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What genes are required to make E. coli photosynthetic?

What genes are required to make E. coli photosynthetic?



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"All" of the genes for bacterial photosynthesis were discovered in a gene cluster almost 40 years ago. Marrs, J.Bact. What more is needed to make E. coli photosynthetic?


I think the crux of the answer is only hinted at in WYSIWYG's answer. The crux of photosynthesis is partitioning coupled reactions.

Photosynthesizing plant cells have sub-cellular structures called chloroplasts; their membranes are especially important for the compartmentalization of different functions and chemical processes for the entirety of photosynthesis to even be possible. Partitioning is difficult to achieve in E. coli, and that to me seems a necessity for achieving true heterologous, truly ectopic photosynthesis… Even photosynthesizing cyanobacteria (photosynthesizing prokaryotes) possess thylakoids, membrane-bound compartments where the magic happens. Photosynthesis though can certainly proceed and be made more efficient in single cells (example).

That leaves us with a simple answer: engineering E. coli with thylakoid-like structures is not really practicable. Why not simple use cyanobacteria instead?

Further reading: review on single cell photosynthesis in single cells.


A very basic kind of photosynthesis can be realized in E.coli by expressing a rhodopsin (Kim et al., 2017). Rhodopsins are membrane proteins that can pump ions (including protons) across the membrane and generate a transmembrane potential. When coupled to ATP synthase, rhodopsins can cause ATP generation in the presence of light. In an earlier work, Hara et al., (2013) had expressed a rhodopsin in mammalian mitichondria to make them synthesize ATP in response to light exposure.

This is not the kind of energy efficient photosynthesis that we see in cyanobacteria and plants. The group of Ron Milo have engineered Calvin-Benson cycle (carbon fixation) in E.coli, and optimized carbon fixation using adaptive laboratory evolution (Antonovsky et al., 2017; Gleizer et al., 2019).

Engineering the entire photosystem in E.coli would be a difficult task because a functional photosystem does not merely involve expression of the pigments but also their 3D assembly into functional sub-cellular "organelle" like structures.


16.2 Prokaryotic Gene Regulation

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

  • Describe the steps involved in prokaryotic gene regulation
  • Explain the roles of activators, inducers, and repressors in gene regulation

The DNA of prokaryotes is organized into a circular chromosome, supercoiled within the nucleoid region of the cell cytoplasm. Proteins that are needed for a specific function, or that are involved in the same biochemical pathway, are encoded together in blocks called operons . For example, all of the genes needed to use lactose as an energy source are coded next to each other in the lactose (or lac) operon, and transcribed into a single mRNA.

In prokaryotic cells, there are three types of regulatory molecules that can affect the expression of operons: repressors, activators, and inducers. Repressors and activators are proteins produced in the cell. Both repressors and activators regulate gene expression by binding to specific DNA sites adjacent to the genes they control. In general, activators bind to the promoter site, while repressors bind to operator regions. Repressors prevent transcription of a gene in response to an external stimulus, whereas activators increase the transcription of a gene in response to an external stimulus. Inducers are small molecules that may be produced by the cell or that are in the cell’s environment. Inducers either activate or repress transcription depending on the needs of the cell and the availability of substrate.

The trp Operon: A Repressible Operon

Bacteria such as Escherichia coli need amino acids to survive, and are able to synthesize many of them. Tryptophan is one such amino acid that E. coli can either ingest from the environment or synthesize using enzymes that are encoded by five genes. These five genes are next to each other in what is called the tryptophan (trp) operon (Figure 16.3). The genes are transcribed into a single mRNA, which is then translated to produce all five enzymes. If tryptophan is present in the environment, then E. coli does not need to synthesize it and the trp operon is switched off. However, when tryptophan availability is low, the switch controlling the operon is turned on, the mRNA is transcribed, the enzyme proteins are translated, and tryptophan is synthesized.

The trp operon includes three important regions: the coding region, the trp operator and the trp promoter. The coding region includes the genes for the five tryptophan biosynthesis enzymes. Just before the coding region is the transcriptional start site . The promoter sequence, to which RNA polymerase binds to initiate transcription, is before or “upstream” of the transcriptional start site. Between the promoter and the transcriptional start site is the operator region.

The trp operator contains the DNA code to which the trp repressor protein can bind. However, the repressor alone cannot bind to the operator. When tryptophan is present in the cell, two tryptophan molecules bind to the trp repressor, which changes the shape of the repressor protein to a form that can bind to the trp operator. Binding of the tryptophan–repressor complex at the operator physically prevents the RNA polymerase from binding to the promoter and transcribing the downstream genes.

When tryptophan is not present in the cell, the repressor by itself does not bind to the operator, the polymerase can transcribe the enzyme genes, and tryptophan is synthesized. Because the repressor protein actively binds to the operator to keep the genes turned off, the trp operon is said to be negatively regulated and the proteins that bind to the operator to silence trp expression are negative regulators .

Link to Learning

Watch this video to learn more about the trp operon.

Catabolite Activator Protein (CAP): A Transcriptional Activator

Just as the trp operon is negatively regulated by tryptophan molecules, there are proteins that bind to the promoter sequences that act as positive regulators to turn genes on and activate them. For example, when glucose is scarce, E. coli bacteria can turn to other sugar sources for fuel. To do this, new genes to process these alternate sugars must be transcribed. When glucose levels drop, cyclic AMP (cAMP) begins to accumulate in the cell. The cAMP molecule is a signaling molecule that is involved in glucose and energy metabolism in E. coli. Accumulating cAMP binds to the positive regulator catabolite activator protein (CAP) , a protein that binds to the promoters of operons which control the processing of alternative sugars. When cAMP binds to CAP, the complex then binds to the promoter region of the genes that are needed to use the alternate sugar sources (Figure 16.4). In these operons, a CAP-binding site is located upstream of the RNA-polymerase-binding site in the promoter. CAP binding stabilizes the binding of RNA polymerase to the promoter region and increases transcription of the associated protein-coding genes.

The lac Operon: An Inducible Operon

The third type of gene regulation in prokaryotic cells occurs through inducible operons, which have proteins that bind to activate or repress transcription depending on the local environment and the needs of the cell. The lac operon is a typical inducible operon. As mentioned previously, E. coli is able to use other sugars as energy sources when glucose concentrations are low. One such sugar source is lactose. The lac operon encodes the genes necessary to acquire and process the lactose from the local environment. The Z gene of the lac operon encodes beta-galactosidase, which breaks lactose down to glucose and galactose.

However, for the lac operon to be activated, two conditions must be met. First, the level of glucose must be very low or non-existent. Second, lactose must be present. Only when glucose is absent and lactose is present will the lac operon be transcribed (Figure 16.5). In the absence of glucose, the binding of the CAP protein makes transcription of the lac operon more effective. When lactose is present, its metabolite, allolactose, binds to the lac repressor and changes its shape so that it cannot bind to the lac operator to prevent transcription. This combination of conditions makes sense for the cell, because it would be energetically wasteful to synthesize the enzymes to process lactose if glucose was plentiful or lactose was not available. It should be mentioned that the lac operon is transcribed at a very low rate even when glucose is present and lactose absent.

Visual Connection

In E. coli, the trp operon is on by default, while the lac operon is off. Why do you think this is the case?

trp receptor is repressed. Lactose, a sugar found in milk, is not always available. It makes no sense to make the enzymes necessary to digest an energy source that is not available, so the lac operon is only turned on when lactose is present.

If glucose is present, then CAP fails to bind to the promoter sequence to activate transcription. If lactose is absent, then the repressor binds to the operator to prevent transcription. If either of these conditions is met, then transcription remains off. Only when glucose is absent and lactose is present is the lac operon transcribed (Table 16.2).

Link to Learning

Watch an animated tutorial about the workings of lac operon here.


Bond booster

Scientists in the Soviet Union were the first to discover Z-DNA, in the late 1970s, in a phage called S-2L, which infects photosynthetic bacteria 4 . They found that the phage DNA behaved oddly when its two helical strands were melted apart. The bond that forms between G and C bases breaks at a higher temperature, compared with that joining A and T, and the phage’s DNA behaved as if it was made primarily from G and C. But further analysis by the Soviet team showed that the phage had replaced A with Z, which formed a stronger bond with T.

“It looked like something transgressive,” says Philippe Marlière, an inventor and geneticist at the University of Evry, France, who led one of the Science studies. “Why did this phage have a special base like this?”

Scientists glimpse oddball microbe that could help explain rise of complex life

Follow-up studies showed that S-2L’s heartier genome was resistant to DNA-chomping enzymes and other anti-phage defences that bacteria wield. But researchers didn’t know how the Z-DNA system worked or whether it was common. Z-DNA is only one of a host of modifications known to exist in phage DNA.

To answer those questions, a team led by Marlière and Pierre-Alexandre Kaminski, a biochemist at the Pasteur Institute in Paris, sequenced the phage’s genome in the early 2000s. They found a gene that’s potentially involved in one step of making Z-DNA, but not in others. But the sequence had no matches in genomic databases at the time, and the team’s quest to understand the basis for Z-DNA hit a dead end.

Marlière and his colleagues patented the S-2L genome, but also made it public, and he continued to scour genomic databases. Finally, in 2015, the team got a hit: a phage that infects aquatic bacteria of the genus Vibrio harboured a gene that matched a stretch of S-2L’s genome. The gene encoded an enzyme that resembled one that bacteria use to make adenine. “It was an exhilarating moment,” says Marlière.

In 2019, Zhao’s team found similar database matches. Both teams showed that the phages all had a gene named PurZ. This codes for an enzyme that plays an early but crucial part in making the Z nucleotide from a precursor molecule that is present in bacterial cells. They then identified additional enzymes — encoded in the genomes of bacteria that the phages infect — that complete the pathway.

DNA's secret weapon against knots and tangles

But a key question lingered. The enzymes that the teams identified produced the raw ingredient for Z-DNA — a molecule called dZTP — but that didn’t explain how phages insert the molecule into DNA strands, while excluding A bases (in the form of a chemical called dATP).

Here, the teams’ conclusions differed slightly. Alongside PurZ in the Vibrio phage’s genome sits a gene that makes an enzyme called a polymerase, which copies DNA strands. Marlière and Kaminski found that the phage polymerase incorporates dZTP into DNA, while cutting out any A bases that were introduced. “This explained to us why A was excluded,” says Kaminski. “This was really spectacular.”

Zhao thinks this isn’t the whole story. Her work suggests that another phage enzyme is needed, one that breaks up dATP but preserves dZTP inside cells. Her team found that increasing dZTP levels relative to those of dATP was enough to trick a cell’s own polymerase into making Z-DNA.


Abstract

Bioremediation has the potential to restore contaminated environments inexpensively yet effectively, but a lack of information about the factors controlling the growth and metabolism of microorganisms in polluted environments often limits its implementation. However, rapid advances in the understanding of bioremediation are on the horizon. Researchers now have the ability to culture microorganisms that are important in bioremediation and can evaluate their physiology using a combination of genome-enabled experimental and modelling techniques. In addition, new environmental genomic techniques offer the possibility for similar studies on as-yet-uncultured organisms. Combining models that can predict the activity of microorganisms that are involved in bioremediation with existing geochemical and hydrological models should transform bioremediation from a largely empirical practice into a science.


Selectable Marker Genes and Reporter Genes

The selectable marker genes are usually an integral part of plant transformation system. They are present in the vector along with the target gene. In a majority of cases, the selection is based on the survival of the transformed cells when grown on a medium containing a toxic substance (antibiotic, herbicide, antimetabolite). This is due to the fact that the selectable marker gene confers resistance to toxicity in the transformed cells, while the non- transformed cells get killed.

A large number of selectable marker genes are available and they are grouped into three categories— antibiotic resistance genes, antimetabolite marker genes, and herbicide resistance genes (Table 49.3).

(a) Antibiotic Resistance Genes:

In many plant transformation systems, antibiotic resistance genes (particularly of E. coli) are used as selectable markers. Despite the plants being eukaryotic in nature, antibiotics can effectively inhibit the protein biosynthesis in the cellular organelles, particularly in chloroplasts. Some of the antibiotic resistance selectable marker genes are briefly described.

Neomycin phosphotransferase II (npt II gene):

The most widely used selectable marker is npt II gene encoding the enzyme neomycin phospho­transferase II (NPT II). This marker gene confers resistance to the antibiotic kanamycin. The trans-formants and the plants derived from them can be checked by applying kanamycin solution and the resistant progeny can be selected.

Hygromycin phosphotransferase (hpt gene):

The antibiotic hygromycin is more toxic than neomycin and therefore can kill non-transformed plant cells much faster. Hygromycin phospho­transferase (hpt) gene thus provides resistance to transformed cells.

Aminoglycoside adenyltransferase (aadA gene):

Aminoglycoside 3′-adenyltransferase (aadA) gene confers resistance to transformed plant cells against the antibiotics streptomycin and spectionomycin.

(b) Antimetabolite Marker Genes:

Dihydrofolate reductase (dhfr gene):

The enzyme dihydrofolate reductase, produced by dhfr gene is inhibited by the antimetabolite methotrexate. A mutant dhfr gene in mouse that codes for this enzyme which has a low affinity to methotrexate has been identified. This dhfr gene fused with CaMV promoter results in a methotrexate resistant marker which can be used for the selection of transformed plants.

(c) Herbicide Resistance Markers:

Genes that confer resistance to herbicides are in use as markers for the selection of transgenic plants.

Phosphinothricin acetytransferase (pat/bar gene):

Bialophos, phosphinothricin and glufosinate are commonly used herbicides. The pat/bar genes code for phosphinothricin acetyltransferase which converts these herbicides into acetylated forms that are non-herbicidal. Thus, pat/bar genes confer resistance to the transformed plants.

Enolpyruvylshikimate phosphate synthase (epsps/aroA genes):

The herbicide glyphosate inhibits photosynthesis. It blocks the activity of enolpyruvylshikimate phosphate (EPSP) synthase, a key enzyme involved in the biosynthesis of phenylalanine, tyrosine and tryptophan. Mutant strains of Agrobacterium and Petunia hybrida that are resistant to glyphosate have been identified. The genes epsps/aroA confer resistance to transgenic plants which can be selected.

Bromoxynil nitrilase (bxn gene):

The herbicide bromoxynil inhibits photosynthesis (photosystem II). Bromoxynil nitrilase enzyme coded by the gene bxn inactivates this herbicide. The gene bxn can be successfully used as a selectable marker for the selection of transformed plants.

Production of Marker-Free Transgenic Plants:

There is a growing concern among the public regarding the use of antibiotic or herbicide resistance genes as selectable markers of plant transformation:

i. The products of some marker genes may be toxic or allergic.

ii. The antibiotic resistance might be transferred to pathogenic microorganisms in the soil.

iii. There is a possibility of creation of super weeds that are resistant to normally used herbicides.

iv. A transgenic plant with selectable marker genes cannot be transformed again by using the same selectable markers.

In light of the apprehensions listed above, the public is concerned about the safety of transgenic technology, particularly related to the selectable marker genes (antibiotic/herbicide resistance genes). There are fears about the safety of consumption of foodstuffs derived from genetically engineered plants. This is despite the fact that so far none of the marker genes have been shown to adversely affect human, animal or environmental safety.

Clean Gene Technology:

The process of developing transgenic plants without the presence of selectable marker genes or by use of more acceptable marker genes is regarded as clean gene technology. And this will result in the production of many marker-free transgenic plants that will be readily acceptable by the public. Some of the approaches for clean gene technology are given.

Avoiding selectable marker genes:

Theoretically, it is possible to totally avoid marker genes and introduce only the transgene of interest. The transformed paints can then be screened by an advanced technique like polymerase chain reaction and the desirable plants selected. This approach is not practicable due to cost factor.

Co-transformation with two DNAs:

The transgenic plants can be produced by employing two separate DNAs — one carrying the desired target gene and the other the marker gene. The transformed plants contain both the genes, but at different sites on the chromosomal DNA. Traditional breeding techniques (a few rounds) can be used to get rid of the transgenic plants with selectable markers.

Removal of selectable markers:

It is possible to selectively remove the selectable marker genes from the plant genome. For this purpose, site-specific recombinase systems are utilized. Several recombinase systems are in fact available which can be used to selectively excise the marker genes from the plant genome.

Cloning of selectable markers between transposable elements:

A selectable marker gene can be cloned between plant transposable elements (Ds elements) and then inserted. The selectable marker is planked by the sequences that increase the intra-chromosomal recombination. This results in the excision of the marker gene.

Type # II. Reporter Genes:

A reporter gene may be regarded as the test gene whose expression can be quantified. The plant transformation can be assessed by the expression of reporter genes (also called as screenable or scoreable genes). In general, an assay for the reporter gene is carried out by estimating the quantity of the protein it produces or the final products formed. A selected list of the reporter genes along with the detection assays is given in Table 49.4, some of the important ones are discussed below.

Opine synthase (ocs, nos genes):

The common opines present in T-DNA of Ti or Ri plasmids of Agrobacterium are octopine and nopaline, respectively produced by the synthase genes ocs and nos. The transformed status of the plant cells can be easily detected by the presence of these opines. Opines can be separated by electrophoresis and identified. Alternately, the enzyme activities responsible for the production of opines can also be assayed.

β-Glucuronidase (gusluidA gene):

β-Glucuronidase producing gene (gusluidA) is the most commonly used reporter gene in assessing plant transformation for the following reasons:

i. β-Glucuronidase assays are very sensitive.

ii. Quantitative estimation of the enzyme can be done by fluorometric method (using substrate 4-methylumbelliferryl P-D-glucuronide which is hydrolysed to 4-methylumbelliferone).

iii. Qualitative data on the enzyme can be obtained by histochemical means (enzyme localization can be detected by chromogenic substance such as substrate X-gluc).

iv. No need to extract and identify DNA.

Green fluorescent protein (gfp gene):

Green fluorescent protein (GFP), coded by gfp gene, is being widely used in recent years. In fact, in many instances, GFP has replaced GUS since assays of GFP are easier and non-destructive. Thus, screening of even the primary transplants can be done by GFP which is not possible with other reporter genes.

Gene for GFP has been isolated from jelly fish Aequorea victoria which is a luminescent organism. The original gfp gene has been significantly modified to make it more useful as a reporter gene. GFP emits fluorescence which can be detected under a fluorescent microscope.

Bacterial luciferase (luxA/luxB genes):

The bacterial luciferase genes (luxA and luxB) have originated from Vibrio harveyi. They can be detected in some plant transformation vectors. The detection assay of the enzyme is based on the principle of bioluminescence. Bacterial luciferase catalyses the oxidation of long-chain fatty aldehydes that results in the emission of light which can be measured.

Firefly luciferase (luc gene):

The enzyme firefly luciferase, encoded by the gene luc, catalyses the oxidation of D-luciferin (ATP dependent) which results in the emission of light that can be detected by sensitive luminometers. The firefly luciferase gene, however, is not widely used as a marker gene since the assay of the enzyme is rather cumbersome.

Chloramphenicol acetyl transferase (cat gene):

The cat gene producing chloramphenicol acetyl transferase (CAT) is a widely used reporter gene in mammalian cells. Due to the availability of GUS and GFP reporter systems for plant trans-formants, CAT is not commonly used. However, some workers continue to use CAT by a sensitive radioactive assay, for the detection of the reporter gene cat.


DNA Replication in Prokaryotes

The prokaryotic chromosome is a circular molecule with a less extensive coiling structure than eukaryotic chromosomes. The eukaryotic chromosome is linear and highly coiled around proteins. While there are many similarities in the DNA replication process, these structural differences necessitate some differences in the DNA replication process in these two life forms. DNA replication in prokaryotes has been extensively studied, so we will learn the basic process of prokaryotic DNA replication, then focus on the differences between prokaryotes and eukaryotes.

How does the replication machinery know where to start? It turns out that there are specific nucleotide sequences called origins of replication where replication begins. E. coli has a single origin of replication on its one chromosome, as do most prokaryotes (Figure 1). The origin of replication is approximately 245 base pairs long and is rich in AT sequences. This sequence of base pairs is recognized by certain proteins that bind to this site. An enzyme called helicase unwinds the DNA by breaking the hydrogen bonds between the nitrogenous base pairs. ATP hydrolysis is required for this process because it requires energy. As the DNA opens up, Y-shaped structures called replication forks are formed (Figure 1). Two replication forks are formed at the origin of replication and these get extended bi-directionally as replication proceeds. Single-strand binding proteins (Figure 2) coat the single strands of DNA near the replication fork to prevent the single-stranded DNA from winding back into a double helix.

Figure 1: DNA replication in prokaryotes, which have one circular chromosome.

The next important enzyme is DNA polymerase III, also known as DNA pol III, which adds nucleotides one by one to the growing DNA chain (Figure 2). The addition of nucleotides requires energy this energy is obtained from the nucleotides that have three phosphates attached to them. ATP structurally is an adenine nucleotide which has three phosphate groups attached breaking off the third phosphate releases energy. In addition to ATP, there are also TTP, CTP, and GTP. Each of these is made up of the corresponding nucleotide with three phosphates attached. When the bond between the phosphates is broken, the energy released is used to form the phosphodiester bond between the incoming nucleotide and the existing chain.

In prokaryotes, three main types of polymerases are known: DNA pol I, DNA pol II, and DNA pol III. DNA pol III is the enzyme required for DNA synthesis DNA pol I is used later in the process and DNA pol II is used primarily required for repair (this is another irritating example of naming that was done based on the order of discovery rather than an order that makes sense).

DNA polymerase is able to add nucleotides only in the 5′ to 3′ direction (a new DNA strand can be only extended in this direction). It requires a free 3′-OH group (located on the sugar) to which it can add the next nucleotide by forming a phosphodiester bond between the 3′-OH end and the 5′ phosphate of the next nucleotide. This essentially means that it cannot add nucleotides if a free 3′-OH group is not available. Then how does it add the first nucleotide? The problem is solved with the help of a primer that provides the free 3′-OH end. Another enzyme, RNA primase, synthesizes an RNA primer that is about five to ten nucleotides long and complementary to the DNA. RNA primase does not require a free 3′-OH group. Because this sequence primes the DNA synthesis, it is appropriately called the primer. DNA polymerase can now extend this RNA primer, adding nucleotides one by one that are complementary to the template strand (Figure 2).

Figure 2 A replication fork is formed when helicase separates the DNA strands at the origin of replication. The DNA tends to become more highly coiled ahead of the replication fork. Topoisomerase breaks and reforms DNA’s phosphate backbone ahead of the replication fork, thereby relieving the pressure that results from this supercoiling. Single-strand binding proteins bind to the single-stranded DNA to prevent the helix from re-forming. Primase synthesizes an RNA primer. DNA polymerase III uses this primer to synthesize the daughter DNA strand. On the leading strand, DNA is synthesized continuously, whereas on the lagging strand, DNA is synthesized in short stretches called Okazaki fragments. DNA polymerase I replaces the RNA primer with DNA. DNA ligase seals the gaps between the Okazaki fragments, joining the fragments into a single DNA molecule. (credit: modification of work by Mariana Ruiz Villareal)

The replication fork moves at the rate of 1000 nucleotides per second. DNA polymerase can only extend in the 5′ to 3′ direction, which poses a slight problem at the replication fork. As we know, the DNA double helix is anti-parallel that is, one strand is in the 5′ to 3′ direction and the other is oriented in the 3′ to 5′ direction. One strand, which is complementary to the 3′ to 5′ parental DNA strand, is synthesized continuously towards the replication fork because the polymerase can add nucleotides in this direction. This continuously synthesized strand is known as the leading strand. The other strand, complementary to the 5′ to 3′ parental DNA, is extended away from the replication fork, in small fragments known as Okazaki fragments, each requiring a primer to start the synthesis. Okazaki fragments are named after the Japanese scientist who first discovered them. The strand with the Okazaki fragments is known as the lagging strand.

The leading strand can be extended by one primer alone, whereas the lagging strand needs a new primer for each of the short Okazaki fragments. The overall direction of the lagging strand will be 3′ to 5′, and that of the leading strand 5′ to 3′. A protein called the sliding clamp holds the DNA polymerase in place as it continues to add nucleotides. The sliding clamp is a ring-shaped protein that binds to the DNA and holds the polymerase in place. Topoisomerase prevents the over-winding of the DNA double helix ahead of the replication fork as the DNA is opening up it does so by causing temporary nicks in the DNA helix and then resealing it. As synthesis proceeds, the RNA primers are replaced by DNA pol I, which breaks down the RNA and fills the gaps with DNA nucleotides. The nicks that remain between the newly synthesized DNA (that replaced the RNA primer) and the previously synthesized DNA are sealed by the enzyme DNA ligase that catalyzes the formation of phosphodiester linkage between the 3′-OH end of one nucleotide and the 5′ phosphate end of the other fragment.

(Lisa’s note: I think this process is almost impossible to visualize from reading text. I strongly recommend that you watch a couple of animations / videos like the one available here. There are additional links in Blackboard)

Once the chromosome has been completely replicated, the two DNA copies move into two different cells during cell division. The process of DNA replication can be summarized as follows:

  1. DNA unwinds at the origin of replication.
  2. Helicase opens up the DNA-forming replication forks these are extended in both directions.
  3. Single-strand binding proteins coat the DNA around the replication fork to prevent rewinding of the DNA.
  4. Topoisomerase binds at the region ahead of the replication fork to prevent supercoiling (over-winding).
  5. Primase synthesizes RNA primers complementary to the DNA strand.
  6. DNA polymerase III starts adding nucleotides to the 3′-OH (sugar) end of the primer.
  7. Elongation of both the lagging and the leading strand continues.
  8. RNA primers are removed and gaps are filled with DNA by DNA pol I.
  9. The gaps between the DNA fragments are sealed by DNA ligase.

Table 1: The enzymes involved in prokaryotic DNA replication and the functions of each.

Prokaryotic DNA Replication: Enzymes and Their Function
Enzyme/protein Specific Function
DNA pol I Exonuclease activity removes RNA primer and replaces with newly synthesized DNA
DNA pol II Repair function
DNA pol III Main enzyme that adds nucleotides in the 5′-3′ direction
Helicase Opens the DNA helix by breaking hydrogen bonds between the nitrogenous bases
Ligase Seals the gaps between the Okazaki fragments to create one continuous DNA strand
Primase Synthesizes RNA primers needed to start replication
Sliding Clamp Helps to hold the DNA polymerase in place when nucleotides are being added
Topoisomerase Helps relieve the stress on DNA when unwinding by causing breaks and then resealing the DNA
Single-strand binding proteins (SSB) Binds to single-stranded DNA to avoid DNA rewinding back.

DNA replication has been extremely well-studied in prokaryotes, primarily because of the small size of the genome and large number of variants available. Escherichia coli has 4.6 million base pairs in a single circular chromosome, and all of it gets replicated in approximately 42 minutes, starting from a single origin of replication and proceeding around the chromosome in both directions. This means that approximately 1000 nucleotides are added per second. The process is much more rapid than in eukaryotes.


Most humans have nearly the same complement of genes, all of which have come from our primate ancestors (Salzberg, 2017). On the other hand, even closely related strains of the bacterium Escherichia coli can differ by hundreds of genes (Touchon et al., 2009) despite having a much smaller genome. These genes have been acquired via a process called horizontal gene transfer (HGT), which is an important driver of adaptation, as it allows bacteria and other prokaryotes to gain the genes they need in order to thrive in certain environments (Koonin et al., 2001). Moreover, this exchanging of genes has resulted in many genetic elements in prokaryotes becoming highly mobile, making it easier for DNA to be transferred to a diverse range of hosts.

HGT has also been observed in animals, plants and other eukaryotes (Husnik and McCutcheon, 2018), but its role in determining genome composition and facilitating adaptation in these species remains unclear (Ku and Martin, 2016). Now, in eLife, Andreas Weber and co-workers at Heinrich Heine University, Arizona State University and Rutgers University – including Alessandro Rossoni as first author – report evidence for HGT between prokaryotes and the red alga Cyanidiales (Rossoni et al., 2019). These are remarkable single-cell organisms that can perform photosynthesis at temperatures up to 56°C, and can live in extreme environments such as hot springs and acid rivers (Schönknecht et al., 2013). Cyanidiales can also be used to investigate HGT over geological timescales because they share a common ancestor that dates back 800 million years to a time before animals had even evolved.

Based on an analysis of ten new and three previously reported Cyanidiales genomes, Rossoni et al. found that 1% of genes had been obtained via HGT. Moreover, many of these genes coded for proteins that were needed to survive in extreme environments (such as proteins involved in detoxifying heavy metals like arsenic or mercury, or removing free radicals Figure 1). Additionally, prokaryotes adapted to the same extreme environment as Cyanidiales were commonly identified as the source of these genes. It seems likely, therefore, that HGT influenced the evolution of Cyanidiales, especially because the criterion used to detect HGT was conservative and the study did not attempt to detect gene transfer from other eukaryotes.

Horizontal gene transfer in the evolution of red algae.

The evolutionary trajectory of the red algae Cyanidiales is shown from top to bottom. Rossini et al. investigated genetic changes that took place before and after the Cambrian explosion 541 million years ago, and found that Cyanidiales obtained 1% of their genes during this time by horizontal transfer. Many of these genes allowed Cyanidiales to adapt to extreme environments, such as genes related to the detoxification of heavy metals including mercury and arsenic (represented by green arrows). Some of the lineages of Cyanidiales that were sequenced by Rossoni et al. are shown in the bottom panels: two of these have the same taxonomic name despite having diverged from one another millions of years ago. Image credit: Andreas Weber (left panel), Debashish Bhattacharya (two middle panels), and Shin-ya Miyagishima (right panel).

Comparing the new Cyanidiales genes to genes found in present-day bacteria and archaea databases did not yield any recent examples of HGT. This absence of recent events is unsurprising, as Rossoni et al. estimated that Cyanidiales acquire just one gene via HGT every 14.6 million years – the same amount of time it took for humans to diverge from the orangutan. Such a low rate makes finding a fresh transfer in a small number of genomes unlikely. Instead, the majority of HGT candidate genes found by Rossoni et al. have acquired introns (non-protein coding segments of DNA), and then persisted over hundreds of millions of years.

Despite there being evidence to show HGT occurred, it still remains unclear how these transfers took place. The best-studied mechanisms by which eukaryotes acquire DNA from other organisms are sexual reproduction and by transferring DNA from symbionts (biological organisms that live cooperatively with other organisms). However, meiotic sex only occurs between closely related species, and therefore cannot explain how Cyanidiales appear to have gained DNA from such a diverse range of prokaryotes: moreover, the evolution of symbiotic transfer is uncommon in most taxonomic groups. Instead DNA was more likely obtained via viral infection or plasmids (circular molecules of double stranded DNA) being transferred between prokaryotes and eukaryotes (Heinemann and Sprague, 1989). Indeed, a recent study has shown that many eukaryotes, including red algae, can acquire plasmids carrying genes derived from plants, viruses and bacteria (Lee et al., 2016).

The work of Rossoni et al. suggests that, in terms of gene content evolution, Cyanidiales are more similar to humans than to E. coli, which is consistent with previous qualitive comparisons of HGT patterns in eukaryotes and prokaryotes (Ku and Martin, 2016). However, a number of mysteries still remain. For example, what are the most common modes of plasmid transmission in Cyanidiales? How do plasmids maintain themselves in populations? How often do they jump between species, and how far do they jump? To answer these questions we should first observe what is happening all around us today (Popa et al., 2017) and, if possible, study events that occur more frequently than once every 14.6 million years.


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Introduction

Bacteria and simple eukaryotic organisms that have adapted to anoxic or hypoxic conditions for all or part of their life-cycle employ a variety of metabolic strategies to cope with such environments [1,2]. One such strategy relies upon fumarate (E°′ = +30 mV) as an electron acceptor in a reversal of the succinate dehydrogenase (SDH) reaction of the citric acid cycle that comprises a fundamental component of aerobic metabolism. While ubiquinone (Coenzyme Q or Q, Fig 1, compound 1) is the electron acceptor in the SDH reaction, a quinone with a lower standard reduction potential is required to make fumarate reduction more favorable. Rhodoquinone (RQ) (Fig 1, compound 2) or menaquinone (MK) (Fig 1, compound 3) are naturally occurring compounds that meet this requirement [3].

The number of isoprene units (n) in the tail varies by species from 6–10. The reduction potentials of the quinones are as follows: Q, E°′ = +100 mV RQ, E°′ = −63 mV MK, E°′ = −80 mV.

The SDH reaction is catalyzed by the Complex II family of integral membrane, multisubunit enzymes. Two homologous forms of Complex II are recognized and characterized: succinate:ubiquinone reductase (SQR) and quinol:fumarate reductase (QFR), that are optimized to function in aerobic and anaerobic metabolism, respectively. In E. coli, the expression of these two Complex II homologs is adjusted in response to oxygen levels, with hypoxic and anoxic conditions promoting QFR predominance, along with a shift in composition of the quinone pool toward MK with which QFR functions most efficiently [4]. The change in Complex II homolog expression and the adjustment to a lower potential quinone is crucial to the thermodynamic and kinetic favorability of fumarate reduction. A similar scenario is played out in the parasitic nematode, Ascaris suum, when migration into a host confronts the organism with lowered oxygen levels. In response, the subunit composition of Complex II is altered in favor of fumarate reduction in this case, RQ is employed as the low potential electron carrier, and it becomes the predominant mitochondrial quinone component [5].

Since the discovery of RQ in the alphaproteobacterium Rhodospirillum rubrum [6], RQ has only been found in a limited number of bacterial and eukaryotic species. Although it fulfills an analogous role, RQ stands in marked contrast to MK in structure, biosynthesis, phylogenetic distribution and evolutionary origins [3]. The occurrence of MK, a naphthoquinone, among prokaryotic organisms is extremely broad, more so than Q (confined to alpha-, beta-, and gamma-proteobacteria), reflecting its earlier evolutionary origins in an anoxic environment. The much more restricted phylogenetic distribution of RQ, which like Q is a benzoquinone, points to a relatively recent origin of RQ from an augmentation of Q biosynthesis, possibly even later than the divergence of the eukaryotic lineage [7].

The study of RQ in R. rubrum has revealed further details about its biosynthetic origins and relationship to Q. As a photosynthetic facultative anaerobe, R. rubrum grows aerobically, utilizing Q in the dark, but also photoheterotrophically with light under anaerobic conditions. While in the latter case, R. rubrum would be expected to utilize RQ and QFR (as is the case in parasitic helminths and a few other eukaryotic species that have adapted to anoxic environments) [2,8], a recent study by Ghosh, et al. supports the conclusion that R. rubrum lacks an orthologous QFR, implying that SQR is capable of functioning in reverse with RQ as an electron donor [9]. A similar phenomenon has been reported in E. coli, where SQR can replace QFR to support anaerobic growth when fumarate is utilized as the terminal electron acceptor using the low potential carrier, MK [10]. We previously reported that Q is a required precursor for RQ biosynthesis in R. rubrum [11]. Investigation of a mutant strain incapable of anaerobic growth and devoid of RQ led to the identification of the rquA gene as the locus of this loss of function. Complementation of the mutant with an intact rquA gene restored both anaerobic growth and RQ production [12].

The phylogenetic distribution of rquA was analyzed in a recent report [13]. In addition to its sparse distribution across eukaryotes and bacteria adapted to hypoxia, this analysis supports the hypothesis that this pattern resulted from multiple lateral gene transfer events. Interestingly, this work highlights the fact that a subset of eukaryotic organisms (such as A. suum and Caenorhabditis elegans) known to produce RQ lack an rquA ortholog, suggesting that there are alternative biosynthetic routes to RQ. Nonetheless, it is clear that rquA performs a necessary role in R. rubrum (and presumably in all other organisms possessing rquA orthologs), a role we seek to further clarify in the present study. We report here the first transcriptome data obtained by RNA sequencing (RNAseq) of R. rubrum under aerobic and anaerobic conditions. This data, in conjunction with comparative genomic analysis, was used to evaluate putative gene candidates involved in RQ biosynthesis or its regulation. We then characterized several such candidates by generating knockouts in R. rubrum and assessing the effect on RQ levels and rquA expression.


It's not easy being green: Scientists grow understanding of how photosynthesis is regulated

Once the energy resources in a seed are depleted, seedlings switch on a "green" photosynthesis program ( Arabidopsis seedling at the top). Credit: Courtesy of Jesse Woodson, Salk Institute for Biological Studies

The seeds sprouting in your spring garden may still be struggling to reach the sun. If so, they are consuming a finite energy pack contained within each seed. Once those resources are depleted, the plant cell nucleus must be ready to switch on a "green" photosynthetic program. Researchers at the Salk Institute for Biological Studies recently showed a new way that those signals are relayed.

In a study published in the May 24, 2011, issue of the journal Current Biology, a team led by Joanne Chory, Ph.D., professor and director of the Plant Molecular and Cellular Biology Laboratory, and including postdoctoral fellows, Jesse Woodson, Ph.D., and Juan Perez-Ruiz, Ph.D., identify a signaling factor sent by plant chloroplasts to turn on photosynthesis-related genes. Their finding may help achieve greater crop yields and better plant health.

"When a seedling establishes a photosynthetic lifestyle, it needs to activate several thousand genes in the nucleus," says Chory, also a Howard Hughes Medical Institute investigator and holder of the Howard H. and Maryam R. Newman Chair in Plant Biology. "One of the signals to do this comes from the organelle in charge of photosynthesis, called the chloroplast. In this study we identified this signaling molecule as heme."

Although in plants and animals most genes reside in the nucleus, small DNA rings of genes are found in other cellular venues such as energy-producing mitochondria. Plant chloroplasts, whose primary function is to turn light and carbon dioxide into energy and carbohydrates required for growth, also contain genes that regulate photosynthesis-related factors encoded in the plant cell nucleus.

"The Chory lab previously identified mutations in five genes in Arabidopsis thaliana plants that were unable to synthesize molecules such as chlorophyll or respond to signals generated by intermediates of the chlorophyll biosynthetic pathway," explains Woodson, the study's first author. "Those studies suggested that when plants undergo stress, an intermediate accumulates that tells the nucleus to stop 'turning green.'"

When plants encounter stressful situations, such as the herbicide norflurazon, the genes that control photosynthesis are turned off leading to a bleached appearance ( Arabidopsis seedling at the bottom). Credit: Courtesy of Jesse Woodson, Salk Institute for Biological Studies

These mutants—called GUN (for genomes uncoupled) 1 to 5—lack proteins necessary to relay these signals to the nucleus. Chloroplasts in normal plants might deploy those signals when plants encounter stress, such as too much heat or too little water. Inhibitory signals could also be sent when germinating sprouts are not yet mature enough to make the leap from relying on the seed energy pack to generating their own energy using sunlight.

Suspecting that positive signals must also govern the process, the team screened Arabidopsis for factors that switched photosynthetic proteins on, rather than off. For that, they employed an experimental approach called activation tagging, in which gene-activating DNA sequences are inserted randomly into the Arabidopsis genome and plants are then subjected to a herbicide shower. The team then looked for any survivor that persisted in making photosynthetic proteins. By definition, in that mutant plant a gene required to sustain a photosynthetic growth response must have been experimentally switched on.

What they found was a gene designated gun 6, which encodes the enzyme ferrochelatase 1 (FC1), the first gun mutant indicating a positive rather than a negative signal. "Gun 6 mutants make too much FC1 protein, an enzyme required to make a signaling molecule called heme," explains Woodson. Although it is a cofactor in numerous plant and animal pathways, heme is most famous as the oxygen-carrying component of hemoglobin.

The study suggests that excess heme drives expression of photosynthesis-related genes. "If a plant makes abnormally high levels of heme, the nucleus could be unaware that the chloroplast is nonfunctional and may keep making growth-related proteins," says Woodson. "Heme is likely the signal sent from a healthy chloroplast to the nucleus saying it is time to make proteins required for photosynthesis."

The team also genetically engineered plants to make too much of an FC1 isoform, known as FC2, and found photosynthesis-related genes were not upregulated, suggesting that heme made from FC2 differs from that made by FC1, and that overall signals regulating plant growth are highly complex.

The mustard plant Arabidopsis is favored by plant biologists for the same reasons that animal biologists rely on mice and fruitflies—it's easy to grow, compact, reproduces rapidly (which for plant biologists means it makes tons of seeds—and fast), its genome is sequenced, and it can be manipulated genetically.

Plus, it is incredibly boring. "There is nothing unusual about Arabidopsis," notes Woodson. "Which is good, because anything we learn from it will be generally true for most plants."

Affirming that "every plant" quality, the team isolated the corn (Zea mays) equivalent of the FC1 gene and engineered Arabidopsis to make artificially high levels of the corn protein. Like gun 6 mutants, those plants continued to make photosynthesis-related genes when subjected to herbicide, demonstrating that FC1 likely does the same thing in a crop plant as it does in a weed. However, analysis of corn, rice or barley would be experimentally less practical, given their longer growth cycles and the space required to grow them.

"Overall, this work answers basic questions regarding how a plant grows, builds chloroplasts and harvests light energy in order to turn into a photosynthetic organism," says Chory. "Understanding how plants coordinate gene expression between the chloroplast and nucleus will ultimately increase crop yields in the field, where plants often encounter multiple stresses during the growing season."