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Why don't cells double gene expression after S-Phase?

Why don't cells double gene expression after S-Phase?



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In the cell cycle (G1-S-G2-M), all of the DNA is replicated during the S or Synthesis stage. The cell may then spend some considerable time in the G2 phase before splitting in the M phase. Since there is double the amount of genetic material in the G2 phase, what mechanism if any, prevents the amount of gene expression from also doubling?


When the double strand is duplicated, the old strand (or, in other terms, the template) is methylated : this modification is enough to prevent the bind by the RNApol system and, by so, the transcription. By methylation, DNA-repair systems are able to detect which is the newer strand, to discover and fix replication errors.

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Genetic engineering or DNA technology has been useful for producing large quantities of a specific protein to treat human diseases. For example, patients with diabetes, hemophilia, or anemia require treatments with insulin, clotting factor, and growth factor proteins. Targeted genes (DNA) can be cut with restriction enzymes and joined with other DNA with the enzyme ligase. A cloning vector is used to carry the recombinant DNA into living cells, so that the cells can synthesize the encoded proteins. The best cloning vectors are small in size, able to replicate its DNA, contain restriction enzyme recognition sites, and have a marker gene (usually antibiotic resistance gene). In this lab, we will use a recombinant plasmid as the cloning vector. This recombinant plasmid contains (1) a promoter that enables transcription of desired gene, (2) a sequence for the initiation of DNA replication (ori site), and (3) an antibiotic resistance gene.

Transforming Bacteria with Recombinant Plasmid

Inserting a gene into a plasmid vector is an important first step in the gene cloning process. However, if the ultimate goal is to produce a large amount of a particular protein, the plasmid must replicate to make sure that there are many copies of the gene and the gene of interest must be expressed, meaning the gene is utilized to produce the encoded protein. Both activities can only occur inside a cell. Therefore, in this lab we will put a recombinant plasmid into E. coli bacteria through a process that is called transformation, so named because it changes the DNA content of the bacteria.

The plasmid will be taken up by bacteria where it replicates, and its genes will be expressed using the bacterial cellular machinery. If a gene of interest has been inserted into the plasmid vector, the bacteria produces the product encoded by that gene.

In this exercise, you will carry out the transformation of E. colibacteria using a recombinant plasmid that contains a gene that produces colored proteins.

Bacterial Transformation

Once a recombinant plasmid is made that contains a gene of interest, such as insulin, the plasmid can enter bacterial cells by a process called transformation. Figure 13.1 illustrates transformation. The uptake of DNA from the environment of a bacterial cell occurs with a very low efficiency in nature. E. coli bacteria have complex plasma membranes that separate the external environment from the internal environment of the cell and carefully regulate which substances can enter and exit the cell. In addition, the cell wall is negatively charged and repels negatively charged DNA molecules.

Cells that have been treated to become competent are more efficient at taking in DNA from their surrounding environment. Competent cells can be made by treating the bacteria with a calcium solution. Calcium ions are positively charged, and will neutralize the negatively charged outer membrane on the E. coli bacteria. With the positive charge now coating the membrane, the inherently negatively charged DNA molecules will move through the plasma membranes and into the cell. The transformation efficiency can be further increased by stressing the cells in a heat shock. By changing the temperature of the cells drastically from cold to warm, the plasma membranes become more fluid and create pores in them. The plasmid DNA can travel from the environment through these pores and enter the cell. The cells are then plunged back into a cold temperature, which causes the pores to close and the plasmid DNA to remain inside the cell.

However, even competent cells do not always uptake the plasmid. For some plasmid DNA molecules, only about 1 in 10,000 cells will be transformed. When so few cells have taken in the plasmid, how will you be able to identify transformed cells? When designing a recombinant plasmid, one of the requirements is to add a gene for an antibiotic resistance. This way, the bacteria can be grown in the media with an antibiotic added to it, and only cells that have the resistance gene, such as those that express the recombinant plasmid, will be able to grow.

Figure 1. Bacterial transformation.

From Plasmid DNA to Protein

After a recombinant plasmid enters a bacterial cell, the cell begins to express the genes on it. DNA polymerase locates the ori- the origin of replication, and starts to replicate the plasmid using the bacterial cell&rsquos machinery. Multiple copies of the recombinant plasmid can enable the bacterial cell to express large amounts of a protein. Usually, a bacterial cell will only make the protein of interest, after it is induced to do so by adding a chemical which will promote the transcription of the gene. Recall that to express the gene encoding the protein on the recombinant plasmid, DNA is transcribed to mRNA, which is then translated to protein (Figure 13.2). The expressed proteins may affect the visible traits when observing the bacteria colonies.

Figure 2. Gene expression from a plasmid in the bacterial cell

Recombinant plasmids and other forms of genetic engineering is possible because all living organisms use DNA as a platform to encode genetic information. Genes from different organisms can be expressed in other organisms like bacteria since they are encoded in DNA. The DNA instructions can be transferred, and other organisms can express foreign traits.

Proteins have many different functions inside and cells. They are made up of smaller subunits, amino acids, which are encoded by DNA nucleotides. A specific three nucleotide sequence that codes for a single amino acid is called a codon. For example, the codon TTG codes for the amino acid tryptophan, whereas the codon AAG codes for the amino acid lysine. In many cases, more than one codon can encode the same amino acid. For example, AAA is also a codon for lysine. In addition, there are informational codons, such as the start codon (ATG) and the stop codon (TTA), which show where in the DNA sequence the code for the protein begins and ends.

Transforming Bacteria with Plasmids

In this laboratory experiment you will transform E. coli bacteria cells with plasmids. You will be using E. coli that has been made competent with a calcium chloride treatment, and form two different testing groups: a negative control cell group that does not have plasmids added to it, and the experimental group that has the plasmids added. After the cells are heat-shocked, they will be grown under various testing conditions:

  • The control group on nutrient agar (a type of growth media that bacteria thrive on).
  • The control group on nutrient agar with an antibiotic added.
  • The experimental group on nutrient agar.
  • The experimental group on nutrient agar with an antibiotic.
  • The experimental group on nutrient agar, antibiotic and an inducer (such as IPTG).

By examining the growth of bacteria under these conditions, you can verify that your procedure worked, and you can identify the bacteria transformed with the added plasmid. How will you know if you are successful? In the examples for plasmids we have recommended for this exercise, the recombinant bacteria will have a new and highly visible trait: It will now produce colored protein, which makes the cells themselves colored! As the bacteria multiply on the media, they form visible collections of cells called colonies. Each colony represents the decedents of the original bacterial cell that landed on that spot on the medium and began to replicate. Thus colonies are clones (exact copies) of the cell that began the replication process. 


The relevant components of your plasmid are the gene for the colored protein, the inducible promoter, and the ampicillin resistance gene (ampR). The ampR gene confers resistance to the antibiotic ampicillin. (Biotechnologists call these genes selectable markers because only bacteria having the gene will survive in the presence of an antibiotic.) If the inducer is present in the bacteria, the promoter will be &ldquoturned on&rdquo so RNA polymerase can transcribe the gene of interest. This will allow protein to be produced.


Biology TEST 2

__3__
DNA replication or S-phase occurs prior to this type of division.

__2__
Used to produce cells during tissue growth and healing.

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Produces daughter cells with the same number of chromosomes as the parent cell.

__1__
Produces daughter cells with half the chromosomes of the parent cell.

Separation of homologous chromosomes into different daughter cells.

1:2:1 ratio of homozygous dominant, heterozygous, and homozygous recessive genotypes

__1__
Transcription of DNA to mRNA

__2__
Translation of mRNA to polypeptide

__2__
Protein folding and processing

__2__
The molecule that binds to the mRNA and brings amino acid monomers to build the polypeptide

__5__
The monomers used to build the polypeptide or protein

__3__
The molecule that carries the genetic information out of the nucleus and interacts with tRNA

__4__
The molecule in the nucleus that contains the genetic information


Introduction

T cells are key components of the adaptive immune system. Mature T cells are generally considered to express either the CD4 or CD8 coreceptor, in addition to their TCR, and consequently, the T cell pool is commonly divided into two subsets, based on expression of either CD4 or CD8. The CD4 molecule, a member of the IgR family, is encoded by a single gene and expressed on the surface as a transmembrane monomer [1, 2]. CD4 interacts with the β2-domain of MHC class II molecules and has also been shown to act as an important chemotactic receptor for IL-16 [3]. In contrast to the CD4 monomer, the CD8 coreceptor exists as a CD8αα homodimer or a CD8αβ heterodimer yet, in both cases, the Ig domains of the CD8 molecule bind MHC class I [4]. No substantial difference in MHC class I affinity binding has been observed between murine CD8αα and CD8αβ molecules [5], despite that the sequences for the CD8 homo- and heterodimer are shown to be <20% identical [4]. The intracellular regions of CD4 and CD8 interact with the tyrosine kinase, lck [6]. By inducing intracellular signaling through lck, the CD8 coreceptor is an important factor affecting TCR-mediated T cell activation and modulation of immune responses, both in terms of immunosuppression and cytotoxicity [7, 8].


Discussion

In this study, we systematically investigated transcriptional and epigenetic dynamics during the cell cycle by analyzing GRO-seq, RNA-seq, and histone marks ChIP-seq data at G0/G1, G1/S, and M phases in the MCF-7 breast cancer cell line. Our study revealed (i) a lag between transcription and steady-state RNA expression at the cell-cycle level (ii) a large amount of active transcription during early mitosis (iii) a global increase in active histone modifications at mitosis (iv) thousands of cell-cycle–regulated eRNAs and (v) dynamic eRNAs bound by transcription factors such as KLF4 that regulate cell-cycle progression.

Steady-state mRNA abundance is influenced by a few factors, including transcription, RNA processing, maturation, and degradation. Therefore, measuring steady-state mRNA levels by microarray or RNA-seq techniques may not accurately reflect active transcription. Indeed, GRO-seq and 4-thiouridine metabolic labeling followed by sequencing (4sU-seq) analyses that measure nascent transcription have revealed a broad inconsistency between transcription rate and mRNA levels (25, 28, 61, 62). Specifically, there is a delay in steady-state expression reflecting the transcription and mass production of rapidly degraded transcripts that are not detectable at the steady-state expression level. Most of the previous nascent transcription analyses were performed with unsynchronized cells or with synchronized cells within a short time window that was insufficient to cover multiple cell-cycle stages (26, 28, 29, 32, 35, 36, 62). Importantly, our GRO-seq and RNA-seq analysis at different cell-cycle stages revealed a lag between active transcription and steady-state expression during the cell cycle. The RNA degradation rate has been considered the most prominent measurable factor that contributes to the lag between transcription and accumulated RNA levels. Recent studies demonstrated that the half-lives of mammalian genes range from less than 1 min to more than 3 h (61, 62). In agreement with these observations, our data showed that mitotic genes are most highly transcribed at G1/S, and the genes most highly transcribed at M phase are more abundant at G0/G1, suggesting that these genes have an extremely long half-life.

Mitotic chromatin is transcriptionally inactive in general, and even ongoing transcriptions are aborted to ensure the integrity of the separating chromosomes (63). However, exceptions have been found in which the promoter of the cyclin B1 gene maintains an open chromatin configuration, and the gene is actively transcribed during mitosis (64). Recently, additional large-scale studies have revealed that part of the mitotic chromatin remains accessible to Pol II and transcription factors such as MLL, BRD4, GATA1, FOXA1, and AR (43 ⇓ ⇓ –46, 65). Our GRO-seq data showed that although CCNB1 transcription peaks at G1/S, strong nascent transcription was observed at M phase. More interestingly, we identified a group of genes with a transcription peak at M phase. The observation that this group was enriched for unusually long genes made us hypothesize that the GRO-seq signal was from the incomplete transcription from previous stages (66). We therefore compared the GRO-seq signal along the gene body to identify the longest quarter of genes with the highest GRO-seq signal at M phase. If the hypothesis is correct, we should be able to observe a GRO-seq signal pattern shifted from the TSS toward the CPA site during the cell-cycle progression from G0/G1 to M phase. Our analysis revealed a uniform distribution of signal along the gene body for most genes. In addition, reanalysis of publically available Pol II ChIP-seq data in early mitotic cells pretreated with and without flavopiridol (47) confirmed that Pol II is actively engaged at the TSS of these genes. Together, the results suggested that the high GRO-seq signal of these genes arose from active transcription at early M phase rather than from incomplete transcription at the G0/G1 and G1/S phases. Importantly, Liang et al. (47) recently reported mitotic transcriptional activation as a mechanism to clear actively engaged Pol II from mitotic chromatin this mechanism is consistent with our observation of active transcription at early mitotic cells.

In support of active transcription at M phase, we observed extremely stable chromatin states marked by active histone modifications H3K4me2 and H3K27ac across different cell-cycle stages. In addition, the total H3K4me2 and H3K27ac levels increased significantly at M phase. Previous studies have identified mitotic-specific H4K20 methylation and the dynamics of H3K36 and H3K27 methylations across the cell cycle (67 ⇓ –69). The functional role of posttranscriptional histone modifications in the cell cycle is still largely unknown and warrants further analysis. It is worth noting that these observations were made in cancer cells with uncontrolled cell division these cells may differ from normal cells with more stringent cell-cycle regulation (70). Future studies are warranted to explore the mechanisms underlying the active transcription during mitosis in normal and cancer cells.

Taken together, our analyses identified thousands of eRNAs and related transcription factors that are highly correlated with cell-cycle–regulated transcription but not with steady-state expression, thus highlighting the importance of transcriptional and epigenetic dynamics during cell-cycle progression. Overall, our study provides a comprehensive view of transcriptional landscape across the cell cycle and deepens our understanding of transcriptional dynamics during cell cycle. Future studies combining transcription, expression, and proteomics data at more detailed time courses are warranted to provide a more comprehensive view of cell-cycle regulation.


'Zombie' genes? Research shows some genes come to life in the brain after death

In the hours after we die, certain cells in the human brain are still active. Some cells even increase their activity and grow to gargantuan proportions, according to new research from the University of Illinois Chicago.

In a newly published study in the journal Scientific Reports, the UIC researchers analyzed gene expression in fresh brain tissue -- which was collected during routine brain surgery -- at multiple times after removal to simulate the post-mortem interval and death. They found that gene expression in some cells actually increased after death.

These 'zombie genes' -- those that increased expression after the post-mortem interval -- were specific to one type of cell: inflammatory cells called glial cells. The researchers observed that glial cells grow and sprout long arm-like appendages for many hours after death.

"That glial cells enlarge after death isn't too surprising given that they are inflammatory and their job is to clean things up after brain injuries like oxygen deprivation or stroke," said Dr. Jeffrey Loeb, the John S. Garvin Professor and head of neurology and rehabilitation at the UIC College of Medicine and corresponding author on the paper.

What's significant, Loeb said, is the implications of this discovery -- most research studies that use postmortem human brain tissues to find treatments and potential cures for disorders such as autism, schizophrenia and Alzheimer's disease, do not account for the post-mortem gene expression or cell activity.

"Most studies assume that everything in the brain stops when the heart stops beating, but this is not so," Loeb said. "Our findings will be needed to interpret research on human brain tissues. We just haven't quantified these changes until now."

Loeb and his team noticed that the global pattern of gene expression in fresh human brain tissue didn't match any of the published reports of postmortem brain gene expression from people without neurological disorders or from people with a wide variety of neurological disorders, ranging from autism to Alzheimer's.

"We decided to run a simulated death experiment by looking at the expression of all human genes, at time points from 0 to 24 hours, from a large block of recently collected brain tissues, which were allowed to sit at room temperature to replicate the postmortem interval," Loeb said.

Loeb and colleagues are at a particular advantage when it comes to studying brain tissue. Loeb is director of the UI NeuroRepository, a bank of human brain tissues from patients with neurological disorders who have consented to having tissue collected and stored for research either after they die, or during standard of care surgery to treat disorders such as epilepsy. For example, during certain surgeries to treat epilepsy, epileptic brain tissue is removed to help eliminate seizures. Not all of the tissue is needed for pathological diagnosis, so some can be used for research. This is the tissue that Loeb and colleagues analyzed in their research.

They found that about 80% of the genes analyzed remained relatively stable for 24 hours -- their expression didn't change much. These included genes often referred to as housekeeping genes that provide basic cellular functions and are commonly used in research studies to show the quality of the tissue. Another group of genes, known to be present in neurons and shown to be intricately involved in human brain activity such as memory, thinking and seizure activity, rapidly degraded in the hours after death. These genes are important to researchers studying disorders like schizophrenia and Alzheimer's disease, Loeb said.

A third group of genes -- the 'zombie genes' -- increased their activity at the same time the neuronal genes were ramping down. The pattern of post-mortem changes peaked at about 12 hours.

"Our findings don't mean that we should throw away human tissue research programs, it just means that researchers need to take into account these genetic and cellular changes, and reduce the post-mortem interval as much as possible to reduce the magnitude of these changes," Loeb said. "The good news from our findings is that we now know which genes and cell types are stable, which degrade, and which increase over time so that results from postmortem brain studies can be better understood."

Fabien Dachet, Tibor Valyi-Nagy, Kunwar Narayan, Anna Serafini and Gayatry Mohapatra of UIC James Brown and Susan Celniker of Lawrence Berkeley National Laboratory Nathan Boley of the University of California, Berkeley and Thomas Gingeras of Cold Spring Harbor Laboratory are co-authors on the paper.

This research was funded by grants from the National Institutes of Health (R01NS109515, R56NS083527, and UL1TR002003).


Contents

Barbara McClintock discovered the first TEs in maize (Zea mays) at the Cold Spring Harbor Laboratory in New York. McClintock was experimenting with maize plants that had broken chromosomes. [5]

In the winter of 1944–1945, McClintock planted corn kernels that were self-pollinated, meaning that the silk (style) of the flower received pollen from its own anther. [5] These kernels came from a long line of plants that had been self-pollinated, causing broken arms on the end of their ninth chromosomes. [5] As the maize plants began to grow, McClintock noted unusual color patterns on the leaves. [5] For example, one leaf had two albino patches of almost identical size, located side by side on the leaf. [5] McClintock hypothesized that during cell division certain cells lost genetic material, while others gained what they had lost. [6] However, when comparing the chromosomes of the current generation of plants with the parent generation, she found certain parts of the chromosome had switched position. [6] This refuted the popular genetic theory of the time that genes were fixed in their position on a chromosome. McClintock found that genes could not only move, but they could also be turned on or off due to certain environmental conditions or during different stages of cell development. [6]

McClintock also showed that gene mutations could be reversed. [7] She presented her report on her findings in 1951, and published an article on her discoveries in Genetics in November 1953 entitled "Induction of Instability at Selected Loci in Maize". [8]

At the 1951 Cold Spring Harbor Symposium where she first publicized her findings, her talk was met with dead silence. [9] Her work was largely dismissed and ignored until the late 1960s–1970s when, after TEs were found in bacteria, it was rediscovered. [10] She was awarded a Nobel Prize in Physiology or Medicine in 1983 for her discovery of TEs, more than thirty years after her initial research. [11]

Approximately 64% of the maize genome is made up of TEs, [12] [13] as is 44% of the human genome. [14]

Transposable elements represent one of several types of mobile genetic elements. TEs are assigned to one of two classes according to their mechanism of transposition, which can be described as either copy and paste (Class I TEs) or cut and paste (Class II TEs). [15]

Retrotransposon Edit

Class I TEs are copied in two stages: first, they are transcribed from DNA to RNA, and the RNA produced is then reverse transcribed to DNA. This copied DNA is then inserted back into the genome at a new position. The reverse transcription step is catalyzed by a reverse transcriptase, which is often encoded by the TE itself. The characteristics of retrotransposons are similar to retroviruses, such as HIV.

Retrotransposons are commonly grouped into three main orders:

  • Retrotransposons, with long terminal repeats (LTRs), which encode reverse transcriptase, similar to retroviruses
  • Retroposons, long interspersed nuclear elements (LINEs, LINE-1s, or L1s), which encode reverse transcriptase but lack LTRs, and are transcribed by RNA polymerase II (SINEs) do not encode reverse transcriptase and are transcribed by RNA polymerase III

(Retroviruses can also be considered TEs. For example, after conversion of retroviral RNA into DNA inside a host cell, the newly produced retroviral DNA is integrated into the genome of the host cell. These integrated DNAs are termed proviruses. The provirus is a specialized form of eukaryotic retrotransposon, which can produce RNA intermediates that may leave the host cell and infect other cells. The transposition cycle of retroviruses has similarities to that of prokaryotic TEs, suggesting a distant relationship between the two.)

DNA transposons Edit

The cut-and-paste transposition mechanism of class II TEs does not involve an RNA intermediate. The transpositions are catalyzed by several transposase enzymes. Some transposases non-specifically bind to any target site in DNA, whereas others bind to specific target sequences. The transposase makes a staggered cut at the target site producing sticky ends, cuts out the DNA transposon and ligates it into the target site. A DNA polymerase fills in the resulting gaps from the sticky ends and DNA ligase closes the sugar-phosphate backbone. This results in target site duplication and the insertion sites of DNA transposons may be identified by short direct repeats (a staggered cut in the target DNA filled by DNA polymerase) followed by inverted repeats (which are important for the TE excision by transposase).

Cut-and-paste TEs may be duplicated if their transposition takes place during S phase of the cell cycle, when a donor site has already been replicated but a target site has not yet been replicated. [17] Such duplications at the target site can result in gene duplication, which plays an important role in genomic evolution. [18] : 284

Not all DNA transposons transpose through the cut-and-paste mechanism. In some cases, a replicative transposition is observed in which a transposon replicates itself to a new target site (e.g. helitron).

Class II TEs comprise less than 2% of the human genome, making the rest Class I. [19]

Autonomous and non-autonomous Edit

Transposition can be classified as either "autonomous" or "non-autonomous" in both Class I and Class II TEs. Autonomous TEs can move by themselves, whereas non-autonomous TEs require the presence of another TE to move. This is often because dependent TEs lack transposase (for Class II) or reverse transcriptase (for Class I).

Activator element (Ac) is an example of an autonomous TE, and dissociation elements (Ds) is an example of a non-autonomous TE. Without Ac, Ds is not able to transpose.

New discoveries of transposable elements have shown the exact distribution of TEs with respect to their transcription start sites (TSSs) and enhancers. A recent study found that a promoter contains 25% of regions that harbor TEs. It is known that older TEs are not found in TSS locations because TEs frequency starts as a function once there is a distance from the TSS. A possible theory for this is that TEs might interfere with the transcription pausing or the first-intro splicing. [20] Also as mentioned before, the presence of TEs closed by the TSS locations is correlated to their evolutionary age (number of different mutations that TEs can develop during the time).

  • The first TEs were discovered in maize (Zea mays) by Barbara McClintock in 1948, for which she was later awarded a Nobel Prize. She noticed chromosomal insertions, deletions, and translocations caused by these elements. These changes in the genome could, for example, lead to a change in the color of corn kernels. About 85% of the maize genome consists of TEs. [21] The Ac/Ds system described by McClintock are Class II TEs. Transposition of Ac in tobacco has been demonstrated by B. Baker (Plant Transposable Elements, pp 161–174, 1988, Plenum Publishing Corp., ed. Nelson).
  • In the pond microorganism, Oxytricha, TEs play such a critical role that when removed, the organism fails to develop. [22]
  • One family of TEs in the fruit fly Drosophila melanogaster are called P elements. They seem to have first appeared in the species only in the middle of the twentieth century within the last 50 years, they spread through every population of the species. Gerald M. Rubin and Allan C. Spradling pioneered technology to use artificial P elements to insert genes into Drosophila by injecting the embryo. [23][24][25]
  • In bacteria, TE's usually carry an additional gene for functions other than transposition, often for antibiotic resistance. In bacteria, transposons can jump from chromosomal DNA to plasmid DNA and back, allowing for the transfer and permanent addition of genes such as those encoding antibiotic resistance (multi-antibiotic resistant bacterial strains can be generated in this way). Bacterial transposons of this type belong to the Tn family. When the transposable elements lack additional genes, they are known as insertion sequences.
  • In humans, the most common TE is the Alu sequence. It is approximately 300 bases long and can be found between 300,000 and one million times in the human genome. Alu alone is estimated to make up 15–17% of the human genome. [19] are another prominent class of transposons found in multiple species, including humans. The Mariner transposon was first discovered by Jacobson and Hartl in Drosophila. [26] This Class II transposable element is known for its uncanny ability to be transmitted horizontally in many species. [27][28] There are an estimated 14,000 copies of Mariner in the human genome comprising 2.6 million base pairs. [29] The first mariner-element transposons outside of animals were found in Trichomonas vaginalis. [30] These characteristics of the Mariner transposon inspired the science fiction novel The Mariner Project by Bob Marr. transposition is the best-known example of replicative transposition.
  • In Yeast genomes, (Saccharomyces cerevisiae) there are five distinct retrotransposon families: Ty1, Ty2, Ty3, Ty4 and Ty5. [31]
  • A helitron is a TE found in eukaryotes that is thought to replicate by a rolling-circle mechanism.
  • In human embryos, two types of transposons combined to form noncoding RNA that catalyzes the development of stem cells. During the early stages of a fetus's growth, the embryo's inner cell mass expands as these stem cells enumerate. The increase of this type of cells is crucial, since stem cells later change form and give rise to all the cells in the body.
  • In peppered moths, a transposon in a gene called cortex caused the moths' wings to turn completely black. This change in coloration helped moths to blend in with ash and soot-covered areas during the Industrial Revolution.

Transposons have coexisted with eukaryotes for thousands of years and through their coexistence have become integrated in many organisms' genomes. Colloquially known as 'jumping genes', transposons can move within and between genomes allowing for this integration.

While there are many positive effects of transposons in their host eukaryotic genomes, there are some instances of mutagenic effects that TEs have on genomes leading to disease and malignant genetic alterations. [32]

Mechanisms of mutagenesis Edit

TEs are mutagens and due to the contribution to the formation of new cis-regulatory DNA elements that are connected to many transcription factors that are found in living cells TEs can undergo many evolutionary mutations and alterations. These are often the causes of genetic disease, and gives the potential lethal effects of ectopic expression. [33]

TEs can damage the genome of their host cell in different ways: [32]

  • A transposon or a retrotransposon that inserts itself into a functional gene can disable that gene.
  • After a DNA transposon leaves a gene, the resulting gap may not be repaired correctly
  • Multiple copies of the same sequence, such as Alu sequences, can hinder precise chromosomal pairing during mitosis and meiosis, resulting in unequal crossovers, one of the main reasons for chromosome duplication.

TEs use a number of different mechanisms to cause genetic instability and disease in their host genomes.

  • Expression of disease causing, damaging proteins that inhibit normal cellular function.
    • Many TEs contain promoters which drive transcription of their own transposase. These promoters can cause aberrant expression of linked genes, causing disease or mutantphenotypes. [34]

    Diseases often caused by TEs include

      A and B
      • LINE1 (L1) TEs that land on the human Factor VIII have been shown to cause haemophilia [35]
      • Insertion of L1 into the APC gene causes colon cancer, confirming that TEs play an important role in disease development. [36]
      • Insertion of Alu element into the PBGD gene leads to interference with the coding region and leads to acute intermittent porphyria [37] (AIP).
      • LINE1(L1) TE's and other retrotransposons have been linked to cancer because they cause genomic instability. [35]
      • Caused by SVA transposable element insertion in the fukutin (FKTN) gene which renders the gene inactive. [35]
      • Transposable element dysregulation can cause neuronal death, leading to neurodegenerative disorders [40]

      One study estimated the rate of transposition of a particular retrotransposon, the Ty1 element in Saccharomyces cerevisiae. Using several assumptions, the rate of successful transposition event per single Ty1 element came out to be about once every few months to once every few years. [41] Some TEs contain heat-shock like promoters and their rate of transposition increases if the cell is subjected to stress, [42] thus increasing the mutation rate under these conditions, which might be beneficial to the cell.

      Cells defend against the proliferation of TEs in a number of ways. These include piRNAs and siRNAs, [43] which silence TEs after they have been transcribed.

      If organisms are mostly composed of TEs, one might assume that disease caused by misplaced TEs is very common, but in most cases TEs are silenced through epigenetic mechanisms like DNA methylation, chromatin remodeling and piRNA, such that little to no phenotypic effects nor movements of TEs occur as in some wild-type plant TEs. Certain mutated plants have been found to have defects in methylation-related enzymes (methyl transferase) which cause the transcription of TEs, thus affecting the phenotype. [4] [44]

      One hypothesis suggests that only approximately 100 LINE1 related sequences are active, despite their sequences making up 17% of the human genome. In human cells, silencing of LINE1 sequences is triggered by an RNA interference (RNAi) mechanism. Surprisingly, the RNAi sequences are derived from the 5' untranslated region (UTR) of the LINE1, a long terminal which repeats itself. Supposedly, the 5' LINE1 UTR that codes for the sense promoter for LINE1 transcription also encodes the antisense promoter for the miRNA that becomes the substrate for siRNA production. Inhibition of the RNAi silencing mechanism in this region showed an increase in LINE1 transcription. [4] [45]

      TEs are found in almost all life forms, and the scientific community is still exploring their evolution and their effect on genome evolution. It is unclear whether TEs originated in the last universal common ancestor, arose independently multiple times, or arose once and then spread to other kingdoms by horizontal gene transfer. [46] While some TEs confer benefits on their hosts, most are regarded as selfish DNA parasites. In this way, they are similar to viruses. Various viruses and TEs also share features in their genome structures and biochemical abilities, leading to speculation that they share a common ancestor. [47]

      Because excessive TE activity can damage exons, many organisms have acquired mechanisms to inhibit their activity. Bacteria may undergo high rates of gene deletion as part of a mechanism to remove TEs and viruses from their genomes, while eukaryotic organisms typically use RNA interference to inhibit TE activity. Nevertheless, some TEs generate large families often associated with speciation events. Evolution often deactivates DNA transposons, leaving them as introns (inactive gene sequences). In vertebrate animal cells, nearly all 100,000+ DNA transposons per genome have genes that encode inactive transposase polypeptides. [48] The first synthetic transposon designed for use in vertebrate (including human) cells, the Sleeping Beauty transposon system, is a Tc1/mariner-like transposon. Its dead ("fossil") versions are spread widely in the salmonid genome and a functional version was engineered by comparing those versions. [49] Human Tc1-like transposons are divided into Hsmar1 and Hsmar2 subfamilies. Although both types are inactive, one copy of Hsmar1 found in the SETMAR gene is under selection as it provides DNA-binding for the histone-modifying protein. [50] Many other human genes are similarly derived from transposons. [51] Hsmar2 has been reconstructed multiple times from the fossil sequences. [52]

      Large quantities of TEs within genomes may still present evolutionary advantages, however. Interspersed repeats within genomes are created by transposition events accumulating over evolutionary time. Because interspersed repeats block gene conversion, they protect novel gene sequences from being overwritten by similar gene sequences and thereby facilitate the development of new genes. TEs may also have been co-opted by the vertebrate immune system as a means of producing antibody diversity. The V(D)J recombination system operates by a mechanism similar to that of some TEs. TEs also serve to generate repeating sequences that can form dsRNA to act as a substrate for the action of ADAR in RNA editing. [53]

      TEs can contain many types of genes, including those conferring antibiotic resistance and ability to transpose to conjugative plasmids. Some TEs also contain integrons, genetic elements that can capture and express genes from other sources. These contain integrase, which can integrate gene cassettes. There are over 40 antibiotic resistance genes identified on cassettes, as well as virulence genes.

      Transposons do not always excise their elements precisely, sometimes removing the adjacent base pairs this phenomenon is called exon shuffling. Shuffling two unrelated exons can create a novel gene product or, more likely, an intron. [54]

      Evolutionary drive for TEs on the genomic context Edit

      There is a hypothesis that states that TEs might provide a ready source of DNA that could be co-opted by the cell to help regulate gene expression. Research showed that many diverse modes of TEs co-evolution along with some transcription factors targeting TE-associated genomic elements and chromatin are evolving from TE sequences. Most of the time, these particular modes do not follow the simple model of TEs and regulating host gene expression. [55]

      Transposable elements can be harnessed in laboratory and research settings to study genomes of organisms and even engineer genetic sequences. Use of transposable elements can be split into two categories: for genetic engineering and as a genetic tool.

      Genetic engineering Edit

      • Insertional mutagenesis uses the features of a TE to insert a sequence. In most cases this is used to remove a DNA sequence or cause a frameshift mutation.
        • In some cases the insertion of a TE into a gene can disrupt that gene's function in a reversible manner where transposase-mediated excision of the DNA transposon restores gene function.
        • This produces plants in which neighboring cells have different genotypes.
        • This feature allows researchers to distinguish between genes that must be present inside of a cell in order to function (cell-autonomous) and genes that produce observable effects in cells other than those where the gene is expressed.

        Genetic tool Edit

        In addition to the qualities mentioned for Genetic engineering, a Genetic tool also:-

        • Used for analysis of gene expression and protein functioning in signature-tagging mutagenesis.
          • This analytical tool allows researchers the ability to determine phenotypic expression of gene sequences. Also, this analytic technique mutates the desired locus of interest so that the phenotypes of the original and the mutated gene can be compared.

          Specific applications Edit

          • TEs are also a widely used tool for mutagenesis of most experimentally tractable organisms. The Sleeping Beauty transposon system has been used extensively as an insertional tag for identifying cancer genes. [56]
          • The Tc1/mariner-class of TEs Sleeping Beauty transposon system, awarded Molecule of the Year in 2009, [57] is active in mammalian cells and is being investigated for use in human gene therapy. [58][59][60]
          • TEs are used for the reconstruction of phylogenies by the means of presence/absence analyses. [61] Transposons can act as biological mutagen in bacteria.
          • Common organisms which the use of Transposons has been well developed are:
            • Drosophila[62]
            • Arabidopsis thaliana[44]
            • Escherichia coli

            De novo repeat identification is an initial scan of sequence data that seeks to find the repetitive regions of the genome, and to classify these repeats. Many computer programs exist to perform de novo repeat identification, all operating under the same general principles. [57] As short tandem repeats are generally 1–6 base pairs in length and are often consecutive, their identification is relatively simple. [56] Dispersed repetitive elements, on the other hand, are more challenging to identify, due to the fact that they are longer and have often acquired mutations. However, it is important to identify these repeats as they are often found to be transposable elements (TEs). [57]

            De novo identification of transposons involves three steps: 1) find all repeats within the genome, 2) build a consensus of each family of sequences, and 3) classify these repeats. There are three groups of algorithms for the first step. One group is referred to as the k-mer approach, where a k-mer is a sequence of length k. In this approach, the genome is scanned for overrepresented k-mers that is, k-mers that occur more often than is likely based on probability alone. The length k is determined by the type of transposon being searched for. The k-mer approach also allows mismatches, the number of which is determined by the analyst. Some k-mer approach programs use the k-mer as a base, and extend both ends of each repeated k-mer until there is no more similarity between them, indicating the ends of the repeats. [57] Another group of algorithms employs a method called sequence self-comparison. Sequence self-comparison programs use databases such as AB-BLAST to conduct an initial sequence alignment. As these programs find groups of elements that partially overlap, they are useful for finding highly diverged transposons, or transposons with only a small region copied into other parts of the genome. [58] Another group of algorithms follows the periodicity approach. These algorithms perform a Fourier transformation on the sequence data, identifying periodicities, regions that are repeated periodically, and are able to use peaks in the resultant spectrum to find candidate repetitive elements. This method works best for tandem repeats, but can be used for dispersed repeats as well. However, it is a slow process, making it an unlikely choice for genome scale analysis. [57]

            The second step of de novo repeat identification involves building a consensus of each family of sequences. A consensus sequence is a sequence that is created based on the repeats that comprise a TE family. A base pair in a consensus is the one that occurred most often in the sequences being compared to make the consensus. For example, in a family of 50 repeats where 42 have a T base pair in the same position, the consensus sequence would have a T at this position as well, as the base pair is representative of the family as a whole at that particular position, and is most likely the base pair found in the family's ancestor at that position. [57] Once a consensus sequence has been made for each family, it is then possible to move on to further analysis, such as TE classification and genome masking in order to quantify the overall TE content of the genome.

            Transposable elements have been recognized as good candidates for stimulating gene adaptation, through their ability to regulate the expression levels of nearby genes. [59] Combined with their "mobility", transposable elements can be relocated adjacent to their targeted genes, and control the expression levels of the gene, dependent upon the circumstances.

            The study conducted in 2008, "High Rate of Recent Transposable Element–Induced Adaptation in Drosophila melanogaster", used D. melanogaster that had recently migrated from Africa to other parts of the world, as a basis for studying adaptations caused by transposable elements. Although most of the TEs were located on introns, the experiment showed the significant difference on gene expressions between the population in Africa and other parts of the world. The four TEs that caused the selective sweep were more prevalent in D. melanogaster from temperate climates, leading the researchers to conclude that the selective pressures of the climate prompted genetic adaptation. [60] From this experiment, it has been confirmed that adaptive TEs are prevalent in nature, by enabling organisms to adapt gene expression as a result of new selective pressures.

            However, not all effects of adaptive TEs are beneficial to the population. In the research conducted in 2009, "A Recent Adaptive Transposable Element Insertion Near Highly Conserved Developmental Loci in Drosophila melanogaster", a TE, inserted between Jheh 2 and Jheh 3, revealed a downgrade in the expression level of both of the genes. Down regulation of such genes has caused Drosophila to exhibit extended developmental time and reduced egg to adult viability. Although this adaptation was observed in high frequency in all non-African populations, it was not fixed in any of them. [61] This is not hard to believe, since it is logical for a population to favor higher egg to adult viability, therefore trying to purge the trait caused by this specific TE adaptation.

            At the same time, there have been several reports showing the advantageous adaptation caused by TEs. In the research done with silkworms, "An Adaptive Transposable Element insertion in the Regulatory Region of the EO Gene in the Domesticated Silkworm", a TE insertion was observed in the cis-regulatory region of the EO gene, which regulates molting hormone 20E, and enhanced expression was recorded. While populations without the TE insert are often unable to effectively regulate hormone 20E under starvation conditions, those with the insert had a more stable development, which resulted in higher developmental uniformity. [63]

            These three experiments all demonstrated different ways in which TE insertions can be advantageous or disadvantageous, through means of regulating the expression level of adjacent genes. The field of adaptive TE research is still under development and more findings can be expected in the future.

            Recent studies have confirmed that TEs can contribute to the generation of transcription factors. However, how this process of contribution can have an impact on the participation of genome control networks. TEs are more common in many regions of the DNA and it makes up 45% of total human DNA. Also, TEs contributed to 16% of transcription factor binding sites. A larger number of motifs are also found in non-TE-derived DNA, and the number is larger than TE-derived DNA. All these factors correlate to the direct participation of TEs in many ways of gene control networks. [64]


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            Acknowledgements

            The authors are grateful to Monica Romero for help with the imaging in the LLUSM Advanced Imaging and Microscopy Core with support of NSF Grant MRI-DBI 0923559 and the Loma Linda University School of Medicine.

            Funding

            This work was supported by the Ministry of Science and Technology of China (2015CB964902, 2013CB966902, and 2012CB966601), the National Natural Science Foundation of China (81500148, 81570164, and 81421002), the Loma Linda University School of Medicine GCAT grant (2015), and Telemedicine and Advanced Technology Research Center (W81XWH-08-1-0697).

            Availability of data and materials

            Authors’ contributions

            XBZ conceived of and supervised the study. JPZ, XLL, GHL, WC, CA, LZ, WW, YWF, JX, and XBZ conducted the experiments. JPZ, XLL, DB, GDB, WY, TC, and XBZ analyzed the results. JPZ, XLL, TC, and XBZ wrote the paper. All authors reviewed the manuscript. All authors read and approved the final manuscript.

            Competing interests

            The authors declare that they have no competing interests.

            Ethics approval and consent to participate

            This study did not require ethics approval.


            Knock-in By Numbers

            Researcher: Greg Findlay, MD/PhD candidate in the lab of Jay Shendure, University of Washington

            Project: Findlay and his colleagues were aiming to improve how clinicians interpret mutations in the breast and ovarian cancer gene BRCA1. That gene has thousands of variants, but researchers don’t know how most of them affect its function. To study the impact of these variants, they used a knock-in technique they developed called saturation genome editing (Nature, 562:217–22, 2018).

            In an immortalized haploid human cell line, they used CRISPR-Cas9 to knock in 4,000 tiny variants in millions of cells at once in vitro. The genome is cut at the same spot in each cell, but each cell’s genome receives a different variant. To promote HDR, they also knocked out the ligase4 gene, disabling the NHEJ repair pathway—a step that yielded a threefold gain in efficiency, Findlay says. Finally, since all the cells’ knock-ins are different, they sequenced the cells deeply, covering the same genomic region millions of times, to make sure they actually knocked in the 4,000 variants they wanted to study. They sequenced at two time points, and deduced that the knock-ins that didn’t come up in the sequencing at the second time point were ones that interfered with the gene’s function, because the cells carrying them must have died.

            Try It: Findlay’s team had the DNA oligos for the 4,000 variants manufactured for them on a microarray. You can buy arrays of 6,000 to 250,000 oligos, so consider getting more bang for your buck by combining multiple experiments on the same array, says Findlay. Their lab pays about $5,000 for 100,000 oligos.

            This strategy comes with limitations: it has so far only been used to knock in single-nucleotide variants, and all the edits need to be in the same gene. The method works best when editing a fairly narrow region of DNA, about 110–120 base pairs, because longer DNA oligos would have too many errors, Findlay says. It’s also important to sequence very deeply to make sure that you account for the full number of variants you intended to knock in.


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