Sequences of nitrogen bases found in rna that are involved in the synthesis of proteins.

RNA and Protein Synthesis

All eukaryotic cells use DNA to direct protein synthesis. Proteins are made in the cytoplasm on the ribosome. These polypeptide-making factories contain more than 50 different proteins, as well as RNA. RNA is similar to DNA, and its presence in ribosomes suggests its important role in protein synthesis (Fig. 10.2). RNA differs from DNA in two ways: RNA contains ribose as sugar rather than the deoxyribose in DNA, and RNA contains the pyrimidine uracil (U in codon designations) instead of thymine.2 In addition, RNA does not have a regular helical structure. The class of RNA present in ribosomes is called ribosomal RNA (rRNA).3 rRNA and ribosomal proteins provide sites at which polypeptides are assembled. Transfer RNA (tRNA) transports the amino acids to the ribosome for the synthesis of polypeptide.4,5 There are more than 40 different tRNA molecules in human cells. tRNA is smaller than rRNA and is present in free form in the cytoplasm. Messenger RNA (mRNA) consists of long strands of RNA molecules that are copied from DNA. mRNA travels to the ribosome to direct the assembly of polypeptides.

RNA is synthesized on a DNA template by a process of DNA transcription in which RNA polymerase enzymes make an RNA copy of a DNA sequence. RNA polymerases are formed from multiple polypeptide chains with a molecular weight of 500,000.6,7 In eukaryotic cells there are three different types of RNA polymerases. RNA polymerase II transcribes the gene whose RNAs will be translated into proteins. RNA polymerase I makes the large rRNA precursor (45S rRNA) containing the major rRNAs. RNA polymerase III makes very small, stable RNAs, including tRNA and the small 5S rRNA. In mammalian cells there are approximately 20,000 to 40,000 molecules of each of the RNA polymerases.

Abstract

Manipulating RNA synthesis rates is a primary method the cell uses to adjust its physiological state. Therefore to design synthetic genetic networks and circuits, precise control of RNA synthesis rates is of the utmost importance. Often, however, a native promoter does not exist that has the precise characteristics required for a given application. Here, we describe two methods to change the rates and regulation of RNA synthesis in cells to create RNA synthesis of a desired specification. First, error-prone PCR is discussed for diversifying the properties of native promoters, that is, changing the rate of synthesis in constitutive promoters and the induction properties for an inducible promoter. Specifically, we describe techniques for generating diversified promoter libraries of the constitutive promoters PLtetO-1 in Escherichia coli and TEF1 in Saccharomyces cerevisiae as well as the inducible, oxygen-repressed promoter DAN1 in S. cerevisiae. Beyond generating promoter libraries, we discuss techniques to quantify the parameters of each new promoter. Promoter characteristics for each promoter in hand, the designer can then pick and choose the promoters needed for the specific genetic circuit described in silico. Second, Chemically Induced Chromosomal Evolution (CIChE) is presented as an alternative method to finely adjust RNA synthesis rates in E. coli by variation of gene cassette copy numbers in tandem gene arrays. Both techniques result in precisely defined RNA synthesis and should be of great utility in synthetic biology.

The σ subunit recognizes promoters

The enzymes that synthesize RNA on a DNA template are calledRNA polymerases. These enzymes do not initiate transcription randomly along the length of the chromosome. They start precisely where the gene starts. The transcriptional start sites are marked bypromoter sequences on the DNA, and the first task for the RNA polymerase is finding the promoter.

The RNA polymerase ofE. coli (Table 6.4) consists of a core enzyme with the subunit structure α2ββ′ω and a σ (sigma) subunit that is only loosely bound to the core enzyme.The σsubunit recognizes the promoter, and the core enzyme synthesizes RNA.

The promoters inE. coli have a length of about 60 base pairs, and they look quite different in different genes. Only two short segments, located about 10 base pairs and 35 base pairs upstream of the transcriptional start site, are similar in all promoters. Even these sequences are variable, although we can define aconsensus sequence of the most commonly encountered bases (Fig. 6.17).

This diversity is required because genes must be transcribed at different rates. Some are transcribed up to 10 times per minute, but others are transcribed only once every 10 to 20 minutes. The rate of transcriptional initiation depends on the base sequence of the promoter. In general,the more the promoter resembles the consensus sequence, the higher is the rate of transcription.

The RNA polymerase then separates the DNA double helix over a length of about 18 base pairs, starting at a conserved A-T–rich sequence about 10 base pairs upstream of the transcriptional start site. Strand separation is essential becausetranscription, like DNA replication, requires a single-stranded template.

The σ subunit separates from the core enzyme after the formation of the first 5 to 15 phosphodiester bonds. This marks the transition from the initiation phase to the elongation phase of transcription. The core enzyme now moves along the template strand of the gene while synthesizing the RNA transcript at a rate of about 50 nucleotides per second.

Making RNA

In bacteria, once the sigma subunit of RNA polymerase recognizes the −10 and −35 regions, the core enzyme forms a transcription bubble where the two DNA strands are separated from each other (Fig. 2.3). The strand used by RNA polymerase is called the template strand (aka noncoding or antisense) and is complementary to the resulting mRNA. The core enzyme adds RNA nucleotides in the 5′ to 3′ direction, based on the sequence of the template strand of DNA. The newly made RNA anneals to the template strand of the DNA via hydrogen bonds between base pairs. The opposite strand of DNA is called the coding strand (aka nontemplate or sense strand). Because this is complementary to the template strand, its sequence is identical to the RNA (except for the replacement of thymine with uracil in RNA).

RNA synthesis normally starts at a purine (normally an A) in the DNA that is flanked by two pyrimidines. The most typical start sequence is CAT, but sometimes the A is replaced with a G. The rate of elongation is about 40 nucleotides per second, which is much slower than replication (∼1000 bp/sec). RNA polymerase unwinds the DNA and creates positive supercoils as it travels down the DNA strand. Behind RNA polymerase, the DNA is partially unwound and has surplus negative supercoils. DNA gyrase and topoisomerase I either insert or remove negative supercoils, respectively, returning the DNA back to its normal level of supercoiling (see Chapter 4).

RNA polymerase makes a copy of the gene using the noncoding or template strand of DNA. RNA has uracils instead of thymines.

Viruses must first synthesize messenger RNA (mRNA)

Viruses contain either DNA or RNA, never both. The nucleic acids are present as single or double strands in a linear (DNA or RNA) or circular (DNA) form. The viral genome may be carried on a single molecule of nucleic acid or on several molecules. With these options, it is not surprising that the process of replication in the host cell is also diverse. In viruses containing DNA, mRNA can be formed using the host's own RNA polymerase to transcribe directly from the viral DNA. The RNA of viruses cannot be transcribed in this way, as host polymerases do not work from RNA. If transcription is necessary, the virus must provide its own polymerases. These may be carried in the nucleocapsid or may be synthesized after infection.

Introduction and Background

RNA synthesis by DNA-dependent RNA polymerases (RNAPs) is processive, requiring a single enzyme molecule to transcribe the full length of a gene regardless of the length. The requirement for RNAP to remain resolutely associated with the DNA template through multiple kilobases necessitates an extremely stable transcription elongation complex that can transcribe through different sequences and protein-bound DNA templates. Despite this stability, cells must be able to halt RNA synthesis after transcription of a complete gene or operon, and stop any RNAP that has initiated transcription aberrantly. Failure to terminate transcription of an upstream gene could allow regulation-independent expression of downstream genes, and synthesis of untranslated or antisense transcripts with detrimental consequences; aberrant transcription is particularly problematic for the gene-dense chromosomes common to Bacteria and Archaea. Two general mechanisms have evolved to efficiently disrupt transcription elongation complexes that release the RNA transcript and recycle RNAP for further rounds of transcription.

Multi-subunit RNAPs from each domain share a near identical core structure that envelopes an 8- or 9-bp RNA:DNA hybrid within a tight-fitting pocket (Figure 1). High-resolution crystal structures and a wealth of biochemical data from many different RNAPs demonstrate that hydrogen bonding within the hybrid and contacts between the enclosed nucleic acids and RNAP provide stability to transcription elongation complexes. Despite similar transcription elongation complex architecture, RNAPs from different domains, and each of the eukaryotic RNAPs, respond to different termination signals and factors, suggesting that several mechanisms of transcription termination are possible, or that a diverse set of factors and sequences use a common mechanism to disrupt the complex. Conserved elongation factors (i.e., NusG and NusA) modify RNAP activities and add an additional level of regulation to the elongation–termination decision. The mechanistic details of transcript release are best understood in Bacteria, although some features are shared in each domain. This article focuses on transcription termination and its regulation in Bacteria, with relevant comments to bring attention to similarities and differences in Archaea and Eukarya.

2.1.5 Nucleotide pool and DNA damage response

RNA and DNA synthesis are increased in proliferating cells, especially in chronically proliferative cancer cells. As a result, maintenance of the nucleotide pool is critical for cancer cells to sustain their proliferative ability. It has been shown that cancer cells tend to use de novo nucleotide synthesis pathways, which require glycolysis metabolites, glutamine and aspartate. Deficiency in these metabolites causes increased radio-sensitivity (Villa et al., 2019). Importantly, ribose-5-phosphate from PPP provides the ribose backbone for DNA stability. For instance, deficiency of in PPP enzymes such as transketolase (TKT), transketolase-like 1 (TKTL1), and 6-phosphogluconate dehydrogenase (6PGDH) affects radio- and chemoresistance (Dong and Wang, 2017; Li et al., 2019; Liu et al., 2019). Additionally, N10-formyl-tetrahydrofolate (THF) from the folate cycle is necessary for purine ring synthesis, which is affected by the serine pathway (Villa et al., 2019). The upregulated glucose metabolism in cancer cells favors the production of above metabolites and supports the nucleotide pool (Hay, 2016). Taken together, these findings support the notion that nucleotide pool significantly affects the metabolic pathways involved in DNA damage responses.

Inhibition of Host Cell RNA and Protein Synthesis

Cellular RNA and protein biosynthesis begin to decline within the first few hours after cardiovirus infection, at about the time viral RNA synthesis enters its exponential phase. The decrease in cellular RNA synthesis, which can fall to ∼10% of normal, depending upon the type of cell infected, may reflect a competition among polymerases for ribonucleoside triphosphates. The decrease in host cell protein synthesis is similar in magnitude, and is caused primarily by the ability of the IRES in the 5′-noncoding region of the viral RNA to outcompete capped cellular mRNAs for ribosomes and initiation factors. There is no virus-induced cleavage of the p220 component of the cap-binding complex, eIF–4F, in cardiovirus-infected cells; this is in contrast to the situation in cells infected with poliovirus, rhinovirus or FMD viruses. Cardioviruses may gain their competitive advantage in part by causing modifications to ribosomes (an inhibitor of the translation of capped mRNAs in vitro can be washed off ribosomes from Mengo virus-infected cells) and/or to eIF2 (partial phosphorylation results in partial inactivation with respect to cellular, but not viral, mRNAs).

4.6 EMCV RNA replication protocol

RNA synthesis in EMCV RNA translation–replication reactions is assayed as described previously (Barton et al., 1996; Svitkin and Sonenberg, 2003).

Set up reactions in a 40-μl total volume without [35S]methionine (substitute l-methionine for [35S]methionine). Use a final concentration of ~10 μg/ml of EMCV RNA. [Note: Excess input RNA inhibits RNA replication (Svitkin and Sonenberg, 2003). We recommend that EMCV RNA be titrated for each extract preparation to determine its optimal concentration.] As a negative control, use the reaction that does not contain EMCV RNA.

Incubate the reaction mixtures at 32° for 4 h.

Add 1 μl [α-32P]CTP to the reaction mixtures and continue the incubation at 32° for 1 h.

Stop the reactions by adding 200 μl of deproteinization solution. Incubate the samples at 37° for 15 min.

Add 240 μl of the phenol/chloroform/isoamyl alcohol mixture. Vortex for 30 sec and centrifuge at 16,000×g for 1 min. Carefully recover the aqueous phases (withdraw 200 μl from each sample).

Precipitate the RNA with 20 μl of 10 M ammonium acetate and 2 vol of 100% ethanol. Store samples at −20° (overnight).

7 Conclusions

Viral RNA or DNA synthesis plays a key role in the viral replication cycle, and the enzymes involved are therefore important targets for the development of antivirals. Ensemble, cell-based, and structural biology approaches have provided a wealth of insight into how various viral replication enzymes work, but these techniques have not been able to reveal protein characteristics that occur on ms timescales or at nm distances. Single-molecule tools have been able to probe these time and spatial domains, and have helped us visualize the behavior of enzymes, uncover transient states, and understand how antivirals act. With single-molecule tools becoming more and more accessible for non-biophysicists, including virologists, we look forward to seeing new discoveries in the virology field in the future.