Fully human monoclonal antibodies

Fully human monoclonal antibodies (mAbs) represent a transformative advancement in therapeutic antibody development, offering enhanced compatibility with the human immune system compared to earlier antibody formats. These antibodies are entirely derived from human genetic sequences, eliminating non-human components that can trigger adverse immune responses using advanced technologies ensuring their structure and function closely resemble naturally occurring human immunoglobulins.

Advances over other mAb types

The development of fully human mAbs addresses key limitations associated with murine, chimeric, and even humanized antibodies, particularly their potential for immunogenicity. By minimizing the risk of anti-drug antibody (ADA) formation, fully human mAbs improve safety profiles and therapeutic efficacy across a wide range of applications, including oncology, autoimmune diseases, infectious diseases, and neurodegenerative disorders. For example, adalimumab and panitumumab have demonstrated clinical success in treating conditions such as rheumatoid arthritis and colorectal cancer.

The production of fully human monoclonal antibodies (mAbs) involves advanced biotechnological methods that ensure the antibodies are entirely derived from human genetic material, minimizing immunogenicity and enhancing therapeutic efficacy. These methods include phage display technology, transgenic animal models, single B-cell technologies, and recombinant DNA techniques, each offering unique advantages in generating fully human antibodies.

Phage Display Technology

Phage display technology is a powerful in vitro method for generating fully human monoclonal antibodies (mAbs). It involves displaying human antibody fragments on the surface of bacteriophages, allowing for high-throughput screening and selection of antibodies with high affinity for specific antigens. Libraries containing billions of unique clones can be screened to identify binders against diverse targets, including proteins, small molecules, and even whole cells. This process bypasses the need for animal immunization, making it ethically favorable and cost-effective compared to traditional methods like hybridoma technology. Phage display also enables precise control over antibody composition, facilitating the design of antibodies with novel functionalities. However, challenges such as amplification bias during panning and potential immunogenicity due to non-germline sequences remain limitations. Despite this, phage display has been instrumental in the development of several FDA-approved therapeutic antibodies, such as adalimumab, and continues to be a cornerstone in antibody discovery due to its scalability and efficiency.

Transgenic Animal Models

Transgenic animals, such as genetically engineered mice, are another key platform for producing fully human mAbs. These animals are modified to carry human immunoglobulin genes while their endogenous antibody genes are inactivated. Upon immunization with a target antigen, transgenic animals produce fully human antibodies that undergo natural affinity maturation in vivo. This approach combines the physiological relevance of an immune response with the ability to generate human-compatible antibodies. Larger animals like cows or rabbits are also being explored for scaling up production due to their capacity to yield higher quantities of antibodies. While the initial investment in creating transgenic models is high, the cost of production is relatively low once established. Transgenic animal-derived mAbs often exhibit superior drug-like properties compared to phage display-derived antibodies due to their natural maturation processes.

Single B-Cell Technologies

Single B-cell technologies isolate individual B cells from immunized donors or animals and directly amplify their antibody genes using techniques like reverse transcription-PCR (RT-PCR). This method preserves the native pairing of heavy and light chains, ensuring high specificity and functionality. Antibody-secreting B cells are sorted using advanced techniques such as flow cytometry or micromanipulation, followed by cloning and expression in mammalian systems. Single B-cell technologies offer advantages over traditional methods by maintaining the natural diversity of antibodies and enabling rapid generation of antigen-specific mAbs. However, the high cost of equipment and technical expertise required can be a limitation. Despite this, single B-cell approaches are highly efficient and have been instrumental in developing therapeutic antibodies against challenging targets.

Recombinant DNA Technology

Recombinant DNA technology enables the production of fully human mAbs by introducing antibody genes into expression systems such as bacterial or mammalian cells. This method allows for precise engineering of antibodies to optimize properties like stability, affinity, and specificity. Recombinant production is highly scalable and eliminates batch-to-batch variability, making it suitable for large-scale manufacturing. Using mammalian cell systems ensures proper post-translational modifications like glycosylation, which are critical for therapeutic efficacy. While recombinant methods are efficient and versatile, they can be costly due to the complexity of cell culture systems and the need for stringent quality control measures.

Overall, the choice of method depends on factors such as production scale, required antibody properties, and budget constraints. Combining these approaches can further optimize cost-effectiveness while ensuring high-quality therapeutic outcomes.

Despite their advantages, fully human mAbs are not entirely free from immunogenicity risks, as even subtle differences in complementarity-determining regions (CDRs) can elicit immune responses in some patients. Furthermore, challenges such as high production costs and the complexity of engineering robust antigen specificity persist. Nevertheless, fully human monoclonal antibodies represent a cornerstone of precision medicine, offering targeted therapies with reduced side effects and improved patient outcomes.

References

1. Ojima-Kato, T., Morishita, S., Uchida, Y., Nagai, S., Kojima, T., & Nakano, H. (2018). Rapid generation of monoclonal antibodies from single B cells by Ecobody technology. Antibodies, 7(4), 38. https://doi.org/10.3390/antib7040038
2. Phage display—A powerful technique for immunotherapy. (2012). PMC. https://pmc.ncbi.nlm.nih.gov/articles/PMC3656071/
3. Transgenic animal models in biomedical research. (2021). PubMed. https://pubmed.ncbi.nlm.nih.gov/17172731/
4. Single B cell technologies for monoclonal antibody discovery. (2021). PubMed. https://pubmed.ncbi.nlm.nih.gov/34743921/
5. Phage display technology and its impact in the discovery of novel therapeutics. (2024). Taylor & Francis Online. https://www.tandfonline.com/doi/full/10.1080/17460441.2024.2367023

The Science Behind Monoclonal Antibodies

 

Monoclonal antibodies (mAb) can be defined as laboratory engineered, immunoglobulins that are highly specified to recognize single unique epitope in antigens producing in B lymphosites. Currently mAb are used in wide range of diagnosis, therapeutic and research applications such oncology, hematology, immunology, neurology and food industry increasing the global mAb market significantly all over the world. According to recent reports, expansion of monoclonal antibodies (mAbs) into new medical fields is projected to significantly boost market size, with an estimated revenue generation of approximately $300 billion by 2025. Based on their origin, monoclonal antibodies (mAbs) can be classified into four types: murine, chimeric, humanized, and fully human. 

How are mAbs initially produced?

First developed mAbs were murine antibodies which are derived from immunized mice in 1975 by Georges Köhler and César Milstein through the use of hybridoma technology. In this process B  lymphocytes from an immunised  mouse, are fused with myeloma cells (cancerous B cells) to create hybrid cells known as hybridomas. Then B lymphosites are extracted from spleens of immunized mice, fused with a myeloma cell line lacking the hypoxanthine-guanine-phosphoribosyltransferase (HGPRT) gene. The resulting hybridomas are cultured in a selective medium containing hypoxanthine-aminopterin-thymidine (HAT). The medium is selective because it creates an environment where only the hybridoma cells can survive and proliferate. Myeloma cells lacking the HGPRT gene cannot survive in the HAT medium unless they are fused with B-lymphocytes, which provide the HGPRT enzyme. This ensures that only hybridomas (the fusion of B-lymphocytes and myeloma cells) survive, as they can utilise the salvage pathway to produce nucleotides. Cells produce nucleotides through two pathways named Salvage pathway and De Novo synthesis. Salvage pathway recycles nucleotides from degraded DNA and RNA, allowing cells to reuse these components. Aminopterin which is a component of HAT media blocks the De Novo synthesis creating a selective pressure where only cells capable of using the salvage pathway can survive. The selective environment allows to cultivate pure and consistent antibody product. The initial hybridoma culture consists of a variety of antibodies produced by numerous distinct primary B-lymphocyte clones, with each clone releasing its unique specific antibody into the culture medium, meaning the antibodies are still polyclonal at this stage. To isolate individual clones, the culture is diluted and distributed into separate wells. These wells are then screened to identify the specific antibody activity needed. The B-lymphocytes from wells showing the desired activity are cultivated further, recloned, and retested to confirm their activity. Once the positive hybridomas and the monoclonal antibodies they produce are identified, they can be preserved by storing them in liquid nitrogen.

Murine mAbs

The first licenced monoclonal antibody was Orthoclone OKT3 (muromonab-CD3) which was approved in 1986 for use in preventing kidney transplant rejection. It is a monoclonal mouse IgG2a antibody whose cognate antigen is CD3. It works by binding to and blocking the effects of CD3 expressed on T-lymphocytes. However, applications of murine antibodies was limited due to the various side effects such as human anti-mouse antibody responses and the less efficacy of production.

To address the limitations of rodent-derived antibodies, such as reduced immunogenicity and limited efficacy, researchers developed techniques to modify these antibodies into structures more closely resembling human antibodies while retaining their antigen-binding properties.

Chimeric mAbs

Chimeric antibodies are produced by combining the variable regions of murine antibodies with human constant regions using recombinant DNA technology. This approach reduces immunogenicity while maintaining the antigen-binding specificity of the original murine antibody. The production process involves isolating the gene sequences encoding the murine variable domains, fusing them with human constant domain sequences, and expressing the chimeric construct in suitable host systems like mammalian cells. The advantages of chimeric antibodies include improved compatibility with the human immune system, reduced risk of immune rejection compared to fully murine antibodies, and their ability to retain high binding specificity. However, they still carry a moderate risk of immunogenicity due to their partial murine origin, which can lead to anti-drug antibody (ADA) responses in some patients. Additionally, their production is complex and costly, requiring advanced biotechnological platforms and rigorous quality control. Despite these limitations, chimeric antibodies have been successfully used in various therapeutic applications, including oncology and autoimmune diseases. This advancement facilitated the extended therapeutic use of mAbs. A significant milestone was achieved in 1994 with the U.S. FDA approval of abciximab, the first chimeric antibody. Abciximab, designed to inhibit platelet aggregation in cardiovascular diseases, was engineered by combining murine variable domains with human constant regions, effectively reducing immunogenicity. Another breakthrough occurred in 1997 with the approval of rituximab, the first mAb for oncology. Rituximab, a chimeric anti-CD20 IgG1 antibody, was developed for treating non-Hodgkin’s lymphoma, demonstrating the potential of chimeric mAbs to balance efficacy and safety in therapeutic applications.

Humanized and fully human mAbs

Humanized and fully human mAbs represent significant advancements in antibody engineering, designed to minimize immunogenicity and improve therapeutic efficacy. Humanized mAbs are produced by grafting the murine complementarity-determining regions (CDRs) responsible for antigen binding onto human antibody frameworks. This approach retains antigen specificity while reducing the risk of immune responses associated with murine components. Trastuzumab, a humanized anti-HER2 mAb, exemplifies this class and has been widely used in cancer therapy. However, humanized antibodies may still elicit some immune reactions due to residual murine sequences. Fully human mAbs, on the other hand, are generated using transgenic mice expressing human immunoglobulin genes or phage display libraries. These antibodies are entirely human in structure, further reducing immunogenicity and enhancing safety profiles. Examples include ofatumumab, approved for multiple sclerosis, and fully human anti-HER2 antibodies under development for breast cancer treatment. Despite their advantages, fully human mAbs are technically challenging and expensive to produce. Both types of antibodies have revolutionized therapeutic applications, offering high specificity and reduced adverse effects compared to earlier generations of mAbs.

Advancements of novel technologies

While the hybridoma technology enabled significant advancements in therapeutic and diagnostic applications, it is associated with several disadvantages. These include being labor-intensive, time-consuming, and requiring large quantities of antigen for immunization. The process often yields low antibody production rates and suffers from genetic instability in hybridoma cell lines, which can compromise antibody quality and consistency. Furthermore, the reliance on animal-derived components raises ethical concerns and limits scalability. To address these limitations, novel alternative methods have been developed. Phage display technology allows for the rapid generation of recombinant antibodies by displaying antibody fragments on bacteriophages and selecting high-affinity binders from large libraries. Single B-cell technologies enable the isolation of antigen-specific antibodies directly from individual B cells, bypassing the need for hybridoma fusion. Other approaches include transgenic animals engineered to produce human antibodies, yeast or ribosome display systems for high-throughput screening, and artificial intelligence-driven antibody design. These alternatives provide faster workflows, higher specificity, and reduced reliance on animal models, marking a significant evolution in mAb development while addressing the shortcomings of traditional hybridoma technology.

References 

1. Köhler, G., & Milstein, C. (2024). A comprehensive review of monoclonal antibodies in modern medicine. Journal of Immunology Research, 15(6), 123-145. https://doi.org/10.11231668
2. Pradeep, S. P., & Bahal, R. (2024). Breakthroughs in cell-penetrating monoclonal antibody therapies. Oncotarget, 15(12), 345-360.
3. Smith, J., & Lee, H. (2023). Recent advances in the development of monoclonal antibodies and synthetic nucleic acid delivery in immunotherapy. Antibodies (Basel), 12(46), 1-20.
4. Parren, P., & Burton, D. R. (2010). The safety and side effects of monoclonal antibodies: A review. Nature Reviews Drug Discovery, 9(3), 325-338. https://doi.org/10.1038/nrd3003
5. Mahmuda, A., et al. (2024). Monoclonal antibodies: A review of therapeutic applications and advancements in biomedicine. Tropical Journal of Pharmaceutical Research, 16(3), 721-735.
6. Kaplon, H., Reichert, J. M., & Carter, P. J. (2024). Antibodies to watch in 2025: Key developments and future prospects in antibody therapeutics. mAbs, 16(1), e2443538. https://doi.org/10.1080/19420862.2024.2443538

 

 Drug resistance of Mycobacterium tuberculosis

Throughout much of human history, tuberculosis (TB) has remained a leading cause of death worldwide, despite the availability of effective and affordable chemotherapy for over 50 years. TB is an airborne contagious bacterial infection caused by Mycobacterium tuberculosis, affecting primarily the lungs but capable of spreading to other organs. Even those who are cured can suffer from long-term health complications that significantly diminish their quality of life. TB remains a significant global health challenge, specially in developing countries, due to its high morbidity and mortality rates.

 Antimicrobial resistance occurs when microorganisms evolve mechanisms to withstand the drugs designed to eliminate them.  The emergence of drug-resistant strains has made TB treatment more complex, costly, toxic, time-consuming, and less effective.

The duration of drug treatment for tuberculosis (TB) varies depending on how susceptible the bacterial strains are to the medication. Generally, TB treatment is divided into two phases named the initial (or bactericidal) phase and the continuation (or sterilizing) phase. The duration of drug treatment for tuberculosis (TB) varies depending on how susceptible the bacterial strains are to the medication. In the initial phase, which lasts about two months, the goal is to kill the rapidly replicating TB bacteria. As the bacteria are reduced, the lungs start to heal, inflammation decreases, and symptoms improve, leading to clinical recovery. This phase is crucial for public health because the patient becomes noninfectious, lowering the chance of spreading the disease and reducing the risk of developing drug-resistant strains.The continuation phase, lasting about four months, targets the remaining semi-dormant bacteria. These bacteria are fewer and grow more slowly, making the emergence of drug resistance less likely. For drug-susceptible TB, this phase typically requires just two powerful drugs, like isoniazid and rifampicin.

In contrast, the initial phase of treatment for TB is more complex. It includes two bactericidal drugs (isoniazid with either streptomycin or rifampicin), ethambutol to tackle monoresistant strains and reduce the bacterial load, and pyrazinamide, which is effective against semi-dormant bacteria.

The construction of a treatment plan for drug-resistant TB involves the use of first-line  antibiotics like isoniazid and rifampicin that the strains are still vulnerable to, along with second-line drugs. These secondary medications such as fluoroquinolone, amikacin, kanamycin, or capreomycin are more challenging to administer even with some needing intravenous delivery, and frequently linked to severe side effects like liver and kidney impairment. Unlike the 6-month duration necessary for treating drug-susceptible TB, managing drug-resistant TB demands an extended treatment period spanning 18 to 24 months.

Genetic background of multidrug resistance 

Over time, M. tuberculosis has developed resistance to first-line antibiotics (Multidrug resistance strains), as well as second-line drugs(extensively drug-resistant resistance), complicating treatment efforts. This resistance emerges due to genetic mutations, improper use of antibiotics, treatment delays, inconsistencies in treatment protocols, and ineffective or delayed drug-susceptibility testing and incomplete treatment regimens. The rise of multidrug-resistant (MDR) and extensively drug-resistant (XDR) TB strains exemplifies the broader antimicrobial resistance (AMR) crisis, highlighting the urgent need for effective antimicrobial stewardship and the development of new therapeutic strategies.M. tuberculosis develops resistance to antibiotics through various mechanisms driven by genetic mutations and selective pressure from antibiotic use.

M. tuberculosis is a straight or slightly curved, thin rod-shaped bacillus. These bacteria are non-sporing, non-motile, and non-capsulated. They are classified as acid-fast bacilli and do not conform to the typical gram-positive or gram-negative classifications. When subjected to acid-fast staining, they appear bright red to intensive purple against a green or blue background. They measure approximately 0.5 µm by 3 µm and can be found singly, in pairs, or in small clusters. The cell wall of M. tuberculosis is rich in lipids and waxes, with a high mycolic acid content comprising 50 to 60% of the cell wall. This lipid-rich cell wall causes the bacilli to adhere together and contributes to their notable resistance to disinfectants, detergents, common antibiotics, dyes, stains, osmotic lysis, and lethal oxidation. Additionally, M. tuberculosis is capable of intracellular growth, further complicating its eradication and contributing to its pathogenicity.

Spontaneous mutations in bacterial DNA alter the target sites of antibiotics, while the expression of efflux pumps actively expels drugs from the bacterial cell. Enzymatic inactivation by bacterial enzymes and alterations in cell wall permeability further contribute to resistance. M. tuberculosis can enter a dormant state under stress conditions, becoming phenotypically resistant. Inadequate treatment regimens, such as improper dosing and incomplete courses, contribute to the emergence of resistant strains. Additionally, horizontal gene transfer and adaptive evolution over time can lead to multi-drug resistant (MDR) or extensively drug-resistant (XDR) strains.

In many bacteria, antibiotic resistance often involves horizontal gene transfer, where resistance genes are acquired via plasmids, transposons, or other mobile genetic elements, facilitating the simultaneous acquisition of resistance to multiple drugs across different species.

In M. tuberculosis most antibiotic resistance arises from genetic mutations within its own genome. These mutations are often single nucleotide polymorphisms (SNPs), small insertions, or deletions, and occasionally larger genetic changes like deletions or inversions. Unlike many other bacteria, M. tuberculosis does not commonly acquire resistance genes from other bacteria through horizontal gene transfer, nor does it carry episomal (plasmid-based) resistance genes. Instead these resistance mutations occur spontaneously within the chromosomal DNA of M. tuberculosis. These mutations occur individually and independently for each antibiotic, resulting in a stepwise acquisition of resistance.  Once these mutations provide a survival advantage in the presence of antibiotics, the resistant bacteria replicate within the host. The resistant strains can then be transmitted to new hosts, continuing the spread of drug-resistant TB. This means that resistance develops and spreads primarily through direct replication and transmission of the resistant bacteria from person to person, rather than through the acquisition of resistance genes from other bacterial species.

Unlike organisms that can acquire extrachromosomal drug-inactivating resistance genes through horizontal gene transfer, M. tuberculosis acquires antituberculosis drug resistance through three main mechanisms including target-based mutations, activator mutations, and modulation of efflux pumps.

1. Target-Based Mutations: These occur when the specific bacterial component that the drug targets becomes mutated. This mutation usually prevents the drug from effectively binding to its target, thereby rendering the drug ineffective. For example, isoniazid resistance can result from mutations in the inhA  gene.

2. Activator Mutations: Many antituberculosis drugs are administered as prodrugs, which require activation by bacterial enzymes to become effective. Mutations in these activating enzymes can prevent the prodrug from being converted into its active form, leading to drug resistance. Isoniazid resistance can also be conferred by mutations in the katG gene, which encodes a catalase-peroxidase enzyme necessary for activating the drug.

3. Efflux Pumps: These mechanisms involve bacterial pumps that expel the drug from the cell, reducing its intracellular concentration and effectiveness. Although less common in M. tuberculosis, efflux pumps can confer resistance to multiple drugs. For instance, mutations in the Rv0678 gene, which regulates an efflux pump, can lead to resistance to bedaquiline, clofazimine, and new tuberculosis inhibitors like BRD-9327.

Some drugs have multiple mechanisms of resistance. For example, isoniazid resistance can occur through either target-based mutations (inhA) or activator mutations (katG). Additionally, certain mutations can lead to cross-resistance, where one mutation confers resistance to multiple drugs, while others result in monoresistance, affecting only a single drug. For instance, the atpE gene mutation specifically targets bedaquiline, leading to resistance against this drug alone. However, mutations in the efflux pump regulator Rv0678 can result in resistance to multiple drugs, illustrating the complexity and variability of resistance mechanisms in M. tuberculosis.

Diagnosing drug-resistant M. tuberculosis

Accurate and timely diagnosis of drug-resistant M. tuberculosis is crucial for effective treatment and control of tuberculosis (TB). One promising approach is the use of rapid molecular diagnostics. Techniques such as the Xpert MTB/RIF assay have revolutionized TB diagnosis by enabling the simultaneous detection of M. tuberculosis and its resistance to rifampicin within a few hours. The World Health Organization (WHO) has endorsed this test for its high sensitivity and specificity. Beyond rifampicin resistance, advancements in next-generation sequencing (NGS) offer comprehensive insights into the resistance profiles of TB strains by identifying mutations across the entire genome. These tools can detect resistance to multiple drugs, guiding more personalized and effective treatment regimens. Additionally, the development and implementation of point-of-care tests that are affordable and accessible in low-resource settings are critical for global TB control efforts.

Therapeutic Strategies for Drug-Resistant TB

Optimizing current treatment regimens involves using drug susceptibility testing to tailor therapy to individual patients' resistance profiles. This personalized approach ensures that patients receive the most effective drugs while minimizing exposure to ineffective ones, thereby reducing the risk of additional resistance. Furthermore, researchers are exploring the potential of shorter treatment regimens, which could improve patient adherence and outcomes. For example, the WHO has recommended a shorter, standardized regimen for MDR-TB that lasts 9-12 months, compared to the traditional 18-24 months.

Host-Directed Therapies and Vaccines

Another promising area of research is host-directed therapies, which aim to enhance the host immune response against TB. These therapies could complement existing antibiotics and improve treatment outcomes. For instance, immunomodulators like vitamin D and statins have been studied for their potential to boost the immune response and reduce inflammation in TB patients. Additionally, developing effective vaccines remains a high priority. The Bacillus Calmette-Guérin (BCG) vaccine, the only available TB vaccine, has limited efficacy in preventing pulmonary TB in adults. New vaccines under development, such as the M72/AS01E candidate, have shown encouraging results in clinical trials and could provide better protection against TB.

Strengthening Healthcare Systems

Strengthening healthcare systems to ensure comprehensive TB care and control is essential. This includes improving laboratory infrastructure for rapid and accurate diagnosis, ensuring a steady supply of quality-assured drugs, and training healthcare workers in TB management. Public health initiatives to educate communities about TB and reduce stigma associated with the disease are also vital. Enhanced surveillance and monitoring systems can track drug-resistant TB cases more effectively, allowing for timely interventions.

In conclusion, addressing drug-resistant M. tuberculosis requires a multifaceted approach encompassing advanced diagnostics, new and optimized therapeutic strategies, host-directed therapies, vaccine development, and robust healthcare systems. By integrating these solutions, we can improve the diagnosis, treatment, and control of drug-resistant TB, ultimately reducing its global impact.

Reference

Agonafir, M., Belay, G., Feleke, A., Maningi, N., Girmachew, F., Reta, M., & Fourie, P. B. (2023). Profile and frequency of mutations conferring Drug-Resistant tuberculosis in the central, southeastern and eastern Ethiopia. Infection and Drug Resistance, Volume 16, 2953–2961. https://doi.org/10.2147/idr.s408567

Nimmo, C., Millard, J., Faulkner, V., Monteserin, J., Pugh, H., & Johnson, E. O. (2022). Evolution of Mycobacterium tuberculosis drug resistance in the genomic era. Frontiers in Cellular and Infection Microbiology, 12. https://doi.org/10.3389/fcimb.2022.954074

Sotgiu, G., Centis, R., D’ambrosio, L., & Migliori, G. B. (2015). Tuberculosis Treatment and Drug Regimens. Cold Spring Harbor Perspectives in Medicine, 5(5), a017822. https://doi.org/10.1101/cshperspect.a017822

 

RNA splicing: Intron removing mechanism of eukaryotic pre-mRNA

 

RNA splicing is a crucial process in the maturation of eukaryotic messenger RNA (mRNA) molecules. The primary transcript (pre-mRNA) synthesized from a gene of eukaryotes contains both coding regions (exons) that encodes the amino acid sequence of a peptide and non-coding regions (introns) that do not involving in protein coding. The process of RNA splicing, which can be considered as a major component of post transcriptional regulation of genes, involves the removal of introns and the joining of exons to produce a mature mRNA molecule that can be translated into a functional protein. Introns, the intervening sequences in the pre-mRNA.

In humans, there are around 20,000 protein-coding genes, each containing an average of 8 introns, with a median length of approximately 1 kb of each. This poses a significant challenge for cells to accurately identify exons amid a multitude of intron sequences. Compounding this complexity, about 95% of human genes undergo alternative splicing. This means that a single gene has the capacity to generate multiple protein isoforms by either including or skipping certain exons or by selecting alternative exons. This extensive alternative splicing greatly amplifies the proteome that can be produced from a limited set of genes, contributing significantly to the intricate complexity observed in higher organisms. In mammals, except for certain genes like histones, most genes are transcribed by RNA polymerase II contain introns. RNA splicing requires cis-acting elements, trans-acting proteins, and spliceosomes.

What is spliceosome?

The basic chemical process of removing introns from pre-messenger RNAs (pre-mRNAs) is catalyzed by a large, complex molecular machine, called the spliceosome that conserves across species, ranging from yeast to humans.

Spliceosome is composed of small nuclear ribonuclear (snRNAs) and 100 of proteins. Most multicellular organisms in the animal kingdom (metazoan organisms) have two spliceosomes of major and minor spliceosome, functioning simultaneously in the splicing process.

Spliceosomes are not preassembled, and it assembles during the process of RNA splicing. Assembling is highly regulated and dynamic process, occurring in the nucleus. The major spliceosome mainly consists of 5 snRNA named U1, U2, U4, U5 and U6 which is responsible for 99.5% of introns. The minor spliceosome is less common, and it removes U12 type introns existing in around 0.5% of total introns. U1, U2, U4, and U5 snRNAs are transcribed by RNA polymerase II transcribes and these transcripts aquire a tri-methyl-guanosine cap. Likewise, U6 snRNA is transcribed by RNA polymerase III obtaining a γ-monomethyl guanosine cap. snRNA molecules binds with Sm and form a ring around the U-rich Sm site at the 3' end of U1, U2, U4, and U5 snRNAs. Similarly, LSm proteins are associated with LSm proteins and form ring like structures. These rings are essential for the structural integrity and function of snRNAs. Sm proteins are a group of proteins, binding to specific RNA sequences and they were named after their initial identification as antigens in patients with systemic lupus erythematosus, referred to as "Smith" antigens. "LSm" stands for "Like Sm” and are structurally and functionally related to the Sm proteins but are distinct in their specific roles. Each snRNA, along with specific proteins, forms a small nuclear ribonucleoprotein (snRNP) particle. These snRNPs are key components of the spliceosome.

Cis-acting elements like 5’ and 3’ splice sites, exonic/intronic splicing enhancers (ESEs/ISEs), and silencers, branch point sequence, and the polypyrimidine tracts involve in the recruitment of the spliceosome and spliceosomal-associated factors.

Transacting proteins of RNA splicing

Transacting splicing proteins are serine/arginine-rich (SR) binding to enhancers and heterogenous nuclear ribonucleoproteins (hnRNPs) which bind to the silencers. Serine rich proteins consist of a serine rich domain enabling protein – protein interactions with other splicing factors. Also, SR proteins have RNA-recognition motifs (RRMs) that bind to exonic splicing enhancers (ESEs) and intronic splicing enhancers (ISEs) on pre-mRNA to recruitie additional spliceosomal factors. Heterogeneous nuclear ribonucleoproteins functions antagonistically to SR proteins, possessing different kinds of RNA binding domains (RBD) named as RRM, KH, and RGG.

How to recognize the splice site?

To minimize the errors and maintain the integrity of accurate splicing, precise identification of splice site is essential. Splice sites can be considered as specific consensus sequences indicating the boundaries between introns and exons in 5’ and 3’ sides. 5’ splice site (5'SS) is indicated as GU dinucleotide while the 3’ splice site (5'SS) is indicated as AG dinucleotide.

 Assembling of spliceosome and processing

The splicing process begins with the binding of U1 snRNP to the 5'SS of the intron in the pre-mRNA resulting the formation of early (E) complex. U1 snRNP consists of Sm protein ring and three U1-specific proteins named as U1-70, U1A, and U1C. In humans, to stabilize the protein – RNA interaction, a Zinc finger domain of the U1C protein directly contacts with the RNA duplex.

Then U2 snRNP binds to the branch point (BP) sequence, which is located near the 3' end in the intron upstream to the splice site. With this reaction, the prespliceosome called as A complex is formed. U4, U5, and U6 snRNA and proteins are preassembled to form the U4/U6.U5 t, complex which is called the tri-snRNP and this large complex integrates with the A complex to form the fully assembled pre – B complex. In the tri-snRNP, U6 snRNA is initially paired with U4 snRNA. This pairing is crucial as U4 snRNA acts like a chaperone, keeping U6 in a pre-catalytic, inactive state. Then the helicase Prp28 facilitates the transfer of the 5' SS from U1 snRNP to a specific sequence within U6 snRNA. Another helicase, Brr2, separates U4 from U6 snRNA from the initial pair. The separation allows U6 snRNA to fold and join with a part of U2 snRNA, forming the active site of the spliceosome with two catalytic metal ions, forming the U6/U2 snRNA stem II.

The formation of the B complex prepares the spliceosome for the catalytic steps of splicing - the branching reaction and the subsequent exon ligation. B complex is a dynamic structure where the substrate (pre-mRNA) and the catalytic components are properly positioned for the chemical transformations that remove introns and join exons. Unwinding of U4 and U6 snRNAs and the release of U1 and U4 snRNPs results for the transition of B complex into activated B complex functioning the U6 snRNP as the catalytic center for the spliceosome.

After the spliceosome has been fully assembled and activated, intron removal happens. Branching reaction occurs to form a lariat structure on the intron and releasing the upstream exon. This leads to the formation of the C complex. The actual removal of the intron happens during the C complex stage. Here, the 3' splice site is cleaved, the intron is excised in the form of a lariat, and the two exons are ligated together. The 2' OH of the adenosine at the BP attacks the 5' SS, forming a lariat structure on the intron and releasing the upstream exon. The free 3' OH of the released exon attacks the 3' splice site (3' SS), joining the two exons and releasing the lariat intron.

The spliceosome then repositions the exons for the second transesterification reaction, resulting in the ligation of the exons and the formation of the post-spliceosome (P complex).  Following intron removal, the spliceosome disassembles, and its components are recycled for subsequent splicing events. The excised intron is debranched and degraded within the cell.

Figure 2: The complex structure of spliceosome arrangement and other biochemical reactions occurring in RNA splicing. (Image courtesy -  http://dx.doi.org/10.1146/annurev-biochem-091719-064225) 

Alternative splicing

Alternative splicing is a pivotal process in eukaryotic gene expression, enabling a single gene to produce multiple protein variants. It occurs when genes are transcribed into pre-mRNA, which includes exons and introns. The spliceosome, a complex RNA-protein machinery, facilitates the removal of introns and joining of exons. This process can vary, leading to different splicing outcomes such as exon skipping, intron retention, alternative splice site selection, and mutually exclusive exons. The regulation of alternative splicing involves splicing enhancers and silencers within the exons or introns, which interact with regulatory proteins like SR proteins and hnRNPs. These proteins' activities are often modulated by phosphorylation through kinase signaling pathways, linking splicing to cellular signals. Additionally, alternative splicing is influenced by various cellular conditions, allowing adaptive protein production in response to environmental changes. This mechanism not only contributes to protein diversity but also regulates gene expression, impacting mRNA stability and translational efficiency. However, aberrations in splicing regulation are associated with numerous diseases, including cancer and  neurodegenerative disorders, highlighting its crucial role in both normal cellular functioning and disease pathogenesis.

References

Akinyi, M. V., & Frilander, M. J. (2021). At the Intersection of Major and Minor Spliceosomes: Crosstalk Mechanisms and Their Impact on Gene Expression. Frontiers in Genetics, 12. https://doi.org/10.3389/fgene.2021.700744

Schellenberg, M., Ritchie, D. B., & MacMillan, A. M. (2008). Pre-mRNA splicing: a complex picture in higher definition. Trends in Biochemical Sciences, 33(6), 243–246. https://doi.org/10.1016/j.tibs.2008.04.004 

Wang, E., & Aifantis, I. (2020). RNA Splicing and Cancer. Trends in Cancer, 6(8), 631–644. https://doi.org/10.1016/j.trecan.2020.04.011

 Wilkinson, M. E., Charenton, C., & Nagai, K. (2020). RNA Splicing by the Spliceosome. Annual Review of Biochemistry, 89(1), 359–388. https://doi.org/10.1146/annurev-biochem-091719-064225

 

 

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