**Abstract:** The cell cycle is the series of events that take place in a cell, which drives it to divide and produce two new daughter cells. The typical cell cycle in eukaryotes is composed of the following phases: G1, S, G2, and M phase. Cell cycle progression is mediated by cyclin-dependent kinases (Cdks) and their regulatory cyclin subunits. However, the driving force of cell cycle progression is growth factor-initiated signaling pathways that control the activity of various Cdk–cyclin complexes. While the mechanism underlying the role of growth factor signaling in G1 phase of cell cycle progression has been largely revealed due to early extensive research, little is known regarding the function and mechanism of growth factor signaling in regulating other phases of the cell cycle, including S, G2, and M phase. In this review, we briefly discuss the process of cell cycle progression through various phases, and we focus on the role of signaling pathways activated by growth factors and their receptor (mostly receptor tyrosine kinases) in regulating cell cycle progression through various phases.

**Keywords:** cell cycle; growth factors; receptor tyrosine kinases; G1 phase; S phase; G2 phase; M phase; Ras/Erk; PI3K/Akt

#### **1. Introduction**

The cell cycle, or the cell division cycle, is the series of events that take place in a cell that drive it to divide and produce two new daughter cells. The typical cell cycle in eukaryotes is composed of four phases including the G1, S, G2, and M phase. G1, S, and G2 together are called interphase. M phase is comprised of mitosis, in which the cell's nucleus divides, and cytokinesis, in which the cell's cytoplasm divides to form two daughter cells. Mitosis and cytokinesis are tightly coupled together. Mitosis is further divided into five subphases including prophase, prometaphase, metaphase, anaphase, and telophase (Figure 1). Each phase of cell cycle progression is reliant on the proper completion of the previous cell cycle phase. A cell could also exit from cell cycle to enter G0 phase, a state of quiescence [1,2].

Cell cycle progression is mediated by cyclin-dependent kinases (Cdks) and their regulatory cyclin subunits. Cdks, such as Cdk4/6, Cdk2, and Cdk1 (also known as Cdc2) are serine/threonine kinases with a wide variety of substrates. Cdks are activated mainly by binding to their cyclin partners, whose expressions rise and fall throughout the cell cycle to mediate the temporal activation of each Cdks. Various cell cycle checkpoints exist to ensure that critical processes are engaged prior to progression to the next phase. There are three major cell cycle checkpoints, including the G1/S checkpoint (also referred as restriction point), the G2/M DNA damage checkpoint, and the spindle assembly checkpoint (SAC) [3–5].

Growth factors (GFs) are a group of proteins stimulating the growth of specific tissues. GF plays important roles in the regulation of cell division that drives cell proliferation. Each GF binds to a specific cell–surface receptor. A specific group of GF receptors possesses tyrosine kinases activity and is termed as receptor tyrosine kinases (RTKs). RTK plays

**Citation:** Wang, Z. Regulation of Cell Cycle Progression by Growth Factor-Induced Cell Signaling. *Cells* **2021**, *10*, 3327. https://doi.org/ 10.3390/cells10123327

Academic Editors: Stephen Yarwood and Ludger Hengst

Received: 9 October 2021 Accepted: 24 November 2021 Published: 26 November 2021

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**Copyright:** © 2021 by the author. 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/).

most important roles in cell cycle regulation. RTKs are classified into 20 families. The most studied RTKs in terms of cell cycle include epidermal growth factor (EGF) receptor (EGFR) family, insulin receptor family, platelet-derived growth factor (PDGF) receptor (PDGFR) family, and nerve growth factor (NGF) receptor (NGFR). GFs drive cell cycle by activating RTKs and downstream signaling pathways, which regulates cyclin-Cdk complexes [6,7].

In this review, we will briefly discuss the process of cell cycle progression through various phases and will focus on the role of GF/RTK-activated signaling cascades in regulating cell cycle.

**Figure 1.** Diagram to illustrate a complete cell cy **Figure 1.** Diagram to illustrate a complete cell cycle progression through four cell cycle phases (G1, S, G2, and M) and three major checkpoints (G1/S, G2/M, and SAC). M phase is further divided into Prophase, Prometaphase, Metaphase, Anaphase, and Telophase.

#### **2. Early History of Cell Cycle Discovery**

Cell theory was developed in the middle of the 19th century. This theory has three main components: (1) Every living organism is composed of one or more cells; (2) cells are the basic unit of life for all living organisms, and (3) cells only arise from pre-existing cells. While the first two components were the contribution of Theodor Schwann and Matthias Jakob Schleiden, the last is the contribution of German scientist and physician Rudolf Virchow. His discovery that all cells arise from pre-existing cells is the start point of cell cycle research [8–10] (Figure 2).

**Figure 2.** Timeline of major discoveries in the early cell cycle research. **Figure 2.** Timeline of major discoveries in the early cell cycle research.

At the turn of the 19th century to 20th century, the cell cycle has been the subject of intense study. The cytology of cell division is described in great detail by microscopists and embryologists, however, the underlying mechanisms driving cell division are mostly unknown. In the late 1970s and 1980s, the advancement of modern molecular biology provided means and knowledge to study the molecular mechanisms regulating cell cycle. Cell biologists, biochemists, and geneticists joined forces and demonstrated that the basic processes and control mechanisms of cell cycle are universal in eukaryotes.

In the late 19th century, early light microscopic studies recognized that cell division follows mitosis, during which cells condensed their chromosomes. Based on his observations of cell division in various stages, German biologist and a founder of cytogenetics Walther Flemming identified the sequence of chromosome movements in mitosis. Flemming's discovery was proven correct decades later by the study of live dividing cells [11]. However, the only observable morphological changes outside of mitosis is the growth of the cell size. Interphase remained a black box and recognized as one phase until the discovery that DNA synthesis occurs only in a short period during interphase [12]. This discovery split interphase into three phases: This DNA synthesis period is termed as S phase, the gap between mitosis and S phase is termed G1 phase, and the gap between S and M phases is termed as G2 phase [13].

Following the recognition of four major cell cycle states G1, S, G2, and M, the focus of cell cycle study shifted to understand the transition between these phases. A major task is to identify the factors driving the transition. In the early 1970s, by fusing cells at different stages of the cell cycle, it was shown that late G2 or M phase cells contained an M phase-promoting factor (MPF) capable of accelerating the onset of mitosis in early G2 cells [14]. It was further shown that S phase cells contains an S phase-promoting factor (SPF) in nuclei, which is able to accelerate S phase [14,15].

While there is no biochemical method available to purify either MPF or SPF at the time, genetic studies of cell cycle related genes are fruitful. At the end of the 1960s, Leland Hartwell realized the possibility of using genetic methods to study cell cycles. He established budding yeast *Saccharomyces cerevisiae* as a highly suitable model system to study cell cycles. In an elegant series of experiments in 1970–1971, he used the temperature sensitive lethal mutants of *S. cerevisiae* to isolate yeast cells with mutated genes, controlling the cell cycle. By this approach, he successfully identified more than one hundred genes which specifically involved in cell cycle control. Among these genes are genes encoding SPF and MPF. Hartwell named these genes Cdc-genes (cell division cycle genes) [16–18]. One particularly important gene identified is Cdc28, which controls the first step of cell cycle progression in G1 phase and was also known as "start".

In the middle of the 1970s, Paul Nurse followed Hartwell's approach to study cell cycle regulation with similar genetic methods but using fission yeast *Schizosaccharomyces pombe* as a model system. Through this research, Paul Nurse discovered the gene Cdc2 in fission yeast. Cdc2 is identical to Cdc28 identified in budding yeast. Nurse found that Cdc2 had a key function in the control of transition from G2 to mitosis during cell cycle [19]. In 1987, Nurse isolated the human version of Cdc2 gene, Cdk1. Cdk1 encodes a protein called cyclin-dependent kinase (Cdk). They found that phosphorylation status of the mammalian Cdc2 protein (p34Cdc2) is closely related to cell cycle progression. It is phosphorylated when cells are stimulated to enter the cell cycle in G1 phase, but dephosphorylated when cells go to quiescence [20,21]. Based on these findings, half a dozen different Cdk molecules have been found in humans.

In the early 1980s, Tim Hunt discovered the first cyclin molecule by studying sea urchins, Arbacia. There are eight very rapid cell divisions during the cleavage in embryos of the sea urchin. To sustain these cell divisions, the continual protein synthesis is required. Hunt found that one protein is always destroyed each time the cells divide. This protein was named cyclin as the level of the protein vary periodically during the cell cycle [22]. In the following years, more cyclins were identified in various species by Hunt and other

groups. Moreover, it was discovered that the cyclins bind to the Cdk molecules to regulate the Cdk activity and determine the substrate specificity of Cdks [23].

Another important concept introduced during this period is "Checkpoint". In the late 1980s, by studying the sensitivity of yeast cells to irradiation, Hartwell developed concept of checkpoint [24,25] (Figure 2). He observed that the cell cycle is arrested at certain point when DNA is damaged. This cell cycle checkpoint concept is then expanded as surveillance mechanisms used by the cells to check the integrity, fidelity, and the sequences of the major cell cycle events. The events being monitored include cell size growth, DNA replication, and integrity, and the accurate chromosome segregation [3].

The historical contribution of Leland H. Hartwell, Paul M. Nurse, and R. Timothy (Tim) Hunt earned them 2001 Nobel Prize in Physiology or Medicine for their discovery of "key regulators of the cell cycle".

#### **3. Cell Cycle Progression through Various Phases**

The cell cycle consists of G1, S, G2, and M phases. In G1 phase, the cell grows and becomes larger. The cell enters S phase when it reaches a certain size. S phase is the period for DNA-synthesis, during which the cell duplicates its DNA. In the following G2 phase, the cell monitors the completion of DNA-replication and prepares for mitosis. Chromosome segregation and cell division are completed in M phase. The proper cell cycle progression ensures that each of the two daughter cells receives identical chromosome from parent cell. After cell division, the cell cycle is completed, and the cells are back in G1 phase. The duration of the cell cycle varies between 10 and 30 h in most mammalian cells. Cells in the G1 can exit from the cell cycle and enter G0 phase, a state of quiescence.

Cell cycle progression is mainly driven and regulated by two classes of proteins, Cdks and cyclins [26] (Figure 3). In yeast, while several Cdks are expressed, including Cdk1, PHO85, and Kin28, only Cdk1 directly regulates cell cycle progression. Cdk1 is equivalent to p34Cdc2 in *S. pombe* and p34Cdc28 in *S cerevisiae*. By associating with different cellcycle stage-specific cyclins Cdk1 regulates diverse cell cycle transitions including G1 to S transition and G2 to M transition. The roles of PHO85 and Kin28 in cell-cycle regulation are indirect [26,27]. Higher organisms possess many yeast Cdk1 functional homologues. These functional homologues are phase-specific Cdks. Each phase-specific Cdk acts in a specific cell phase to perform the function of Cdk1 in yeast. Approximately 20 Cdkrelated proteins are discovered, which leads to the concept that cell cycle events in higher eukaryotic cells are regulated by complex combinations of Cdks and cyclins in various cell cycle phases. For Cdk/cyclin complexes, cyclins confer substrate specificity and determine the regulatory consequence of the substrates such as activation, inactivation, and localization. Based on this hypothesis, the classical model of cell cycle regulation is established through extensive research in eukaryotic cells.

According to this model, Cdk4 and/or Cdk6 form complexes with D-type cyclins, which activates Cdk4/6 and initiates phosphorylation of the retinoblastoma protein (Rb) family in early G1 phase [28,29]. Rb phosphorylation stimulates the release of transcription factor E2F, which then stimulates the transcription of early E2F responsive genes required for the progression of the cell cycle [30,31]. Early E2F responsive genes include A- and E-type cyclins [28,32]. In the late G1 phase, cyclin E binds to and activate Cdk2, which leads to the full Rb phosphorylation and the further activation of E2F mediated transcription [28,29]. Together, the above events drive the passage of the cell through the restriction point at the boundary of the G1/S phase and initiate the S phase. At the onset of the S phase, A-type cyclins are synthesized and form complex with Cdk2, which phosphorylates proteins involved in DNA replication and drive the cell progression to G2 phase [33,34]. At the late G2 phase, Cdk1/cyclin A is formed and activated, which is required for the G2/M transition and the initiation of prophase [35]. Finally, Cdk1/cyclin B complexes are formed in M phase and drive the completion of mitosis [36] (Figure 3).

**Figure 3.** Regulation of cell cycle progression by Cdks and Cyclins.

#### *3.1. G1 Phase*

Cells enter G1 either from the preceding M phase or from G0 phase. The transition of cells between G0 and G1 phase is determined by extracellular mitogenic signals [37,38]. G1 phase is the growth phase. The biosynthetic activities of the cell are slowed down considerably in M phase; however, it resumes at a high rate in G1 phase. In G1 phase, the cells synthesize many proteins, amplify organelles including ribosomes and mitochondria, and grow in size. The duration of cell cycle phases varies considerably in different types of cells. For a typical proliferating human cell, if we assume the total cycle time is 24 h, the duration of G1 phase is approximately 11 h, S phase duration last 8 h, G2 phase last 4 h, and the duration of M phase is approximately 1 h.

During G1 phase, diverse signals, including environmental cues, stress, and metabolic cues intervene to influence cell's developmental program. These signals are integrated and interpreted by the cells. Based on these inputs, the cell decides whether to self-renew, differentiate, or die; however, to enter S phase for starting its renewal, all cells must fulfill one essential requirement: activation of Cdks [38,39].

#### *3.2. S Phase*

S phase is marked by DNA synthesis. In S phase, each chromosome consists of two sister chromatids following replication to double the amount of DNA. However, S phase also marked with low activities of gene expression and protein synthesis. A noticeable exception is the production of histone. Most histones are produced in the S phase [40].

It is suggested that an intra-S phase checkpoint exists to control S phase progression. Intra-S phase checkpoint turns off Cdk2 in response to DNA damage and other replication stress, which blocks origin firing to avoid replication of damaged DNA [41]. S phase to G2 phase transition is regulated by the active checkpoint kinase ATR (ataxia-telangiectasia and Rad3-related) [42].

#### *3.3. G2 Phase*

The cell enters G2 phase after successful completion of S phase. G2 phase ends with the onset of mitosis. The major task of cells in G2 phase is to prepare itself for mitosis. G2 phase is marked by significant protein/lipid synthesis and cell growth [43]. While it is known that protein synthesis inhibitor arrests cells at G2 phase, a recent study suggests

that this may be due to the inhibition of p38, and the protein synthesis is not absolutely required for mitosis entry [44]. Interestingly, some cell types, including certain cancer cells and *Xenopus* embryos, lack the G2 phase. Cell cycle proceeds directly from S phase to M phase. It is hypothesized that cell size controls the growth in G2 phase, however, this is only demonstrated in fission yeast [45]. Another process that occurs during G2 phase is to repair DNA double-strand breaks. During and after DNA replication, DNA double-strand breaks accumulate in the cell and need to be repaired before cell can move to pass G2/M checkpoint [24,46,47].

#### *3.4. Mitosis and Cytokinesis*

M phase is comprised of mitosis, in which the cell's nucleus divides, and cytokinesis, in which the cell's cytoplasm divides to form two daughter cells. Mitosis is further divided into prophase, prometaphase, metaphase, anaphase, and telophase (Figure 1).

Prophase is characterized with chromatin/chromosome condensation, centrosome separation, and nuclear membrane breakdown. The migration of centrosome to two opposite poles is important for the later formation of the bipolar mitotic spindle apparatus. A recent detailed study shows that the interphase organization is rapidly lost in prophase by a condensin-dependent manner [48]. Observations with a microscope indicate that chromosomes become recognizable as linearly organized structures in early prophase [49]. Sister chromatids are mixed in early prophase, but they are separated in late prophase. Each chromatid is shown as an array of loops radiating from an axial core that contains topoisomerase II alpha and condensin complexes [50]. The rise of cyclin B-Cdk1 activity is a defining molecular event of prophase [51].

Prometaphase starts from the nuclear envelope breakdown, which marks the end of prophase, ends when chromosome alignment at the spindle equator completes, which defines the beginning of metaphase. For faithful chromosome segregation, it is essential to establish a metaphase plate in which all chromosomes aligned at the cell equator attach to mitotic spindle microtubules. The achievement of this configuration depends on the precise coordination of several mitotic events including nuclear envelope breakdown, connection between chromosome kinetochores, and microtubules of the mitotic spindle assembly, and the congression of all chromosomes to the spindle equator. A kinetochore is a disc-shaped protein structure in duplicated chromatids [52]. During prometaphase the chromatids shorten and become thicker [49] and ultimately form fully condensed metaphase chromosomes [53].

Metaphase starts when the duplicated chromosomes are aligned along the metaphase plate in the middle of the cell. During metaphase, the sister chromatids are pulled back and forth by the kinetochore microtubules until they align along the equatorial plane. The chromosome segregation process is monitored by SAC pathway to ensures that all kinetochores are attached to microtubules of the opposite poles before segregation proceeds after metaphase-to-anaphase transition. Once all the chromosomes are properly aligned and the kinetochores are correctly attached, the cohesion between sister chromatids is dissolved, leading to the migration of the separated chromatids towards opposite sides of the cell by the pulling force of spindle microtubules. The cell now enters the anaphase [54].

Anaphase involves two mechanistically distinct steps, the shortening of kinetochore microtubules and the spindle elongation in the midzone. The shortening of kinetochore microtubules causes the migration of each chromatid towards its respective pole. The disjointed sister chromatids are further separated through spindle elongation in the midzone. These two steps may be temporally divided in some organisms while occurring simultaneously in other organisms. These two steps are called anaphase A and anaphase B, respectively [55]. In human mitotic cells, anaphase B usually starts 30–50 s later than the start of the anaphase A [46,56]. During anaphase, the spindle elongates 8 µm and additional 3 µm in telophase [46,55].

Telophase follows anaphase and starts at the onset of the chromosome recondensation and the nuclear envelope reformation [47]. During telophase the duplicated chromosomes in the nucleus of a parent cell separate into two identical daughter cells. A nuclear membrane forms around each set of chromosomes to divide the nuclear DNA from the cytoplasm. Simultaneously, the chromosome decondensation begins [55].

Cytokinesis results the physical separation of the cytoplasm of a mother cell into two daughter cells [57,58]. The segregation of chromosomes and cytoplasm needs to be tightly coordinated to generate offspring with the right complement of chromosomes [59]. Cell cytokinesis is initiated in anaphase, when lower Cdk1 activity causes the reorganization of the mitotic spindle and the stabilization of microtubules. The assembly of the central spindle is the key early event, which provides the template for the midbody and contributes to division plane specification. The division plane is positioned between the two sets of segregated chromosomes. The precise position of the plane is critical to prevent segregation errors. Cytokinetic furrow ingression of the attached plasma membrane is then initiated by the contraction of the actomyosin ring, which partitions the cytoplasm into two domains of emerging daughter cells. The last step of cytokinesis is abscission [60]. Abscission is the physical separation of the plasma membrane of the two daughter cells. During abscission, cells remove the cytoskeletal structures from the intercellular bridge, followed by constriction of the cell cortex, and finally the division of the plasma membrane [61,62].

#### **4. The Regulation of Cell Cycle by GF-Initiated Signaling Pathways**

GFs play vital role in driving cell proliferation by activating RTK and the downstream signaling cascades. The aberrant activity of these signaling cascades frequently leads to cancer [63,64]. Cell proliferation can only be realized through cell division and early studies have demonstrated that GF stimulation is the driving force of cell cycle initiation and progression. However, after extensive research in the late 1990s and early 2000s, little research directly studies the role of GF/RTK in the regulation of cell cycle. While the role of GF receptor in G1 phase has been well studied, very little is known regarding their role in other phases of cell cycle.

GFs regulate diverse functions of the cells, and the effects of most GFs are mediated by RTKs (Figure 4). Signal transduction starts when a GF binds to its receptor at the cell surface, which stimulates the dimerization of RTK and the activation of its kinase. Two monomers of the RTK dimer phosphorylate each other to fully activate RTK and the phosphorylated Tyr residues in the C-terminus of the receptor become the binding sites for recruiting multiple downstream signaling molecules. The formation of the RTK-signaling protein complex initiates the activation of multiple signaling pathways including Ras/Erk, PI3K/Akt, Src/Jak/Stat, and PLC-γ1. These signaling pathways interact each other to form a signaling network. Signals from various signaling pathways are eventually integrated at the level of transcription. There are several major transcriptional nodes: (1) The p53 node mediates cytostasis and apoptosis in response to high level of mitogenic signals or DNA damage; (2) The SMAD node mediate the expression of cytostatic (arrest of cell growth and multiplication) and apoptotic factors in response to TGF-β; (3) The FOXO node also mediates cytostasis and apoptosis, but in response to oxidative stress and starvation; and (4) ID and Myc node that suppress Cdk inhibitors to favor cell proliferation. By controlling the transcription of specific genes, GF-initiated cell signaling regulates diverse cellular functions including cell migration, cell survival, cell cycle progression, and differentiation (Figure 4) [7,64].

**Figure 4.** Diagram to illustrate signaling pathways initiated by GF throug **Figure 4.** Diagram to illustrate signaling pathways initiated by GF through their membrane receptors, mostly RTKs.

β

#### *4.1. EGFR-Mediated Signaling Pathways*

As a prototypical receptor of all RTKs, EGFR signaling network has been extensively studied and very well understood [7,64]. Here, we are using EGFR as an example to illustrate the signaling pathways of GF/RTK. Following its identification in the 1970s [65], EGFR is shown to possess intrinsic kinase activity. The full-length receptor is cloned in 1984 [66]. EGFR is a single polypeptide chain transmembrane glycoprotein. The following is a brief description of the two major signaling pathways activated by EGFR that is most relevant to cell cycle progression.

#### 4.1.1. Ras/Erk Pathway

A major signaling pathway downstream of EGFR is the Ras/Erk pathway. The phosphorylated EGFR interacts with SHC/Grb2, which recruits Sos to cell membrane to activate Ras. Activated Ras then stimulates the activation of Raf. Mitogen-activated protein kinase (MEK) is phosphorylated and activated by Raf, which activates Erk. Erk activation stimulates the activation and nuclear translocation of RSK. In the nucleus, RSK stimulates the activation of transcription factors including c-Fos and SRF. On the other hand, following its activation, Erk also translocates into the nucleus where it stimulates c-Fos and Elk1 (Figure 5A) [67–73].

#### 4.1.2. PI3K/Akt Pathway

EGFR stimulates PI3K either by binding to its p85 subunit directly or indirectly by activating Ras [74,75] (Figure 5B). The function of PI3K is to generate phosphatidylinositol-3,4,5 trisphosphate (PIP3) by phosphorylating phosphatidylinositol-4,5-bisphosphate (PIP2). However, this process is reversed by the phosphatase and tensin homologue deleted on chromosome 10 (PTEN). PTEN acts as a direct antagonist of PI3K and provides important negative control over the PI3K pathway. It is also reported that PI3K directly interacts with PTEN to stimulate PTEN activity [76,77]. PIP3 interacts with pleckstrin homology (PH) domains to recruit PH-domain containing proteins to the plasma membrane. Among the recruited PH-domain containing protein is the serine threonine kinase Akt. In the plasma membrane, phosphoinositide-dependent kinase 1 (PDK-1) phosphorylates Akt Thr 308 to partially activate Akt. The following phosphorylation of Akt in Ser 473 by the

rapamycin complex 2 (mTORC2) results the full activation of Akt. Activated Akt activates multiple downstream substrates, including the forkhead box O transcription factors (FoxO), the BCL2-associated agonist of cell death (BAD), and glycogen synthase kinase 3 (GSK3), to promote cell cycle entry and cell survival. Akt also activates the small G-protein ras homologue enriched in the brain (Rheb), leading to the activation of mTORC1. mTORC1 phosphorylates the eukaryotic translation initiation factor 4E binding protein 1 (4EBP1) and the p70S6 kinase (S6K1), which promotes protein translation and protects the cell from apoptosis [78–80].

**Figure 5.** Activation of Ras/Erk (**A**) and PI3K/Akt (**B**) signal **Figure 5.** Activation of Ras/Erk (**A**) and PI3K/Akt (**B**) signaling pathways downstream of EGFR.

#### *4.2. GF-Activated Ras/Erk Pathway Activation in Cell Cycle Progression*

An earlier study indicates that Erk is rapidly phosphorylated in response to GFs including EGF and PDGF [81]. The first evidence regarding the role of Erk in cell proliferation comes from the Erk-inhibition experiments. Inhibition of Erk by various means block the GF-induced cell proliferation, which indicates that GF activation of p42mapk and p44mapk is an absolute requirement for triggering the proliferative response [82]. However, Erk1/2 activation is not sufficient to drive cells into S phase [83–85].

Accumulated evidence indicates that Erk activation promotes G1 progression with multiple mechanisms [86] (Figure 6). One mechanism is by induction of cyclin D and assembly of cyclin D–Cdk4 complexes. Expression of activated Ras in different cell types is sufficient to induce the accumulation of cyclin D1 [87–89]. As a downstream signaling protein of Ras, Erk activation is both necessary and sufficient for transcriptional induction of the Cyclin D1 gene [90]. In order for cells to enter S phase, sustained Erk activation is essential to maintain the level of cyclin D1 in G1 phase [91]. The precise mechanism that connects Erk signaling to Cyclin D1 transcription is not clear. However, it is known that the Cyclin D1 promoter contains a functional AP-1 binding site. It is also shown that Erk signaling stimulates the expression of c-Fos and c-Jun, the critical AP-1 components in

response to GFs [91,92]. Sustained Erk activation also stabilizes AP-1 proteins by stimulating the phosphorylation of their C-terminal residues [86]. The Erk pathway also regulates the cellular cyclin D1 expression level through the post-transcriptional regulation [86,93]. There is also evidence to support the role of Erk in the induction of cyclin D2 and cyclin D3 [94,95].

**Figure 6.** Regulation of cell cycle progression in G1 phase by GF-induced activation of **Figure 6.** Regulation of cell cycle progression in G1 phase by GF-induced activation of Ras/Erk signaling pathway.

The second mechanism is to stabilize transcription factor c-Myc. C-Myc regulates cell cycle progression and apoptosis [96]. Activation of Erk1 by GF strongly enhances the stability of c-Myc protein by direct phosphorylating c-Myc at Ser 62 [97]. C-Myc regulates many proteins that function directly in cell cycle control. These proteins include Cdk4 [98], cyclin D2 [99], Cdc25A [100], and p21 [99]. It is shown that both Erk activation and c-Myc expression are required to drive cells from G0 to late G1 phase [85].

Another mechanism is through the regulation of p21Cip1 and p27Kip1 expression. While p21Cip1 was initially thought to be a cyclin-dependent kinase inhibitor (CKI), further study indicates that p21Cip1 plays multiple roles in regulating Cdk activity. p21Cip1 interact with both Cdk4–cyclin D and Cdk6–cyclin D complexes under physiological conditions, which induces their kinase activation from early G1 to middle S phase [28]. In response to GFs p21Cip1 is transiently accumulated in early G1 phase by an Erk-dependent and p53-independent mechanism [101,102]. A transient activation of Erk is sufficient to induce the expression of p21Cip1 [103], which contributes to the stabilization of cyclin D/Cdk4 complexes in G1 [104]. On the other hand, p21Cip1 also inhibits the activation of Cdk2–cyclin A and Cdk2–cyclin E complexes from late G1 to S phase [105]. Thus, p21Cip1 regulates cell cycle progression by controlling the activation of various Cdks.

Ras/Erk signaling pathway also mediates the GF-induced downregulation of p27Kip1 [84,106–108]. It is possible that Erk1/2 downregulates p27Kip1 by stimulating the activation of cyclin D/Cdk4/6 complexes, which induces the degradation of p27Kip1 at the G1/S transition. It is also suggested that Erk signaling downregulates p27Kip1 expression by a Skp2- and Cdk2-independent mechanism [109]. Finally, Erk signaling also contributes indirectly to p27Kip1 regulation by stimulating the synthesis of autocrine GFs [86].

The fourth mechanism is to downregulate antiproliferative genes. As revealed by a gene profiling analysis, 173 antiproliferative genes are downregulated by an Erk-dependent mechanism during G1 phase [110]. It is further shown that to maintain decreased expression levels of antiproliferative genes, continuous activation of Erk throughout G1 is required.

While the activation of the Ras–Erk pathway stimulates cell cycle progression in G1 phase in general, it is interesting to note that some studies suggest that too strong Erk activation results reversible or permanent cell cycle arrest [86] by stimulating the expression of the Cdk inhibitor p21Cip1 [111–114].

#### *4.3. GF-Activated PI3K/Akt Pathway Activation in Cell Cycle Progression*

GFs also regulate cell cycle progression through the activation of PI3K/Akt pathway. It was shown in the early 1990s that PI3K mediated the mitogenic signals of PDGFR [115,116]. In the following years, the role of PI3K in mediating PDGF-induced cell cycle progression were revealed [117,118]. The activation of Akt by PI3K or inhibition by PTEN has been shown to play critical role in GF-induced cell cycle progression [119–121]. These studies firmly established roles of PI3K in GF induced cell cycle progression. Extensive research during this period also revealed various mechanisms by which PI3K/Akt pathway regulate in GF-induced cell cycle progression (Figure 7).

**Figure 7.** Regulation of cell cycle progression in G1 phase by GF-induced act **Figure 7.** Regulation of cell cycle progression in G1 phase by GF-induced activation of PI3K/Akt signaling pathway.

Stimulation of p21Cip1 is an important mechanism underlying PI3K/Akt regulation of cell cycle progression. The PI3K/Akt signaling is required for the accumulation of p21Cip1 in human ovarian carcinoma cells. While expression of a constitutively active mutant Akt increases the expression of p21Cip1, expression of dominant negative Akt decreases p21Cip1 expression [122]. Further research about the mechanisms underlying the PI3K/Akt pathway in regulating p21Cip1 expression indicates that Akt directly phosphorylates T145 and S146 near the c-terminus of p21Cip1. These phosphorylations stimulate DNA synthesis and Cdk activity, increasing cellular proliferation [123]. Akt also phosphorylates p21Cip1 at S146, which enhances p21Cip1 protein stability and significantly prolongs the half-life of p21Cip1. High level of p21Cip1 induces assembly and activation of p21Cip1/Cdk4/cyclin D and p21Cip1/Cdk6/cyclin D complexes [124].

The inhibition of p27Kip1 is another mechanism. The PI3K/Akt pathway decreases the expression of p27KIP1. Expression of dominant negative Akt caused transcriptional induction of p27Kip1 in mesenchymal cells, inhibiting both Cdk2 activity and DNA synthesis [125]. It is shown simultaneously by three groups that Akt directly phosphorylates

p27Kip1 on T157, which leads to the retention of p27Kip1 in the cytoplasm. Cytoplasmlocalized p27Kip1 cannot bind to nuclear Cdk2, thus cannot inhibit Cdk-2 [126–128].

The third mechanism is to stimulate cyclin D1 by inhibiting GSK-3. GSK-3 is constitutively active in unstimulated cells and phosphorylates numerous proteins including cyclin D, which triggers the degradation of Cyclin D through the ubiquitin-dependent proteolysis pathway, thus maintains cyclin D in an inactive state by reducing its expression level. Phosphorylation of GSK-3 by PI3K/Akt pathway causes the inactivation of GSK-3, thus promoting the accumulation and activation of cyclin D [129–132].

The fourth mechanism is the regulation of c-Fos. Expression of a constitutively active PI3K p110 mutant in NIH-3T3 cells induced c-Fos transcription [125]. Moreover, expression of a dominant negative Akt mutant rat mesenchymal cells decreases c-Fos transcription. As discussed above, c-Fos transcription stimulates cell proliferation. It is further shown that PI3K/Akt-induced c-Fos transcription is mediated by Elk-1 [116].

It is notable that the role of PI3K/Akt in the regulation of the cell cycle is heavily influenced by PTEN [133]. As a lipid phosphatase, PTEN antagonizes the function of phosphoinositide 3-kinase (PI3K) by converting phosphatidylinositol-3, 4, 5-trisphosphate (PIP3) to phosphatidylinositol-4, 5-biphosphate (PIP2) [134]. Thus, PTEN inactivates Akt and suppresses cell cycle progression [133].

#### *4.4. Co-Regulation of Cell Cycle Progression by PI3K/Akt Pathway and Ras/Erk Pathway*

Upon GF stimulation, the PI3K/Akt and Ras/Erk pathways co-regulate cell cycle progression by interacting closely [135]. Ras/Erk pathway and PI3K/Akt pathway interact at multiple point to coordinately regulate cell cycle progression. Many crosstalk mechanisms have been revealed between these two pathways. These include cross-activation, cross-inhibition, and pathway convergence on substrates [136] (Figure 8).

**Figure 8.** Interplay between Ras/Erk pathway and PI3K/Akt pathway in terms of cell cycle regulation. Green arrow indicates stimulation and red line indicates inhibition. Bottom box listed cell cycle related protein regulated by both signaling pathways.

First, Ras itself is an upstream activator of PI3K, thus activation of Ras leads to the activation of both Erk and PI3K [137]. In addition, Raf may activate Akt in hematopoietic cells [131].

The second mechanism is through mutual inhibition. Ras/Erk signaling may negatively regulate PI3K/Akt signaling and vice versa [138,139]. It is known that activated Erk phosphorylates GAB1, which inhibits GAB1-mediated recruitment of PI3K to the EGFR for activation [140]. It is shown that Runx2 expression relieves Erk-mediated negative regulation of EGFR and Akt [141]. It is also reported that MEK suppresses PI3K signaling by promoting membrane localization of the phosphatase PTEN [142]. Cross-inhibition

between Akt and Raf by IGF1 stimulation is also reported [143]. Akt inhibits Raf1 by phosphorylating Raf N-terminal inhibitory sites, which leads to the inhibition of Erk [144,145].

The most striking feature of the crosstalk is that both Ras/Erk pathway and PI3K/Akt pathway regulate same proteins involved in the regulation of cell cycle. As discussed above, both pathways regulate p21Cip1, p27Kip1, GSK-3, Cyclin D, and c-Fos, which control cell cycle progression [86,131].

#### **5. Two Waves of GF Signaling Drives Cell Cycle in G1 Phase**

Cell cycle progression is controlled at multiple stages. During the early time of cell cycle research, a key regulatory step identified was the restriction point in cultured cells. GF is needed to drive the cell cycle to pass the restriction point, and once the restriction point is passed, GF is no longer required for the continuation of cell cycle progression [146]. What we know now is that the restriction point is G1/S checkpoint, which represents commitment to subsequent DNA replication (S phase). As discussed above, at a restriction point a signaling activity threshold must be reached to allow the continuation of the cell cycle progression. Mitogenic signals generated from GFs/RTKs are sufficient to drive the cells pass the restriction point.

The earliest studies of the restriction point in mammalian cells indicated that different GFs function differently in regulating cell cycle [147,148]. PDGF was shown as a competence factor, and EGF and IGF-I were defined as progression factors. However, later research shows that different GF may function similarly in driving cell cycle progression [85,149]. As revealed by live imaging in a study using dual-reporter cell model, when compared with serum-free medium, insulin-like GF-I (IGF-I) enhances cell cycle entry by more than 5-fold. Similar results are also obtained when GFs such as EGF, PDGF-AA, and PDGF-BB are used [150]. Another early observation is that the continuous presence of the GF for at least 6–8 h is required for cell cycle progression to pass restrict point [151,152]. Later research finds that the continuous presence of GF could be substituted with two short pulses of GF, one at early G1 phase and one at late G1 phase [6,153]. It is revealed that even with the continuous presence of GFs, the downstream signaling proteins including Ras/Erk and PI3K show two waves of activation (Figure 9).

**Figure 9.** Two waves of GF-induced cell signaling in G1 phase drive the **Figure 9.** Two waves of GF-induced cell signaling in G1 phase drive the cell cycle progression to pass the restriction point (R).

In the continuous presence of GFs, Ras shows two waves of activation, an early transient activation and a more sustained activation approximately 4 h later [154]. Interestingly, it has been shown by earlier study that Ras activity is needed for cell cycle progression in two distinct times during the G1 phase [155,156], which corresponds to the two waves of Ras activation. In addition to functioning as a molecular switch for re-entry into the cell cycle from the G0 to G1 phase, Ras functions late in G1 to induce p27kip1 downregulation and cell cycle progression to pass the restriction point in response to EGF [157].

Similarly, GF also stimulates two waves of PI3K activation during G1 phase. It is shown that inhibition of PI3K catalytic subunit p110α (but not p110β) several hours after PDGF stimulation suppresses PDGF-dependent DNA synthesis [158]. In the mid-G1 phase, treatment with PI3K inhibitors inhibits GF-dependent cell-cycle progression and the inhibition is due to impaired degradation of p27Kip1 [159]. Indeed, a second, prolonged wave of accumulation of PI3K products is detected 3–7 h after PDGF stimulation [117]. By using an inhibition–rescue approach, this research further shows that, unlike the initial wave, the second wave of PI3K activation is required for PDGF-dependent DNA synthesis.

While prolonged presence of EGF is required to stimulate cell proliferation, we show that two short pulses of EGF are also sufficient to stimulate cell proliferation. We also show that two pulses of endosomal EGFR signaling are sufficient to stimulate cell proliferation [153,160]. The first pulse of EGFR signaling stimulates the exit from G0 into G1 phase. The second pulse, required 4–8 h later, drives the transition of cells from late G1 into S phase [153]. We also show that two waves of PDGFR signaling from endosome are sufficient to drive cell cycle progression [161].

The effects of the two short pulse of GF exposure on cell signaling network is further studied by a comprehensive transcriptomic and proteomic analyses. It is revealed that three processes are responsible for regulating restriction point crossing. In addition to activating essential metabolic enzymes, the first pulse also relieves p53-related restraining processes. The study also shows that the second pulse eliminates the suppressive action of p53 by activating the PI3K/Akt pathway. Finally, the second pulse uses the Erk-EGR1 threshold mechanism to digitize the graded external signal into an all-or-nothing decision-making obligation into the S phase [149]. Recent research shows that Ras signals mainly through Erk in G1, however, it also functions through PI3K/Akt to induce Cyclin D, driving S-phase entry [162].

Together, the findings to date highlight the importance of GF signaling for the progression of the cell cycle. These studies also demonstrated the presence of two action period for GF signaling in G1 phase. The first is in the early stage of G1 phase, which is critical for the G0/G1 transition to commit the cells to enter cell cycle, instead of going into a quiescent state. The second is right before the R-point, which is critical to drive the cells to pass the G1/S checkpoint and to start DNA synthesis.

#### **6. GF Signaling in S, G2, and M Phase of Cell Cycle**

The study regarding the role of GF-mediated signaling in other cell cycle phases are limited and controversial. The initial studies in the early 1990s to establish the role of GF-mediated cell signaling in cell cycle regulation suggests that GF-activated signaling cascades especially Ras/Erk and PI3K/Akt signaling pathways are required for the cell cycle progression from G1 to S phase by passing through restriction point [86,131]. The role of GF-activated signaling pathways in other phases only began to emerge in the late 1990s to early 2000s.

While it is proposed that GFs only regulate cell cycle progression during G0 to S interval [85], sporadic publications have emerged and shown the involvement of GFmediated cell signaling in S and G2 phase [163–165]. It is shown that the presence of hepatocyte GF in S phase induces G2 delay by sustaining the activation of Erk [164]. On the other hand, it is reported that activation of EGFR by EGF in S phase induces centrosome separation, which promotes mitotic progression and cell survival [166]. The data regarding the role of EGFR activation in G2 phase are also limited and controversial. While all data indicate that EGF stimulates the activation of MEK-Erk signaling pathways [165,167], the findings regarding the effects of activated MEK-Erk pathway are quite different.

It is reported that activation of MEK-Erk signaling pathway by okadaic acid is required for entry into M phase and cell survival [168] and activation of both PI3K/Akt and MEK/Erk pathways by FBS is required for mitotic entry [169]. However, other studies indicate that activation of MEK-Erk signaling pathway by EGFR or phorbol 12-myristate 13-acetate (PMA) in G2 phase delays the M phase entry through p21Cip1 [166]. Activation of Erk signaling by EGF or PMA also induces the G2 phase delay and the blocked exit from G2 checkpoint by destabilizing Cdc25B [170]. By following live cells in which ERK1/2 activity was inhibited through late G2 and mitosis, it is shown that, for a timely entry into mitosis, Erk activity in early G2 is essential. However, Erk activity from late G2 through mitosis does not directly affect cell cycle progression [171].

The role of GF-induced cell signaling in mitosis has been less studied and poorly understood. It is reported that while the level of EGFR expression is the same between M phase and interphase, EGFR-mediated cell signaling pathways is tightly suppressed in M phase as EGF fails to bind to EGFR with high affinity to induce EGFR dimerization [170]. It is reported that EGFR, PLC-γ1, GTPase-activating protein, and Erk2 are less phosphorylated in M phase than in interphase [168]. A further study shows that Cdc2 inhibits EGF-induced Erk activation in M phase [169]. These studies argue that inhibition of GF signaling in M phase shelters the cell from extracellular signals during cell division, which helps the cell to preserve the precious energy needed for mitotic structural changes [168–170].

However, we recently showed that in M phase, EGFR is both expressed at the same level and activated to the same level by EGF as in interphase [171]. We further show that, in mitosis, EGFR is phosphorylated at all the major tyrosine residues in the C-terminus to the level similar to that in interphase, suggesting that EGFR is fully activated. However, the fully activated EGFR regulates downstream signaling pathways differently from interphase. It selectively activates some downstream signaling pathways while avoid others. Two major differences include the activation of Akt2, not Akt1, and inability to activate Erk despite of strong activation of Ras [172]. The activation level of other signaling proteins including PLC-γ1, PI3K, Cbl, and Src are all similar between mitosis and interphase. While EGF promotes cell survival in mitosis, it does not alter mitosis progression significantly. The only effect observed is the longer mitosis with the inhibition of EGFR by inhibitor AG1478 [172,173].

By using a cell line that is defective in EGFR downregulation, and thus maintains sustained EGFR signaling, it is shown that EGF-induced activation of EGFR signaling is required in G2 phase to drive the transition of cell cycle from G2 to M phase [174].

A novel method combining quantitative time-lapse fluorescence microscopy and microinjection is developed to examine cell cycle progression without cell synchronization [175–177]. By using this method, series research analyzes the role of Ras in the regulation of cyclin D expression and cell cycle progression. The results show that cyclin D1 is induced in a Ras-dependent manner in asynchronous NIH3T3 cells from S to G2 transition during cell cycle. Interestingly, the expression of cyclin D1 is Ras independent during the next G1 phase once induced in G2 phase. It is further shown that the Rasdependent induction of cyclin D1 in the S/G2 transition is mediated by post-transcriptional mechanisms [178]. While endogenous Ras is active in all cell cycle phases, cyclin D1 is only induced during G2 phase in cycling cells, which indicates that the function of Ras is regulated by cell cycle phase [179]. Constitutively activated mutant Ras accelerates the cell cycle transition through G2/M and renders the G2/M checkpoint and SAC ineffective [180].

The downstream signaling proteins of Ras have also been shown to be involved in the cell cycle phase rather than G0–G1. For example, the MAPK kinase 1 (MAPKK1) activity in synchronized NIH 3T3 cells affects the kinetics of the cell cycle progression through both the G1 and G2 phases. Inhibition of MAPKK1 by dominant negative mutant is also found to delay progression of cells through G2. Moreover, inhibition of MAPKK1 in cells synchronized to S phase arrests the cell in G2 phase, which demonstrates a role for MAPKK1 in G2/M transition [181]. It is interesting to note that some studies suggest that too much Erk activity at the G2/M transition blocks entry in mitosis. Cells lacking VHR arrest at the G1-S and G2-M transitions of the cell cycle, which is dependent on the hyperactivation of Jnk and Erk. Moreover, this arrest is reversed by Jnk and Erk inhibition [182]. Hyperactivation of the Erk pathway due to expression of activated Ras or Raf mutants arrest cell cycle progression by promoting the accumulation of cyclindependent kinase inhibitors [86]. Wentilactone B (WB), a tetranorditerpenoid derivative induces G2/M phase arrest in human hepatoma SMMC-7721 cells via the Ras/Raf/Erk and Ras/Raf/JNK signaling pathways [183].

Inhibition of the PI3K/Akt pathway with PTEN or a PI-3 kinase inhibitor results in the cell cycle arrest in G2 phase and overexpression of constitutively active Akt kinase relieves

this inhibition [184–186]. A PI3K inhibitor shortens the IR-induced G2 arrest [187]. These findings suggest a role of PI3K/Akt pathway G2 phase of cell cycle. Further study suggests that Chk1 kinase mediates the function of Akt in G2 phase. Akt kinase activity increase in G2/M, which coincides with the fall in Chk1 kinase activity [186]. Inhibition of PI3K by inhibitor LY294002 increase Chk1 kinase activity by decreasing Akt activity. Moreover, constitutively active Akt inhibits hydroxyurea-induced activation of Chk1 activation, which relieving DNA damage-induced G2 arrest [186]. Multiple mechanisms underly the ability of Akt to inhibit Chk1. These mechanisms include phosphorylation, ubiquitination, and reduced nuclear localization [188]. It is also reported that blocking PI3K/Akt signaling prolongs progression through S/G2 [189].

By using a Time-Resolved Single-Cell Imaging method, EGF-induced signal processing in individual cells is quantitated over time, which reveals the dynamic contribution of various signaling pathways [190]. It is shown that both PI3K and Erk activity are required for initial cell cycle entry, however, only PI3K activity regulates the duration of S phase. Importantly, if PI3k activity is blocked for 10–20 h after EGF treatment, the durations of S and G2 phase is increased dramatically. A likely underlying mechanism is the phosphorylation and subcellular translocation of Cdk2 by Akt during S phase [190].

#### **7. Conclusions**

′

Mitogenic signals of GFs are mediated by their transmembrane receptors, mostly RTKs. RTK activation initiates the activation of multiple downstream signaling cascades. Both Ras/Erk signaling pathway and PI3K/Akt signaling pathway play major role in regulating cell cycle progression. While the mechanism underlying the role of GF signaling in G1 phase of cell cycle progression has been largely revealed due to early extensive research, little is known regarding the function and mechanism of GF signaling in regulating other phases of cell cycle including S, G2, and M phase. Accumulated results so far suggest that GF signaling my regulate cell cycle progression throughout the cell cycle, but further research is needed to sustain these findings and to uncover the underlying molecular mechanisms (Figure 10).

.

**Figure 10.** Regulation of cell cycle progression by GF-induced cell signaling in variou **Figure 10.** Regulation of cell cycle progression by GF-induced cell signaling in various phases of cell cycle. Question mark indicates that the data are limited and controversial.

**Author Contributions:** Z.W. is the sole author of this review. The author has read and agreed to the published version of the manuscript.

**Funding:** Our research was funded in part by grants from the Canadian Institutes of Health Research (CIHR) to Z.W.

**Acknowledgments:** This work was supported by CIHR to Z.W.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**

## *Review* **Cell Cycle, Telomeres, and Telomerase in** *Leishmania* **spp.: What Do We Know So Far?**

**Luiz H. C. Assis 1,† , Débora Andrade-Silva 1,† , Mark E. Shiburah 1 , Beatriz C. D. de Oliveira 1 , Stephany C. Paiva 1 , Bryan E. Abuchery 2 , Yete G. Ferri 2 , Veronica S. Fontes 1 , Leilane S. de Oliveira 1 , Marcelo S. da Silva 2, \* and Maria Isabel N. Cano 1, \***

**Abstract:** Leishmaniases belong to the inglorious group of neglected tropical diseases, presenting different degrees of manifestations severity. It is caused by the transmission of more than 20 species of parasites of the *Leishmania* genus. Nevertheless, the disease remains on the priority list for developing new treatments, since it affects millions in a vast geographical area, especially low-income people. Molecular biology studies are pioneers in parasitic research with the aim of discovering potential targets for drug development. Among them are the telomeres, DNA–protein structures that play an important role in the long term in cell cycle/survival. Telomeres are the physical ends of eukaryotic chromosomes. Due to their multiple interactions with different proteins that confer a likewise complex dynamic, they have emerged as objects of interest in many medical studies, including studies on leishmaniases. This review aims to gather information and elucidate what we know about the phenomena behind *Leishmania* spp. telomere maintenance and how it impacts the parasite's cell cycle.

**Keywords:** *Leishmania* spp.; leishmaniases; cell cycle; telomeres; telomerase

#### **1. Introduction**

Leishmaniases are among the poverty-related endemic diseases. They are well-known to cause a wide spectrum of clinical manifestations and their harsh incidences in East Africa, the Indian subcontinent, and Latin America, where approximately one million new diagnostics are expected yearly [1]. The disease is vector-induced and caused by more than twenty species of the *Leishmania* genus, protozoan parasites that belong to the Trypanosomatidae family [1]. The invertebrate host is a phlebotomine insect that is infected during a blood meal with amastigote forms. Inside the insect digestive system, amastigotes transform into procyclic promastigotes, which are noninfective but highly proliferative forms. Procyclics eventually migrate to the proboscis and differentiate into infective metacyclic promastigote forms [2,3]. The transmission to humans occurs when a preinfected insect (female phlebotomines) regurgitates during its blood meal infective but nonproliferative forms (metacyclic promastigotes) into the mammalian host skin. Afterward, metacyclics are phagocytosed by neutrophils or macrophages, and further inside the phagolysosomes, the parasites undergo a series of morphogenetic modifications, leading to the formation of amastigotes. The amastigotes then multiply and reach the bloodstream, causing the initiation of clinical manifestations. A new cycle of infection can occur when infected macrophages are ingested by other female phlebotomines (Figure 1).

**Citation:** Assis, L.H.C.; Andrade-Silva, D.; Shiburah, M.E.; de Oliveira, B.C.D.; Paiva, S.C.; Abuchery, B.E.; Ferri, Y.G.; Fontes, V.S.; de Oliveira, L.S.; da Silva, M.S.; et al. Cell Cycle, Telomeres, and Telomerase in *Leishmania* spp.: What Do We Know So Far? *Cells* **2021**, *10*, 3195. https:// doi.org/10.3390/cells10113195

Academic Editor: Zhixiang Wang

Received: 16 October 2021 Accepted: 14 November 2021 Published: 16 November 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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/).

**Figure 1.** Schematic illustration of the *Leishmania* spp. infective cycle showing different parasite stages of development. Phlebotomine female sandflies get infected with amastigote forms during bloodmeals (1). Amastigotes transform into procyclic promastigotes, which will proliferate inside the invertebrate midgut (2 and 3). Promastigotes then migrate to the sandflies' pharyngeal valve (4) and differentiate into metacyclic forms. The metacyclic forms are transferred to the mammalian hosts' bloodstream during a new blood meal and infect macrophages and other cells from the mononuclear phagocyte system (5). Inside the macrophages, there is a metacyclic change into amastigote forms, which multiply, lyse the macrophage, and reinfect new macrophages (6 and 7). In a new cycle of infection, the infected macrophages are ingested by new phlebotomines (8). The silhouettes of man and dog and the sandfly clipart are free for use and were withdrawn from the websites HiClipart (https://www.hiclipart.com, accessed on 8 November 2021) and Gratispng (https://www.gratispng.com, accessed on 8 November 2021), respectively.

> Different species of *Leishmania* associated with the host biology and vector factors can lead to different clinical symptoms, ranging from light and self-cured cutaneous lesions (e.g., *Leishmania major*) to life-threatening visceral complications (e.g., *Leishmania infantum*) [2]. The disease is endemic in more than 60 countries, with East Africa and the Indian subcontinent remarkably impacted. Although a significant decrease in the number of cases has been reported in the past years, mainly due to efforts in vector control, leishmaniasis is still on the top priority list for developing new treatments. Such a scenario is typified, because toxic antiparasitic drugs (e.g., antimonial pentavalent and amphotericin B) are still in use nowadays. Additionally, other therapeutic approaches such as vaccination have been shown to be not adequate in eradicating the disease [4]. Thus, there is urgency for new specific and efficient treatments against leishmaniasis. For this purpose, several studies on *Leishmania* spp. have been conducted to elucidate the different aspects of parasite biology, including genetic studies and full sequencing of the parasite genome [4–6]. Among these studies, trypanosomatid telomere biology has awakened great interest in the scientific community, as telomeres are essential for genome stability and cell cycle progression. Furthermore, some components involved in parasite telomere maintenance, such as the telomerase RNA component and members of the shelterin-like telomeric complex (e.g., TRF, RPA-1, and Rbp38), are unique and present parasite-specific features relative to their hosts [7–14]. Therefore, new advances in understanding and exploring the parasite telomeric environment may reveal how these and other unknown parasite factors could be used as potential and specific targets for drug development [8–19].

Telomeres are the physical ends of eukaryotic linear chromosomes. Structurally, they are an association between tandemly repeated noncoding DNA sequences and nucleoproteins forming complexes at the end of chromosomes [20]. Telomeric DNA is composed of a double-stranded sequence (one of them rich in cytosine, the other in guanine) and a G-rich single strand that protrudes toward the ends of the chromosome, known as the 3 ′ overhang [20]. In humans and other vertebrates, the repeated telomeric sequence is 5 ′ -TTAGGG-3′ , approximately 3–15 kb in length and associated with a six-telomeric protein complex called shelterin [21–25]. These arrangements of DNA and proteins influence cell homeostasis. They are crucial to cell cycle maintenance, being decisive in important cellular processes such as cell aging, genome integrity, and maintenance of the nuclear arrangement [26–28]. It is known that, lengthwise, telomeres tend to shorten after each cell division due to the inability of DNA polymerase to complete replications in the lagging strand of linear chromosomes [29], culminating in the progressive loss of telomere repeats. Thus, telomeres act as a molecular clock. If its size reaches critical levels, it can lead to early/unprogrammed cell senescence or even activate local DNA damage repair, eventually triggering a mitotic catastrophe [14,30–32]. In most organisms, including *Leishmania* spp., telomeric DNA is elongated by telomerase. This specialized reverse transcriptase forms a ribonucleoprotein (RPN) complex [31], whose function is strictly regulated throughout the cell cycle [17,21,32]. In that sense, telomerase, telomeres, and their associated proteins are now recognized as potential drug targets [33]. In the face of the critical medical relevance of leishmaniases, peculiarities associated with the cell cycle, telomeres dynamics, and telomerase have reached the mainstream studies on parasite biology. This review aims to shed light on what we know about the phenomena behind telomere maintenance and how it impacts parasites' cell cycle and survival.

#### **2.** *Leishmania* **spp. Cell Cycle**

The cell cycle comprises a repeating series of events encompassing growth, DNA synthesis, and cell division. For most eukaryotes (including *Leishmania* spp.), the cell cycle follows a single pattern of organization consisting of the following phases: G1 (gap 1), S (DNA synthesis), G2 (gap 2), M (mitosis), and C (cytokinesis). In addition, the G0 state is sometimes included as part of the cell cycle. However, it is noteworthy that cells in the G0 state are not stimulated to proliferate [34]. In other words, cells in the G0 state are not yet committed to genome replication and cell division. Interestingly, the infective forms of most parasitic microorganisms are at the G0 state (e.g., metacyclic promastigotes of *Leishmania* spp.) [35,36] (Figure 2), which makes us wonder if, in these organisms, proliferation and infection are mutually exclusive events.

In *Leishmania* spp., as in most trypanosomatids, the G1 phase corresponds to the larger proportion of the cell cycle, while the other phases vary slightly in duration [42,43]. No studies have evidenced the metabolic changes and main events during the G1 phase in *Leishmania* spp. However, compared with the other trypanosomatids, we can infer that, in the G1 phase, there is an increase in the transcription rate and intense protein synthesis of the factors related to DNA replication [44,45]. Moreover, in the G1 phase occurs the establishment of a divergent prereplication protein complex at specific sites on the chromosomes called replication origins, which can give rise to a replication bubble [45].

The firing of replication origins starts the S phase, which briefly consists of the reliable replication of DNA molecules. Usually, eukaryotes have many replication origins per chromosome. However, in *Leishmania* spp., the number of replication origins per chromosome is an issue that generates debate. Six years ago, Marques et al., (2015) [46] used a marker frequency analysis coupled with deep sequencing (MFA-seq) to evidence that *L. major* replicates each of its chromosomes during the S phase using a single replication origin. However, in a more recent study, da Silva et al., (2020) [47] used mathematical equations to reveal that it is improbable that *L. major* uses only a single origin per chromosome during the S phase. The explanation for these discrepancies is that the MFA-seq approach is not sensitive enough to identify all replication origins [47]. Nevertheless, further sensitive

assays are needed to establish how many origins are used per chromosome during the S phase in *Leishmania* spp.

**Figure 2.** Scheme showing the two distinct patterns of kinetoplast segregation relative to the nucleus in promastigotes of *Leishmania* spp. throughout the cell cycle. Most *Leishmania* species presents one of the two kinetoplast segregation patterns presented: kinetoplast segregates after the nucleus (light red) or kinetoplast segregates before the nucleus (light blue). For instance, *L. mexicana* segregates its kinetoplast predominantly after the nucleus [37], while *L. major* and *L. tarentolae* do the opposite [38,39]. However, *L. donovani* and *L. amazonensis* exhibit these two patterns distributed in the population [40,41].

In model eukaryotes, the G2 phase is characterized by the duplication of centrioles and other cytoplasmic organelles [48]. In trypanosomatids, homologs of the proteins described in model eukaryotes involved with centriole biogenesis are associated with the basal body and flagellum biogenesis [49,50]. Furthermore, based on studies with other organisms [51], we can infer that, in the G2 phase, *Leishmania* spp. increase the rate of transcription and resumption of intense protein synthesis, which are necessary for the completion of cell division. This entire process results in an increase in cell volume and size [37,40].

During the M phase, *Leishmania* spp. and all other trypanosomatids do not disassemble their nuclear envelope and perform a closed mitosis process, which is organized by spindle pole body-like structures [52]. Moreover, due to the absence of the N-terminal portion and globular domain of histone H1 and the absence of phosphorylation on serine 10 of histone H3 (H3S10), *Leishmania* spp. and other trypanosomatids are unable to condensate their chromosomes into 30-nm fibers [53,54].

In mammals, a curious feature worth being highlighted is that mitosis and cytokinesis overlap, since cytokinesis begins before the mitotic chromosome segregation is complete. However, *Leishmania* spp. seem not to strictly follow this premise. Once mitosis ends, *Leishmania* spp. undergo a rapid remodeling in shape, first growing in length and then in width prior to cytokinesis, which ends cell division [52,55]. Although it is challenging, all these peculiarities related to the cell cycle phases may provide new routes toward the search for suitable targets for parasite cell cycle interventions aiming at their elimination.

A set of events that deserve being highlighted during the *Leishmania* spp. cell cycles refers to the replication and segregation of the kinetoplast. The coordination of these events throughout the *Leishmania* cell cycle does not follow those equivalents in model eukaryotes, where mitochondrial DNA replicates at any cell cycle stage [56,57]. The nuclear and kinetoplast S phase occurs almost simultaneously, but the effective segregation of these organelles can occur at different periods according to the species analyzed. Many studies have established a pattern of segregation for the kinetoplast relative to the nucleus in some species of *Leishmania* [37–41].

One of these studies characterized the main morphological events of the cell cycle of *L. mexicana* promastigotes [37]. Wheeler et al., (2011) [37] described the cell cycle phase durations and established a duplication order for the kinetoplast and nucleus. Two years later, da Silva et al., (2013) [40] characterized the length of the *L. amazonensis* cell cycle phases and evidenced that the promastigote forms present two distinct modes of kinetoplast segregation relative to the nucleus, following a distinct temporal order in different proportions of the cells. Using DAPI staining and EdU (5-ethynyl-2′ -deoxyuridine) to monitor, respectively, the segregation of DNA-containing organelles and DNA replication, the authors found a curious feature: 65% of the dividing promastigotes duplicate the kinetoplast before the nucleus, and the remaining 35% do the opposite or duplicate both organelles concomitantly. This finding corroborates another study carried out with *L. donovani* promastigotes, where the authors found that about 80% of the cells divide the nucleus before kinetoplast, and the remaining 20% do the opposite [41]. In other words, *L. donovani* and *L. amazonensis* exhibit a nonfixed pattern of nucleus and kinetoplast segregation. In fact, when we compare the cell cycle among different *Leishmania* spp., the order and timing of the kinetoplast and nucleus division are not consensual and cannot be generalized [37–41].

For instance, *L. mexicana*, *L. major*, and *L. tarentolae* exhibit fixed patterns of kinetoplast and nucleus segregation. However, *L. mexicana* segregates its kinetoplast predominantly after the nucleus [37], while *L. major* and *L. tarentolae* do the opposite [39,41] (Figure 2). A possible explanation for these different behaviors resides in the fact that, although belonging to the same genus, these parasites show considerable phylogenetic distance [58]. In other words, this phylogenetic divergence may reflect possible species-specific differences relative to kinetoplast segregation, suggesting that some *Leishmania* spp. have less stringent control over the order of division of their DNA-containing organelles (nucleus and kinetoplast). More studies are needed to uncover the potential players involved in controlling cell division and organelle segregation, since some of them could be explored for precise interventions related to the parasite cell cycle.

#### **3.** *Leishmania* **spp. Telomeres**

In *Leishmania* spp., telomeres constitute short noncoding repetitive sequences of 5′ TTAGGG 3′ [16,59], except for *Leishmania braziliensis,* which, in addition to the conventional sequence, is also observed the presence of 5′ -CCTAACCCGTGGA-3′ sequences at the ends of some chromosomes [60]. While, in *L. amazonensis*, the 3′ G overhang has an approximate size of 12 nt long (5′ -GTTAGGGTTAGG-3′ ), in *L. donovani* and *L. major*, the 3′G overhang is a 9-nt sequence composed of 5′ GGTTAGGGT 3′ [61]. The studies performed by Genest & Borst [62] described that the length of the *L. tarentolae* and *L. major* telomeres increase over time by approximately 1 bp per population doubling. Similar results were obtained by Oliveira et al., (2021) [15], where telomeres of the *L. amazonensis* procyclic promastigotes increased during continuous passages. It was also observed that amastigote telomeres (Figure 1, numbers 1, 6, and 7) are shorter compared to procyclic (Figure 1, number 3) and metacyclic promastigotes (Figure 1, numbers 4 and 5).

Near telomeres, there are the subtelomeric sequences, which, in *Leishmania*, are composed of 100-bp-long conserved telomere-associated sequences (LCTAS) that contain two conserved domains, the conserved sequence block 1 (CSB1) and conserved sequence block 2 (CSB2) [60]. These sequences have a high degree of conservation among the different *Leishmania* species, which may indicate their importance in chromosome segregation or as binding sites for telomeric proteins [60]. However, despite the CSB sequences and depending on the species, LCTAS also contains some nonconserved sequences and a high degree of polymorphism [59,60,63,64]. Furthermore, telomeres are commonly associated with proteins that interact directly with both telomeric strands and other telomeric pro-

teins that can influence the telomere size by regulating the telomerase access, providing chromosome stability and protecting telomeres from degradation [65]. So far, within the *Leishmania* genus, the *L. amazonensis* telomeric complex (Figure 3) is the best-characterized.

′ ′

′ ′ ′

′ ′

′

′ **Figure 3.** *Leishmania* spp. telomeres are formed by subtelomeric regions (LCTAS); a double-stranded region; a singlestranded protrusion (3′ -G overhang); and associated proteins, such as TBP-1, TRF, Rbp38, RPA-1, and LCalA. Of note, it is not clear if TBP1 binds DNA as a monomer or dimer. The telomeres are elongated by telomerase, a ribonucleoprotein composed minimally by TERT and TER. From the C-rich strand of the telomeric region, ncRNAs called TERRAs are transcribed and appear to be involved in telomere length regulation. Adapted from da Silva et al., 2012 [66].

> Lira et al., (2007) [67] described a 45-kDa telomere-binding protein named LaTBP1 as a *Leishmania* double-stranded DNA-binding protein that interacts with GT-rich and telomeric DNA sequences. LaTBP1 has at least a central Myb-like DNA-binding domain containing a conserved hydrophobic cavity involved in DNA binding, a feature conserved among the proteins that bind the telomeric double-stranded DNA [67]. However, it is unclear if LaTBP1 binds DNA as a monomer like RAP1 (Repressor Activator Protein 1) in mammals or as a dimer like the TRFs. It is worth highlighting that the LaTBP1 Myb-like DNA-binding domain is related closer to the two centrally located RAP1 Myb domains than the single C-terminal TRF (Telomere Repeat-binding Factor) Myb domain [67].

> Later, a TRF homolog was described in *L. amazonensis* (LaTRF) by da Silva et al., 2010 [68]. This protein presents 82.5 kDa, shares structural features with mammalian TRF1 and TRF2, and is highly similar to *Trypanosoma brucei* TRF. It also contains a Myb-like DNA-binding domain that allows it to bind double-stranded telomeric DNA. Apparently, LaTRF is little abundant, and it is localized in the nucleus [68].

> Some proteins that bind single-stranded telomeric DNA were identified in *L. amazonensis* [69]. RNA-binding protein 38 (LaRbp38) is one of those. It is a 38-kDa protein with a noncanonical binding site with an affinity to both single- and double-stranded G-rich telomeric DNA and GT-rich kinetoplast DNA (kDNA) [11,70]. LaRbp38 has dual cellular localization, and in synchronized *Leishmania* promastigotes, it shuttles from the mitochondria to the nucleus at the late S and G2 phases via importin alpha [11]. LaRbp38 was first identified in a telomerase-positive extract together with RPA-1, LCalA, and DNA polymerase alpha [69]. The LaRbp38 ability to bind telomeres and kDNA, and the fact that this protein is involved with kDNA stability and replication, suggest that it may also be involved with telomere elongation. However, further assays are necessary to check the veracity of this hypothesis.

Replication protein A1 (LaRPA-1) was another protein pulled down with LaRbp38 [71]. LaRPA-1 is a nuclear protein of 51 kDa that presents a single N-terminal OB-fold domain. This protein binds the telomeric G-rich ssDNA, protecting it from 3′ -5′ exonucleolytic degradation. Thus, LaRPA1 is considered a potential trypanosomatid telomere end-binding protein (TEBP) that shares structural and functional features with other TEBPs described in model eukaryotes. It binds at least one telomeric repeat using an OB-fold domain, protects telomeres from exonuclease degradation, and unfolds the telomeric Gquadruplex [12,71,72]. The G-quadruplex structure is known to impair telomerase activity in humans [73,74]. Although not tested against *Leishmania* spp., drugs that specifically bind these structures could be a new source for antiparasitic therapies.

LCalA (MW 16 kDa) was the third protein identified in the parasite telomerasepositive extracts [75]. It is the first reported protozoa calmodulin-like nuclear protein that binds in vivo to the G-rich telomeric strand and the 3′ G overhang. The binding of LCalA to telomeres is calcium-dependent. Biophysical assays showed structural changes in LCalA in the presence of calcium ions, increasing the affinity of this protein to telomeres. Additionally, LCalA partially colocalizes with telomeres throughout the parasite's cell cycle. LCalA resembles human KIP1 protein, since both share with calmodulins two EF hand domains and the affinity for calcium. Furthermore, KIP1 was shown to be involved with telomere homeostasis by its interactions with the telomerase TERT component and with shelterin member TRF2 through the EF hand domains. Whether LCalA is a functional homolog of KIP1 is a question that remains open.

Long noncoding RNAs called TERRA (telomeric repeat-containing RNA) were also described at *Leishmania* telomeres. TERRA is transcribed from the C-rich subtelomeric strand towards the end of chromosomes. Its main function is to regulate the telomere length [76]. In *L. major*, TERRA was shown to be polyadenylated and processed by transsplicing [77]. It was also observed that the number of TERRA transcripts was higher in the infective forms of the parasite (metacyclic promastigotes) relative to the procyclic promastigote and amastigote forms [77]. Moreover, Morea et al., (2021) [77] showed that TERRA could form R-loops, suggesting that *L. major* TERRA is engaged with various cellular processes, including telomere maintenance and the regulation of telomere lengths.

Another interesting finding related to *Leishmania* telomeres is the presence of base J (β-d-glucosyl-hydroxymethyluracil). Base J is a modified thymine that can interfere with RNA polymerase II function, and its presence abrogates DNA cleavage by restriction endonucleases [78]. This hypermodified DNA base was first described in the bloodstream form of *T. brucei* [79,80]. Later, base J was characterized in many other trypanosomatids, and in most of them, except the *Leishmania* spp., it was found spread out in different chromosome regions [81,82]. In *Leishmania* spp., ~98% of base J is found in telomeres [62], suggesting that it can be related to telomere function [83]. Furthermore, the recent results of Morea et al. (2021) [77] suggest that differences in the telomeric base J levels may control TERRA transcription in the *L. major* developmental forms and during continuous in vitro passages.

#### **4.** *Leishmania* **spp. Telomerase**

#### *4.1. Structure and Function*

Telomerase is the ribonucleoprotein responsible for telomere elongation. The enzyme adds 5′ -TTAGGG-3′ repetitions at the single-stranded 3′G overhangs of telomeres [83]. Thus, telomerase is thought to be essential to the regulation of telomere lengths and maintenance of genomic stability. Telomerase presents two main components, protein Telomerase Reverse Transcriptase (TERT) and Telomerase RNA (TER), minimally required for enzyme activity in vitro [84,85]. TERT is the catalytic component, and TER contains the template used by TERT to synthesize telomeric repeats. These two components form a complex with accessory proteins and are necessary for in vivo biogenesis, enzyme activity, and nucleotide addition processivity [13,32,86]. *Leishmania* spp. telomerase activity was first reported by Cano et al., (1999) [87], and enzyme purification and biochemical characterization were

further described by Giardini et al., 2011 [88]. Parasite enzyme activity was detected in all three parasite life forms presenting the canonical properties of other telomerases. However, the catalysis was shown to be temperature and life-stage dependent [15]. The genes encoding the *Leishmania* spp. TERT was characterized by Giardini et al., (2006) [89], and the TER component was described later by Vasconcelos et al., 2014 [7].

In trypanosomatids, as in most eukaryotes, the TERT component is structurally composed of four domains [13]: The Telomerase Essential N-terminal (TEN), Telomerase RNA-Binding Domain (TRBD), Reverse Transcriptase domain (RT), and the C-terminal extension (CTE). The TEN domain is involved in telomerase recruitment to the telomeres and enzyme processivity [90]. The TRBD interacts with TER and is connected to the TEN domain by an unstructured linker and creates the RNA-binding pocket that binds single stranded and paired RNA [91]. Both domains can also interact with proteins that stabilize the complex and help to recruit telomerase to telomeres and regulate enzyme activity [92–97]. The RT is the catalytic core of the enzyme, which interacts with TER through the pseudoknot region [98], and it is involved in the interactions with hybrid RNA–DNA. The CTE domain stabilizes the RNA–DNA duplex, and differently from the other three domains, it is less conserved among different species [13]. *Leishmania* TERT preserves all the canonical domains found in other TERTs but shows some amino acid substitutions that are specific to the genus [89]. The knockout of *Leishmania* TERT seems to be very harmful to the parasite, because it induces a gradual decrease in cell density in the culture, apparent during G1/G0 cell cycle arrest, morphological alterations, and telomere shortening (unpublished data).

The RNA component TER varies in length and sequence, presenting a conserved secondary structure in most eukaryotes. Variations of the TER size and sequence are observed among different organisms, and they are more prominent than TERT [99], which is conserved even among different taxa. In *Leishmania* spp., TER (LeishTER) is about ~2100 nucleotides long, and the mature molecule modified by trans-splicing contains a 5 ′ cap, a spliced leader sequence (SL), and a 3′ polyA tail. It also presents a C/D box snoRNA domain found in other TER [7]. LeishTER is expressed at similar levels in its procyclic and metacyclic promastigote forms. The mature molecule coimmunoprecipitates and colocalizes with the TERT component in a cell cycle-dependent manner. Its secondary structure prediction shows the template sequence (5′→3 ′ ) in a single-stranded form localized near the 5′ end of the RNA molecule. LeishTER also presents a conserved TBE (Template Boundary Element) motif C[U/C]GUCA in helix II, which is responsible for interacting with the TERT TRB domain [7]. Its double knockout led to partial cell cycle arrest and increased apoptosis in procyclic promastigotes. TER KO also triggers a progressive telomere shortening during continuous parasite passages (unpublished data). A similar effect was observed in *T. brucei* TER knockouts [100].

#### *4.2. Biogenesis and Mechanisms*

The biogenesis of the telomerase ribonucleoprotein complex is an intricate process involving the assembly of proteins and TER with the subcellular localization of the ribonucleoprotein. Besides TERT and accessory proteins, other proteins interact transiently with telomerase. These other proteins are important players in maturation, stability, and subcellular localization [101,102].

The telomerase biogenesis starts with the transcription of TERT and TER mRNAs by RNA polymerase II. While TERT mRNA is addressed to the cytoplasm to be translated, the TER remains in the nucleus and is assembled with the accessory proteins at the 3 ′ end—specifically, at the C/D box motif [7]. Unfortunately, no 3′ end-binding protein has yet been identified in *Leishmania* spp. Still, considering the other trypanosomatids, it is known that the C/D box motif of *T. brucei* TER interacts with SNU13, a protein known to interact with the C/D box in other eukaryotes. Together with SNU13, in humans, three other proteins recognize and bind to this RNA motif: fibrillarin, NOP56, and NOP58. However, the evidence for homologs of these proteins is still missing in these parasites.

In the case of TERT mRNA, in most organisms, after translation, the protein migrates from the cytoplasm to the nucleus, where the telomerase is assembled. In humans, the main proteins involved in TERT traffic are HSP90, its cochaperone p23, and two AAA<sup>+</sup> ATPases (Pontin and Reptin) [101–103]. However, a complete understanding of the TERT traffic in *Leishmania* spp. remains to be determined. In these parasites, the HSP90 ortholog is the protein HSP83. Recently, Oliveira et al., (2021) [15] revealed an interaction with the TERT component and the importance of HSP83 for telomerase activity and telomere maintenance in *L. amazonensis*. These findings suggest that *Leishmania* spp. TERT traffic to the nucleus may resemble the human telomerase process. Furthermore, in this study, the HSP90 inhibitor 17AAG affected both the *L. amazonensis* telomere length and telomerase activity using a thousand-times less of the drug needed to inhibit HSP90 in host cells [104].

Considering the assemblance of human telomerase as an example, the location of telomerase in Cajal bodies is the key to ribonucleoprotein trafficking and recruitment to the telomeres [101,102]. This subcellular localization is derived by the CAB box sequence at the 3′ end of human TER and its interaction with the TCAB1 protein. Otherwise, in the context of *Leishmania*, nothing is known about the existence of Cajal bodies [105]. The closest information is related to the identification of the MTAP protein, a TCAB1 protein ortholog of *T. brucei* [106] that can interact with TER [107]. Therefore, the biogenesis of telomerase in *Leishmania* spp. still has many gaps to be replenished.

Once the telomerase RNP complex is assembled, it is recruited preferentially to the shortest telomeres. The ability to add more than one repeat at one telomere is called repeat addition processivity (RAP) [108]. In humans, telomerase activity is regulated by shelterin subcomplex POT1-TPP1, with TPP1 being the key player [109–111]. These two proteins are important to decrease the dissociation rate of telomerase and help the translocation step. In *Leishmania* spp., RPA-1 seems to play the same role as POT1 at telomeres [12]. However, no ortholog of TPP1 was described in the parasite. The identification of other parasite telomerase cofactors and the characterization of this multistep process are under investigation.

Interestingly, in mammals, it is known that telomerase is possibly involved in other roles not related to telomere maintenance, such as cellular proliferation, mitochondrial activity, and gene expression regulation [112]. In *Leishmania* spp., for example, Campelo et al., (2015) [113] studied the TERT component of *Leishmania major* and observed that, besides the nucleus, it could also be found in its unique mitochondrion. In addition, parasites exposed to hydrogen peroxide showed increased TERT levels, especially in the mitochondria compartment. However, it is important to remember that parasite mitochondrion contains circular DNA (kDNA), which does not have telomeres. Therefore, it is important to demonstrate if parasite telomerase presents extra telomeric functions, since it could increase the druggable potential of this important ribonucleoprotein.

Our belief in telomerase as a good target for antiparasitic drug development is also related to its essential role in cell homeostasis and genome stability. We must also consider the structural differences between the telomerase complex (TERT and TER) of mammals and *Leishmania* spp. [7,89] and the fact that telomerase is inactive in most mammalian somatic cells. Thus, it is reasonable to hypothesize that impairing the infective parasite forms to elongate telomeres would directly affect parasites without harming host cells.

#### *4.3. Phylogenetic Context of Leishmania spp. Telomerases among Other Pathogenic Trypanosomatids*

Our very little knowledge about telomerases in *Leishmania* spp. reflects the challenges involved in the genome manipulation of these parasites [114,115]. However, the recent advent of genome editing using CRISPR-Cas9 and its variations should help resolve some of these functional questions that are still lingering, as has been done for other genes and in other species [116–118]. In addition, the available evidence on the subject matter shows some differences in the TERT between members of the trypanosomatid family and may be due to the enzyme′ s usefulness to the parasite [89,119].

Considering the importance of a high proliferative rate for the cell cycle of these parasites, the question of whether the parasite depends on an enzyme for longevity, as has been reported in other cells, is still unanswered. The knockdown of *T. brucei* telomerase induced telomere shortening without effects on the parasite growth [119]. However, the existent differences between trypanosomatids cannot be confidently extrapolated. In *Leishmania* spp., for example, the absence of the TERT and the TER minimal telomerase components seems to be very harmful to the cells (see Section 4.1).

Between the members of the *Leishmania* genus, Giardini et al., (2006) [89] found that the TERT component of *L. donovani*, *L. amazonensis*, *L. major*, and *L. braziliensis* has a nucleotide identity exceeding 90%, while sharing a little similarity with *T. cruzi* and *T. brucei*. Concerning amino acids, their sequence analysis shows over 70% identity between the *Leishmania* spp. and under 40% similarity with *T. cruzi* and *T. brucei* [89]. The *Leishmania* TER component also does not share nucleotide sequence similarities with other trypanosome TERs. They only share similarities in their secondary structures [7].

In other eukaryotes, the TERT component is also involved with noncanonical and extratelomeric functions, such as cell proliferation and apoptosis [120–124]. However, these functions may more likely result from evolutional adaptation events, which sometimes result in the loss and gain of certain protein domains with organism-specific functions. Hopefully, with the new approaches based on the CRISPR/Cas system, the many gaps of the *Leishmania* spp. telomerase will begin to be covered, and more details will probably be available to elucidate the telomere biology of these parasites.

#### **5. Conclusions Remarks**

The telomeres dynamics throughout the cell cycle are an essential phenomenon for all eukaryotic cells. Additionally, for most eukaryotes, the holoenzyme telomerase is the key behind these dynamics. When we expand this fact for single-cell parasites, we assume that understanding the differences and similarities between the pathogen and the host is an essential pathway for specific and successful treatment development. Here, we covered the knowledge available so far on *Leishmania* spp. cell cycle and telomere homeostasis, unveiling the remaining gaps and the advances reached in the last years. Among the impressive progress on the biology of these parasites are the facts that they remain in the G0 state during the infective stages and the remarkable divergence of their telomeric shelterin-like complex relative to mammals. However, these aspects are only a few examples of how this subject can be future linked to leishmaniases treatment and how far scientists are from a deeper knowledge of these peculiar eukaryote parasites.

**Author Contributions:** Conceptualization, M.I.N.C. and M.S.d.S.; writing—original draft preparation, L.H.C.A., D.A.-S., B.C.D.d.O., M.E.S., S.C.P., V.S.F., L.S.d.O., B.E.A. and Y.G.F.; writing—review and editing, L.H.C.A., D.A.-S., M.S.d.S. and M.I.N.C.; supervision, M.I.N.C. and M.S.d.S.; project administration, M.I.N.C. and M.S.d.S.; and funding acquisition, M.I.N.C. and M.S.d.S. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by the São Paulo Research Foundation (FAPESP) under grants 2018/04375-2 (M.I.N.C.), and 2019/10753-2 and 2020/10277-3 (M.S.d.S.). L.H.C.A. and D.A.-S. are postdoctoral fellows from FAPESP (grants 2021/04253-7 and 2021/05523-8, respectively). B.C.D.d.O., and M.E.S. are Ph.D. students from FAPESP (grants 2019/25985-6 and 2020/00316-1, respectively). B.E.A. and Y.G.F. are M.Sc. students from FAPESP (grants 2020/16480-5 and 2020/16465-6, respectively). S.C.P. is a M.Sc. student from CAPES (Coordenação de Aperfeiçoamento de Pessoal de Nível Superior—Brazil), and V.S.F. is an undergraduate student (FAPESP grant 2020/08162-3).

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable since this work is a review of published data.

**Conflicts of Interest:** The authors declare no conflict of interests, and the funders had no role in the design of the study; in the collection, analyses, or interpretation of the data; in the writing of the manuscript; or in the decision to publish the results.

#### **References**

## *Article* **M2 Muscarinic Receptor Activation Impairs Mitotic Progression and Bipolar Mitotic Spindle Formation in Human Glioblastoma Cell Lines**

**Maria Di Bari 1 , Vanessa Tombolillo 1 , Francesco Alessandrini 1 , Claudia Guerriero 1 , Mario Fiore 2 , Italia Anna Asteriti 2 , Emilia Castigli 3 , Miriam Sciaccaluga 4 , Giulia Guarguaglini 2 , Francesca Degrassi 2 and Ada Maria Tata 1,5, \***

**Abstract:** Background: Glioblastoma multiforme (GBM) is characterized by several genetic abnormalities, leading to cell cycle deregulation and abnormal mitosis caused by a defective checkpoint. We previously demonstrated that arecaidine propargyl ester (APE), an orthosteric agonist of M2 muscarinic acetylcholine receptors (mAChRs), arrests the cell cycle of glioblastoma (GB) cells, reducing their survival. The aim of this work was to better characterize the molecular mechanisms responsible for this cell cycle arrest. Methods: The arrest of cell proliferation was evaluated by flow cytometry analysis. Using immunocytochemistry and time-lapse analysis, the percentage of abnormal mitosis and aberrant mitotic spindles were assessed in both cell lines. Western blot analysis was used to evaluate the modulation of Sirtuin2 and acetylated tubulin—factors involved in the control of cell cycle progression. Results: APE treatment caused arrest in the M phase, as indicated by the increase in p-HH3 (ser10)-positive cells. By immunocytochemistry, we found a significant increase in abnormal mitoses and multipolar mitotic spindle formation after APE treatment. Time-lapse analysis confirmed that the APE-treated GB cells were unable to correctly complete the mitosis. The modulated expression of SIRT2 and acetylated tubulin in APE-treated cells provides new insights into the mechanisms of altered mitotic progression in both GB cell lines. Conclusions: Our data show that the M2 agonist increases aberrant mitosis in GB cell lines. These results strengthen the idea of considering M2 acetylcholine receptors a novel promising therapeutic target for the glioblastoma treatment.

**Keywords:** M2 muscarinic receptor; glioblastoma; cell cycle; aberrant mitosis; mitotic spindle

#### **1.  Glioblastoma (GB) cells have a highly infiltrative capacity, being able to spread into surrounding brain tissue by using the perivascular space [2]. Despite the great advances made in understanding the molecular alterations that occur in GB, there is not a definitive cure and mortality is still very high. Eighty-eight percent of patients affected by GB succumb to the disease within

**Citation:** Di Bari, M.; Tombolillo, V.; Alessandrini, F.; Guerriero, C.; Fiore, M.; Asteriti, I.A.; Castigli, E.; Sciaccaluga, M.; Guarguaglini, G.; Degrassi, F.; et al. M2 Muscarinic Receptor Activation Impairs Mitotic Progression and Bipolar Mitotic Spindle Formation in Human Glioblastoma Cell Lines. *Cells* **2021**, *10*, 1727. https://doi.org/10.3390/ cells10071727

Academic Editor: Zhixiang Wang

Received: 31 May 2021 Accepted: 6 July 2021 Published: 8 July 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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/).

one to three years [3]. GB is normally treated with surgery, followed by radiotherapy and chemotherapy with temozolomide [4,5]. Therefore, research focused on the development of new therapeutic agents is clinically relevant today.

Muscarinic acetylcholine receptors (mAChRs) are G protein-coupled receptors widely distributed in the nervous system and in several mammalian organs [6]. mAChRs are expressed in several primary and metastatic tumors, such as breast [7], ovarian [8], and lung cancers [9] and in astrocytoma and glioblastoma cells [10–12]. In particular, M2 mAChRs appear to be involved in tumor behavior, negatively modulating the proliferation and the migration of cancer cells [13–15].

Recently, by in vitro studies, we showed that the selective stimulation of M2 mAChRs by the orthosteric agonist arecaidine propargyl ester (APE) inhibits cell cycle progression and decreases cell survival, both in GB cell lines [14,16] and in GB stem cells [17]. Furthermore, the activation of M2 mAChRs induces apoptosis and oxidative stress, predominantly in p53-mutated glioblastoma cell lines [18].

In recent years, a wide range of clinically employed and experimental anticancer agents arresting the cell cycle and triggering apoptosis have started emerging [19]. Like other solid tumors, GB cells show an aberrant cell cycle and an increased rate of proliferation [20], as well as marked aneuploidy [21], suggesting that one of the causes of chromosomal instability could be determined by deregulated checkpoints.

In the last decade, the phenomenon of mitotic catastrophe has achieved increasing prominence and, in recent years, this mechanism has gained importance as a possible therapeutic target in cancer treatment [22]. In 2012, mitotic catastrophe was defined by the International Nomenclature Committee on Cell Death as an oncosuppressive mechanism that prevents the genomic instability of cells through the induction of mitosis-related cell death or permanent cell cycle arrest [23]. At least three mechanisms of mitotic catastrophe have been described: (I) Activation of the death machinery while the cell is still in mitosis; (II) "mitotic checkpoint adaptation," i.e., arrested cells enter the interphase without chromosome segregation, and cell death is triggered at the following interphase; (III) development of the senescent phenotype after aberrant mitosis [24,25]. Depending on the status of cell cycle checkpoints, several cytotoxic agents could induce aberrant mitosis/mitotic catastrophe, activating different pathways. Moreover, mitotic catastrophe could also occur after abnormal re-entry of tumor cells into the cell cycle following prolonged growth arrest [26,27].

Previous results have shown that M2 mAChR activation by APE causes the proliferative arrest of U251MG and U87MG cells [16]. Given the ability of APE to increase ROS levels and to cause cytotoxic damage, including chromosomal aberrations [17], in the present work, we investigated in depth the cell cycle progression upon M2 receptor activation in these two cell lines. The data obtained show that M2 agonist treatment causes an arrest in cell cycle progression with an accumulation of cells during pro-metaphase/metaphase transition, resulting in a significant increase in abnormal mitosis and multipolar mitotic spindle formation.

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

#### *2.1. Cell Cultures*

U251MG and U87MG cell lines were maintained at 37 ◦C in an atmosphere with 10% CO2. The cells were grown in DMEM (Sigma-Aldrich, St. Louis, MO, USA) plus 10% fetal bovine serum (Sigma-Aldrich, St. Louis, MO, USA), 50 µg/mL of streptomycin, 50 IU/mL of penicillin, 2 mM of glutamine (Sigma-Aldrich, St. Louis, MO, USA), and 1% no essential amino-acids (Sigma-Aldrich, St. Louis, MO, USA).

#### *2.2. Pharmacological Treatment*

The M2 agonist arecaidine propargyl ester hydrobromide (APE) was used as a preferential agonist of the M2 muscarinic receptor subtype. Pharmacological binding experiments

and silencing of the receptor by short interference RNA demonstrated the selectivity of this agonist for the M2 receptor subtype [16,17].

#### *2.3. Western Blot Analysis*

Cells were homogenized in lysis buffer (Tris-EDTA 10 mM, 0.5% NP40, and NaCl 150 mM) containing protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO, USA). After protein extraction, the total amount of protein was determined by a Pierce BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer's protocol. The protein extracts were run on SDS-polyacrilamide gel (SDS-PAGE) and transferred to polyvinylidene difluoride (PVDF) sheets (Merck Millipore, Darmstadt, Germany). Membranes were blocked in 5% non-fat milk powder (Sigma-Aldrich, St. Louis, MO, USA) in PBS containing 0.1% Tween-20, and then incubated with the primary antibodies overnight at 4 ◦C. The primary antibodies used were SIRT-2 (Santa Cruz Biotechnologies, Dallas, TX, USA) and acetylated alpha-tubulin (Lys 40) (Sigma-Aldrich, St. Louis, MO, USA). The blots were washed three times with PBS + 0.1% Tween-20, then incubated with secondary antibodies conjugated to horseradish-peroxidase for 1 h at room temperature. The immunoreaction was revealed by ECL chemiluminescence reagent (Immunological Science, Rome, Italy). The bands were detected by exposition to Chemidoc (Molecular Imager ChemiDoc XRS + System with Image Lab Software; Biorad, CA, USA).

#### *2.4. Flow Cytometry Analysis*

The cells were plated at a density of 5 × 10<sup>5</sup> cells/dish. The day after, the cells, excluding the control samples, were treated either with 100 µM of APE (24 and 48 h of treatment) or with nocodazole (0.2 µg/mL) used as a positive control (24 h treatment). At the end of the treatments, cells were collected by trypsinization and fixed in PBS/ethanol (1:1; *v*/*v*). Samples were then incubated with monoclonal antibodies against phosphohistone H3 (p-HH3) (Ser10) (Merck Millipore, Darmstadt, Germany) for 60 min at room temperature, washed twice with 0.5% Tween-20 in PBS and incubated for 30 min with anti-mouse Alexa fluor 488-conjugated antibody (Invitrogen, Monza, Italy). Samples were washed with PBS and finally stained with 20 µg/mL propidium iodide for 15 min at room temperature. Flow cytometry analysis was performed using a flow cytometer Coulter Epics XL with 488 nm wavelength excitation. For each sample, at least 10<sup>4</sup> events were measured.

#### *2.5. Immunocytochemistry*

Cells were fixed with 4% paraformaldehyde for 20 min, permeabilized in PBS containing 0.1% Triton X-100 and incubated in 10% normal goat serum (NGS) for 1 h at room temperature. Coverslips were then incubated overnight at +4 ◦C with a human CREST antibody (Biolegend, San Diego, CA, USA), diluted 1:50 in PBS containing 0.1% Triton X-100 5% NGS, and then incubated in a Red-X anti-human antibody (1:1000, Jackson Immunoresearch Europe, Cambridge, U.K.) after extensive washing. After a 45 min incubation in a FITC conjugated anti-alpha tubulin antibody (1:100, Sigma-Aldrich, St. Louis, MO, USA), coverslips were washed three times with PBS + 1% BSA and twice with PBS, incubated with Hoechst 33,342 (1:15,000; *v*:*v* in PBS) for 10 min and mounted with glycerol/PBS (3:1; *v*/*v*). In parallel experiments, an anti-acetylated tubulin antibody (1:500, Sigma-Aldrich, St. Louis, MO, USA) and a Red X anti-mouse antibody (1:1000, Jackson Immunoresearch Europe, Cambridge, UK) were used.

#### *2.6. M2 Receptor Knock-Down*

U251 cells were transfected with a pool of siRNAs selective for human M2 receptors (CHRM2), as previously described [14].

The siRNA duplexes were synthesized by Riboxx Life Sciences. The following sequences of siRNAs (Riboxx Life Sciences, Dresden, Germany) used were:

(siRNA1129-1) sense, 5′ -AUUUACUACUAAAUCCUCCCCC-3′ antisense 5′ -GGGGGAGGAUUUAGUAGUAAAU-3′ ;

(siRNA1129-2) sense 5′ -AUGUAGCCCAUUUCUUCCCCC-3′ antisense 5′ -GGGGGAAGAAAUGGGCUACUA-3′ ; (siRNA 1129-3) sense 5′ -UCCUUUGAGUUUCAGGCUGCCCCC-3′ antisense 5′ - GGGGGCAGCCUGAAACUCAAAGGA-3′ ; (siRNA 1129-4) sense 5′ -AGUUACACCUUGACCUAACCCCC-3′ antisense 5′ -GGGGGUUAGGUCAAGGUGUAACU-3′

#### *2.7. Time-Lapse Live Cell Imaging*

Cells seeded in four-well micro-slides (ibiTreat, cod. 80426, Ibidi, Planegg, Germany) were observed under an Eclipse Ti inverted microscope (Nikon, Tokyo, Japan), using a 40× (Plan Fluor, 0.60 N.A., DIC) objective; during the whole observation, cells were kept in a microscope stage incubator (Basic WJ, Okolab, Pozzuoli, Italy), at 37 ◦C and 5% CO2. DIC images were acquired every 7 min over 72 h using a DS-Qi1Mc camera and NIS-Elements AR 3.22 software (Nikon). Image and movie processing were performed with NIS-Elements HC 4.2.

#### *2.8. Statistical Analysis*

Data presented are the average ± SEM obtained from three independent experiments. Statistical analysis was performed by Student's *t*-tests, Mann–Whitney tests, and one way ANOVAs followed by Tukey's comparison post-hoc tests. For the evaluation of aberrant mitosis, the ratio between total abnormal metaphases/mitotic cells was calculated. Ten photographic fields for each sample were considered. Each sample was produced in triplicate. Data from time-lapse experiments were statistically analyzed using Mann– Whitney or *χ* 2 tests, as indicated. The results were considered statistically significant at *p* < 0.05 (\*), *p* < 0.01 (\*\*), *p* < 0.001 (\*\*\*), and *p* < 0.0001 (\*\*\*\*).

#### **3. Results**

#### *3.1. M2 Receptor Activation Caused Accumulation of the GB Cell Lines in the M Phase*

Our previous data indicated that APE induces G2/M arrest in the U251 glioblastoma cell line [16]. In order to verify whether APE was able to induce an arrest in the G2 or M phase, we evaluated—by FACS analyses—the expression of histone H3 phosphorylated at serine 10 (p-HH3), a specific mitotic marker (Figure 1A). Nocodazole, a drug that interferes with microtubule polymerization [28], was used as a positive control. As expected, nocodazole treatment (0.2 µg/mL) caused a significant increase in the percentage of cells positive for p-HH3. Interestingly, APE treatment (100 µM) increased the percentage of p-HH3-positive cells in a comparable manner to nocodazole, at least in terms of 24 h of treatment. As shown in Figure 1C, 30% of treated U251 cells appeared to have accumulated in the M phase. Forty-eight hours after treatment, the percentage of p-HH3-positive cells significantly decreased (18%), albeit remaining higher than untreated cells.

cate. Student's **Figure 1.** FACS analysis showing the expression of phospho-H3 (ser10) in the U251 (**A**) and U87 (**B**) cell lines. The graphs report the percentage of phospho-H3-positive cells upon treatment with 100 µM of APE or 0.2 µg/mL of nocodazole in the U251 (**C**) and U87 (**D**) cells. Data presented are the average ± SEM of three independent experiments conducted in triplicate. Student's *t*-test was used to statistically compare the different experimental conditions (\*\*\* *p* < 0.001, \*\* *p* < 0.01, and \* *p* < 0.05).

As previously observed, U87 cells appeared mainly accumulated in the G1 phase after M2 receptor activation [16]. However, an increase in p-HH3-positive cells after APE treatment was observed in U87 cells. In this cell line, only 22% of cells were accumulated in the M phase, mainly 24 h after APE treatment (Figure 1B,D).

In order to confirm cell accumulation in the M phase, we performed a microscopic analysis on the GB cells after staining with the nuclear dye Hoechst 33342 to assess the frequency of cells in the mitotic stage. Figure 2A is a representative image of the U251 nuclei after APE treatment. Evaluating the nuclei organization and the chromosome distribution, we counted the number of the cells in the mitotic stage at both 24 and 48 h after M2 agonist treatment. As shown in Figure 2B,C, a progressive increase in cells in mitosis was evident after 24 h of APE treatment in both cell lines (U251 and U87). This increase appeared more consistent in U251 than in U87 cells. After longer treatment (48 h), we observed a significant decrease in the percentage of dividing cells.

To verify the direct involvement of the M2 receptor in this phenomenon, we evaluated the percentage of dividing cells in the U251 cell line, treated for 24 h with 100 µM of APE after M2 receptor silencing by siRNA transfection (Figure 2D). After M2 silencing, the percentage of dividing cells in the presence of APE was comparable to that observed in untreated cells, albeit the number of the cells resulted significantly reduced compared to non-transfected cells, most likely as a consequence of the toxicity caused by the transfection.

's **Figure 2.** (**A**) Nuclei staining by Hoechst 33,342 in the U251 cells untreated or treated with APE (100 µM) for 24 and 48 h. The arrows indicate the cells in mitosis (bar = 10 µm). The graphs indicate the percentage of cells in mitosis 24 and 48 h after treatment with 100 µM of APE: (**B**) U251 cells and (**C**) U87 cells. (**D**) Percentage of cells in mitosis in the U251 cell line in untreated and APE-treated cells in the presence or absence of M2 receptor knock-down by siRNA transfection. An ANOVA test was used, followed by Tukey's test, to statistically compare all of the experimental conditions (\*\*\* *p* < 0.001; ### *p* < 0.001).

#### *3.2. Analysis of Mitosis Progression*

In order to directly follow the entrance to the mitotic stage, as well as the progression through mitosis, we videorecorded U251 cells for 72 h after treatment with 100 µM of APE. More than 80% of the cells entered mitosis during videorecording, both in the control and the APE-treated cultures, confirming that no G2 arrest was induced by the treatment. Nonetheless, mitotic entry may be delayed by APE, since we noticed that approximately 30% of the dividing cells entered mitosis between 24 and 72 h after treatment, while the majority (>95%) of control cells progressed to mitosis within the first 24 h (see video in Supplementary Materials). A strong delay/arrest in the pro-metaphase was observed (Figure 3A): While the average time from rounding-up to chromosome segregation onset was approximately 50 min in the control cells, APE-treated mitoses remained for approximately 22 h in a prometaphase-like state and were not able to divide normally. Only 15% of those cells were eventually able to originate two daughter cells through an abnormal division (second row of Figure 3B,C) (see video in Supplementary Materials). The majority of mitoses (>70%) were still arrested in a prometaphase-like state after 48 h and displayed membrane blebbing movements often associated with an elongated shape (third row of Figure 4B,C). At the end of the videorecording (72 h), more than 50% of the analyzed cells had exited mitosis without segregating chromosomes (fourth row of Figure 4B,C), suggesting a failure of cell division after the long blebbing phase (see video in Supplementary Materials).

– – Student's **Figure 3.** Time-lapse analysis of APE-treated cultures revealing abnormal mitotic phenotypes. The U251 cultures treated with 100 µM of APE were recorded by time-lapse for 72 h upon M2 agonist treatment, and 60–90 mitoses per condition were analyzed. (**A**) Time from mitotic round-up to chromosome segregation (\*\*\*\* *p* < 0.0001; Mann–Whitney test). (**B**) Representative single photograms are shown for a normal mitotic division (first row) and the different observed defects (second to fourth rows); minutes from round-up are indicated. Scale bar: 10 µm. (**C**) Histograms showing the percentage of abnormal mitoses in the APE-treated cultures, divided according to their fate at 48 and 72 h from the beginning of the treatment (\*\*\*\* *p* < 0.0001; Student's *t*-test comparing 48 vs. 72 h). All mitoses in the control cultures were normal.

#### *3.3. APE-Arrested Glioblastoma Cells during Prometa-Metaphase Transition and Induced Multipolar Mitotic Spindles*

To further investigate the effect of APE on mitosis progression, cells in the mitotic stage were classified as pro-metaphase/metaphase or anaphase, considering the position the chromosomes and their organization on the metaphase plate (Figure 4A,B). We chose to analyze the cells 30 h after treatment, since this is a time before the severe cell death observed after 48 h of APE treatment [16,18]. The data obtained indicate that the number of cells in the pro-metaphase/metaphase was higher in the APE-treated cells compared to the untreated cells (control) and was associated with a concomitant decrease in cells in the anaphase. This trend was observed in both cell lines, but it was statistically significant only in the U251 cell line (Figure 4B). Thus, our data indicate that APE consistently arrested glioblastoma cells prior to anaphase chromosome segregation.

. Student's – α Student's Student's **Figure 4.** Percentage of pro-metaphases/metaphases and anaphases in the U251 (**A**) and in U87 (**B**) cells after treatment with 100 µM of APE for 30 h. Student's *t*-test was used to statistically compare the different experimental conditions (\* *p* < 0.05). (**C**–**F**) Immunocytochemistry analysis of U251 cells untreated or treated with APE (100 µM) for 30 h. Cells were immunostained with anti-CREST (**red**) and anti-α-tubulin (**green**) antibodies and counterstained with Hoechst 33,258 (**blue**). Scale bars: 5 µm. (**G**,**H**) Percentage of the cells presenting abnormal mitosis in the control condition and after 30 h of treatment with 100 µM of APE, in the U251 and U87 cells, respectively. Student's *t*-test was used to statistically compare the different experimental conditions (\*\* *p* < 0.01 and \* *p* < 0.05). (**I**,**J**) Percentage of the cells with multipolar spindles in the control condition and 30 h after treatment with 100 µM of APE, in the U251 and U87 cells, respectively. Student's *t*-test was used to statistically compare the different experimental conditions (\*\* *p* < 0.01 and \* *p* < 0.05).

– The effect of APE treatment on mitotic spindle structure was additionally examined in pro-metaphase/metaphase cells by immunofluorescence microscopy (Figure 4C–F). We used an antibody against alpha-tubulin to detect the microtubules forming the mitotic spindles and an antibody against kinetochore protein (CREST antibody) to visualize the centromere. The presence of bipolar, well-organized, overall fusiform spindles was evident in the untreated cells. Within these spindles, the highly condensed chromosomes appeared to be aligned along the equatorial plate (Figure 4D). On the contrary, strong abnormalities in mitosis were found in the cells treated with APE for 30 h. This treatment caused a significant increase in the percentage of abnormal mitoses in both the U251 and U87 cell lines compared to the control condition, as shown in Figure 4G,H. Thirty hours after APE treatment, some of the chromosomes appeared not aligned at the equatorial plate or with the spindles themselves, being asymmetric and often of multipolar origin (i.e., bipolar spindles) with clustered centrosomes (Figure 4E) or showing several centrosomes and a multipolar organization of chromosomes (Figure 4F). The graphs in Figure 4I,J show a significant increase in multipolar spindles in the APE-treated cells of both glioblastoma cell lines compared to the untreated cells.

#### *3.4. Modulation of SIRT2 and Acetylated Tubulin Expression after APE Treatment*

Several pieces of evidence have connected the activity of deacetylase SIRT2 to the control of mitotic progression. Indeed, SIRT2 appears to act in the mitotic checkpoint, blocking chromosome condensation in response to mitotic stress [29]. In light of these findings, we evaluated SIRT2 protein expression by Western blot analysis after APE treatment. As shown in Figure 5A–E, SIRT2 expression significantly increased in the U251 (Figure 5A,B) and U87 cells (Figure 5E,F), particularly 30 h after treatment with 100 µM of APE. Considering that SIRT2 deacetylates tubulin [30], we assessed the expression of acetylated tubulin by Western blot analysis, showing that its levels significantly decreased only in the U87 (Figure 5G,H) cells 24 and 30 h after APE (100 µM) treatment. Apparently, no significant modulation of acetylated tubulin was evident in the U251 cells (Figure 5C,D). Nevertheless, high-resolution immunodetection of acetylated microtubules on mitotic cells demonstrated that the spindle microtubules were extremely abnormal in the U251 treated cells. Meanwhile, the mitotic spindles of the untreated cells consisted of several arrays of acetylated microtubules, and the mitotic spindles in APE-treated cells were characterized by long fibers. These fibers were composed of bundled acetylated microtubules that passed over the chromosomes and did not attach the kinetochores (Figure 5I). –

**Figure 5.** Western blot analysis of SIRT2 and acetylated tubulin expression in U251 and U87 cells treated with APE (100 µM) for 24 and 30 h. Representative blot for SIRT2 (**A**,**E**) and acetylated tubulin (**C**,**G**) in U251 and U87 cells, respectively. GAPDH was used as an internal reference protein. (**B**,**D**,**F**,**H**) Densitometric analysis obtained from three independent experiments in U251 (**B**,**D**) and U87 (**F**,**H**) cells, respectively. Student's *t*-test was used to statistically compare the different experimental conditions (\*\* *p* < 0.01 and \* *p* < 0.05). (**I**) Immunocytochemistry analysis of U251 cells untreated or treated with APE (100) µM for 30 h. Cells were immunostained with an anti-acetylated tubulin (**red**) antibody and counterstained with Hoechst 33,258 (**blue**). Scale bars: 5 µm.

#### **4. Discussion**

Glioblastoma cells are known to respond to cholinergic stimulation [6,31,32]. The effects of M2 receptor subtype activation on cell growth and survival have been largely characterized using two different glioblastoma cell lines (U251MG and U87MG) in primary cultures obtained from different human biopsies and in glioblastoma cancer stem cells [17]. In particular, we previously demonstrated that M2 receptor activation causes a cell cycle arrest in the G1/S and at G2/M phases in U87MG and U251MG cells, respectively [16]. In addition, cell viability analysis showed that the M2 orthosteric agonist APE causes cytotoxic effects (i.e., ROS production, double-strand DNA breaks, and chromosomes aberrations) and severe apoptosis, especially in p53-mutated cells such as U251MG [18].

Starting from these already described observations, in the present work we investigated in depth the modalities through which the M2 agonist APE triggers the cytostatic effects in glioblastoma cells.

Since M2 receptor activation arrests cell cycle progression, we analyzed the progression of the mitotic phase. In both glioblastoma cell lines (U251MG and U87MG), we observed an arrest at the mitotic stage, as shown by a significant increase in the percentage of p-HH3 (ser10)-positive cells after M2 agonist treatment. Mitotic arrest was much less evident in the U87MG cell line, in accordance with the higher levels of cells accumulated in the G1 phase than in the G2/M phase, as previously observed in this cell line [16]. Instead, the U251MG cells showed a larger increase in p-HH3-positive cells upon APE treatment, confirming an accumulation of cells in the M phase rather than in the G2 phase, as previously suggested by FACS analysis results [16]. This different behavior may be dependent on the genetic background of two the different glioblastoma cell lines; in fact the presence of wildtype or mutated p53 in U87 and in U251 cells, respectively, may be responsible for the cell accumulation in the different phased of the cell cycle.

Nuclei staining and immunocytochemistry analysis also confirmed that APE caused an accumulation of cells in the M phase. Moreover, a significant increase in abnormal mitosis and an accumulation of cells during pro-metaphase/metaphase transition were observed, accompanied by a reduction in the number of cells in the anaphase. Additionally, the untreated cells formed normal bipolar spindles and the chromosomes appeared aligned along the equatorial plate during the metaphase. Conversely, in the APE-treated cells, asymmetric spindle formation was observed, together with incorrect chromosome alignment along the equatorial plate. In addition, M2 agonist treatment increased the formation of pseudobipolar, tripolar, and multipolar spindles.

To demonstrate the direct involvement of the M2 receptor in abnormal mitosis induction, we silenced *CHRM2* expression by siRNA transfection in the U251 cells, with the cell line showing a higher percentage of abnormal mitosis after M2 agonist treatment. In this condition, the number of abnormal mitoses in the untreated and APE-treated cells was comparable. These results, in accordance with previous studies [16,17], confirm the selectivity of the M2 agonist APE for M2 receptors and show that APE is unable to cause a mitotic arrest in the absence of M2 receptors.

Several studies have shown that sirtuins, particularly Sirtuin 2 (SIRT2), are among the factors involved in mitotic progression in the normal cell cycle [30]. SIRT2 is an NAD+-dependent histone deacetylase [33]. SIRT2 overexpression has been shown to prolong the M phase and delay mitotic exit [34]. It has been reported that SIRT2 mRNA expression is reduced in approximately 70% of human gliomas [35]. Like SIRT1, SIRT2 confers protection against genotoxic and oxidative stress by reducing cellular ROS levels through the induction of the mitochondrial anti-oxidant enzymes manganese superoxide dismutase (MnSOD) [36] and by blocking chromosome condensation, acting at the level of the mitotic checkpoint [29].

According to the literature data [35], SIRT2 appears to be strongly downregulated, both in U87 and U251 cell lines. Interestingly, APE treatment induced a significant upregulation of SIRT2 expression 30 h after treatment, in line with the increased accumulation of the cells in the M phase. Moreover, SIRT2 activity may contribute to the formation of deacetylated unstable microtubules [37]. In our experiments, we observed a decrease in acetylated tubulin expression, particularly in the U87 cells. This phenomenon may lead to spindle fragmentation, which contributes to the formation of spindle extra poles and ab-

normal chromosome segregation. Although U251 did not show a significant modulation of acetylated tubulin after M2 agonist treatment, by immunocytochemistry analysis for acetylated tubulin, we observed that in U251 cells treated with APE, the acetylated microtubules passed over the chromosomes and did not attach the kinetochores. Previous data have demonstrated that APE treatment induces oxidative stress, DNA damage, and chromosomal instability, particularly in the U251 cell line [18]. These results suggest that the inability of microtubules to bind chromosomes may also be dependent on the possible alterations that may occur on DNA or other proteins associated with the kinetochores that prevent the interaction between chromosomes and microtubules. Altogether, these alterations may negatively influence chromosomes' organization and their attachment to microtubules of the mitotic spindle and their inability of divide cells. The dramatically extended mitotic arrest and the highly abnormal phenotypes observed by time-lapse microscopy showed that glioblastoma cells are unable to further proliferate after APE treatment. This suggests that mitotic catastrophic and apoptotic events may occur in subsequent mitosis.

#### **5. Conclusions**

Glioblastoma cells are characterized by an aberrant cell cycle [20] and marked aneuploidy [21]. Therefore, mitotic catastrophe induction may be an interesting oncosuppressive mechanism with the aim of inducing mitosis-related cell death or permanent cell cycle arrest [23]. The data obtained in the present work, together with previous data [14,16–18,38,39], may allow to hypothesize that the activation of the M2 receptor through selective agonists causes two significant effects in glioblastoma cells: (1) A cytostatic effect, impairing cell proliferation and inducing cell cycle arrest; (2) a cytotoxic effect, increasing apoptosis and cell death. The two effects should be strictly correlated and both contribute to impairment of glioblastoma cell proliferation and survival.

These results, together with the data obtained in other tumor types (i.e., neuroblastoma, breast cancer, and urothelial cancer) [13,15,40,41], highlight the M2 muscarinic receptor as a new strategic therapeutic target in cancer therapy. Therefore, a special effort to identify new selective ligands able to bind this receptor with more efficacy and to reduce the possible side effects appears clinically relevant and may open new therapeutic perspectives for glioblastoma treatment, as well as for the treatment of other tumors.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/cells10071727/s1, Video S1: Abnormal cell division in U251 cells.

**Author Contributions:** Conception and design: A.M.T. and E.C.; acquisition of data: M.D.B., F.A., V.T., M.F., C.G., M.S. and F.D.; time-lapse experiments: I.A.A. and G.G.; analysis and interpretation of data: A.M.T., M.D.B., F.D., I.A.A. and G.G.; drafting of the article: M.D.B., C.G. and A.M.T.; critical revision of the article: A.M.T., F.D., G.G., E.C. and F.A. All authors have read and agreed to the published version of the manuscript.

**Funding:** No external funds were received.

**Informed Consent Statement:** Not applicable.

**Acknowledgments:** This work was supported by Ateneo Sapienza and Network CIB 2018 funds to A.M.T. Time-lapse experiments were performed at the IBPM Nikon Reference Centre.

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

#### **References**

#### *Review* **Incorporation of 5-Bromo-2** ′ **-deoxyuridine into DNA and Proliferative Behavior of Cerebellar Neuroblasts: All That Glitters Is Not Gold**

**Joaquín Martí-Clúa**

Unidad de Citología e Histología, Departament de Biologia Cellular, de Fisiologia i d'Immunologia, Facultad de Biociencias, Institut de Neurociències, Universidad Autónoma de Barcelona, Bellaterra, 08193 Barcelona, Spain<EMAIL_ADDRESS>Tel.: +34-93-581-1666