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Why do mutations not take place in mRNA of higher eukaryotes?

Why do mutations not take place in mRNA of higher eukaryotes?


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Is it because it is too short-lived to be mutated? Both DNA and RNA are nucleic acids so how is mRNA protected? RNA viruses undergo mutations to evolve so I guess it is not immune to mutations


The premise of the questions suggests that mutations cannot take place in the mRNAs of higher eukaryotes. To answer your question I think it is important to consider two viewpoints:

First, from a theoretical point of view, since DNA and RNA are as you pointed out composed of nucleic acids, they both can be mutated if enough energy is provided (UV light, chemicals, etc) which invalidates the premise of the questions.

Now, from a practical point of view, as you mentioned most mRNAs molecules have a short half life typically in the minute to day range whereas DNA molecules exist during the whole existence of the organism.

While it can occurs that mRNAs are mutated people are not interested to study this aspect for the following reasons:

You pointed out in the comment section, a mutation in a mRNA molecule might lead to translated malformed protein which can easily be degraded. It will be only one mRNA transcript from thousands transcripts. With a short half life, the mRNA an proteins will be degraded which will not have a long standing impact of the cell/organism. As such it will be very difficult to observe a phenotype which will affect the whole host.

Thus RNAs mutations have only transient effects which will not affect the host in the long term. RNAs molecules are not more protected than DNA molecules just that they are short lived so the host is protected from the effects of RNA mutations.

Hope this helps!


Additionally DNA is inherently more stable due to the removal of the hydroxyl group from the C2 carbon on the ribose making it less reactive. This could be pointed to as an example of the limiting factor in the inherent complexity of any organism that uses RNA instead of DNA for long term data storage. That would be a discussion for another thread though.


Your supposition is incorrect. RNA does accumulate "mutations" because the RNA polymerase also makes errors. The misincorporation rate of bacterial RNA polymerase is ~10-5 per nucleotide (Traverse & Ochman, 2016). Compared to that, DNA polymerase has much higher fidelity and has misincorporation of ~10-10 per nucleotide per generation (Zhu et al., 2014).

As pointed out in the other answer, RNA mutational effects are short-lived as RNAs are short-lived. However, RNA mutations can contribute to phenotypic heterogeneity which can, in some cases, be beneficial to the organism (Ackermann, 2015).


There is nothing special about higher eukaryotes as replication and transcription process is quite similar across different domains of life. Viruses are special, however. They rely on higher mutation rate for faster adaptation.


Mutations do occur in mRNA. This can occur during transcription, when RNA polymerase incorporates a non-cognate nucleotide into the nascent RNA strand (this may be because the DNA forms a short lived tautomer from quantum tunnelling, or because ). Alternatively, just like DNA bases, RNA bases are inherently unstable due to things like deamination events, but RNA is also capable of base catalyzed hydrolysis. To say that mutations are not interesting in RNA is also wrong as RNA editing is something that may be used to treat numerous diseases. RNA editing is transient, which is useful for treating diseases caused by temporary changes in cell state and modifying disease-related signal transduction through editing serine, threonine and tyrosine residues to affect phosphorylation sites. mRNA mutations are also important in understanding the affects of splicing on phenotypes and I'm sure there are more reasons, but these are just off the top of my head. For more information about RNA editing see: DOI: 10.1126/science.aaq0180, a really cool potential of CRISPR


15.4: RNA Processing in Eukaryotes

  • Contributed by OpenStax
  • General Biology at OpenStax CNX
  • Describe the different steps in RNA processing
  • Understand the significance of exons, introns, and splicing
  • Explain how tRNAs and rRNAs are processed

After transcription, eukaryotic pre-mRNAs must undergo several processing steps before they can be translated. Eukaryotic (and prokaryotic) tRNAs and rRNAs also undergo processing before they can function as components in the protein synthesis machinery.


15.4 RNA Processing in Eukaryotes

By the end of this section, you will be able to do the following:

  • Describe the different steps in RNA processing
  • Understand the significance of exons, introns, and splicing for mRNAs
  • Explain how tRNAs and rRNAs are processed

After transcription, eukaryotic pre-mRNAs must undergo several processing steps before they can be translated. Eukaryotic (and prokaryotic) tRNAs and rRNAs also undergo processing before they can function as components in the protein-synthesis machinery.

MRNA Processing

The eukaryotic pre-mRNA undergoes extensive processing before it is ready to be translated. Eukaryotic protein-coding sequences are not continuous, as they are in prokaryotes. The coding sequences (exons) are interrupted by noncoding introns, which must be removed to make a translatable mRNA. The additional steps involved in eukaryotic mRNA maturation also create a molecule with a much longer half-life than a prokaryotic mRNA. Eukaryotic mRNAs last for several hours, whereas the typical E. coli mRNA lasts no more than five seconds.

Pre-mRNAs are first coated in RNA-stabilizing proteins these protect the pre-mRNA from degradation while it is processed and exported out of the nucleus. The three most important steps of pre-mRNA processing are the addition of stabilizing and signaling factors at the 5' and 3' ends of the molecule, and the removal of the introns (Figure 15.11). In rare cases, the mRNA transcript can be “edited” after it is transcribed.

Evolution Connection

RNA Editing in Trypanosomes

The trypanosomes are a group of protozoa that include the pathogen Trypanosoma brucei, which causes nagana in cattle and sleeping sickness in humans throughout great areas of Africa (Figure 15.12). The trypanosome is carried by biting flies in the genus Glossina (commonly called tsetse flies). Trypanosomes, and virtually all other eukaryotes, have organelles called mitochondria that supply the cell with chemical energy. Mitochondria are organelles that express their own DNA and are believed to be the remnants of a symbiotic relationship between a eukaryote and an engulfed prokaryote. The mitochondrial DNA of trypanosomes exhibit an interesting exception to the central dogma: their pre-mRNAs do not have the correct information to specify a functional protein. Usually, this is because the mRNA is missing several U nucleotides. The cell performs an additional RNA processing step called RNA editing to remedy this.

Other genes in the mitochondrial genome encode 40- to 80-nucleotide guide RNAs. One or more of these molecules interacts by complementary base pairing with some of the nucleotides in the pre-mRNA transcript. However, the guide RNA has more A nucleotides than the pre-mRNA has U nucleotides with which to bind. In these regions, the guide RNA loops out. The 3' ends of guide RNAs have a long poly-U tail, and these U bases are inserted in regions of the pre-mRNA transcript at which the guide RNAs are looped. This process is entirely mediated by RNA molecules. That is, guide RNAs—rather than proteins—serve as the catalysts in RNA editing.

RNA editing is not just a phenomenon of trypanosomes. In the mitochondria of some plants, almost all pre-mRNAs are edited. RNA editing has also been identified in mammals such as rats, rabbits, and even humans. What could be the evolutionary reason for this additional step in pre-mRNA processing? One possibility is that the mitochondria, being remnants of ancient prokaryotes, have an equally ancient RNA-based method for regulating gene expression. In support of this hypothesis, edits made to pre-mRNAs differ depending on cellular conditions. Although speculative, the process of RNA editing may be a holdover from a primordial time when RNA molecules, instead of proteins, were responsible for catalyzing reactions.

5' Capping

While the pre-mRNA is still being synthesized, a 7-methylguanosine cap is added to the 5' end of the growing transcript by a phosphate linkage. This functional group protects the nascent mRNA from degradation. In addition, factors involved in protein synthesis recognize the cap to help initiate translation by ribosomes.

3' Poly-A Tail

Once elongation is complete, the pre-mRNA is cleaved by an endonuclease between an AAUAAA consensus sequence and a GU-rich sequence, leaving the AAUAAA sequence on the pre-mRNA. An enzyme called poly-A polymerase then adds a string of approximately 200 A residues, called the poly-A tail . This modification further protects the pre-mRNA from degradation and is also the binding site for a protein necessary for exporting the processed mRNA to the cytoplasm.

Pre-mRNA Splicing

Eukaryotic genes are composed of exons , which correspond to protein-coding sequences (ex-on signifies that they are expressed), and intervening sequences called introns (int-ron denotes their intervening role), which may be involved in gene regulation but are removed from the pre-mRNA during processing. Intron sequences in mRNA do not encode functional proteins.

The discovery of introns came as a surprise to researchers in the 1970s who expected that pre-mRNAs would specify protein sequences without further processing, as they had observed in prokaryotes. The genes of higher eukaryotes very often contain one or more introns. These regions may correspond to regulatory sequences however, the biological significance of having many introns or having very long introns in a gene is unclear. It is possible that introns slow down gene expression because it takes longer to transcribe pre-mRNAs with lots of introns. Alternatively, introns may be nonfunctional sequence remnants left over from the fusion of ancient genes throughout the course of evolution. This is supported by the fact that separate exons often encode separate protein subunits or domains. For the most part, the sequences of introns can be mutated without ultimately affecting the protein product.

All of a pre-mRNA’s introns must be completely and precisely removed before protein synthesis. If the process errs by even a single nucleotide, the reading frame of the rejoined exons would shift, and the resulting protein would be dysfunctional. The process of removing introns and reconnecting exons is called splicing (Figure 15.13). Introns are removed and degraded while the pre-mRNA is still in the nucleus. Splicing occurs by a sequence-specific mechanism that ensures introns will be removed and exons rejoined with the accuracy and precision of a single nucleotide. Although the intron itself is noncoding, the beginning and end of each intron is marked with specific nucleotides: GU at the 5' end and AG at the 3' end of the intron. The splicing of pre-mRNAs is conducted by complexes of proteins and RNA molecules called spliceosomes.

Visual Connection

Errors in splicing are implicated in cancers and other human diseases. What kinds of mutations might lead to splicing errors? Think of different possible outcomes if splicing errors occur.

Note that more than 70 individual introns can be present, and each has to undergo the process of splicing—in addition to 5' capping and the addition of a poly-A tail—just to generate a single, translatable mRNA molecule.

Link to Learning

See how introns are removed during RNA splicing at this website.

Processing of tRNAs and rRNAs

The tRNAs and rRNAs are structural molecules that have roles in protein synthesis however, these RNAs are not themselves translated. Pre-rRNAs are transcribed, processed, and assembled into ribosomes in the nucleolus. Pre-tRNAs are transcribed and processed in the nucleus and then released into the cytoplasm where they are linked to free amino acids for protein synthesis.

Most of the tRNAs and rRNAs in eukaryotes and prokaryotes are first transcribed as a long precursor molecule that spans multiple rRNAs or tRNAs. Enzymes then cleave the precursors into subunits corresponding to each structural RNA. Some of the bases of pre-rRNAs are methylated that is, a –CH3 methyl functional group is added for stability. Pre-tRNA molecules also undergo methylation. As with pre-mRNAs, subunit excision occurs in eukaryotic pre-RNAs destined to become tRNAs or rRNAs.

Mature rRNAs make up approximately 50 percent of each ribosome. Some of a ribosome’s RNA molecules are purely structural, whereas others have catalytic or binding activities. Mature tRNAs take on a three-dimensional structure through local regions of base pairing stabilized by intramolecular hydrogen bonding. The tRNA folds to position the amino acid binding site at one end and the anticodon at the other end (Figure 15.14). The anticodon is a three-nucleotide sequence in a tRNA that interacts with an mRNA codon through complementary base pairing.

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    76 RNA Processing in Eukaryotes

    By the end of this section, you will be able to do the following:

    • Describe the different steps in RNA processing
    • Understand the significance of exons, introns, and splicing for mRNAs
    • Explain how tRNAs and rRNAs are processed

    After transcription, eukaryotic pre-mRNAs must undergo several processing steps before they can be translated. Eukaryotic (and prokaryotic) tRNAs and rRNAs also undergo processing before they can function as components in the protein-synthesis machinery.

    MRNA Processing

    The eukaryotic pre-mRNA undergoes extensive processing before it is ready to be translated. Eukaryotic protein-coding sequences are not continuous, as they are in prokaryotes. The coding sequences (exons) are interrupted by noncoding introns, which must be removed to make a translatable mRNA. The additional steps involved in eukaryotic mRNA maturation also create a molecule with a much longer half-life than a prokaryotic mRNA. Eukaryotic mRNAs last for several hours, whereas the typical E. coli mRNA lasts no more than five seconds.

    Pre-mRNAs are first coated in RNA-stabilizing proteins these protect the pre-mRNA from degradation while it is processed and exported out of the nucleus. The three most important steps of pre-mRNA processing are the addition of stabilizing and signaling factors at the 5′ and 3′ ends of the molecule, and the removal of the introns ((Figure)). In rare cases, the mRNA transcript can be “edited” after it is transcribed.


    The trypanosomes are a group of protozoa that include the pathogen Trypanosoma brucei, which causes nagana in cattle and sleeping sickness in humans throughout great areas of Africa ((Figure)). The trypanosome is carried by biting flies in the genus Glossina (commonly called tsetse flies). Trypanosomes, and virtually all other eukaryotes, have organelles called mitochondria that supply the cell with chemical energy. Mitochondria are organelles that express their own DNA and are believed to be the remnants of a symbiotic relationship between a eukaryote and an engulfed prokaryote. The mitochondrial DNA of trypanosomes exhibit an interesting exception to the central dogma: their pre-mRNAs do not have the correct information to specify a functional protein. Usually, this is because the mRNA is missing several U nucleotides. The cell performs an additional RNA processing step called RNA editing to remedy this.


    Other genes in the mitochondrial genome encode 40- to 80-nucleotide guide RNAs. One or more of these molecules interacts by complementary base pairing with some of the nucleotides in the pre-mRNA transcript. However, the guide RNA has more A nucleotides than the pre-mRNA has U nucleotides with which to bind. In these regions, the guide RNA loops out. The 3′ ends of guide RNAs have a long poly-U tail, and these U bases are inserted in regions of the pre-mRNA transcript at which the guide RNAs are looped. This process is entirely mediated by RNA molecules. That is, guide RNAs—rather than proteins—serve as the catalysts in RNA editing.

    RNA editing is not just a phenomenon of trypanosomes. In the mitochondria of some plants, almost all pre-mRNAs are edited. RNA editing has also been identified in mammals such as rats, rabbits, and even humans. What could be the evolutionary reason for this additional step in pre-mRNA processing? One possibility is that the mitochondria, being remnants of ancient prokaryotes, have an equally ancient RNA-based method for regulating gene expression. In support of this hypothesis, edits made to pre-mRNAs differ depending on cellular conditions. Although speculative, the process of RNA editing may be a holdover from a primordial time when RNA molecules, instead of proteins, were responsible for catalyzing reactions.

    5′ Capping

    While the pre-mRNA is still being synthesized, a 7-methylguanosine cap is added to the 5′ end of the growing transcript by a phosphate linkage. This functional group protects the nascent mRNA from degradation. In addition, factors involved in protein synthesis recognize the cap to help initiate translation by ribosomes.

    3′ Poly-A Tail

    Once elongation is complete, the pre-mRNA is cleaved by an endonuclease between an AAUAAA consensus sequence and a GU-rich sequence, leaving the AAUAAA sequence on the pre-mRNA. An enzyme called poly-A polymerase then adds a string of approximately 200 A residues, called the poly-A tail . This modification further protects the pre-mRNA from degradation and is also the binding site for a protein necessary for exporting the processed mRNA to the cytoplasm.

    Pre-mRNA Splicing

    Eukaryotic genes are composed of exons , which correspond to protein-coding sequences (ex-on signifies that they are expressed), and intervening sequences called introns (int-ron denotes their intervening role), which may be involved in gene regulation but are removed from the pre-mRNA during processing. Intron sequences in mRNA do not encode functional proteins.

    The discovery of introns came as a surprise to researchers in the 1970s who expected that pre-mRNAs would specify protein sequences without further processing, as they had observed in prokaryotes. The genes of higher eukaryotes very often contain one or more introns. These regions may correspond to regulatory sequences however, the biological significance of having many introns or having very long introns in a gene is unclear. It is possible that introns slow down gene expression because it takes longer to transcribe pre-mRNAs with lots of introns. Alternatively, introns may be nonfunctional sequence remnants left over from the fusion of ancient genes throughout the course of evolution. This is supported by the fact that separate exons often encode separate protein subunits or domains. For the most part, the sequences of introns can be mutated without ultimately affecting the protein product.

    All of a pre-mRNA’s introns must be completely and precisely removed before protein synthesis. If the process errs by even a single nucleotide, the reading frame of the rejoined exons would shift, and the resulting protein would be dysfunctional. The process of removing introns and reconnecting exons is called splicing ((Figure)). Introns are removed and degraded while the pre-mRNA is still in the nucleus. Splicing occurs by a sequence-specific mechanism that ensures introns will be removed and exons rejoined with the accuracy and precision of a single nucleotide. Although the intron itself is noncoding, the beginning and end of each intron is marked with specific nucleotides: GU at the 5′ end and AG at the 3′ end of the intron. The splicing of pre-mRNAs is conducted by complexes of proteins and RNA molecules called spliceosomes.


    Errors in splicing are implicated in cancers and other human diseases. What kinds of mutations might lead to splicing errors? Think of different possible outcomes if splicing errors occur.

    <!– <link window=”new” target-id=”fig-ch15_04_02″ document=””/>Mutations in the spliceosome recognition sequence at each end of the intron, or in the proteins and RNAs that make up the spliceosome, may impair splicing. Mutations may also add new spliceosome recognition sites. Splicing errors could lead to introns being retained in spliced RNA, exons being excised, or changes in the location of the splice site. –>

    Note that more than 70 individual introns can be present, and each has to undergo the process of splicing—in addition to 5′ capping and the addition of a poly-A tail—just to generate a single, translatable mRNA molecule.

    See how introns are removed during RNA splicing at this website.

    Processing of tRNAs and rRNAs

    The tRNAs and rRNAs are structural molecules that have roles in protein synthesis however, these RNAs are not themselves translated. Pre-rRNAs are transcribed, processed, and assembled into ribosomes in the nucleolus. Pre-tRNAs are transcribed and processed in the nucleus and then released into the cytoplasm where they are linked to free amino acids for protein synthesis.

    Most of the tRNAs and rRNAs in eukaryotes and prokaryotes are first transcribed as a long precursor molecule that spans multiple rRNAs or tRNAs. Enzymes then cleave the precursors into subunits corresponding to each structural RNA. Some of the bases of pre-rRNAs are methylated that is, a –CH3 methyl functional group is added for stability. Pre-tRNA molecules also undergo methylation. As with pre-mRNAs, subunit excision occurs in eukaryotic pre-RNAs destined to become tRNAs or rRNAs.

    Mature rRNAs make up approximately 50 percent of each ribosome. Some of a ribosome’s RNA molecules are purely structural, whereas others have catalytic or binding activities. Mature tRNAs take on a three-dimensional structure through local regions of base pairing stabilized by intramolecular hydrogen bonding. The tRNA folds to position the amino acid binding site at one end and the anticodon at the other end ((Figure)). The anticodon is a three-nucleotide sequence in a tRNA that interacts with an mRNA codon through complementary base pairing.


    Section Summary

    Eukaryotic pre-mRNAs are modified with a 5′ methylguanosine cap and a poly-A tail. These structures protect the mature mRNA from degradation and help export it from the nucleus. Pre-mRNAs also undergo splicing, in which introns are removed and exons are reconnected with single-nucleotide accuracy. Only finished mRNAs that have undergone 5′ capping, 3′ polyadenylation, and intron splicing are exported from the nucleus to the cytoplasm. Pre-rRNAs and pre-tRNAs may be processed by intramolecular cleavage, splicing, methylation, and chemical conversion of nucleotides. Rarely, RNA editing is also performed to insert missing bases after an mRNA has been synthesized.

    Visual Connection Questions

    (Figure) Errors in splicing are implicated in cancers and other human diseases. What kinds of mutations might lead to splicing errors? Think of different possible outcomes if splicing errors occur.

    (Figure) Mutations in the spliceosome recognition sequence at each end of the intron, or in the proteins and RNAs that make up the spliceosome, may impair splicing. Mutations may also add new spliceosome recognition sites. Splicing errors could lead to introns being retained in spliced RNA, exons being excised, or changes in the location of the splice site.

    Review Questions

    Which pre-mRNA processing step is important for initiating translation?

    What processing step enhances the stability of pre-tRNAs and pre-rRNAs?

    A scientist identifies a pre-mRNA with the following structure.


    What is the predicted size of the corresponding mature mRNA in base pairs (bp), excluding the 5’ cap and 3’ poly-A tail?

    Critical Thinking Questions

    Chronic lymphocytic leukemia patients often harbor nonsense mutations in their spliceosome machinery. Describe how this mutation of the spliceosome would change the final location and sequence of a pre-mRNA.

    Nonsense spliceosome mutations would eliminate the splicing step of mRNA processing, so the mature mRNAs would retain their introns and be perfectly complementary to the entire DNA template sequence. However, the mRNAs would still undergo addition of the 5’ cap and poly-A tail, and therefore each has the potential to be exported to the cytoplasm for translation.

    Glossary


    Eukaryotic mRNA Decay: Methodologies, Pathways, and Links to Other Stages of Gene Expression

    mRNA concentration depends on the balance between transcription and degradation rates. On both sides of the equilibrium, synthesis and degradation show, however, interesting differences that have conditioned the evolution of gene regulatory mechanisms. Here, we discuss recent genome-wide methods for determining mRNA half-lives in eukaryotes. We also review pre- and posttranscriptional regulons that coordinate the fate of functionally related mRNAs by using protein- or RNA-based trans factors. Some of these factors can regulate both transcription and decay rates, thereby maintaining proper mRNA homeostasis during eukaryotic cell life.


    Polyuridylation in Eukaryotes: A 3′-End Modification Regulating RNA Life

    In eukaryotes, mRNA polyadenylation is a well-known modification that is essential for many aspects of the protein-coding RNAs life cycle. However, modification of the 3′ terminal nucleotide within various RNA molecules is a general and conserved process that broadly modulates RNA function in all kingdoms of life. Numerous types of modifications have been characterized, which are generally specific for a given type of RNA such as the CCA addition found in tRNAs. In recent years, the addition of nontemplated uridine nucleotides or uridylation has been shown to occur in various types of RNA molecules and in various cellular compartments with significantly different outcomes. Indeed, uridylation is able to alter RNA half-life both in positive and in negative ways, highlighting the importance of the enzymes in charge of performing this modification. The present review aims at summarizing the current knowledge on the various processes leading to RNA 3′-end uridylation and on their potential impacts in various diseases.

    1. Introduction

    RNA 3′-end processing or modification plays an important role in determining their biological fate [1–3]. One major type of modification encountered by mRNAs is the addition of nontemplated nucleotides [3–7]. The main functional consequence of this nucleotide addition is to protect newly transcribed mRNAs from degradation. More generally, tail addition to RNAs regulates cellular RNA content by influencing RNA steady-state levels. Nuclear polyadenylation is essential to degrade various classes of noncoding RNAs (ncRNAs) in the nucleus [8–11]. However, once in the cytoplasm, RNAs carrying a 3′-poly(A) tail are protected from 3′ to 5′ exonucleases. Polyuridylation is another 3′ modification that involves the addition of uridines at the 3′-end of RNA molecules. This modification is found on various types of RNAs such as mRNAs, small RNAs, miRNAs, or guide RNAs (gRNAs) [7, 12–22]. This modification is known to have a major impact in multiple aspects of RNA turnover and metabolism, which are reviewed hereafter [7, 13–15, 20, 21].

    1.1. Polyadenylation

    Eukaryotic mRNAs start to be modified during their transcription, where capping and polyadenylation take place at their 5′- and 3′-ends, respectively, except for histone and some viral mRNAs [23]. Pre-mRNAs are first cleaved by the cleavage and polyadenylation machinery at the polyadenylation site located near the potential 3′-end. This cleavage is followed by the addition of the poly(A) tail by nuclear poly(A) polymerases (PAPs). This event will determine the 3′ untranslated region (UTR) of the RNA, which is crucial for the regulation of gene expression processes [24]. Mutations and changes in the length of this region will immediately affect a variety of processes such as mRNA stability, mRNA localization, and mRNA translation efficiency [25–29]. Once the mRNAs are exported to the cytoplasm, they may undergo several additional modifications such as methylation, editing, deadenylation, decapping, and polyuridylation, which again influence the stability or degradation of the RNA [7, 14, 17, 20–22, 30–35]. Polyadenylation regulates RNA degradation, which is one of the most important gene expression mechanisms not only for the removal of mRNAs that should not be translated anymore, but also for the disposal of the incorrectly transcribed mRNAs that have escaped the nuclear surveillance mechanisms. The general basis of RNA degradation is well conserved throughout eukaryotes, from yeast to mammals, and has two major directions: the 5′-3′ degradation by Xrn1 exoribonuclease and the 3′-5′ degradation catalyzed by the exosome complex (for recent review, see [36]). However, before degrading the mRNA bodies, cells must first identify the mRNAs to degrade. The cellular cues initiating mRNA degradation are still poorly understood for mRNAs encoded by the so-called “house-keeping” genes, while physiological inputs that trigger mRNA decay such as proinflammatory responses, heat shock, or differentiation are far better characterized [37, 38]. Deadenylation is generally the rate-limiting event in the cytoplasmic mRNA degradation and is catalyzed by the PAN2/PAN3 complex followed by the CCR4/NOT complex [31, 35]. Once the poly(A) tail has been removed, the Dcp1-Dcp2 decapping complex will withdraw the 7-methylguanylate cap from the 5′-end of the mRNA allowing the trimming of this RNA in a 5′ to 3′ manner by Xrn1 exonuclease [31–33, 39, 40]. Following deadenylation, the cytoplasmic exosome complex may cut down deadenylated RNAs as the 3′-5′ mRNA decay pathway [41, 42].

    1.2. Polyuridylation

    Recently, another player in the mRNA decay pathways has come into focus: the cytoplasmic poly(U) polymerases. These enzymes add uridine residues to the 3′-end of either coding RNAs or ncRNAs. Even though this modification has been known since the late fifties, its significance had been underestimated [43–45]. In the middle of the eighties, the importance of uridylation increased with the discovery and the characterization of the uridine insertion/deletion editing mechanisms in the mitochondria of kinetoplastids. This process was subsequently shown to be crucial for generating functional mRNA sequences as well as for increasing translation efficiency of local mRNAs [14, 30, 34]. Studies from the Aphasizhev laboratory on poly(U) polymerase family members present in trypanosomal species demonstrated additional roles for these enzymes, not only in the uridine insertion/deletion mechanism (generally known as the RNA editing process) but also during the processing of gRNA molecules and during mitochondrial mRNA translation [46, 47]. During the last decade, evidence showed that polyuridylation also existed in higher eukaryotes. The team of C. Norbury was the first to show that cells overexpressing a cytoplasmic poly(U) polymerase named Cid1 were less sensitive to hydroxyurea treatment, although the exact molecular mechanism was not fully understood [48]. Further studies demonstrated that polyuridylation was a critical step for the degradation of nonpolyadenylated mRNAs encoding histone proteins in mammals [20]. This new enzymatic step occurring at the 3′-end of nonpolyadenylated and polyadenylated mRNAs added another level of complexity to the known mRNA decay pathways [7, 20, 21, 49]. Finally, polyuridylation has also been found to occur in other types of RNA molecules such as miRNAs, siRNAs, and piRNAs with various functional consequences described hereafter [12, 16–19, 22].

    In this review, we focus on the latest research about the terminal polyuridylation by a specific group of noncanonical ribonucleotidyl transferases, a long time underestimated 3′-end posttranscriptional modification found in various RNAs and influencing RNA half-life and functions. The review will be divided in the following sections including a brief overview of the nucleotidyl transferase family followed by a review of the functional consequences of RNA polyuridylation in the different cell compartments. Finally, we will touch upon the multiple implications of polyuridylation mechanisms in diseases.

    2. The Noncanonical Ribonucleotidyl Transferase Family

    Enzymes performing terminal polyuridylation belong to the polymerase β- (Pol β-) like nucleotidyl transferase superfamily and more specifically to the group of template-independent polymerases that covalently add nucleotides to the 3′-end of RNA molecules. This protein family was precisely defined a few years ago [5]. Briefly, proteins from this family are named RNA-specific nucleotidyl transferases (rNTrs) and classified in three subgroups: (i) The canonical group, which corresponds to the nuclear poly(A) polymerases α, β, and γ. These are found in eukaryotes and share similar enzymatic and RNA-binding domains. (ii) The noncanonical rNTrs regroup a variety of proteins such as the Gld-2-, Trf4/5- and Cid1-type of poly(A) or poly(U) polymerases, the 2′-5′-oligo(A) synthetases, and the trypanosomal terminal uridylyl transferases. (iii) The third group is the one of the CCA-adding enzymes. In this review, we will only focus on the noncanonical rNTrs group as previously defined in [5].

    Every member from the noncanonical rNTrs group is characterized by an enzymatic domain made of two lobes named the catalytic and the central domains. The catalytic domain is made of four or five β-strands. The second β-strand contains a DxD or DxE motif (aspartate “D” or glutamate “E” residues separated by one hydrophobic residue “x”). A third aspartic residue is found in the third β-strand of the catalytic domain. The catalytic reaction is similar to the one described for the Pol β enzyme that includes a nucleophilic attack on the alpha phosphate of the bound nucleotide triphosphate by the 3′-OH group of the RNA substrate. The three aspartate residues interact with the incoming RNA and two metal ions necessary to stabilize the reaction intermediate as described previously [50]. The central domain contains the nucleotide recognition motif (NRM), which corresponds to a 10–15 amino acid long loop forming one end of the nucleotide triphosphate binding pocket. The residues located in the NRM stabilize the base of the substrate nucleotide triphosphate via water-mediated and/or direct hydrogen bonds with their side chain atoms [51–53]. Subclassification of the rNTrs was attempted based on the local amino acid sequence conservation of the NRM. However, in light of recent crystal structures from members of the rNTrs in complex with their natural substrate NTP, it appears that using the NRM sequence identity may not be sufficient to precisely predict the type of nucleotide accepted in the active site. In fact, it is not fully clear whether these proteins are not able to add various types of nucleotide in vivo as recent sequencing studies specifically designed to identify 3′-addition of nontemplated nucleotides highlighted the diversity of the cellular RNA tails [54].

    Furthermore, a RNA recognition motif (RRM) is also found in all canonical and a few noncanonical rNTrs. Its likely role is to bind RNA substrates in a non-sequence-specific manner [55, 56]. The RRM domains are differentially located in the sequence, that is, near the C-terminus for the canonical PAPs, at the N-terminus, or in the central domain in some noncanonical rNTrs. The RBD is absent in numerous noncanonical rNTrs enzymes, indicating that either these proteins can act on any RNA or that their activity is restricted via a protein partner that targets them to specific RNAs or both. In at least one case, the enzyme ZCCHC11 is targeted to one specific pre-miRNA species through interactions with the Lin28 proteins [57–59].

    From a phylogenetic point of view, several models have been proposed to explain the evolution of the Pol β-NTrs family. The hypothesis of Aravind and Koonin [60] is that the Pol β-NTrs family members have rapidly and independently diverged from a common ancestor presenting a very general and nonspecific nucleotidyl transferase activity. The different family members would have acquired distinct functional domains to occupy vacant evolutionary niches. Then, horizontal gene transfer and lineage-specific gene loss could have explained the actual distribution of the different groups in the three domains of life. Some evidences like the discovery of the archaeal and bacterial minimal nucleotidyl transferases (MNT family) and the restricted phylogenetic distribution of most of the Pol β-NTrs family members support this model [60]. However, it has recently been shown that a bacterial poly(A) polymerase that possesses the RBD of a CCA-adding enzyme is able to act as a CCA-rNTrs [61]. This suggests that the CCA-adding enzymes could be the ancestors of the poly(A) polymerases and possibly the founders of all the remaining rNTrs, which would have adopted different RNA binding domains mediating different target specificity.

    The noncanonical rNTrs is divided into two main groups based on their specific activities: the Cid1-like family and the RNA editing enzymes.

    (i) The Caffeine-induced death suppressor protein 1 (or Cid1) from Schizosaccharomyces pombe is the pioneer of cytoplasmic poly(U) polymerases [62]. Many other proteins are part of this group with highly similar enzymatic properties but limited sequence homology such as the trypanosomal protein RNA editing TUTase1 (RET1). Despite its name, RET1 modifies specifically the 3′-end of both the gRNAs and the mRNAs in kinetoplast without any involvement in the RNA editing process itself [13, 15, 46, 47, 63]. Seven proteins from this group are found in human. Evidences start to accumulate for some of these human proteins but, globally, their precise action still requires a more detailed characterization [58, 62, 64].

    (ii) The RNA editing enzymes, on the other hand, are responsible for mitochondrial mRNA editing by U-insertion/deletion [65–70]. Mainly, two proteins have been studied extensively: RNA editing TUTase 2 (RET2) and the mitochondrial editosome-like complex associated TUTase 1 (MEAT1). RET2 and MEAT1 are found with the 20S editosome complex of trypanosomes and are crucial for the U insertion-type of editing in this organism [70, 71]. Crystal structures of RET2 and MEAT1 showed a conserved domain organization except for the middle domain [51, 52]. The lack of sequence similarity within this middle domain suggests divergent functions.

    3. Polyuridylation according to Cell Compartments

    Until a few years ago, polyuridylation had been only reported in the mitochondria of the parasitic protist trypanosome [14, 30]. More recently, noncanonical rNTrs were found in the cytoplasm of various eukaryotic species and were shown to modify a wide range of nontranslated and translated RNAs [7, 17, 20, 21, 72–75]. Details of the different substrates and the responsible enzymes in the cell nucleus, cytoplasm, and mitochondria are described hereafter and summarized in Figure 1.

    3.1. In the Nucleus

    Until now, the only substrate of uridylation reported in the nucleus is the U6 snRNA (Figure 1). This RNA is uridylated by the U6 TUTase, which is an essential enzyme for cell survival in mammals [64]. SiRNA-mediated silencing of the U6 TUTase leads to U6 snRNA decay, confirming the necessity of uridylation for U6 snRNA stability [64]. U6 TUTase is responsible for the addition or restoration of at least four uridine residues at the 3′-end of U6 snRNA since 3′-end of U6 snRNA is constantly subjected to exonucleases activity [64, 76]. These four U residues form an intramolecular double strand with a stretch of adenines within the U6 snRNA molecule, which is important for mRNA splicing [64]. This uridylation event specifically in the nucleus allows the proper production of a splicing-competent U6 snRNP (Figure 2(a)). Mammalian U6 snRNA uridylation in vivo has been reported with up to 20 nucleotides added at the 3′-end of the RNA molecule [77, 78]. It is important to note that U6 snRNA is also subjected to adenylation and this event inhibits its uridylation (Figure 2(a)) [79]. Moreover, the 3′-end of U6 snRNA is recognized specifically by the Lsm2-8 complex, a doughnut-like heteroheptameric complex related to the Sm complex found on the snRNPs.

    3.2. In the Organelles (Mitochondria)

    Uridylation events in the organelles have been reported in mitochondria [13, 63]. So far, no polyuridylation events have been found in the chloroplast of plants and algal cells. It is apparently absent, although proteins from the rNTrs family are present such as the poly(A) polymerase [80]. One possible reason is the close evolutionary conservation of the RNA processing pathways found in the chloroplast and in bacteria where poly(A) tail present at the 3′-end of mRNAs is the major regulatory modification [81–84].

    Poly(U) tails have been reported mostly in kinetoplastid-containing organisms. Two main substrates are targeted in these organisms: gRNAs and locally transcribed mRNAs (Figure 1). The gRNAs are specific to kinetoplastid species and are crucial for cell survival as they are in charge of guiding the RNA editing machinery to its mRNA targets [14]. RET1, the first characterized ncNTrs, acts on both types of RNAs with strikingly different functional consequences.

    For the gRNAs, uridylation represents their final maturation step [13, 46]. In order to be matured, pre-gRNAs need to pass through an exonucleolytic process followed by stabilization by the gRNA binding complex (GRBC) and RET1 uridylation (Figure 2(b)) [13, 15, 85]. Mature gRNA is thus composed of a 5′ phosphate from the transcription followed by an anchor region complementary to a target unedited mRNA, a guiding region that directs the editing of its mRNA target and a final poly(U) tract at the 3′-end. In RET1-depleted cells, gRNAs are stable but not able to perform their editing function suggesting a crucial role of the oligo(U) tail in the editing event in the mitochondria. This oligo(U) tract may stabilize the gRNA-mRNA hybrid through binding with the purine-rich preedited region [15]. The uridylated gRNA bound to its mRNA target recruits the 20S editosome. This gRNA-mediated mRNA editing in kinetoplastid trypanosomes is crucial for the parasite survival, as these editing events are needed for the proper establishment of the coding sequence of the mitochondrial mRNAs [15]. Currently, it is not yet fully understood how RET1 enzyme recognizes its gRNA substrate nor how the pre-gRNA processing step takes place [13, 46].

    After editing, mRNAs need to be further modified at the 3′-end in order to be translationally competent in trypanosomal mitochondria. This modification is the addition of a long 3′ A/U tail (Figure 2(b)) [47]. This nucleotide addition is due to RET1 which works in concert with the kinetoplast poly(A) polymerase 1 (KPAP1). The RET1/KPAP1 complex adds approximately 200 alternated adenines and uridines to the 3′-end of the targeted mRNAs [47]. Therefore, polyuridylation and polyadenylation are necessary to trigger the translation of edited as well as never edited mRNAs (Figure 2(b)). RET1 and KPAP1 actions are coordinated by the kinetoplast polyadenylation/uridylation factors 1 and 2 (KPAF1 and KPAF2) complex [47]. Currently, our molecular understanding of the sequence of events taking place at the 3′-end of mitochondrial mRNAs is poor and awaits further structural and biochemical characterization [47, 63].

    It is noteworthy that RNAs with poly(U) tails have also been observed in human mitochondria under certain conditions [86–89]. In spite of this, how this process is achieved in this compartment and its implication(s) for human mitochondrial RNA metabolism still remain to be characterized.

    3.3. Polyuridylation in the Cytoplasm

    Cytoplasmic polyuridylation occurs on a variety of RNA molecules ranging from polyadenylated to nonpolyadenylated RNA molecules including mRNAs, small RNAs, miRNAs, or piRNAs (Figure 1) [7, 17, 20, 21, 73–75]. The various functional outcomes of polyuridylation in this compartment offer new insights into RNA turnover and small RNA biogenesis (Figure 3).

    Several eukaryotic mRNAs were shown to be uridylated in the cytoplasm of S. pombe by the poly(U) polymerase Cid1 (Figure 1) [7, 62]. RNA cRACE studies in fission yeast revealed a role of uridylation in a new deadenylation-independent decapping-mediated degradation pathway (Figure 3(a)) [7]. Until now, only a handful of mRNAs has been identified to be specifically uridylated such as act1, urg1, and adh1 [7]. Recent studies looked at the 3′-end sequence of mRNAs at a genome-wide level and revealed that U tails are apparently attached to short poly(A) tracks rather than to the mRNA body [49, 54]. Interestingly, while some mRNAs like the one encoding the poly(A) binding protein 4 are polyuridylated in more than 25% of the cases, about 80% of mRNAs have an uridylation frequency comprised between 2 and 5%. Overall, the functional relevance of those low-level of uridylation is currently unknown. Factors such as ZCCHC6 or ZCCHC11 (also known as TUT7 and TUT4 resp.) have been shown to be responsible for the human cytoplasmic mRNA uridylation activity and the consequence is apparently to induce mRNA degradation [49]. Furthermore, a single uridine at the 3′-end of a RNA molecule is sufficient to be recognized by the Lsm1-7 complex, known to link 3′-end deadenylation and 5′-end decapping, clearly supporting the relationship between uridylation and mRNA degradation [90].

    Nonpolyadenylated mRNAs are also uridylated in the cytoplasm (Figure 1). This is the case of the histone-encoding mRNAs [20]. Upon inhibition of DNA replication or conclusion of S-phase, histone proteins are not necessary anymore and, so, histone mRNAs must be rapidly degraded in order to avoid their accumulation and their interference with other cellular pathways [91]. Histone mRNAs are not polyadenylated but possess a stem loop structure at their 3′-end crucial for pre-mRNA processing, export, and proper translation [92–94]. Studies aiming to understand the mechanism by which histone mRNA degradation was triggered found that histone mRNAs were targeted to decay by uridylation (Figure 3(b)) [20, 21]. The nature of the responsible enzyme(s) is still the subject of conflicting results as different groups found different enzymes [20, 21]. These studies systematically found Cid1 orthologous enzymes such as TUTase1 (PAPD1), TUTase3 (Trf4-2), and ZCCHC11 (TUT4) to be responsible for the uridylation. It is not fully clear however, how PAPD1 enzyme would either switch between cell compartments as PAPD1 is reported as a mitochondrial protein or how they could select which substrates to uridylate and which one to adenylate in vivo as both PAPD1 and Trf4-2 proteins do have reported poly(A) polymerizing activities [20]. One could not exclude that several pools of PAPD1 differentially located in the cell may exist. More data are definitely required to fully apprehend rNTrs role during regulated histone mRNA degradation in particular regarding the factors bringing together the histone mRNAs and the rNTrs. Interestingly, as for polyadenylated mRNAs in fission yeast, uridylation of histone mRNAs was shown to promote decapping followed by 5′-3′ degradation [20]. The Lsm1-7 protein complex was shown to be responsible for the promotion of the decapping activity. More recently, 3′-5′ degradation of histone mRNAs by the exonuclease ERI1 has been reported (Figure 3(b)) [95]. Again, the Lsm1-7 complex was involved in the recruitment of the exonuclease ERI1 to the terminal stem loop. The Lsm1-7 complex apparently binds both the uridylated histone mRNAs and the exonuclease ERI1 [95].

    A variety of ncRNAs from diverse organisms have recently been shown to carry mono- or multiple non-templated uridine residues at their 3′-end (Figure 1) [19, 73, 96–98]. The major functional consequence associated with uridylation is to trigger RNA degradation but is not limited to it. 3′ uridylation of various miRNAs has been observed in multiple sequencing studies suggesting a wide role of uridylation during miRNA biogenesis [99–101]. Mono- or polyuridylation events have been found in both pre-miRNAs and mature miRNAs [73, 96, 98, 102]. In C. elegans and H. sapiens, polyuridylation of pre-let-7-miRNA has been reported and is performed by the proteins PUP-2 and ZCCHC11, respectively [98, 103, 104]. Association between the pre-miRNA and the Lin28 protein induces a conformational change in the pre-miRNA loop, which possibly favors modification by ZCCHC11 [105, 106]. However, the presence of a single 3′-overhanging nucleotide appears critical for the uridylation process therefore excluding the so-called “group I” or canonical miRNAs from being subject to uridylation [96]. Furthermore, in the same study, ZCCHC6 enzyme was found to be responsible for the monouridylation of group II let-7 pre-miRNAs and this modification is independent of the Lin28 protein but is critical for the production of this particular miRNA [96]. So, uridylation of pre-miRNAs can influence the miRNA production both positively and negatively (Figure 3(c)) [21]. Finally, ZCCHC11 has also been involved in the uridylation of specific mature miRNA such as miR-26 [107]. Further biochemical and biophysical studies are needed in order to identify the specific enzymes responsible for the uridylation of other miRNAs in higher organisms as well as the target-specific effects induced by this 3′-end modification. Interestingly, mammalian Dis3L2 exonuclease was also shown to specifically degrade uridylated pre-let-7-microRNA discriminating them from 3′-unmodified RNAs [108]. Recently, Dis3L2 protein was shown to preferentially degrade mRNAs with 3′-end uridylation and its deletion together with the one of Lsm1 led to the accumulation of uridylated mRNAs in fission yeast (Figures 3(a) and 3(c)) [109]. Further studies between the Dis3L2 exonuclease and TUTases will be necessary to better understand their respective functions and the link existing between these enzymatic activities.

    Other types of ncRNAs subject to 3′ uridylation are siRNA and piRNAs (Figure 1). In nematodes and in plants, these particular types of RNA substrate are modified by the protein CDE-1 (cosuppression defective 1) and HESO-1 (Hen1 suppressor1) respectively [110]. In the green algae Chlamydomonas, MUT68 has been implicated in this event [73]. Studies in plant and animal species have demonstrated an antagonistic role of uridylation and 2′-O-methylation in these organisms [12, 17–19, 22]: Hen1 (HUA ENHANCER 1) and its homologs methylate sRNAs in plants, piRNAs in vertebrates, and Ago2-associated siRNAs in flies, protecting these RNAs against 3′ uridylation (Figure 3(d)) [12, 17–19, 22]. In C. elegans, CSR-1 is an Ago protein necessary for proper chromosome segregation rather than regulation of mRNA levels [74, 111, 112]. CDE-1, a C. elegans PUP, uridylates unmethylated siRNAs of the CSR-1 pathway [74]. Mutation of this CDE-1 enzyme leads to accumulation of CSR-1 siRNAs, which promotes erroneous chromosome segregation and defective gene silencing [74]. Uridylation is then a destabilizing factor against CSR-1 siRNAs, which regulates CSR-1-dependent and specific siRNA levels in this organism. In Chlamydomonas reinhardtii, MUT68 was first known to adenylate 5′ cleavage fragments of mRNAs targeted by the RNA-induced silencing complexes (RISC), thereby promoting their decay [97]. Further studies showed an important role of MUT68 in miRNA and siRNA degradation through its 3′ uridylation activity [73]. 3′ uridylation of piRNAs have been observed in zebrafish and drosophila, but the enzymes responsible for this modification are currently unknown [12, 17]. In zebrafish Hen1 mutants, piRNAs derived from retrotransposons are found uridylated and their levels are decreased suggesting a sensitivity of these uridylated piRNAs to degradation. Interestingly, a mild repression of retransposons is observed in these mutants thus highlighting a destabilizing role for uridylation of piRNAs and a stabilizing role for methylation [17]. Taken together, these data highlight the crucial roles of small ncRNA uridylation within diverse biological processes and in several organisms. Defects in the regulation of this phenomenon can have important consequences on gene expression (Figures 2 and 3(d)).

    The HESO-1 enzyme, like MUT68, is also shown to act on atypical substrates, that is, the product of the miRNA-directed mRNA cleavage [113]. In this case, the uridine nucleotides are apparently added to the 5′-fragment of the cleaved mRNA when it is still bound by the Ago1 complex [113]. Further studies will help determining the generality of this mechanism as HESO-1 does not seem to be conserved in higher eukaryotes.

    At last, polyuridylation has also been reported to stabilize RNAs, rather than destabilize them. In Arabidopsis thaliana, uridylation of oligoadenylated mRNAs has been suggested to prevent their 3′ trimming and rather establish a preferential 5′-to-3′ mRNA degradation manner [114]. Indeed, URT1 (UTP:RNA uridylyl transferase 1) was shown to uridylate oligo(A)-tailed mRNAs in vivo and its absence contributed to the degradation of oligoadenylated mRNAs highlighting a new role of uridylation in mRNA stabilization. The ZCCHC11 enzyme, besides its role in histone mRNA and pre-miRNA decay, has also been implicated in indirect mRNA stabilization by uridylation of mature miRNAs (Figure 3(c)). ZCCHC11-dependent uridylation of mature cytokine-targeting miRNAs is known to lead to the stabilization of cytokine transcripts and hence regulates cytokines gene expression. Mature miR-26 can bind interleukin IL-6 mRNA in its 3′ UTR and targets this cytokine-encoding mRNA to degradation [107]. Upon miR-26 uridylation by ZCCHC11, the miRNA is unable to bind the 3′ UTR of the mRNA and thus the transcript is stabilized with no associated degradation of miR-26. This is further confirmed by ZCCHC11 knockdown experiments where several cytokine mRNAs are downregulated in the absence of uridylation [107, 115].

    These data together support a crucial role of cytoplasmic polyuridylation in the regulation of gene expression and stability control of both coding and noncoding RNAs in diverse eukaryotic species.

    4. Polyuridylation and Diseases

    RNA uridylation in the cytoplasm has been shown to induce tumorogenesis in mammals. Uridylation at the 3′-end of the tumor suppressor pre-let-7 microRNA by cytoplasmic ZCCHC11 and ZCCHC6 enzymes blocks let-7 miRNA maturation, which in turn stimulates tumor growth [58]. Lin28 is a factor of pluripotency in stem cells and once it is expressed, it helps the maintenance of an undifferentiated and proliferative state by blocking the expression let-7 miRNA by recruiting ZCCHC11 for uridylation-mediated decay [57–59]. In adult somatic cells, Lin28-let-7 pathway is normally silenced even though we still observe expression of LIN28A or LIN28B in a wide variety of human cancers [116, 117]. Inhibition of this oncogenic pathway blocks the tumorigenicity of cancer cells [116]. It has recently been shown that modified let-7 microRNAs are degraded by Dis3L2 exonuclease [118]. Furthermore, Dis3L2, which preferentially trims uridylated cytoplasmic RNAs, has been found mutated in patients with Perlman syndrome and in some cases this mutation lead to the development of Wilm’s tumor at early stages of child’s growth [118]. Even though RNA uridylation has been linked to tumor growth, the biological significance of such event is still poorly understood and as such is being studied. In order to better understand tumorigenesis, it is necessary to identify the RNA targets as well as the protein partners that recruit either the RNA substrates or the poly(U) polymerases. Such information will allow the in-depth studies of the link between PUPs and diseases. Furthermore, structural and biochemical studies of substrate recognition by rNTrs will provide a rational foundation for therapeutic purposes. In kinetoplastid organisms, this information will bring new insights into U-insertion/deletion, gRNA biogenesis, and translational control required for parasite survival. Thus, it may provide a new avenue for the design of new trypanocides, important to treat various trypanosomal diseases including the fatal human sleeping sickness.

    5. Conclusions and Perspectives on Polyuridylation

    Polyuridylation was for a long time an underestimated 3′-end modification most probably because sequencing techniques were focused on polyadenylated RNAs. With the development of new and adapted techniques to detect 3′ uridylation, this event is starting to gain strength with impacting roles in RNA degradation and stability [99, 119]. RNA sequencing analysis of mammalian cells, not depending on oligodT primers but rather using 3′ ligated linkers specific for small RNAs of 200 nt or less, showed a widespread tendency of 3′-end uridylation of small RNAs [99]. Interestingly, besides the already known uridylated targets, they also found this 3′ modification on transcriptional start-site-associated RNAs along with spliced introns. This suggests a larger role of polyuridylation in RNA metabolism in mammals, despite the fact that PUPs are mostly localized in the cytoplasm. Optimized RNA sequencing methods in different backgrounds, such as DNA replication inhibition and stress conditions, and refinements in these methods, are necessary to understand the global biological consequences of uridylation in RNA metabolism. With RNA-Seq development, more and more RNAs are found to be uridylated in various organisms, but the enzymes responsible for this process are still unknown. The identification of polyuridylating enzymes becomes now critical for obtaining a larger picture of uridine tail addition in eukaryotes, its evolution, and its functional implication in the cell. Finally, 3′ uridylation is involved in several key aspects of RNA biology and all the proteins implicated in this process in eukaryotes are not yet known. It thus brings into focus the importance of multiplying studies concerning this particular process and the relevant players. Several research groups nowadays started to focus their work on identifying new rNTrs along with their targets and possible protein partners. We will most likely hear a lot more about rNTrs and their influence on RNA metabolism and turnover during the coming years.

    Conflict of Interests

    The authors declare that there is no conflict of interests regarding the publication of this paper.

    Authors’ Contribution

    Paola Munoz-Tello and Lional Rajappa contributed equally to the review.

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    Copyright © 2015 Paola Munoz-Tello et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


    Why do mutations not take place in mRNA of higher eukaryotes? - Biology

    AP Biology Learning Objectives Name:

    Understand the concept of heredity by being able to explain in general terms how traits are inherited from parents to offspring.

    Understand that mitosis results in identical daughter cells and that mitosis is the primary form of reproduction in prokaryotes.

    Mitosis alternates with interphase in the cell cycle and that the cell cycle is regulated by a molecular control system.

    Distinguish between asexual and sexual reproduction by describing the different modes of creating the next generation and what types of organisms undergo each type (or both).

    Distinguish between the following pairs of terms: somatic cell and gamete, autosome and sex chromosome.

    Follow the chromosome count (n) in cells of the human body by describing how haploid and diploid cells differ from each other and which cells in the human body are diploid and which are haploid.

    Understand the concept of alternation of generations by explaining why it is crucial that fertilization and meiosis must alternate in all sexual life cycles.

    Describe the process of synapsis during prophase I and explain how genetic recombination occurs by drawing representations of tetrads crossing over and the resulting recombinant gametes.

    Appreciate key differences between mitosis and meiosis by being able to describe three events that occur during meiosis I but not during mitosis.

    Understand how random alignment (also known as independent assortment), crossing over, and random fertilization contribute to genetic variation in sexually reproducing organisms by discussing when these events take place and how they increase genetic variation in populations.

    Define the following terms: true-breeding, hybridization, monohybrid cross, P generation, F 1 generation, and F 2 generation.

    Use a Punnett square to predict the results of a monohybrid cross, stating the phenotypic and genotypic ratios of the F2 generation.

    Distinguish between the following pairs of terms: dominant and recessive heterozygous and homozygous genotype and phenotype.

    Explain how a testcross can be used to determine if an individual with the dominant phenotype is homozygous or heterozygous.

    Use a Punnett square to predict the results of a dihybrid cross and state the phenotypic and genotypic ratios of the F2 generation.

    Connect your understanding of Mendel’s law of independent assortment to the behavior of chromosomes during meiosis.

    Use Multiplication and Addition Rules to calculate the probability that a particular F 2 individual will be homozygous recessive, homozygous dominant, or heterozygous.

    Use the laws of probability to predict, from a trihybrid cross between two individuals that are heterozygous for all three traits, what expected proportion of the offspring would be:

    a. homozygous dominant for the three traits

    b. heterozygous for all three traits

    Explain why it is important that Mendel used large sample sizes in his studies.

    Explain how phenotypic expression of the heterozygote differs with complete dominance, incomplete dominance, and codominance.

    Explain why dominant alleles are not necessarily more common in a population. Illustrate your explanation with an example.

    Describe the inheritance of the ABO blood system and explain why the I A and I B alleles are said to be codominant.

    Explain why most polygenic characters are along a spectrum or continuum of phenotypes.

    Describe how environmental conditions can influence the phenotypic expression of a character.

    Explain why studies of human inheritance are not as easily conducted as Mendel’s work with his peas.

    Given a simple family pedigree, deduce the genotypes for some of the family members.

    Explain how a lethal recessive allele can be maintained in a population.

    Describe the inheritance and expression of autosomal recessive diseases such as cystic fibrosis, Tay-Sachs disease, and sickle-cell disease.

    Explain why lethal dominant genes are much rarer than lethal recessive genes.

    Give an example of a late-acting lethal dominant gene in humans and explain how it can escape elimination by natural selection.

    Explain why Drosophila melanogaster is a good experimental organism for genetic studies.

    Explain why linked genes do not assort independently.

    Distinguish between parental and recombinant phenotypes.

    Explain how crossing over can unlink genes.

    Describe how sex is genetically determined in humans and explain the significance of the SRY gene.

    Distinguish between linked genes and sex-linked (X-linked) genes.

    Explain why sex-linked recessive traits are always expressed in males, but not all females.

    Describe the inheritance patterns and symptoms of color blindness, cystic fibrosis, Huntington's and hemophilia.

    Explain how nondisjunction can lead to aneuploidy.

    Define trisomy, triploidy, and polyploidy. Explain how these major chromosomal changes occur and describe possible consequences.

    Distinguish among chromosomal mutation categories: deletions, duplications, inversions, and translocations.

    Describe the type of chromosomal alterations responsible for the following human disorders: Down syndrome, Klinefelter's syndrome, Turner syndrome, and chronic myelogenous leukemia (CML).

    Explain why mitochondrial-determined traits are maternally inherited in animals.

    Explain why researchers originally thought protein was the genetic material.

    Summarize the experiments performed by the following scientists that provided evidence that DNA is the genetic material:

    a. Contributions of Watson, Crick, Wilkins, and Franklin on the structure of DNA

    b. Avery-MacLeod-McCarty experiments

    c. Hershey-Chase experiment

    Describe the structure of DNA. Explain the base-pairing rule and describe its significance.

    Understand the DNA synthesis is a semiconservative process: that is, one strand serves as the template for a new, complementary strand.

    Describe the process of DNA replication, including the role of the origins of replication and replication forks.

    Explain the role of DNA polymerase in replication.

    Define antiparallel and explain why continuous synthesis of both DNA strands is not possible.

    Explain the bidirectionality of DNA synthesis and how the production of the leading strand and the lagging strand differs.

    Explain the roles of key enzymes in DNA replication: DNA polymerase, ligase, RNA polymerase and helicase.

    Explain how RNA differs from DNA.

    Explain how information flows from gene to protein.

    Distinguish between transcription and translation.

    Compare where transcription and translation occur in prokaryotes and in eukaryotes.

    Define codon and explain the relationship between the linear sequence of codons on mRNA and the linear sequence of amino acids in a polypeptide.

    Explain the significance of the reading frame during translation.

    Explain the evolutionary significance of a nearly universal genetic code.

    Explain how RNA is modified after transcription in eukaryotic cells.

    Describe the functional and evolutionary significance of introns.

    Describe the structure and functions of tRNA.

    Explain the significance of wobble.

    Describe the molecular make-up (structure) and function of the ribosome.

    Understand that the processes of transcription and translation are endergonic events.

    In eukaryotic cells, the mRNA transcript undergoes a series of enzyme-regulated modifications, for example, addition of a poly-A tail, addition of a GTP cap and the excision of introns.

    Translation of the mRNA occurs in the cytoplasm on the ribosome.

    Describe two properties of RNA that allow it to perform so many different functions.

    Compare protein synthesis in prokaryotes and in eukaryotes.

    Define point mutations. Distinguish between base-pair substitutions and base-pair insertions. Give examples of each and note the significance of such changes.

    Describe several examples of mutagens and explain how they cause mutations.

    REGULATION OF GENE EXPRESSION

    Regulation of Prokaryotic Gene Expression

    • Briefly describe two main strategies that cells use to control metabolism.

    • Explain the adaptive advantage of bacterial genes grouped into an operon.

    • Using the trp operon as an example, explain the concept of an operon and the function of the operator, repressor, and corepressor.

    • Explain how repressible and inducible operons differ and how those differences reflect differences in the pathways they control.

    • Describe how the lac operon functions and explain the role of the inducer, allolactose.

    • Distinguish between positive and negative control. Give examples of each from the lac operon.

    Regulation of Eukaryotic Gene Expression

    • Define differential gene expression. At what level is gene expression generally controlled?

    • Distinguish between heterochromatin and euchromatin.

    • Explain how DNA methylation and histone acetylation affects chromatin structure and the regulation of transcription.

    • Define epigenetic inheritance.

    • Describe the role of the transcription initiation complex.

    • Define control elements and explain how they influence transcription.

    • Distinguish between general and specific transcription factors.

    • Explain the role of promoters, enhancers, activators, and repressors in transcriptional control.

    • Describe the process and significance of alternative RNA splicing.

    • Describe the processing of pre-mRNA in eukaryotes.

    • Describe factors that influence the lifespan of mRNA in the cytoplasm. Compare the longevity of mRNA in prokaryotes and eukaryotes.

    • Explain how gene expression may be controlled at the translational and post-translational level.

    Viral replication results in genetic variation, and viral infection can introduce genetic variation into the hosts.

    Viruses have highly efficient replicative capabilities that allow for rapid evolution and acquisition of new phenotypes.

    Viruses replicate via a component assembly model allowing one virus to produce many progeny simultaneously via the lytic cycle.

    RNA viruses (retroviruses) lack replication error-checking mechanisms, and thus have higher rates of mutation.

    The retrovirus HIV is a well-studied system where the rapid evolution of a virus within the host contributes to the pathogenicity of viral infection.

    Some viruses are able to integrate into the host DNA and establish a latent (lysogenic) infection. These latent viral genomes can result in new properties for the host such as increased pathogenicity in bacteria.

    Practical Applications of DNA Technology

    • Explain how gel electrophoresis is used to analyze nucleic acids and to distinguish between two alleles of a gene.

    • Be able to retell the steps of cloning a gene:

    • Describe the natural function of restriction enzymes and explain how they are used in recombinant DNA technology.

    • Explain how the creation of sticky ends by restriction enzymes is useful in producing a recombinant DNA molecule.

    • Outline the procedures for cloning a eukaryotic gene in a bacterial plasmid.

    • Describe two techniques to introduce recombinant DNA into eukaryotic cells.

    • Describe the polymerase chain reaction (PCR) and explain the advantages and limitations of this procedure.

    • Define a single nucleotide polymorphism. Explain how an SNP may produce a restriction fragment length polymorphism (RFLP).

    • Describe an example of a transgenic animal used as a pharmaceutical factory.

    • Explain how DNA technology can be used to improve the nutritional value of crops and to develop plants that can produce pharmaceutical products.


    What Is Prokaryotic Transcription?

    Prokaryotic organisms do not have an organized nucleus. The genetic materials are suspended in the cell cytoplasm. All the precursors needed for transcription are located in the cytoplasm. Prokaryotic transcription usually happens in the cytoplasm. The process needs an RNA polymerase enzyme for transcription to be successful.

    The RNA polymerase enzyme contains five subunits. The enzyme usually binds to the stigma and promoter to initiate transcription in the holoenzyme.

    Keep in mind that the prokaryotes’ DNA is not bound to histone. The transcription process is not initiated at all since it happens directly. It is ideal for those prokaryotes with overlapping genes.

    Prokaryotic transcription starts at the promoter region, elongates via the coding region, and ends when the RNA polymerase enzyme reads the termination signal.

    Prokaryotes have two types of termination signals. Rho dependent and independent termination signal. Transcribe mRNA will be translated during the transcription process.


    • Protein synthesis is the process in which cells make proteins. It occurs in two stages: transcription and translation.
    • Transcription is the transfer of genetic instructions in DNA to mRNA in the nucleus. It includes three steps: initiation, elongation, and termination. After the mRNA is processed, it carries the instructions to a ribosome in the cytoplasm.
    • Translation occurs at the ribosome, which consists of rRNA and proteins. In translation, the instructions in mRNA are read, and tRNA brings the correct sequence of amino acids to the ribosome. Then, rRNA helps bonds form between the amino acids, producing a polypeptide chain.
    • After a polypeptide chain is synthesized, it may undergo additional processing to form the finished protein.
    1. Relate protein synthesis and its two major phases to the central dogma of molecular biology.
    2. Explain how mRNA is processed before it leaves the nucleus.
    3. What additional processes might a polypeptide chain undergo after it is synthesized?
    4. Where does transcription take place in eukaryotes?
    5. Where does translation take place?

    Relating expression differences to underlying regulatory networks

    The recent explosion of genomic resources and high throughput techniques is accelerating the elucidation of regulatory networks in model systems. Comparative studies of gene expression are an important tool for interpreting the function of network components and for understanding their stability and susceptibility to change. To understand how changes in regulatory networks contribute to phenotypic diversity within and between species, the following issues should be addressed.

    (1) Many changes in gene expression are often induced by an environmental change. Some of the changes may be directly involved in the physiological adjustment while others may be secondary consequences of the regulatory network. Comprehensive descriptions of regulatory connections will help disentangle these types of changes by revealing connections between functional modules. However, genetic mapping and functional tests will ultimately be needed to identify the subset of genes for which expression changes impact the phenotype.

    (2) Regulatory changes may be most stable when located in particular parts of a pathway. To test this hypothesis, complete pathways controlling divergent traits should be surveyed to locate all independent regulatory changes within the network. Locating regulatory variants will also make it possible to determine whether the connectivity of a gene within the network influences its propensity for change.

    (3) The distribution of new regulatory mutations within a network appears to differ from the distribution of regulatory variants in the wild(Denver et al., 2005). Network architecture is expected to influence how regulatory variation arises, while the pleiotropic side effects of individual regulatory mutations are expected to influence which changes survive the test of time. To fully appreciate the impact of network architecture on evolutionary trajectories, properties that promote particular changes within regulatory networks must be identified.

    (4) Some functional classes of genes may be more susceptible than others to regulatory mutations affecting their expression. Analyzing the distribution of regulatory variants among genes with different gene ontology designations will test this hypothesis. Such an analysis may also identify specific biological functions with a propensity for regulatory changes (e.g. sperm expressed genes in C. elegans) (Denver et al.,2005). However, any analyses using current gene ontology designations should be interpreted cautiously. At present, for most genes,gene ontology assignments of functional classes and biological processes are predicted solely based on sequence similarity and are awaiting genetic and/or biochemical verification.

    As discussed in this review, existing case studies provide some insight into these issues. However, we have a long way to go toward understanding how regulatory variation is distributed within genomic regulatory networks and how network structure influences patterns of variable gene expression. A combination of genetic and biochemical dissection of regulatory networks in model systems, computational analyses of network properties, and comparative studies of gene expression among non-model species will be needed to resolve these issues. Given the recent growth in these research areas, a comprehensive understanding of regulatory variation in the context of regulatory networks may soon be achieved.


    Watch the video: Protein Synthesis Updated (December 2022).