent experiments. (**B**) TB exclusion assay. Samples of *G-AtBAG2* and *G* transformants incubated at 55 ◦C for 1 h in (**A**) were visualized by fluorescence microscopy after staining with TB. White and black arrowheads indicate TB-negative and -positive cells, respectively. Scale bars, 10 μm. Data represent the mean ± SD of at least three independent experiments.

#### *3.3. AtBAG2 Functions as a Molecular Chaperone*

Since *AtBAG2* conferred thermotolerance in both Arabidopsis and yeast, we explored the mechanism of action of the AtBAG2 protein by examining whether it exhibits molecular chaperone activity. To study the molecular chaperone activity of AtBAG2, we examined its surface hydrophobicity using bis-ANS, which binds to hydrophobic patches on proteins and fluoresces at an emission maximum of ~470 nm [24], and explored its ability to prevent the heat-induced aggregation of CS and MDH or dithiothreitol (DTT)-induced aggregation of insulin by measuring light scattering at 340 nm. The hydrophobicity of GST-fused AtBAG2 (G-AtBAG2) and its ability to prevent protein aggregation were compared with those of GST-fused Ypt1p (G-Ypt1p), a previously characterized molecular chaperone [19] (positive control), and GST alone (G; negative control). The fluorescence signal of bis-ANS-bound G-AtBAG2 was greater than that of G-Ypt1p (Figure 3A), suggesting that the chaperone activity of G-AtBAG2 is higher than that of G-Ypt1p. High surface hydrophobicity is a typical characteristic of molecular chaperones [25]. Based on these results, we investigated the chaperone activity of G-AtBAG2, along with those of G-Ypt1p (positive control) and G (negative control). In light scattering experiments, when CS (2 μM), a substrate protein [19,20], was heated alone or in the presence of 5-fold molar excess (8.35 μM) of the G protein, light scattering increased rapidly; however, when 2 μM CS was heated in the presence of 5-fold molar excess of G-AtBAG2 and G-Ypt1p, its aggregation was successively inhibited, confirming that G-AtBAG2 and G-Ypt1p function as molecular chaperones (Figure 3B). Interestingly, the aggregation prevention activity of G-AtBAG2 was greater than that of G-Ypt1p (Figure 3B). Furthermore, when MDH (1.67 μM) was heated alone or along with a two-fold molar excess (3.34 μM) of G, light scattering increased rapidly; however, when 1.67 μM MDH was heated in the presence of increasing amounts of G-AtBAG2 (0.5-, 1-, and 2-fold molar excess), its aggregation was inhibited in a concentration-dependent manner, confirming that G-AtBAG2 exhibits chaperone activity (Figure 3C). When G-AtBAG2 was present at two-fold molar excess (3.34 μM), approximately 90% of the MDH protein was protected against heat-induced aggregation during the 15 min incubation period (Figure 3C). G-AtBAG2 also inhibited the DTT-induced aggregation of insulin in a concentration-dependent manner (Figure 3D). These observations clearly demonstrate that AtBAG2 functions as a molecular chaperone.

#### *3.4. BD Is Required for the Molecular Chaperone Activity of AtBAG2*

The analysis of the protein domain structure on the InterPro website (http://www. ebi.ac.uk/interpro/) revealed that the amino acid sequence of AtBAG2 contains one UBL domain and one BD. Therefore, we investigated the roles of both of these domains in the molecular chaperone function of AtBAG2. First, we generated GST fusion constructs of three serial deletion derivatives of AtBAG2 (G-AtBAG2-Nt, G-AtBAG2-M, and G-AtBAG2- Ct) (Figure 4A). Among the three AtBAG2 deletion variants, G-AtBAG2-Nt contains the UBL domain, G-AtBAG2-M contains the BD, and G-AtBAG2-Ct contains no unique domain. The recombinant GST fusion proteins were then produced in *E. coli* and purified, and their hydrophobicities were compared with that of G-AtBAG2 using bis-ANS. Upon binding to bis-ANS, G-AtBAG2-M showed high fluorescence, as with G-AtBAG2, whereas G-AtBAG2- Nt and G-AtBAG2-Ct showed very low fluorescence (Figure 4B). We then compared the chaperone activities of G-AtBAG2-Nt, G-AtBAG2-M, and G-AtBAG2-Ct with that of G-AtBAG2 by measuring their ability to prevent the heat-induced denaturation of the substrate protein MDH. Light scattering increased rapidly when MDH (1.67 μM) was heated alone or with a 5-fold molar excess (8.35 μM) of G-AtBAG2-Nt and G-AtBAG2-Ct; however, when MDH (1.67 μM) was heated in the presence of a five-fold molar excess of G-AtBAG2 and G-AtBAG2-M, its aggregation was successively prevented (Figure 4C), confirming the molecular chaperone activity of G-AtBAG2-M and G-AtBAG2. Based on this information, we compared the effects of the overexpression of *G-AtBAG2*, *G-AtBAG2-Nt*, *G-AtBAG2-M*, and *G-AtBAG2-Ct* on the thermotolerance of yeast cells. The *G-AtBAG2-Nt*, *G-AtBAG2-M*, and *G-AtBAG2-Ct* genes were cloned under the control of the *GAL1* promoter

to conditionally overexpress the recombinant proteins, and the constructs were named as *pYES2-G-AtBAG2-Nt*, *pYES2-G-AtBAG2-M*, and *pYES2-G-AtBAG2-Ct*, respectively. These three constructs as well as *pYES2-G-AtBAG2* and *pYES2-G* were separately transformed into the W303 cells. A spot assay was then performed using the heat-shocked *G*, *G-AtBAG2*, *G-AtBAG2-Nt*, *G-AtBAG2-M*, and *G-AtBAG2-Ct* transformants grown in media supplemented with 2% galactose (Gal) and sampled at the mid-exponential stage. The heat-shocked yeast cells overexpressing *G*, *G-AtBAG2-Nt*, and *G-AtBAG2-Ct* showed little difference in viability; by comparison, those overexpressing *G-AtBAG2* and *G-AtBAG2-M* showed significantly greater viability (Figure 4D). Taken together, these results suggest that the BD is critical for the molecular chaperone activity of AtBAG2, and overexpression of the BD-containing *AtBAG2* gene sequence significantly improves the viability of yeast cells under heat stress.

**Figure 3.** AtBAG2 acts as a molecular chaperone. (**A**) Results of bis-ANS-binding assay used to investigate the effect of heat shock on the hydrophobic domains of GST alone (G), GST-AtBAG2 fusion (G-AtBAG2), and G-Ypt1p fusion (G-Ypt1p; control). The fluorescence spectra of 10 μM bis-ANS (---) and 10 μM bis-ANS plus 30 μM G-AtBAG2 (---), G-Ypt1p (--), or G (-•-) were measured at 380 nm (excitation wavelength) and 400–600 nm (emission wavelengths). (**B**–**D**) Molecular chaperone activity assay of AtBAG2 using CS (**B**), MDH (**C**), and insulin (**D**). Light scattering was monitored at 340 nm. In (**B**), 1 μM CS was incubated either alone (---) or with 2 μM G (-•-), G-Ypt1p (--), or G-AtBAG2 (---) in 50 mM HEPES buffer (pH 7.0) in a spectrophotometer cell at 43 ◦C. In (**C**), 1.67 μM MDH was incubated alone (---), with 0.84 μM (---), 1.67 μM (---), or 3.34 μM (---) G-AtBAG2, or with 3.34 μM G (-•-) in 50 mM HEPES buffer (pH 7.0) in a spectrophotometer cell at 45 ◦C. In (D), 1 μM insulin was incubated alone (---), with 0.5 μM (---), 1 μM (---), or 2 μM (---) G-AtBAG2, or with 2 μM G (-•-) in 50 mM HEPES buffer (pH 8.0) containing 10 mM DTT in a spectrophotometer cell at 25 ◦C. Data represent the mean of at least three independent experiments.

**Figure 4.** Identification of the functional domain responsible for the molecular chaperone activity of AtBAG2. (**A**) Schematic representation of AtBAG2 and its serial deletion variants (AtBAG2-Nt, -M, and -Ct). The circle and box indicate the UBL domain and BD, respectively. The amino acid positions of each deletion variant are indicated. G-AtBAG2 and G-AtBAG2-Nt, -M, and -Ct represent the four GST (G)-fusion constructs containing the indicated fragments of AtBAG2. (**B**) Results of bis-ANS-binding assays used to study the effect of heat shock on the hydrophobic domains of G-AtBAG2 and G-AtBAG2-Nt, -M, and -Ct. The samples used were 10 μM bis-ANS (---), 10 μM bis-ANS plus 30 μM G-AtBAG2 (---), G-AtBAG2-Nt (---), G-AtBAG2-M (---), and G-AtBAG2-Ct (---). (**C**) Molecular chaperone activity assay. Solutions containing either 1.67 μM MDH alone (---) or with 2.5 μM G-AtBAG2 (---), G-AtBAG2-Nt (---), G-AtBAG2-M (---), and G-AtBAG2-Ct (---) in 50 mM HEPES buffer (pH 7.0) were incubated in a spectrophotometer cell at 45 ◦C. (**D**) Yeast spot assay. Yeast cells transformed with pYES2-G-AtBAG2 (*G-AtBAG2*), pYES2-G-AtBAG2-Nt (*G-AtBAG2-Nt*), pYES2-G-AtBAG2-M (*G-AtBAG2-M*), pYES2-G-AtBAG2-Ct (*G-AtBAG2-Ct*), or pYES2-G (*G*) were grown in YPD media supplemented with 2% Gal. The transformed cells (5 × 107 cells/mL) were incubated at 55 ◦C for 1 h. Next, 6 μL aliquots of 10-fold serial dilutions of each cell suspension were spotted on YPD plates. The plates were incubated at 27 ◦C for 3 days and then photographed.

#### **4. Discussion**

The BAG family proteins, known to act as co-chaperones, share a well-conserved BD at their C-terminus, which facilitates their interaction with HSP70/HSC70 [1,5]. For example, the most representative BAG protein, human BAG1, functions as a nucleotide exchange factor (NEF) for the chaperone HSP70, and triggers the release of chaperone-bound proteins as a co-chaperone [26,27]. AtBAG1 also interacts with HSC70 via the BD [3], thus acting

as a cofactor of HSC70 for the proteasomal degradation of unimported plastid proteins. However, it is difficult to find literature on BAG protein activity. According to the results of this study, AtBAG2 performs its own biochemical function without associating with other proteins. In particular, AtBAG2 was found to function as a molecular chaperone that prevents the heat- or chemical-induced aggregation of substrate proteins. AtBAG2 exhibited superior molecular chaperone activity with CS, which has frequently been used as a general substrate in molecular chaperone studies compared with Ypt1 (Figure 3B), which was previously identified as a molecular chaperone. In addition, AtBAG2 prevented the aggregation of thermally denatured MDH and chemically denatured insulin proteins, and its activity increased in a concentration-dependent manner (Figure 3C,D). Molecular chaperones exhibit high surface hydrophobicity [19,28], and AtBAG2 was no exception (Figure 3A). Based on these results, AtBAG2 was characterized as a novel molecular chaperone in this study.

To determine the region of AtBAG2 essential for its molecular chaperone activity, three deletion variants of the protein were generated, AtBAG2-Nt (UBL domain-containing N-terminal region), AtBAG2-Ct (domain-less C-terminal region), and AtBAG2-M (BDcontaining middle region), and the activity of each variant protein was tested. Among the three derivative proteins, only AtBAG2-M exhibited molecular chaperone activity (Figure 4C), suggesting that the BD is responsible for the molecular chaperone function of AtBAG2. This suggests the possibility that the BD can act as a functionally active site in molecular chaperones, proposing new research directions and the applicability of BAG proteins in the future. In humans, the first and second helices of BAG1 BD are important for its binding to the serine/threonine kinase Raf-1 [29], while the second and third helices are known to mediate its binding to the ATPase domain of Hsc70/Hsp70 [30]. In the future, it will be interesting to find out which helices and amino acid residues in the BD of AtBAG2 are essential for the high hydrophobicity and molecular chaperone activity of the protein.

The maintenance of intracellular protein homeostasis is one of the most basic processes required for survival. Under stress conditions, intracellular proteins are often damaged and lose function, and molecular chaperones play a major role in maintaining life by preserving protein structure and function. For example, in humans, small heat shock proteins (sHsps), such as Hsp27, maintain cytoplasmic protein function and protect cells under stress conditions [31]. Ypt1p, a small GTPase in yeast, is generally responsible for intracellular trafficking; however, upon exposure to heat shock, Ypt1p undergoes structural changes which amplify its molecular chaperone function to protect cells [19]. In Arabidopsis, AtPP5 usually participates in cell signal transduction as a phosphatase; however, under thermal stress, it protects plants by acting as a molecular chaperone and preventing protein aggregation [32]. AtP3B, a ribosomal protein, exhibits high molecular chaperone activity, and its gene expression increases at high temperatures to protect Arabidopsis plants from heat-induced damage [28]. Similarly, AtBAG2 was found to exhibit excellent molecular chaperone activity in this study (Figure 3). Moreover, the *atbag2* mutant seedlings were sensitive to heat shock (Figure 1), whereas yeast cells overexpressing the *AtBAG2* gene showed enhanced survival under heat stress conditions (Figure 2), indicating that AtBAG2 enhances thermotolerance. Unfortunately, *AtBAG2* complementation lines in the *atbag2* mutant background could not be obtained, and therefore phenotypically characterized, in this study. Given that AtBAG2 enhances the tolerance to multiple abiotic stresses by regulating the expression of downstream genes involved in abscisic acid (ABA) response and stress-induced reactive oxygen species (ROS) production [13], revealing the relationship between its stress response mechanism and molecular chaperone activity is a research task that has yet to be accomplished. Our results, which identified and characterized AtBAG2 as a novel molecular chaperone, are expected to expand the research horizon for key factors such as HSP70 that regulate essential cellular functions by associating with BAG proteins.

**Supplementary Materials:** The following supporting information can be downloaded at: https:// www.mdpi.com/article/10.3390/life13030687/s1, Figure S1: The original image for the Figure 1B (top); Figure S2: The original image for the Figure 1B (bottom); Figure S3: The original image for the Figure 1C (top); Figure S4: The original image for the Figure 1C (bottom).

**Author Contributions:** Conceptualization, C.H.K. and C.O.L.; data curation, C.H.K. and J.H.L.; formal analysis, C.H.K. and J.H.L.; funding acquisition, C.H.K., J.C.H. and C.O.L.; investigation, C.H.K., J.H.L., C.Y.K. and S.I.L.; methodology, C.H.K., Y.-J.K. and S.I.L.; project administration, C.H.K., J.C.H. and C.O.L.; resources, C.Y.K., J.C.H., Y.-J.K.; software, C.H.K. and C.Y.K.; supervision, C.O.L.; validation, C.H.K. and C.Y.K.; visualization, C.H.K.; writing—original draft, C.H.K. and C.O.L.; writing—review and editing, C.H.K. and C.O.L. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was supported by a grant from the BioGreen21 Agri-Tech Innovation Program (Project PJ015862), Rural Development Administration, and by grants from the Basic Science Research Program funded by the Ministry of Education (grant numbers, 2022R1I1A1A01066231 and 2020R1A6A1A03044344) through the National Research Foundation of Korea (NRF), Republic of Korea.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Data are contained within the article.

**Conflicts of Interest:** The authors declare no conflict of interest.


*2.1. Study Area*

The Limpopo River Basin part of Botswana is ecologically and economically significant to indigenous people and surrounding communities. Our case study, Bobirwa sub-district is situated between 28◦09 10 E to 29◦21 42 E and 22◦35 17 S to 21◦35 56 S and lies entirely within the Limpopo River Basin part of Botswana. The sub-district has an altitude ranging between 590 and 886 m making it the lowest part of Botswana hence has a network of channels that drain into the Limpopo River [8]. The sub-district boundary forms the national boundary with Zimbabwe to the north-east and South Africa to the south-east where the Limpopo River marks the boundary. Figure 1 below shows the location map of Bobirwa sub-district in Botswana, the villages and settlements in Bobirwa sub-district, as well as the location of Botswana in Africa. Although local ecosystems in the Limpopo Basin are essential to livelihoods and human well-being, they are greatly threatened by adverse climatic conditions as well as anthropogenic pressure. According to the 2011 National Population and Housing Census report, the population of Bobirwa sub-district was 71,936, comprising of 34,247 males and 37,689 females from 19,213 households with an average household size of 3.74 and a population density of 5.05 people/km2 [27].

The study area is highly susceptible to droughts, erratic rainfall fluctuating well below 400 mm/year in most years [20]. The recent (2010–2016) average minimum and maximum winter temperatures were 7.1 ◦C and 24.5 ◦C while for summer they were 17.6 ◦C and 31.3 ◦C, respectively. Thus, the average winter and summer temperatures were 15.8 ◦C and 24.5 ◦C, respectively. However, summer temperatures have often exceeded 38 ◦C in the last 5 years with occurrence of heatwaves [20]. Average potential evapotranspiration of 1400 mm has been estimated in Bobirwa sub-district which reduces rainfall effectiveness. Bobirwa sub-district experiences frequent droughts (once every 2–4 years) and extreme weather events such as heatwaves, strong storms, strong winds, and flash floods; therefore, it is considered a semi-arid hotspot [20,28]. High variability in both rainfall amount and intensity and recurrence of extended droughts and dry spells is characteristic of the study area. Nonetheless, Bobirwa sub-district experiences a less severe climate than other parts of the country which allows crop and livestock production as well as supporting considerable biodiversity which supports local livelihoods through the delivery of several timber and non-timber forest products [8,29]. The main crops grown under rain-fed conditions are maize, sorghum, millet, cowpeas, groundnuts, round nuts, and watermelons. Subsistence livestock and poultry production, a mainstay of the local economy, is mainly characterized by the rearing of cattle, goats, and chickens under free-range [8].

**Figure 1.** Map of Bobirwa sub-district showing study villages.


**Effects of Low-Dose Radon Therapy Applied Under Hyperthermic Conditions (RnHT) on Inflammatory and Non-Inflammatory Degenerative Disease Conditions**

Angelika Moder, Heidi Dobias and Markus Ritter

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/51401


*Edited by Mohammadreza Aghaei*

This book includes up-to-date and detailed information on fundamental principles, measurement, modeling, and forecasting of solar radiation for technologies and applications of photovoltaic (PV) solar energy. The book also presents basic, modern, and contemporary knowledge and techniques of reliability and performance assessment for PV systems. It includes eleven chapters in four sections: "Introduction", "Fundamentals, Measurements and Modeling of Solar Radiation", "Forecasting and Characterization of Solar Radiation", and "Solar Photovoltaic Technologies and Applications". Chapters address such topics as fundamental principles, accurate measurements, and modeling of solar radiation; innovative methods for forecasting and characterization of solar radiation; PV technology and application, from conventional and emerging PV technologies to bifacial PV cells and modules and concentrator PV systems; and recent advances in reliability and performance assessment for PV systems, including outdoor characterization, degradation analysis, and PV power prediction.

ISBN 978-1-83968-858-4

Solar Radiation - Measurement, Modeling and Forecasting Techniques for Photovoltaic

Solar Energy Applications

Published in London, UK © 2022 IntechOpen © coffeekai / iStock