Neocarzinostatin Synthesis Essay

Genomes are continuously bruised by products of normal cell metabolism and by errors in DNA replication. The frequency of DNA lesions increases when cells are exposed to UV light, gamma rays or toxic chemicals (Figure 1). While DNA damage impacts nearly every aspect of gene expression, including transcription and translation, more attention has recently been devoted to studying how DNA damage affects post-transcriptional events, with several excellent reviews published on this topic [1,2,3,4,5]. Here we review the interconnections that exist between DNA damage and pre-mRNA splicing. Following a brief overview of the basic principles of the DNA damage response (DDR), splicing and alternative splicing control, we outline how DNA damage affects the post-translational modification, localization, expression and activity of splicing factors. We then present examples showing that DNA damage often disrupts splicing by interfering with its coupling to transcription. Finally, we summarize a growing body of data that document the impact of DNA damage on the splicing and alternative splicing of genes intimately associated with the DDR and with cell fate.

1.1. The DNA Damage Response

DNA damage is a cause of cancer, and its accumulation is associated with organismal aging. Paradoxically, provoking DNA damage by irradiation, DNA intercalating drugs, crosslinking agents or topoisomerase inhibitors is a common strategy to treat patients suffering from cancer. This anti-cancer approach is based on the expectations that (1) an excessive number of lesions will overwhelm the DNA repair machinery of cancer cells and trigger apoptosis; and (2) that the normal and cancer cells that acquire a lesser load of mutations will not become cancerous or evolve toward more aggressive behavior, respectively.

Distinct repair mechanisms are used to correct different types of DNA lesions. Mismatch repair and base-excision repair act on simple lesions, while nucleotide excision repair, non-homologous end-joining and homologous recombination (HR) deal with more complex lesions. DNA lesions, if not adequately repaired, impede transcription and replication and result in mutations or chromosomal rearrangements. To deal with DNA lesions, the cell must mount a response that is based first on sensing the DNA lesions themselves or their direct consequences, such as blocks in replication and transcription. DNA damage triggers the phosphorylation of the histone variant H2AX, the modifications of canonical histones, and the recruitment of poly(ADP-ribose) enzymes. Depending on the type of lesions, different axes of the DDR are activated. For example, double-strand breaks (DSBs) recruit the ATM and DNA-PK kinases, while single-stranded breaks (SSBs) recruit the ATR kinase. Following their activation, the ATM/ATR kinases activate other kinases, including CHK1 and CHK2 [6] (Figure 1). These early events ramify into a signal transduction cascade that mobilizes downstream pathways to mediate cell-cycle arrest, DNA repair, and apoptosis, if the intensity of the damage is excessive. Damage sensing and many steps of the DDR rely on a variety of post-translational modifications (e.g., phosphorylation, poly(ADP-ribosylation), acetylation, methylation, ubiquitylation, sumoylation) to guide the interactions between DDR factors and components of the repair and cell-cycle machinery [7]. In addition to these initial stages, the DDR initiates a slower route aimed at modulating transcription, of which the main contributor in the ATM/CHK2 axis is p53. The p53 protein regulates expression of cyclin-dependent inhibitor protein 1A (CDKN1A or p21), apoptotic proteins (e.g., BAX and PUMA) and DNA repair components. Importantly, the E3 ubiquitin ligase MDM2 represses p53 as part of a cyclic strategy that senses the progress of damage repair [8]. When p53 activity is defective, as is often the case in cancer cells, the cell-cycle checkpoint response to damage is rewired through the p38/MK2 pathway [9]. Thus, DNA damage elicits a complex signaling cascade that coordinates cell-cycle progression and DNA repair, and reconfigures gene expression at multiple levels.

Figure 1. The RNA splicing response to DNA damage. Several early and late steps of the DNA damage response alter processes that impact the activity of splicing factors, ultimately affecting the production, through splicing, of components that maintain genome integrity and control cell fate.

Figure 1. The RNA splicing response to DNA damage. Several early and late steps of the DNA damage response alter processes that impact the activity of splicing factors, ultimately affecting the production, through splicing, of components that maintain genome integrity and control cell fate.

While the expression of genes that encode sensing factors, and components of the DNA repair, cell-cycle and apoptotic machineries is controlled by the p53 activation branch of the DDR, these genes also produce splice variants that harbor potentially distinct, and sometimes, opposite activities. For example, the recruitment to chromatin of cyclin D1a, but not its splice variant cyclin D1b, is sufficient to activate the DDR [10], and Bcl-x produces variants with pro-survival or pro-death activities [11]. A change in splicing control elicited by the DDR therefore has the potential to provide feedback on every step of the DDR and regulate repair and cell fate.

1.2. Splicing and Alternative Splicing

Precursor (pre)-mRNA splicing is the process by which introns are removed from a pre-mRNA and exons are joined to produce a mature mRNA. Removal of introns from pre-mRNAs occurs in eukaryotes from yeast to human. The majority of introns in the budding yeast Saccharomyces cerevisiae are found in ribosomal protein genes, which produce approximately 90% of the pre-mRNAs in growing cells [12]. In mammals, except for histones and a few other genes, nearly all RNA polymerase II-transcribed genes contain introns. Splicing is performed by the spliceosome, a large nuclear macromolecular complex that contains five small nuclear ribonucleoproteins (snRNPs) (U1, U2, U4, U5 and U6) and more than 150 accessory proteins [13,14,15,16,17]. Fewer than 0.5% of human introns are processed by a minor form of spliceosome that uses the functionally homologous U11, U12, U4atac and U6atac snRNPs. The U5 snRNP is used in both spliceosome types [18]. Spliceosome assembly is a dynamic process initiated by the recognition of splice sites (Figure 2A,B); the U1 snRNP recognizes the 5' splice site, while the U2AF proteins and U2 snRNP interact with the 3' splice site and the branch site, respectively [14]. Once the borders of the intron are defined, the pre-assembled U4/U6.U5 tri-snRNP is recruited and, with the help of auxiliary proteins, the U1 and U4 snRNPs are displaced to allow U6 and U2 snRNPs to form a catalytically competent core that positions the branch point adenosine for the first of two cleavage steps. The first step produces a free upstream exon and a lariat intron covalently linked to the downstream exon. Following further spliceosome rearrangements, the second step of splicing leads to the excision of the lariat intron and the ligation of both exons. The efficiency of spliceosome assembly is increased when it is coupled to transcription [19], at least in part because the CTD of RNA polymerase II recruits spliceosome components to facilitate their deposition on the nascent pre-mRNA [20] (Figure 2C).

A single type of pre-mRNA can be spliced in different ways (i.e., by inclusion of specific exons in the final spliced product) to generate distinct mRNAs (Figure 2D). This process is named alternative splicing, and is a major contributor to transcriptomic and proteomic diversity in higher eukaryotes. In humans, nearly all multi-exon primary transcripts are alternatively spliced [21,22]. On average, a human gene is made up of 8–10 exons [23], and examples of the diversifying power of alternative splicing range from two to several thousand variants from a single gene (e.g., the Drosophila DSCAM gene produces over 38,000 splice variants [24]). Although much remains to be done to document the remarkable diversity of functions resulting from alternative splicing, examples of functionally relevant splice variants are continuously being reported, and are found in all cellular processes [25]. The production of proteins displaying different functions is expected to be tightly controlled. Indeed, profiles of alternative splicing vary in a tissue-specific manner [26], and are often altered in diseases, including cancer [27,28,29].

Figure 2. Basic principles of pre-mRNA splicing. (A) Schematic structure of a pre-mRNA with the position of core signal sequences that define exons and introns. ss: splice site; (B) A snRNP-biased view of spliceosome assembly leading to two catalytic steps that produce the mRNA and the excised intron. U2AF is a heterodimer made of the U2AF2 (U2AF65) and U2AF1 (U2AF35) proteins that respectively recognize the polypyrimidine tract and the AG dinucleotide at the 3' splice site [15]; (C) Spliceosome assembly is often coupled with transcription, with the carboxyl-terminal domain (CTD) of RNA polymerase II recruiting splicing components that are deposited on the nascent pre-mRNA; (D) In contrast to constitutive splicing, alternative splicing produces different mRNAs from a single kind of pre-mRNA.

Figure 2. Basic principles of pre-mRNA splicing. (A) Schematic structure of a pre-mRNA with the position of core signal sequences that define exons and introns. ss: splice site; (B) A snRNP-biased view of spliceosome assembly leading to two catalytic steps that produce the mRNA and the excised intron. U2AF is a heterodimer made of the U2AF2 (U2AF65) and U2AF1 (U2AF35) proteins that respectively recognize the polypyrimidine tract and the AG dinucleotide at the 3' splice site [15]; (C) Spliceosome assembly is often coupled with transcription, with the carboxyl-terminal domain (CTD) of RNA polymerase II recruiting splicing components that are deposited on the nascent pre-mRNA; (D) In contrast to constitutive splicing, alternative splicing produces different mRNAs from a single kind of pre-mRNA.

Sophisticated mechanisms that regulate alternative splicing profiles are also emerging (Figure 3). Alternative splicing units usually have weak splice sites whose utilization is controlled by sequence elements recognized by RNA binding proteins (RBPs) that act positively or negatively to recruit spliceosome components or prevent spliceosome assembly [30,31]. For example, SR proteins can interact with exonic enhancer sequences to antagonize the activity of a nearby splicing silencer element [32]. The activity of exonic silencers is often mediated by hnRNP proteins of which A1, L and PTBP1 have received most of the attention [33,34,35,36]. The respectively positive and negative functions of SR and hnRNP proteins bound to exons are often reversed when they bind to introns. For example, the binding of an SR protein near the branch site prevents U2 snRNP binding [37], while the binding of hnRNP A1 and H in introns stimulates splicing [38]. Transcript-specific studies and global analyses of alternative splicing indicate that positive and negative interactions between splicing factors with a wide range of sequence specificities play an important role in splicing regulation [30] (Figure 3A–C). In addition, since most introns are removed in a cotranscriptional manner, the control of alternative splicing is often coupled to the local state of the chromatin. Histone marks and chromatin remodeling factors impact the speed of transcription elongation and the recruitment of splicing regulators to alter splice site selection (Figure 3D,E) [39]. Thus, while great progress has been achieved in documenting the function of individual regulatory proteins, we need to better understand how these splicing factors combine their activity to control specific splicing decisions, how these activities are integrated with transcription and chromatin structure, and how they are affected by various cellular inputs and environmental insults. As is often the case in biology, insight can be provided by disrupting homeostasis. Below we will present examples of how DNA damage, by modifying the expression, localization and activity of spliceosomal proteins and splicing regulators, is providing precious information on the rules and mechanisms that control distinct steps of splice site selection.

Figure 3. Molecular mechanisms controlling splice site selection. (A) A variety of splicing regulators, including hnRNP and SR proteins, bind to exon or intron splicing enhancers (ESE or ISE, respectively) and to exon or intron splicing silencers (ESS or ISS, respectively) to control splice site recognition and utilization; (B) The activity of splicing regulators is often position-dependent. For example, hnRNP A1 often acts as a repressor when bound to exons but can enhance splicing when bound to introns. RBFOX2 is associated with exon skipping when bound upstream of that exon, but triggers exon inclusion when bound downstream of the exon; (C) Depending on the identity of the factors that recognize them, the combinatorial configuration of regulatory elements leads either to synergy (left) or to antagonism (right) [30]. Inhibitory and stimulatory factors are shown as black and white, respectively; (D) The structure of the chromatin and the phosphorylation status of the CTD of RNA polymerase II affect the speed of transcription, which in turn impacts the time given for the binding and assembly of regulatory complexes, hence affecting splice site selection [39]; (E) Post-translational modifications of chromatin components and chromatin remodeling activities alter the recruitment of adaptor proteins and splicing regulators to modulate alternative splicing.

Figure 3. Molecular mechanisms controlling splice site selection. (A) A variety of splicing regulators, including hnRNP and SR proteins, bind to exon or intron splicing enhancers (ESE or ISE, respectively) and to exon or intron splicing silencers (ESS or ISS, respectively) to control splice site recognition and utilization; (B) The activity of splicing regulators is often position-dependent. For example, hnRNP A1 often acts as a repressor when bound to exons but can enhance splicing when bound to introns. RBFOX2 is associated with exon skipping when bound upstream of that exon, but triggers exon inclusion when bound downstream of the exon; (C) Depending on the identity of the factors that recognize them, the combinatorial configuration of regulatory elements leads either to synergy (left) or to antagonism (right) [30]. Inhibitory and stimulatory factors are shown as black and white, respectively; (D) The structure of the chromatin and the phosphorylation status of the CTD of RNA polymerase II affect the speed of transcription, which in turn impacts the time given for the binding and assembly of regulatory complexes, hence affecting splice site selection [39]; (E) Post-translational modifications of chromatin components and chromatin remodeling activities alter the recruitment of adaptor proteins and splicing regulators to modulate alternative splicing.

2.1. Small Molecules Targeting DNA Duplex

2.1.1. Small Molecules Forming Covalent Bonds with DNA Duplex

Psoralen or psoralene has been used as mutagen to treat skin diseases including psoriasis and vitiligo. Psoralen interacts with standard DNA duplex, through the 5,6 double bond in pyrimidine ring, which can stop replication and transcription. 4′-(Hydroxymethyl)-4,5′,8-trimethylpsoralen, short for HMT, has reasonable water solubility and high DNA binding affinity, as shown in Figure 1A. Figure 2B shows the crystal structure of a short DNA complex with HMT reported by Spielmann et al. [13]. The HMT structure crosses both the minor and major grooves through the 5,6 double bonds of pyrimidine ring and two thymines. There are apparent stacking interactions from the ring system of HMT and the bases nearby. The high binding affinity of HMT and duplex results from aromatic rings stacking and hydrophobic interactions. The crystal study of the complex of DNA and HMT showed the Holliday junction was formed in the adduct of DNA-HMT. The interaction between DNA and HMT is related with sequence, which may play a role in repairing the psoralen damage in chemotherapy treatment. The conformation of DNA is extremely twisted at the bases of thymine connected with the hexatomic pyrone of the molecule.

Aflatoxin B1 (AFB) is a common contaminant in a variety of foods including peanuts, cottonseed meal, corn, and other grains as well as animal feeds. Aflatoxin B1 is considered the most toxic aflatoxin and it is highly implicated in hepatocellular carcinoma (HCC) in humans [14]. The epoxide metabolite of AFB can react with duplex DNA to produce a cationic adduct (Figure 2A). An NMR structure of this adduct has been resolved and reported by Stone [15], as shown in Figure 2B. This NMR structure shows that Aflatoxin B1 substructure inserts into DNA duplex strand. The adduct acts as a covalent binder, which can crosslink to duplex DNA conformation inducing the modification and change in the DNA conformation. The changed DNA will stop interacting with related proteins, which can block the process of replication or transcription.

Another DNA duplex adduct, a synthetic N4C–ethyl–N4C (Figure 3A), was discovered to interact with DNA by Miller [16]. The DNA study indicated that the ethyl cross-linked the base pairs in the structure solution. However, the ethyl linker does not remarkably change the B-form DNA conformation [17], as shown in Figure 3B.

Similar interactions were reported in the trimethylene related structures (Figure 4A) and DNA duplex conformation [18]. Two DNA duplexes structures were resolved and submitted to Protein Data Bank. Although the stacking between the linked bases and neighbor bases was different, both of the two structures indicated that the duplexes could maintain the hydrogen bonds and the B-form geometry (Figure 4B,C).

Mitomycin C (Figure 5A) is a member of mitomycins, which contains aziridine substructure and are extracted from natural products isolated from streptomycetes. The molecule is used as a chemotherapy drug for cancer. Mitomycin C alkylates the guanine in C–G base pairs of the DNA conformation [19]. The crystal complex of a short DNA with mitomycin C was reported by Patel et al. [20]. Unlike other standard DNA duplexes, the mitomycin alkylated DNA duplex shows A-form base pair stacking and B-form sugar puckers. The ring of mitomycin locates in the minor groove. And the ring of indoloquinone forms a 45° to the helix axis, as shown in Figure 5B.

As mentioned, these small molecules can modify and change the overall conformation of the conformation by cross-linking to DNA duplexes. The changed DNA conformations will stop interacting with the corresponding biological partners, thus the transcription or replication process will be blocked. This is how this kind of molecules works in the chemotherapeutics treatment. These molecules showed high toxicity in normal cells, just because of its low selectivity. To decrease resistance and promote selectivity, the comprehension of the interaction mechanism between DNA and drugs will be of significant importance.

2.1.2. Small Molecules Targeting with DNA Duplex in Minor Groove

To affect the gene expression, the small molecules can not only insert into the DNA strand, but also bind to the major or minor grooves of high binding affinity. For minor groove, typical small molecules are heterocylic dications and polyamides, including netropsin, berenil, and pentamidine, as shown in Figure 6. Originally, these molecules were found to bind AT-rich areas preferentially. Then, these molecules appeared near G–C and C–G base pairs [21,22]. The molecules targeting minor groove have been studied as a series of therapeutics with anti-viral, anti-tumor and anti-bacterial activities [23,24,25]. Some structural studies have been investigated as the basis of the design of small molecules [26,27,28,29,30]. The crystal complex of DNA duplex and Hoechst 33528 is shown in Figure 7. The Hoechst 33528 is clamped in the minor groove of the DNA structure.

Except for the drug-like molecules, some unusual compounds were also identified to locate in the minor groove of DNA. An 8 ring molecule (Figure 8A) was reported that it can bind to a DNA duplex [31]. The resolved structure indicates that the DNA conformation has been remarkably changed. The minor groove has been widened, and the major groove has been narrowed. The minor and major grooves are both approximate 4 Å (Figure 8B). Additionally, the helix center direction is twisted more than 18° to the major groove, in comparison with the inherent counterparts. The distortion provides a fundamental element for supporting the concept of allosteric change of the DNA conformation with inhibitors locating in minor grooves.

The small molecules can bind to nonstandard DNA duplex as well. Hoogsteen base pairs may occur in alternating A–T sequences, which are abundant in eukaryotic animals. Because of the lack of structural information, however, there is little analysis in this field. Recently, the complex of Hoogsteen DNA duplex and pentamidine (Figure 9A) was determined and reported [32]. The X-ray DNA structure presents a mixture of Watson–Crick and Hoogsteen pattern (Figure 9B). Unlike previous minor binders, pentamidine does not bind to the minor groove totally. Only the center of the pentamidine is bound to the minor groove, leaving the positively charged ends detached from the DNA and free to interact with phosphate groups from adjacent duplexes in the crystal. This new binding pattern has potent inspiration for antibacterial design of pentamidine.

2.1.3. Small Molecules Targeting with DNA Duplex in Major Groove

For major groove studies, there are less reports of small molecule targeting information. Due to the requirement of larger compounds, few complexes of molecules and DNA major grooves are determined. Although majority of carbohydrates bind to the DNA minor grooves, some of them can show binding affinity to DNA major groove. The reason for this major groove binding is the large size of carbohydrates and their hydrophilic and hydrophobic substructures. Some classic major groove binders are listed in Figure 10 [33].

Neocarzinostatin and related derivatives have been used as anti-tumor agents for decades [34,35]. This kind of molecule can form a complex with protein and damage DNA by hydrogen abstraction [36]. In order to understand the mechanism and interaction between DNA duplex and molecule, the complex of the DNA duplex and neocarzinostatin were reported. Neocarzinostatin-gb and neocarzinostatin-glu were used as ligand to bind to the DNA duplex. As shown in Figure 11, both of the two complex structures are observed [37,38]. The majority of the neocarzinostatin-gb binds to the major groove. The two rings of the molecule are close to each other in the structure. However, the neocarzinostatin-glu binds to the minor groove. The differences of the two binding modes provide significant inspiration into small molecule-DNA specific targeting.

2.1.4. Small Molecules Intercalating into DNA Duplex

Inserting a molecule between the base pairs of DNA is another binding pattern of small molecules targeting DNA duplexes. Inserting to the base pairs can interrupt DNA replication and transcription. The insertion results from hydrophobic, hydrogen bonding and van der Waals forces [39]. The insertion lengthens the length of DNA and reduces the helical twist [40,41]. In previous study, only molecules with fused ring systems were found to insert into the base pairs of DNA duplexes. Later, molecules with electrophilic or cationic groups behaved the same insertion pattern. However, the ring systems were not required [42,43]. Various compounds have been identified as intercalators, including ditercalinium, dactinomycin, daunomycin and adriamycine. Intercalators can be used as anti-fungal, anti-tumor, and anti-neoplastic agents. However, due to the toxicity, most of the compounds were failed during the clinical trial [44].

Daunomycin (trade name Cerubidine) is used to treat various cancers as chemotherapy drug [45]. Specifically, the most common use is treating some types of leukemia. Daunomycin (Figure 12A) consists of a planar ring, an amino sugar structure and a fused cyclohexane ring system. A lot of structural studies have been investigated to understand the interaction between DNA duplex and molecule [28,46,47,48,49,50,51]. Most of the structures indicate that two daunomycin molecules insert into the G–C steps with the sugar moiety in duplex, as shown in Figure 12B. Through sequence selectivity investigation by different biophysical methods, it was revealed that this ligand prefers a purine-pyrimidine step and the DNA duplexes are usually elongated or twisted after binding to the small molecule.

Adriamycin (Figure 12C) contains an extra hydroxyl group, compared to daunomycin. Although, the daunomycin and adriamyc in have nearly the same structure, their functions are remarkably diverse. Adriamycin is used to treat the tumor, while daunomycin is an effective drug in leukemias [47]. The complex structures of DNA duplex and compounds have been determined. In order to understand the differences between the two compounds, the complexes of the two molecules with a same DNA were resolved. The two complex structures showed a little differences between each other. Moreover, the hydroxyl group of the adriamycin interacts with a water molecule in the complex (Figure 12D).

Ditercalinium, as shown in Figure 13A, is regarded as a DNA intercalator in treatment of cancer. The structure of ditercalinium is a dimer of pyridocarbozole. Thus, the molecule can bind to DNA duplex by bis-intercalation and then induce DNA repair in eukaryotic or prokaryotic cell [52,53,54]. Afterwards, pyridocarbozole derivatives and related bioactivity have been studied [55,56,57]. The structure of DNA duplex and ditercalinium showed that the dimer of the molecule inserted to two G–C steps of the conformation (Figure 13B). The duplex of the DNA maintains a right-handed helix and has a binding site in major groove. The nitrogens with positive charge of the molecule make the charge interaction toward major groove of DNA [58]. In the complex structure, the helix is twisted at approximate 15° to minor groove of DNA. The mentioned structural features give inspiration on further research of optimizing this kind of molecules.

Actually, the specificity of DNA intercalators is the reason for drug toxicity. Molecules can bind to unexpected DNA sequence for new treatment. Cryptolepine (Figure 14A) is a good example for occasional drug discovery. Cryptolepine is isolated from Cryptolepis triangular and used as anti-malarial and cytotoxic agent [59]. This compound can insert with C–G rich sequences [60]. The crystal complex structure indicates that the drug interacts with the duplex in a base stacking insertion pattern (Figure 14B). The molecule and two consecutive C–G base pairs form a sandwich conformation with two C–G rich sequences. And the hexatomic ring stacks in the middle of cytosine and guanine. Structurally, cryptolepine was mildly twisted at about 6.8° between two aromatic rings. Besides, the charged nitrogen can improve the stability of the complex with the interaction between molecule and DNA duplex. The insertion into base pair steps and the molecular dissymmetry were important elements for this interaction.

2.1.5. Small Molecules Targeting with DNA Duplex through Multiple Binding Patterns

Small molecules can not only bind to DNA duplex in a single mode, but also interact with DNA duplex in multiple binding patterns, which will make the complex more stable [61,62]. PBD-BIMZ (Figure 15A) binds to a DNA duplex in a covalent link to a guanine base. The complex shows that the hybrid molecule orients in the minor groove (Figure 15B) [63]. Although the covalent binding distorts the DNA helix, the overall duplex still maintains the standard B-form conformation. It is revealed that covalent binding site in local region possesses specific twisted helix. By comparison, the piperazine ring shows a high flexibility with various conformations. Later, the same group reported another ligand (Figure 15C) in complex with the same DNA duplex. This hybrid compound binds to a guanine in a covalent link and the naphthalimide substructure embeds to (A–T)2 steps, as shown in Figure 15D [64]. The covalent binding will remarkably make the DNA and ligand stable. Moreover, because many molecules have low specificity for sequence, the multiple patterns can make the molecules sequence targetable.

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