Genetic modification: exploiting the potential of bacteria

Main source: J.W.Dale, Molecular genetics of bacteria, Wiley Verlag, 2004

Bacteria, especially E.Coli strains have been utilized for production of a wide range of chemical substances, including their natural products such as antibiotics or amino acids, as well as non-bacterial products such as human growth hormones and vaccines. Though natural wild-type strains are sometimes used, strains for industrial production are often optimized to minimize the production cost and  time. Apart from optimization of temperature, medium conditions, etc. , genetic modification is an important mean for strain development. In this review, the most essential aspects of bacterial genetic modification will be discussed.

1. Strain development is based on two processes: generation of variation and selection of variants with desirable properties, often via evolution.

Variation within a bacterial culture is continuously generated by means of mutations. This processes are normally slow, but they can be accelerated if mutagenic agents are applied to the culture. Ultraviolet irradiation is a mild, easy to control method and thus used widely to create mutation. Giving that the following screening methods are sufficiently powerful, few or knowledge of the genes is required. Another method is gene cloning, which inserts the gene of interest into a bacterial genome. This method often makes it easier for the next screening step but requires more knowledge of genes concerned.

Selection of bacterial strains that grow faster or survive selective conditions such as application of antibiotics, is fairly easy, but it is not always the case. Aside of fitness, other desired characteristics in strains used for industrial production are: high level of production of desired products, substrate utilization, absence of undesirable by-products, etc.

2. Metabolite overproduction

Very often, high level of production of desired products is the ultimate aim of strain optimization. It is usually to distinguish between overproduction of primary metabolites, which are formed as the part of normal growth and play certain roles in the metabolism, and overproduction of secondary metabolites, which are formed in stationary phase and don’t seem to be essential to the cell growth.

a) Overproduction of primary metabolites

– Simple pathways: Since the desired products are products of enzymatic reactions, their yield can be improved either by increasing the concentration of initial substrates, or by improving the enzyme activity. The former is often not the case, since bacteria has developed complex regulation machinery to control their metabolism and increasing the substrate concentration often fails to improve the yield, and in extreme cases, even lowering it. The latter is feasible, but such mutations that alter the enzyme structure and improve its activity are very rare. A third method (and often the most important one) targets the regulation machinery.

The biosynthesis is often regulated by the feedback mechanism, in which the accumulation of products inhibits their further production. Mutants which are deficient in the feedback regulation are therefore capable of overproduction. A widely used method to screen out such mutants is to use antimetabolites. Antimetabolites are structurally similar to the end-product of the pathway, but cannot be utilized by the cell. In normal cells, antimetabolites repress the production of essential products via feedback regulation and thus inhibit the cell growth. However, mutants with defective feedback machinery are not affected by antimetabolites.

– Branched pathways: the biosynthesis of many amino acids is often branched, which means two or more different products are derived from the same precursor. It is therefore expected that if the production of one amino acid is expressed, available materials will be diverted to the other branch, leading to the increasing in yield here. Another advantage is that feedback regulation requires sufficient concentration of both products and thus does not work if one product is not synthesized.

A typical example is the industrial production of the amino acid lysine with a strain of Corynebacterium glutamicum. Lysine is synthesized in a branched pathway, in which another amino acids: threonine, methionine and isoleucine are also produced. A branch point in the pathway is the formation of lysine from the precursor aspartic semialdehyde, which is converted into homoserine by the enzyme homoserine dehydrogenase in the other branch. Mutants that are defective in the enzyme homoserine dehydrogenase can only produced lysine. In addition, since feedback regulation requires accumulation of both serine and threonine, such mutants are not affected by the overproduction of lysine.

The antimetabolite for mutant selection is S-(2-aminoethyl)-L-cystein. Selected strains are cultivated in medium containing small amount of homoserine (or mixture of threonine and methionine), which is sufficient to maintain the cell growth but not enough to trigger the feedback regulation.

b) Overproduction of secondary metabolites

Compared to the synthesis pathways of primary metabolites, those of secondary metabolites are often more complex and diverse. Knowledge about the pathways and their regulation is not always readily available. Therefore, strain improvement is still largely empirical process, involving numerous repeated cycles of mutation generation and screening.

3. Gene libraries

a) Construction of gene libraries

Gene library contains many recombinant E.Coli clones, each carries a piece of DNA from organism of interest inserted into its genome. The first step in library construction is the random fragmentation of the DNA from organism of interest, resulting in a mixture of numerous DNA fragment. Next step is to clone all these fragments into a vector and use it for transformation of E.Coli.  Each individual clone carries a different piece of DNA, and the whole collection of clones constitutes the gene library.

b) Screening of gene libraries

Library construction is quite straightforward, but screening step to find the clones carrying genes of interest is challenging, giving the huge number of clones involved. Following screening methods have been developed:

– Screening with gene probes: The clones are put on a plate and incubated with a radiolabeled gene probe, which is complementary to a sequence within the gene of interest. The gene probe binds specifically to the clones carrying gene of interest, giving out its position. The sequence of gene probe is deduced either from the already known sequence of related gene or of the same gene from related organisms, or from back-translating the peptide sequence to DNA sequence.

– Screening with PCR: if the flanking sequences of the gene of interest are known, PCR can be used to specifically amplify it.

– Screening with antibodies: requires that the gene of interest must be expressed, the antibody should recognize the linear (denatured) form rather than the folded form, and it must recognize the direct product of translation rather than post-translationally modified one.

– Screening with complementation: clones containing genes conferring prototrophy, such as leucine-producing gene leu, can be discovered when cultivated in medium lacking needed substrate (e.g. leucine).

c) cDNA libraries are commonly used in eukaryotes. The reason is eukaryotic transcription mechanism differs significantly from that of bacteria, involving RNA splicing and many other post-transcriptional modifications. cDNA libraries are constructed from mature mRNA rather than from DNA. The central part of this process is the utilization of the enzyme named reverse transcriptase, which produces DNA (termed cDNA or complementary DNA) from a RNA template. cDNAs are next cloned into vector and used for E.Coli transformation.

Auxin: the growth hormone

Main source: Taiz, Zeiger, Plant physiology, Springer Verlag, 2007

Hormones are chemical substances that are synthesized in one cell or tissue but modulate the cellular processes in another cell or tissue by interacting with specific proteins called receptors. Plant development is regulated by six different families of hormones: auxins, gibberelins, cytokinins, ethylene, absciscic acids and brassinosteroids. Further hormones or hormone-like signalling agents which play roles in resistance to pathogens, defense agains herbivores… have been also identified. Two most important plant hormones are auxin and cytokinin. This review will focus on auxin, the first plant hormone to be discovered.

1. Discovery

The discovery of auxin traces back to the experiment of Charles Darwin and his son Francis on a well-known phenomenon in plants called phototropism: plants tend to grow toward the light. Coleoptile, a modified leaf that protects the emerging shoot in monocotyledons, was found very sensitive to blue light: a short pulse of dim blue light is sufficient to induce the phototropic bending.  Darwin covered the tip of the coleoptile with foil and found that it would not bend. The conclusion drawn out was that the region several millimeters below the tip, termed the growth zone, was responsible for the bending.

It was suggested that some sort of signalling agent was produced in the tip, transported to the growth zone. Unilateral light caused the concentration gradient, and the shaded side where the concentration was greater would grow faster than the illuminated side, causing the bending.

Early efforts to isolated the signalling agent from the tip were not successful, since destroying the cells released many inhibitive substances that were normally contained in cell compartments. In 1926, a major breakthrough was achieved by F.W.Went, as he used a gelatin block to absorb the hormone from the cleaved  tip.The block was then placed asymmetrically on top of a decapitated coleoptile. The hormone diffused back to one side, causing faster elongation there and eventually a bent shape. The mechanism proposed early by Darwin was thus confirmed. This hormone is name auxin.

There are many substances that have auxin activity, both natural and synthetic. By far, the most physiologically important and abundant auxin in higher plants is indole-3-acetic acid (IAA). Another popular ones are: indole-3-butyric acid (IBA), 2,4-dichlorophenoxyacetic acid (2,4-D), 2-methoxy-3-dichlorobenzoic acid (dicamba). The former is found in mustards and corn, while the two latter are synthetic auxin and used as herbicides in agriculture.

2. IAA biosynthesis and metabolism

As said above, auxin induces rapid cell growth. Thus, IAA biosynthesis is normally associated with rapidly dividing and growing tissues, especially shoots. Though virtually all plant cells are producing IAA at low level, shoot apical meristem and young leaves are major sites of IAA biosynthesis. Root apical meristem is also able to synthesize IAA, but much of its auxin comes from the shoot.

Apart from sites of biosynthesis, a significant proportion of auxin is also found in seed and storage organs such as cotyledons. In these sites, auxin is covalently bound to both high and low MW compounds and therefore considered hormonally inactive. Such forms of auxin are termed conjugated auxins. Auxin can thus either rapidly released or rapidly removal by enzymatic processes. Such conjugation / deconjugation processes provide a mean to control the level of free auxin. They are hence regulated by environmental stimuli such as light and gravity. Conjugation also protects auxin from oxidative degradation.

3. Auxin transport

Auxin is found unidirectionally transported from shoot (the primary source) to the root. Auxin is the only hormone that is transported that way. Such transport is termed polar transport. Polar transport is found in almost all plants. The auxin transport is gravity-independent but requires metabolic energy.

Auxin transport proceeds in cell-to-cell fashion: auxin escapes the cell through plasma membrane, diffuses across the middle lamella and enters the next cell also through its plasma membrane. The generally accepted model for auxin transport is the chemiosmotic model, in which auxin efflux is driven by membrane potential, while auxin influx is driven by proton motive force (PMF).

a) Auxin influx

Two mechanisms are proposed for the auxin influx. The first one is the passive diffusion of non-dissociated, hydrophilic form of auxin (IAAH) across the phospholipid bilayer. In the apoplast (free diffusional space outside the plasma membrane), pH is mantained from 5 to 5.5, which means about 25% of auxin is in protonated form IAAH and can diffuse readily through the lipid bilayer. Lowering pH in apoplast results in more IAA being uptaken into the cell.

The second mechanism is the active transport of dissociated form (IAA-) facilitated by a 2H+-IAA- symporter: two protons are imported together with IAA-, using PMF as driving force. A permease-type auxin uptake carrier AUX1, which is related to bacterial amino acid carrier,  is probably the auxin influx carrier. It is found in leaf vascular tissues and root apices of Arabidopsis and other plants. aux1 Mutants exhibit agravitropic growth (fail to sense gravity), which can be corrected by treatment with 1-naphtalyene acetic acid (1-NAA), a synthetic hydrophilic auxin.

b) Auxin efflux

In the cytosol, which has pH of about 7.2, all auxin is converted into dissociated form IAA-, which cannot diffuse across the membrane as readily as IAAH. As the result, IAA efflux is solely driven by the inside negative membrane potential.

A family of integral membrane proteins known as PIN proteins is found responsible for the auxin efflux. These proteins are concentrated on one end of the cell, defining the general direction of auxin transport. Different PIN family members mediate auxin efflux in different tissues. Another ATP-dependent mechanism is also involved: the P-glycoproteins (PGP) are capable of carrying hydrophoic IAA- anions out of the cell at cost of ATP, overcoming the effects of back diffusion. In contrast to PIN, PGP are uniformly distributed on the plasma membrane. PIN and PGP can function both independently or synergistically to catalyze auxin transport.

Some compounds have been found being capable of blocking the auxin transport by inhibiting either the auxin efflux (AEIs: auxin efflux inhibitors) or the auxin influx. The mechanisms of inhibition are: competing with auxin at the efflux carrier sites, binding to the regulatory sites of auxin efflux carriers or interfering with the protein trafficking.

c) Auxin transport in phloem

Auxin can also be transported via phloem. Transport in phloem is much faster than polar transport, because translocation in phloem is largely driven by source-sink force.

d) Auxin transport regulation

– Protein phosphorylation: mutations in genes encoding for protein kinases and phosphatases, or overexpression of flavonols (plant compounds that inhibits certain kinases and phosphatases) lead to decrease in auxin transport, which implies that phosphorylation is a key mechanism to control the auxin transport.

– Protein trafficking: Auxin polar transport requires the localization of both efflux facilitator PIN and influx carrier AUX1 proteins in specific sites in the plasma membrane. Studies reveal that it is achieved by an actin-dependent vesicle cycling between plasma membrane and an unidentified endosomal compartment. Treatment with brefeldin A  (BFA, a vesicle transport inhibitor) prevents the proper asymmetric localization of protein PIN1, which is restored after BFA is washed out.

4. Actions of auxin

a) Cell elongation

As said above, auxin promotes the cell elongation. This role is demonstrated in an experiment, in which the endogenous auxin source is removed as sections containing the elongation zones are cut out and cultivated in sucrose medium. As the result, the growth rate rapidly decreases to minimum rate. Growth rate is restored back to the level in intact plant when exogenous auxin is added into the medium. It is also found out that an optimal concentration of auxin is required for the maximal growth rate. As we shall see, that is the result of the auxin-induced biosynthesis of ethylene, which inhibits the cell elongation. On the other hand, studies reveal that roots require only a minimum concentration of auxin to growth, and larger auxin level, which normally promotes the elongation of stems and coleoptiles, inhibits the root growth.

In another experiment, a growing section is split lengthwise and incubated in buffer. In absence of auxin, two halves bend outward, indicating that inner tissues grow faster than outer tissues. However, if auxin is added into the buffer, two halves bend inward, suggesting that the primary target of auxin are the outer tissues. Studies in roots also confirm that root epidermal cells are principal target of auxin action.

To understand the mechanism, in which auxin promotes the cell elongation, it is advisable to take a look at the cell enlargement process in plant. It consists generally of three steps:

– Osmotic uptake of water across the plasma membrane

– Turgor pressure builds up

– Biochemical wall loosening occurs

Experiments have proven that auxin does not increase the turgor pressure when it stimulates growth. Therefore, it is generally accepted that auxin causes an increase in the wall extensibility with a mechanism similar to that of a phenomenon in plants known as acid growth (plant cells expand quicker at lower pH).

In this mechanism, auxin increases the rate of proton extrusion, lowering the pH. At acidic pH, the hydrogen bonds between polysaccharide components of the cell walls are weakened by the action of expansin, a protein family also found in the cell wall. Ultimately, it leads to the weakening of the cell wall and the increase of cell wall extensibility.

Current models for auxin-induced proton extrusion envolve activation of H+-ATPase genes, protein trafficking which increase the transport of H+-ATPase to the membrane and H+-ATPase stabilization.

b) Phototropism and Gravitropism

Auxin is also found regulating the orientation of the plant axis.  This process involves phototropism (growth in response to light), gravitropims (growth in response to gravity) and thigmotropism (growth around obstacles). Though few is known about the mechanism of thigmotropism, it is suggested that auxin gradients may be involved.

– Phototropism: the phototropic bending in response to the unilateral light is the result of the lateral redistribution of auxin, in which the shaded side receives more auxin and grows faster. The zone of photosensing is within the upper 5mm of the tip. The blue light stimulus is sensed by two proteins, phototropis 1 and 2. They are autophosphorylating protein kinases, whose activity is stimulated by blue light. The following blue light response ultimately leads to the lateral redistribution of auxin.

Experiments in Arabidopsis reveal that the protein PIN3 is responsible for the lateral auxin localization. PIN3 also destabilizes the basally-localized efflux complex characterized by PIN1, effectively inhibiting the auxin downward movement.

– Gravitropism: when dark-grown Avena (oat) seedlings are oriented horizontally, their coleoptiles will bend upward in order to grow vertically. This phenomenon can also be explaineb by an asymmetric distribution of auxin, in which the lower side receives more auxin and grows therefore faster. Gravity is the stimulus in this case.

A mechanism to explain how plant cells sense gravity is proposed: large, dense amyloplasts (starch-storing plastids) function as gravity sensors (referred as statoliths) in specialized gravity-sensing cells termed statocytes. How cells utilize statoliths to sense gravity is still poorly understood.

Gravity sensing triggers a signal transduction cascade, which may involve second messengers such as proton or calcium.

– Gravitropism in the root is achieved by auxin redistribution in the root cap.Auxin is synthesize in the shoot, transported to the root in the stele (central part of the root containing vascular tissues), distributed to all sides of the root cap and transported ultimately  to the elongation zone, where it regulates cell elongation. If the root is horizontally oriented, statoliths inside the root cap senses the gravity and trigger an asymmetric distribution of auxin. Since auxin is normally inhibitory to the root elongation, it is actually the side which receives less auxin that grows faster.

c) Developmental effects of auxin

– Apical dominance is the name for the phenomenon that the growing apical bud inhibits the growth of lateral (axillary) buds. Removal of apical bud results in the outgrowth of one or more of the lateral buds, but this effect can be repressed by IAA (applied in lanolin paste as carrier). In another experiment, if an auxin transport inhibitor, for example TIBA, is placed below the shoot apex, the inhibitory effect is repressed.

Studies also reveal that the auxin level doesn’t decrease after the decapitation of the shoot apex, contradicting the earlier proposed mechanism, in which the normal endogenous level of auxin promotes the outgrowth of apical bud but inhibits the outgrowth of axillary buds. Furthermore, experiments with radiolabelled auxin also show that auxin does not enter the bud.

Recent researches propose that apical dominance may be the result of interaction between auxin and the protein AXR1 (auxin resistance), which is related to the ubiquitin-activating enzyme and found exclusively in the xylem and sclerenchyma (supporting tissues in plants) cells. The research for a detailed mechanism of interaction is however still ongoing.

– Floral bud development and phyllotaxy (leaves arrangement)

Application of auxin transport inhibitors such as NPA leads to the abnormal development of leaves and flowers, suggesting that the polar auxin transport may play an important role in these processes. In another experiment, pin1 mutant of Arabidopsis fails to produce leaf primordia, since the PIN1 auxin transport protein is defective. However, by applying auxin in lanolin paste on a tiny spot on the flank of the shoot apical meristem, leaf primordium can be induced to grow.

– Formation of lateral and adventitious roots

Lateral roots extend horizontally and serve as plant’s anchor in soil. They are produced from sites above the elongation and root hair zones. Adventitious roots are derived from normally nonroot tissues. The production of adventitious roots is used in cutting, a very old but still useful vegetative propagation method.

The formation of both lateral and adventitious roots is induced by high auxin levels.

– Vascular differentiation

Auxin plays an important role in the formation of new vascular tissues. If an apical bud (site of auxin biosynthesis) is cut off and grafted on to a callus, xylem and phloem are formed beneath the graft. Vascular differentiation is also a part of the plant wounding response: auxin produced in the young leaves induces the regeneration of vascular tissues in the wounded site.

– Delaying the leaf abscission

In most plants, leaf abscission is preceeded by the formation of a region called abscission zone near the leaf basis. Within the abscission zone is the abscission layer, where the cell walls are digested during the leaf senescence, making it very soft and weak. Auxin also plays an important role in this process: in young leaves, auxin levels are high. In senescing leaves, auxin is no longer produced and auxin levels are relatively low, triggering the leaf abscission.

– Fruit development

5. Auxin signal transduction pathways

The principial auxin receptors are identified as F-box soluble proteins, which was first discovered as a subunit of a specific type of ubiquitin E3 ligase complex called SCF. Like other ubiquitin ligases, SCF complex facilitates the ATP-dependent ubiquitination of proteins targeted for proteolytic degradation. The primary function of F-box in SCF complex is to bind the substrates for ubiquitin-mediated proteolysis.

A F-box protein in Arabidopsis, named TIR1 (transport inhibitor respsonse 1), is essential for auxin-dependent hypocotyl elongation. TIR1 is a subunit of a ubiquitin ligase complex named SCF-TIR1, which plays a crucial role for auxin signalling in cells.

Two families of regulators participating in TIR1-pathway have been identified: auxin response factors (ARFs) and AUX/IAA proteins. ARFs bind specifically to the TGTCTC sequence in the promoter region of many auxin-response genes. In most cases, two ARF proteins dimerize on the DNA, causing a further stimulation (a process referred as potentiation). AUX/IAA proteins regulate the gene expression indirectly by binding to ARF protein bound to DNA, thus reverting its effect. If ARF is a repressor, AUX/IAA will activate the gene expression and vice versa.

AUX/IAA proteins  are targets for SCF-TIR1, a process requiring auxin.  The binding of auxin to TIR1 enhances not only its affinity to AUX/IAA but also the E3 ligase activity. As the result, AUX/IAA proteins are ubiquitinated and targeted for proteolytic degradation at proteasome.

In several very rapid processes such as the increasing in activity of plasma membrane H+-ATPase, the above mentioned mechanism involving transcriptional regulation may be too slow. Instead, other auxin-binding receptor proteins are involved, which work independently of the SCR-TIR1 pathways.

Bacterial genome plasticity: movable genes and phase variation

Main source: J.W.Dale, Molecular genetics of bacteria, Wiley Verlag, 2004

Mutations and recombination with foreign DNA are not the only mechanisms, with which bacterial DNA can be changed. In fact, the presence of transposable elements inside the genome offers a much larger range of possible alterations including insertions, deletions, transpositions and inversions. This plasticity generates a high level of diversity and enables bacteria to survive in many hostile and ever-changing environments.

1. Mobile genetic elements are genetic element that are sometimes inserted into a gene and therefore responsible for the gene inactivation. In contrast to plasmids or phage DNA, some don’t have an independent existence, but exist as a part of another DNA molecule. The most simple elements among them are the insertion sequences (IS). Many ISs have been already identified, which differ in size and other details, but share a similar overall structure: they consist of a central region encoding for a transposase, which is required for the transposition of IS from one site to the other, flanked by two almost perfect inverted repeats (IR).

Further sequence inspection of DNA region containing IS reveals the presence of short direct repeats, which are not part of IS but derived from the duplication of target sequence (the detailed mechanism will be discussed later).

ISs are found in almost all bacterial genera, though the presence as well as the number of copies vary from strains to strains. This phenomenon is termed restriction fragment length polymorphism (RFLP), which can be detected by Southern blot.

Regions containing IS are hot spots for homologous recombination. ISs therefore contribute significantly to the bacterial genetic diversity. ISs are also considered parasitic to the bacteria, since they only carry transposase gene and no gene that brings any benefit to the host.

Transposon was discovered in a search for the mechanism of the dissemination of the antibiotic-resistance genes among bacteria. These genes are found in plasmids, and it has been found that the incubation of two plasmids containing two different antibiotic-resistance genes results in a new plasmid that carries both of them.It is suggested that a mobile genetic element carrying antibiotic-resistance gene, termed transposon, is the cause of this phenomenon.

Transposons also consist of a central regions containing several genes, among them a transposase, an identifiable genetic marker (e.g. antibiotic-resistance gene) and other genes required for promotion and regulation of the transposition. The central region is flanked by two inverted repeats. Like the case of IS, directed repeats are also found on the both ends of the transposon. The more complex transposon, termed composite transposons, have their central regions flanked by two insertion sequence, either as direct repeats or inverted repeats. Transposition behaviors of such composite transposons tends to be complicated, since either only a part or the whole transposons can be transposed. Composite transposons are often termed class I transposons, non-composite transposons are referred as class II.

Integrons are big and complex transposons, derived from the insertion of many genes into an existing transposon. The gene set, termed gene cassette consists of many genes and their recombination sites and . Interestingly, gene cassette normally doesn’t contain promoter. There is however a promoter region within the integron itself.

2. Mechanism of transposition

a) Replicative transposition is a transposition mechanism, in which a copy of the transposed element (either IS or transposon) is inserted at a different site, while the original copy is retained. The target site depends on type of transposons, ranging from more or less random to an unique site in the whole genome.

The first stage is the cointegration of two plasmids: the donor plasmid A containing the transposon and the recipient plasmid B that doesn’t have it. This process is basically a fusion of  two plasmid into a bigger circular DNA, which have two copies of the transposon in the same orientation. The details mechanism is as follows: both strands of the plasmidA are nicked at the 3′-end of its transposon, both strands of the plasmid B are also nicked at the 3′-end of its target sequence. The nicked strands are now crossed over and rejoined in a manner that both target sequence and transposon are found in the new strand. DNA synthesis seals the remaining gaps. Under the consideration that both plasmids are circular, the net result is a bigger circular DNA molecule that has two copies of transposon (and also two copy of the target sequence) in the same orientation. The cointegration is facilitated by the transposon-encoded transposase.

The second step is the recombination, resulting in the resolution of the big DNA molecule, returning plasmid A to its native form and releasing plasmid B with a copy of transposon flanked by two direct repats, which are two copies of the original target sequence. The recombination is ususally the homologous recombination between two transposon copies, facilitated by host recombination machinery. An exception is the case of the transposon Tn3, where it is a site-specific recombination between two sites within the transposon, facilitated by the transposon-encoded protein TnpR.

b) Non-replicative (conservative) transposition

Replicative transposition is not the only mechanism of transposition. Some transposons and insertion elements exhibit another mechanism called non-replicative or conservative transposition. In this mechanism, while the recipient plasmid B is still cut at both 3′-ends of the target sequence, the donor plasmid is cut at both 5′- and 3′-ends of the transposon, releasing the transposon which is rejoined to the recipient plasmid. DNA repair seals the gaps, while the donor plasmid degrades.

3. Regulation of transposition

Excessive level of transposition is harmful to the host cell since it causes many gene inactivation through insertion mutation. There are some mechanisms to contain the transposition. One is to suppress the expression of the transposase gene. Another less conventional one is that a key protein for the transposition is translated from two different reading frames on the mRNA, therefore a one-base frameshifting is required so mRNA is accurately translated. Since frameshifting occurs only occasionally, only a small amount of produced protein is functional, which helps contain the transposition.

4. Exceptions

– Gene activation by transposable elements: so far, transposable elements are shown being able to inactivate a gene by being inserted within it. With some transposable elements, it is the converse effect that is the case: gene is activated by insertion of a transposable element to an adjacent site. It is explained that the inserted DNA provides the promoter that drives the gene expression.

– Mu, a transposable bacteriophage: Mu, standing for mutator, is a bacteriophage that causes an abnormal rate of mutation among E.Coli cells upon infection. The reason lies on the Mu DNA, a 38kb linear structure that acts as a transposon after being inserted (more or less randomly) into the host chromosome.

– Conjugative transposons are transposons that are able to be transferred from one cell to the other by means of conjugation. The mechanism differs significantly to the ones of transposition, involving the excision of transposon as a small circular DNA molecule.

5. Phase variation

In addition to the regulatory mechanisms in transcription and translation, the presence of transposable elements offers further mechanisms to turn genes on/off. In a bacterial population, ON- and OFF-versions of genes co-exist, ensuring that at least some bacteria have the needed genes to survive the environmental changes. Furthermore, it is also the mechanism to maintain the genetic characteristics that are under normal circumstances disadvantageous, but may be offer advantages under other conditions.  In bacteria, the genetic diversity caused by reversible but inherited genetic changes is termed phase variation. Some examples of phase variation in bacteria will be discussed below.

a) Variation mediated by simple DNA inversion

One of the most simple phase variations is the system controlling the expression of type 1 fimbriae in E.Coli, which is the protein that facilitates the adhesion to the host cell, but also the target for the host immune system. The protein is encoded in fimA gene, whose expression is driven by a promoter. The promoter is located in an upstream 314-bp region flanked by 9-bp inverted repeats, practically an insertion sequence. However, the integrases are not encoded within this region, but in other genes: fimB and fimE.  FimB and FimE facilitate the gene inversion, which flips the promoter over and again. In ON-state, the promoter is directed towards fimA and can properly initiate its transcription In OFF-state, the promoter faces in the  wrong orientation and gene expression doesn’t take place.

Most species of Salmonella apply same strategy, albeit more complex.Salmonella produces two types of flagellin protein (termed H-antigens): H1 and H2, which are encoded in two genes: H1 and H2. Between these two genes is a 996-bp region of DNA, containing the promoter and gene for a site-specific recombinase Hin (short of H inversion), and flanked by two 14-bp inverted repeats. Hin promotes the gene inversion, flipping the promoter over and hence turning either H1 or H2 on. The specificity is further improved by another protein rh1, which is downstream of H2 and thus can be expressed from the same promoter as H2. Rh1 inhibits the expression of H1.

b) Variation mediated by nested DNA inversion

Campylobacter fetus is an important Gram-negative pathogen in farm animals. It produces a surface layer (S-layer) of protein SapA, which protects the bacteria from host immune system. However, SapA is also antigenic and provides target for antibody attack. To cope with this problem, Campylobacter fetus possesses up to nine different variants of SapA, which are antigenically distinct and encoded in nine different genes. Each cell expresses only one gene, while the others remain silent due to the lack of promoter. The strategy is: two adjacent genes are always in opposite directions, and recombination between two consevered regions causes gene inversion, bringing the promoter to the 5′ upstream region of the next gene.

c) Antigenic variation in the gonococcus

The Gram-negative bacterium, Neisseria gonorrhoeae, often known as gonococcus, is the pathogen of gonorrhoea (a STD). This bacterium also produces pili, enabling it to attach to the host mucosal surface during the early stage of infection. However, pili are also targets for host immune attack. The bacterium evades this by antigenic variation: they produce a large variety of pili proteins, which share about 50 common amino acids at N-terminus , but the other about 100 varies significantly.

As in the case of Campylobacter fetus, the pili proteins are encoded in many genes. Silent copies of those genes are maintained in region known as pilS where they are not expressed due to the lack of promoter, while only one gene at an expression locus, pilE, where it is expressed. Genes are swapped between two regions by means of recombination.

d) Phase variation by slipped strand pairing

Sequences containing multiple repeats of one (e.g. AAAAAA) or more than one nucleotides (e.g. ATATATAT) are susceptible to deletion mutation during the replication, resulting in the lost of one or several repeats. If the deletion is within the coding sequence, the result can be premature termination during translation and the newly formed protein is defective. If the deletion is within the regulatory sequence, the effect can be not only switching gene on and off, but also the change in level of expression.

e) Phase variation mediated by differential DNA methylation

This is the case of the gene agn43, encoding for the membrane protein antigen 43 in E.Coli, whose roles are still unclear. The regulator, OxyR, binds to an upstream sequence containing 3 GATC repeats, which can be methylated on the N-6 position of adenine by the enzyme deoxyadenosine methylase (DAM). Normally, OxyR binds to DNA and inhibits the gene expression. However, if all three GATC sites are methylated, OxyR is no longer able to bind to DNA, and agn43 is expressed.

Phytochrome and light control of plant development

Main source: Taiz, Zeiger, Plant physiology, Springer Verlag, 2007

Seedlings, such as of corn or bean, burried in the soil have to grow in the dark, where they undergo a special kind of development termed skotomorphogenesis (skotos means darkness in Greek). The seedlings growing in the dark show symptom of etiolation (i.e. pale): elongation of the stems, folded cotyledons (failure of the leaves to unroll), absence of greening (due to the inability to accumulate chlorophyll), the presence of the apical hook… When exposed to the light, the seedlings enter the next stage of development: the photomorphogenesis. The transition is brief but extremely complicated. Among pigments that induce this transition, the most important ones are those that absorb red light and blue light. While the blue light responses will be discussed later, this review is focused on the red light absorbing protein termed phytochrome and their mechanism of action.

1. Phytochrome is a blue (hence red light absorbing) protein pigment with a MW of about 125kDa. Phytochrome is responsible for many red light induced morphogenic responses, especially during the seed germination. Its important roles in plant development had been well identified long before the protein itself was isolated and characterized.

One of the most prominent phytochrome-involving phenomenon is the antagonistic relationship between responses to red (650-680nm) and to far-red (710-740) irradiation. This phenomenon is observed in many processes: stem and leaf growth, floral induction and particularly, seed germination. Experiments on lettuce (an vegetable often used in salad) show that red light induces the seed germination, but a successive treatment with far-red light inhibits this effect. A further second treatment with red light restores the ability to germinate, which is again repressed by the final far-red light exposure. The conclusion drawn out of this experiment is: phytochrome exists in two interconvertible forms: a red light-absorbing form and a far-red light-absorbing form. These two forms act antagonistically in the regulation of seed germination. Later studies confirm this conclusion.

In dark-grown or etiolated plants, these two forms are termed Pr and Pfr. Red light turns Pr into Pfr, while far-red light turns Pfr back into Pr.  Since their absorption spectra overlap, Pr is never 100% converted into Pfr and vice versa. Instead, they co-exist in an equilibrium termed photostationary state. Another thing that need to be mentioned is that both forms also absorp blue light (though not as much as red light), hence the phytochrome effect (the interconversion between Pr and Pfr) can also be elicited (induced) by blue light.

There are a lot of phytochrome-induced responses, which differ significantly in many aspects. Some aspects are used to group phytochrome-induced responses into sub-types:

-Lag time (time between stimulation and response) ranges from a few minutes to weeks.

-Amount of light requires (referred as fluence, the number of photons impinging on a surface area): based on this aspect, responses are grouped in three types: very low fluence responses (VLFR), low fluence responses (LFR) and high irradiance responses (HIR).

In the VLFR, the amount of Pfr needed for induction is very small, often well below the already existing amount of Pfr in equilibrium. As the result, VLFR cannot be inhibited by far-red light (in other words, VLFR is not FR-reversible).

Both VLFR and LFR are promoted if the total fluence meets the requirement. Since fluence is the product of fluence rate and time of irradiation, VLFR and LFR can be induced either by brief exposure to the light source with sufficient brightness, or by very dim light with sufficiently long time of irradiation. They are said to obey the law of reciprocity.

In contrast, HIR can only be induced by continuous exposure to light of realitvely high irradiance. HIR don’t obey the law of reciprocity, they are not photoreversible either. HIR is not triggered by the interconversion between Pr and Pfr, but rather by the photoequilibrium between them.

It should be mentioned that the same effect can be either LFR or HIR, depending on the exposure time. For example, the anthocyanin in seedlings of white mustard is a LFR, induced by red light and can be reversed by far-red light. However, if the seedlings are exposed to high irradiance light for several hours, the effect is now HIR and thus no longer photoreversible.

2. Phytochrome structure

a) Structure: Native phytochrome is a soluble protein with a MW of about 250kDa. It exists as a dimer, consisting of two subunits, each has a light absorbing pigment molecule called chromophore and a long peptide chain called apoprotein. In higheer plants, the chromophore of phytochrome is a linear tetrapyrole termed phytochromobilin. Phytochromobilin alone cannot absorp red or far-red light, the ability of light absorption exists only it is covalently linked to the peptide chain. Phytochromobilin is biosynthesized inside plastids in a pathway which branches from the biosynthesis of chlorophyll. Attachment of phytochromobilin to the apoprotein takes place in the cytosol, the process is autocatalytic and no further protein nor co-factor is required.

The difference between Pr and Pfr is the conformation at C15 of the phytochromobilin: Pr is a cis-isomer while Pfr is a trans-isomer. Light induces the conformation change and thus the interconverstion between two forms.

Each phytochrome monomer consists of two halves, linked by a hinge region. The N-terminal half contains the chromophore-binding bilin lyase domain and the PHY domain, which stabilizes the Pfr form. Downstream in the C-terminal half are two PAS domains, which mediate the phytochromed dimerization. They also contain nuclear localization sequences (NLS), which (when exposed) direct the active Pfr form to the nucleus. At the C-terminus is a serine/threonine kinase domain, which is related to the ligh-regulated histidine kinase domains found in some bacteria. Phytochrome is found capable of autophosphorylation (i.e. the kinase domain phosphorylates the serine side chain in another domain, which in turn phosphorylates other proteins).

b) Genetics

Phytochrome apoproteins are encoded by a family consisted of five structurally related genes: phyA, phyB, phyC, phyD and phyE, of the most importance are phyA and phyB. phyA encodes for the photoreceptor involved in the perception of continuous far-red light. Responses involving phyA are: VLFR germination in response to brief pulse of light, FR-HIR hypocotyl (shoot region in seedling, between the cotyledon and the radicle) elongation.  phyB mediates responses to continuous red or white light. It is involved in R-HIR/LFR hypocotyl elongation, R-HIR/LFR flowering and LFR seed germination.

phyA codes for phytochrome type I, which is light-labile and mostly found in etiolated tissues. phyB, phyC, phyD and phyE code for type II light-stable phytochrome.

c) Mechanism of action

The phytochrome holoprotein is assembled and dimerizes in the inactive Pr form. Red light exposure leads to the conformation change not only at C15 of the chromophore, but also in the hinge region, converting Pr into the active form Pfr. As the result, the NLSs are now exposed, which directs the active Pfr to nucleus, where it changes the gene expression and induces the photomorphogenesis. A small pool of Pfr stays in cytosol, where it mediates the rapid responses by affecting the ion fluxes.

3. Circadian rhythms

It is a well known fact that plants change many of their metabolic processes in accordance to the day-night change, with a regular periodicity of about 24 hours. These rythmic changes are termed circadian rythms. Circadian rythms are considered endogenous, since they persist even in absence of external factors. Light condition is the cause of this phenomenon.

The circadian rhythms are governed by an internal mechanism called the oscillator, which is composed of several clock genes. Three major clock genes have been identified in Arabidopsis: TOC1, LHY and CCA1. The mechanism of action is proposed as follows:

At dawn, light and the TOC1 (which is accumulated during the night, as seen below), activate the expression of LHY and CCA1, which in turn activate other morning genes. LHY and CCA1 also inhibit the expression of TOC1 and other evening genes, which results in the reduction in the levels of all three genes (since LHY and CCA1 require TOC1 for their expression) as the day progresses. At the end of the day, LHY and CCA1 reach their minimum level and can no longer inhibit TOC1 and other evening genes. The genes are expressed during the night, and the cycle begins again as the first light of the next morning activates the expression of LHY and CCA1.

Further proteins are involved in this mechanism. For example, a F-box protein ZTL facilitates the ubiquitination of TOC1, directing it to the 26S proteasome for degradation. The level of ZTL is highest at dusk and lowest at dawn, perhaps as the result of the negative regulation by blue light (ZTL is found structurally similar to phototropin, a blue light receptor).

4.Ecological functions

a)Sleep movement of the leaves

The sleep movement of the leaves (termed nyctinasty) is a well known phenomenon in many legumes (bean-family). The leaves are extened horizontally to face the light during the day, and fold together at night. This movement is caused by the change in turgor pressure in cells located on opposite sides of the pulvinus (the region that containing vascular tissues): the ventral motor cells and dorsal motor cells.

Red light stimulates the close movement: the phytochrome is turned into active form Pfr by red light, which interacts with the proton pumps and potassium channels, changing the ion fluxes and causing the change in turgor pressure.

b) Plant adaption to light quality changes

The change in light quality is characterized by the red light to far-red light ratio (R:FR ratio). This ratio is high during the day and low at night or in shade.

An important function of phytochrome is that it help plants to sense and avoid shading by other plants. This response, termed shade avoidance respsonse, is a response to R:RF ratio. Low R:RF ratio causes elongation.This correlation is easy to observe in sun plants but not as strong in shade plants.

Genetic analyses of Arabidopsis indicates that PhyB plays the predominant role in many shade avoidance responses, while PhA plays the antagonistic role. Transgenic plants in which PhyA is overexpressed, fail to display shade avoidance responses, allowing them to be planted in much higher density.

3.Phytochrome’s ecological functions

a)Sleep movement of the leaves

Plant growth and development

Main source: Taiz, Zeiger, Plant physiology, Springer Verlag, 2007

1.Ovierview of plant growth and development

Almost all land plants are characterized by their sedentary (i.e. the inability to move). This lack of mobility largely simplifies the plant structure, but also gives rise to many significant problems, in particular the need of adaption to local environment. To cope with this challenge, plants maintain a constant reservoir of undetermined cells, which allow the cell proliferation, differentiation and recruitment not only during embryogenesis, as in the case of animal, but also in vegetative development. The structures responsible for these functions are called meristems.

In all seed plants, the life cycle, termed sporophytic development, is divided into three stages: embryogenesis, vegetative development and reproductive development.

– Embryogenesis is the process, by which a single cell is transformed into a multicellular entity having a characteristic but typically rudimentary organization.It takes place normally inside the ovule of the flower.

The ovule is also the site where the development of female gametophyte (haploid phase in early plant development) takes place. The female gametophyte, also known as embryo sac, derived from several rounds of mitotic divisions, consists of only a small number of cells, including the  egg cells and two polar nuclei. Fertilization (syngamy) occurs when the (haploid) pollen sperm cell meets and fuses with the egg cell to create the diploid zygote. Meanwhile, the fusing of another sperm cell with two (also haploid) polar nuclei gives rise to the formation of the triploid endosperm, which supports the embryo development. Together, these two coordinated syngamy events are termed double fertilization.

The next step is the seed formation, which involves cell proliferation and differentiation, establishment of polarity and many physiological changes to protect the seed from harsh environmental conditions.

– Vegetative development begins with the germination. The embryo breaks it dormant state (dormancy is the state where all metabolic activities are kept minimal) and mobilizes its stored reserves to build on its rudimentary form, through activities of root and shoot apical meristems. Photomorphogenesis and shoot development make the seedling photosynthesis competent, which enables further vegetative growth. In contrast to the animals, where it is predeterminate, the vegetative growth in plants is indeterminate.

– Reproductive development: after a period of vegetative development, plants enter the reproductive development stage. In flowering plants, it involves the formation of specialized floral meristems, which eventually give rise to flowers.

2. Embryogenesis is the process that transforms a single cell into a considerably more complex structure, the mature seed. It can be further divided into following sub-processes:

– Morphogenesis: the elaboration of basic forms

– Organogenesis: the associated formation of functionally organized structures

– Histogenesis: the cell differentiation inside various tissues.

An essential feature of embryogenesis is the present of meristems at both shoot and root axes, the so-called shoot and root apical meristems. The embryogenesis also involves the development of mechanisms to withstand the long period of dormancy and to recognize and response to germination signals.

a) Embryogenesis pattern

– In dicots: (Example of Arabidopsis) Embryogenesis consists of five stages:

1) Zygotic stage: single-cell stage derived the fusion of haploid egg and sperm, followed by the first asymmetric zygotic division.

2) Globular stage: The first zygotic division gives rise to the apical and the basal daughter cell. The apical embryo undergoes many rounds of division to form a globular structure called protoderm.

3) Heart stage: The rapid cell divisions on two side of the future shoot apex (tips) form a heart-shaped structure.

4) Torpedo stage: is resulted from cell elongations throughout the embryo axis and the further development of dicotyledons.

5) Mature stage: Toward the end of embryogenesis, the embryo loses water and becomes  metabolically inactive (dormant). Stored compounds needed for germination are also accumulated in this stage.

– In monocots (example of rice): embryogenesis also consists of five stages but in a quite distinct pattern:

1) Zygotic stage: following the fusion of haploid egg and sperm

2) Globular stage: takes place 2-4 days after pollination (DAP), involving the initial horizontal division to create the apical and basal cells, and series of variable cell divisions to form a globular structure.

3) Coleoptile stage:  takes place 5 DAP, involving the formation of coleoptile (specialized tubular first leaf), shoot and root apical meristems and radicle (embryonic root).

4) Juvenile vegetative stage: takes place 6-10 DAP, as the shoot apical meristem initiates several vegetative leaves.

5) Maturation: takes place 11-20 DAP

Both embryogenesis patterns share some common superficial features, most particularly these related to polarity. Beginning from a unicellular zygote, the embryo is progressively polarized along two axes: the apical-basal axe and the radical axe. In the following sections, the basic pattern of these polarizations as well as their regulation in Arabidopsis will be discussed in details.

b) Axial polarity: is the arrangement of tissues and organs along a linear axis stretched from one end, the shoot apical meristems (SAM) to the other one, the root apical meristem (RAM). This asymmetry can be traced back to the first, asymmetric divison of the zygote into two daughter cells, a bigger basal cell inheriting the large zygotic vacuole, and the smaller apical cell which is densely cytoplasmic.

Nearly the entire embryo is initiated from the smaller apical cell. Two vertical divisions and one horizontal division of apical cell give rise to a eight-celled structure called (octant) globular proembryo, which after further divisions becomes the protoderm.

The bigger basal cell also divides, but all of its divisions are horizontal, perpendicular to the shoot axis, forming a filamentous structure of six to nine cells called suspensor, which attaches the embryo to the vascular system of the mother plant. Only the uppermost basal cells, known as hypophysis,  contribute to the embryo. They will eventually become parts of the root called the quiescent center and the central root cap.

reproducible pattern during the early embryogenesis in Arabidopsis suggests that a fixed sequence of divisions is essential, or in other words, the fate of individual cells within embryo is determined. It is also found out that cells differentiate rather according to their position than to their clonal origin (in contrast to animal case). A position-dependent signalling mechanism is therefore suggested to explain this behavior. The basic points of this mechanisms are:

– There must be a coordinate system, by which unique positions within developing structure can be specified.

– Individual cell must have means to assess their positions within this coordinate system

– Cell must somehow respond to this information in an appropriate way

Spatial information is provided via a concentration gradient of chemicals termed morphogens. In plant, the morphogen that acts during the embryogenesis is hormone auxin.

Auxin is a class of plant growth hormones consisting of indole-3-acetic acid and its analogs. The distribution of auxin within the cell is studied either by measuring the promoter strength of auxin-inducible genes (which are fused with a reporter gene, gus for example) or by monitoring the distribution of PIN proteins which are responsible for the auxin intercellular movement (using immunolocalization). These approaches lead to the finding of auxin polar transport during the embryogenesis. The role of auxin is elucidated by looking at the examples of two genes: monopteros mp and gnom gn.

MP protein is necessary for the normal formation of basal elements such as root and hypocotyl. Mutation of mp gene leads to their absence. Analyzing MP’s peptid sequence reveals that it is an auxin response factor (ARF), which is responsible for the transcriptional activation of many auxin-inducible genes. The mechanism of activation is explained with the presence of other enzymes called transcription repressor proteins. Normally, in absence of auxin, these proteins bind to ARF and inhibit its action. Auxin triggers the degradation of repressor proteins and thus activates the genes.

GN protein is a guanine nucleotide exchange factor (GEF), required for the establishment of both apical and basal terminal elements of the embryo. Defect in gn gene upset the cellular distribution of PIN proteins, resulting in the deletion of both regions. GN is therefore thought to facilitate the targeting of vesicles responsible for the polar deposition of PIN proteins.

c) Radial patterning is first observed during the globular stage and divides the embryo into three radically defined regions: the outer layer, a one-celled shell is termed protoderm, which later differentiates into the epidermis. Lying beneath the protoderm are cells which later become the ground meristem, which  ultimately  becomes the cortex (ground tissue between the vascular system and the epidermis) and in root and hypocotyl, the suberized (suberin is a wax-like waterproof substance) endodermis. The other region, the procambium, later generates the vascular tissues and in the root, the pericycle.

Compared to the case of axial polarization, the molecular basis for radical patterning is much less known, though it is quite likely that the position-dependent signalling mechanisms are also involved.

3. Shoot apical meristem (SAM)

Meristem are groups of cells which retain the capacity to proliferate while their ultimate fate remains undetermined. Meristem tissues play important role during the vegetative development, providing a source of undifferentiated cells that can be recruited to form various tissues and organs.

a) SAM formation:

SAM is inititated from the upper terminal domain of the embryo, where the cells are maintained in a pluripotent stage (i.e. undetermined cells that can differentiate into many cell types). The auxin level plays a crucial role in the establisment of SAM. In fact, SMA forms at positions where auxin level is low.

A model of auxin-dependent patterning is suggested as follows: during the transition stage and the early heart stage, the PIN proteins draw auxin out of the central region to its flanking regions, resulting in an asymmetrical expression of mp gene (an ARF, as discussed earlier) and other genes alike, in particular genes coding for the transcription factors named CUP-SHAPED COTYLEDON (CUC). These proteins define the centrally positioned region that will later develop into SAM. Mutations of cuc genes lead to the lack of separation between the cotyledons, preventing the SAM formation and resulting in a cup-shaped structure (hence the name).

The SAM formation also requires activities of other proteins, such as the SHOOT MERISTEMLESS (STM) and the WUSCHEL (WUS) proteins for SAM maintainance.

b) SAM size and structure

SAM provides the precursors that will develop into the stem and later organs such as leaves and branches. SAM is normally concealed by leaf primordia and immature leaves, which form at its base and envelope it. The shoot apical meristem plus the most recently formed leaf primordia is sometimes (more inclusively) termed shoot apex.

The size of SAM varies among species, stages of development and growth conditions. The largest one is about 3mm in diameter, while the SAM of Arabidopsis is less than 50 µm in diameter.

Studies of SAM structure reveal a zonation pattern (i.e. it is divided into zones). The central zone contains the stem cells, which only divide slowly. It is flanked by the peripheral zone, which produces the leaf primordia and where rapid cell division takes place. A rip zone lies below the central zone and generates central tissues of the stem.

c) Maintainance of the stem cell population

Stem cells lost for new formation of tissues or organs are replaced with an increased rate of cell division, a process which involves maintainance proteins such as WUS. The stem cell population is also monitored by the activity of a group of related proteins: CLAVATA (CLV) 1,2, and 3. CLV1 and CLV2 are both membrane-spanning proteins, The intracellular domain of CLV1 also has kinase activity. CLV3 is a soluble protein which directs the signal molecules to the CLV1-CLV2 complex. The binding triggers a signalling cascade, which ultimately inhibits the wus gene expression and stops the cell division.

The meristem growth is self-limited: WUS expression on one hand promotes the meristem growth, but on the other hand also facilitates the expression of CLV proteins, which eventually repress it. As the result, the stem cell population is kept in a dynamic equilibrium.

4.  Root apical meristem (RAM)

Plant roots are derived from the root apical meristem, which shares many common features to its counterpart in the shoot. However, some differences do exists: The regions of root, where branches form, are well separated from the tip to avoid damage when the root is pushed through the soil. RAM is also covered by the root cap, unlike the SAM where epidermal initials are located on the surface of the meristem.

a) RAM formation

In contrast to the SAM, which only forms in regions where auxin level is low, formation of RAM requires high concentration of auxin. The high auxin concentration is the result of the PIN-mediated polar transport of auxin, in which auxin is concentrated to the basal region. There it inhibits the expression of normal ARF transcription factors such as MP or its close relative NPH4, while also activates two other genes: the plt (PLENTHORA) 1 and 2, which in turn activate another set of genes. As the result, a central region of RAM, with low mitotic activity, is formed. It is called the quiescent center (QC). QC is the ultimate source of all cells in the root.

b) Root structure:

Based on characteristic cellular behaviors, root can be divided into 4 developmental zones: the root cap, the meristematic zone, elongation zone and the mature zone.

– Root cap protects the RAM from being damaged when the root is pushed through the soil. Root cap cells are derived from specialized root cap stem cells. Root cap is able to perceive the gravity and to secrete slimy mucopolysaccharides which is used as lubricant as the root penetrates the soil.

– Meristematic zone lies beneath the root cap. It produces only one organ, the primary root. Central region of the meristematic zone is the quiescent center, from which almost all cells in the roots are derived.

– Elongation zone: is the site of rapid and extensive cell division and elongation. The rate of division decreases progressively to zero with increasing distance from the meristem.

– Maturation zone: is where cells get their differentiated characteristics. Cells enter this zone when their division and elongation have ceased.

5. Vegetative organogenesis

In addition to the axial growth from root and shoot, the 3D-structure of plants is also elaborated by the production of lateral organs. In this section, the formation of leaf primordia on the SAM will be discussed.

Leaves are initiated from the bulges on the flank of the shoot apex. It is found out that auxin induces the leaf initiation. The accumulation of auxin is achieved by activities of PIN proteins.

The leaf shape is defined by three developmental axes_

– The proximal – distal axis: from leaf base to the tip

– the axis stretched from edge to edge across the breadth of the leaf

– the adaxial – abaxial axis: from lower side to the upper side

Leaf development involves a complex pattern of cell division. The first stage is the periclinal (parallel to the surface) divisions, which take place in the subepidermal layer and define the proximal-distal axis.

Bacterial gene transfer

Main source: J.W.Dale, Molecular genetics of bacteria, Wiley Verlag, 2004

That bacteria can exchange genetic information, not only in labour but also in nature.  is a well-known phenomenon in microbiology. Three fundamental mechanisms of gene transfer are identified:

– Transformation: the uptake of isolated DNA from the surrounding medium

– Conjugation: the direct transfer of DNA from one cell to the other

– Transduction: the gene transfer via bacteriophages

It should be noted that these mechanisms are not always found in all bacteria. For example, conjugation is popular among Gram-negative bacteria but occurs only in some Gram-positive genera such as Streptomyces or Streptococcus. The fundamental aspects of each mechanism will be discussed in details below.

1. Transformation is the uptake of DNA from surrounding medium by bacterial cells. It was first discovered by Fred Griffith(1879-1941) in 1928, that an avirulent strain of Streptococcus pneumoniae (pathogen of the disease pneumonia or lung inflammation) became virulent after incubation with extract of killed virulent bacteria. 16 years later, it was proven that the phenomenon was caused by the DNA uptake of the bacteria cells.

Cells that are able to uptake DNA are termed competent. Natural competence generally occurs when cells are about to enter stationary phase or the early stage of sporulation (B.subtilis), perhaps as a response to cell density. Natural competence invariably involves the uptake of linear DNA, which limits its usefulness in genetic modification. Upon entering the cell, the DNA fragment has to be incorporated into the cell genome (often by means of recombination) to be replicated and expressed.

Artificial transformation is induced either by treatment with calcium chloride, followed by heat shock, or by electroporation. These methods are of immense practical importance in many regions of molecular biology.

2. Conjugation: is the direct DNA transfer from one cell to the other. In most cases, this transfer is one-way from the donor cell to the recipient one, and requires a special structure called pilum. Conjugation is most readily demonstrated among members of Enterobacteriacea and other Gram-negative bacteria (including E.Coli).

a) Mechanism of conjugation

The donor cell contains genes encoded for a special structure called pili, which binds to a specific receptor on surface of the recipient cell, thus forming a mating pair. A pore or channel is formed through the cell membrane, and DNA is passed to the recipient cell.

The DNA transfer is initiated by a protein that makes a single-strand break (nick) at a specific site of the DNA termed oriT (origin of transfer). A plasmid-encoded helicase unwinds the DNA and the single-strand nicked DNA is transferred to the recipient, starting with the 5′-end  generated by the nick, which is attached to the nicking protein. Replication machinery of the recipient cell seals the nick and synthesizes the complementary strand, completing the process. This mechanism is referred as rolling cycle replication. After gene transfer, each cell retains a copy of the plasmid.

Plasmids that are able to achieve this kind of transfer unaided are termed conjugative plasmids. Not all plasmids are conjugative, but some non-conjugative plasmids can still be transferred with help from another conjugative plasmid in the same cell. For example, the plasmid ColE1 needs for its mobilization both the gene mob, which codes for the nicking nuclease, and the bom site, where it is nicked by the Mob nuclease.  However, it doesn’t have the genes needed for the mating-pair formation, therefore it is non-conjugative. In present of another conjugative plasmid, ColE1 can promote its own transfer.

Sometimes, the transferred hereditary material is not plasmid but chromosomal DNA. This phenomenon is termed chromosomal transfer. In contrast to plasmid transfer, chromosome is rarely completely transferred.

b) F-plasmid is a plasmid that confers the conjugativity. It was first found promoting the conjugation from donor (“male”) cells F+ to recipient (“female”) ones F-, thus converting “female” to “male”.

In the so-called Hfr (high frequency of recombination) strains, F plasmid is found integrated into the chromosome. In contrast to normal chromosomal transfer, which takes place in a more or less random manner, the chromosomal transfer in Hfr strains begins exactly at the oriT site of the F-plasmid. Hfr strains provide an important tool for bacterial chromosomal mapping, which will be discussed in other review.

The integration of F-plasmid into bacterial chromosome is achieved by means of recombination. One of the recombination sites is adjacent to the lac operon. The process is reversible, i.e. F-plasmid can be excised also via recombination. Sometimes, the excision occurs inaccurately, leading to the formation of the so-called F’-plasmid and the deletion mutation of lac. Before the advent of cloning vector, F’-plasmid is a method to transfer a specific gene to a new host strains. The result is the so-called partial haploids, since the recipient cell has two copies of the same genes: one in the F’-plasmid and one in its chromosome.

c) Conjugation in other bacteria

The above discussed mechanism is mainly applied to Gram-negative bacteria such as E.Coli or Pseudomonas. The conjugation in Gram-positive bacteria, such as Streptomyces or Enterococcus, is comparable, though there are still some substaintial differences:

The first difference is, that conjugation in Gram-positive species requires much fewer genes, since pilus is no longer needed. It reflects the differences in structure and composition between two types.

Another difference is the mating-pair mechanism. For example, some strains of Enterococcus faecalis secrete diffusible peptides that work like pheromones, binding to specific sensors on surface of other bacteria. Upon binding, the pheromone-like peptide is transfered into the cytoplasm, where it activates the tra gene, which ultimately leads to the formation of the mating aggregate and the DNA transfer machinery. Interestingly, the peptide secreting cell is the recipient. This mechanism has many advantages, such as the economical aspect (since the mating aggregate is only synthesized when the recipient cell is nearby), and lower risk of being detected by host immune system (due to the lack of some characteristic surface antigenes such as the pilus).

Enterococcus faecalis provides another exception, this time to the rule that conjugation is plasmid-mediated. Enterococcus faecalis contains a transposon named Tn916 that is able to be transferred from one cell to the other via conjugation. The transfer process shares many common features with the standard plasmid transfer. The first step is the excision of transposon from the chromosome using transposon-encoded protein Int and Xis which are related to those responsible for the integration or excision of bacteriophage lambda. The newly formed circular molecule has oriT site and tra gene, which are needed for the conjugation, but no oriV site. Therefore, it cannot be replicated and thus be considered a plasmid. The next steps are identical to those found in plasmids: one strand is transferred to the recipient cell, where its complementary strand is synthesized. The new transposon now is integrated into the chromosome, finishing the transfer process.

Another intriguing feature of Tn916 is that, though it is in the chromosome, it is never found partially transferred via chromosomal transfer. In fact, transfer after excision seems to be the only mechanism. It is explained that the promoter that drives the expression of tra gene is on the wrong side, so the expression of tra gene cannot be initiated when it is still in the chromosome. Only after the excision that RNA polymerase can go around due to the circular structure and transcribes tra.

3. Transduction is the phage-mediated gene transfer. It is discussed in details in the “Bacteriophage” review.

4. Recombination

All gene transfer processes between bacteria, excluding the plasmid transfer,  share a same feature, in which a DNA fragment is inserted into the chromosome by breaking both strands, crossing them over and rejoining. This process is termed recombination.

a) Homologous recombination: is the most common type of recombination. This mechanism requires the presence of highly similar (though not necessarily identical) regions of DNA.It involves an X-shaped structure called Holliday junction. Other recombination mechanisms are site-specific recombination and non-homologous recombination, though they are not discussed in this review.

b) Consequences of recombination

Recombination between a linear DNA fragment, introduced by transformation, transduction or conjugation and the recipient chromosome at two different sites results in the integration of the DNA fragment into the chromosome, replacing the corresponding region between two combination sites in the chromosome.

On the other hand, recombination between plasmid and chromosome or two plasmids leads to the formation of a bigger circular molecule which is the fusion of two participating DNA.

Results of two intramolecular recombination (due to the presence of repetitive sequences inside the molecule) depend on their relative orientation. If two homologous regions are on the same direction (direct repeats), recombination will result in the separation into two smaller circular DNA molecule. On the other hand, if two homologous regions are on opposite directions (inverted repeats), recombination leads to inversion.

The presence of repetitive elements and hence deletion or inversion by recombination are the cause of the phenomenon called plasmid rearrangement.

 

Plasmids

Main source: J.W.Dale, Molecular genetics of bacteria, Wiley Verlag, 2004

1. Roles of plasmids

Plasmids are bacterial extrachrosomal DNA elements. Plasmids and bacteriophages share many common features, though they are in fact two different phenomenons. Some phages do not integrate their genome into bacterial chromosome, the prophage exists instead as a normal bacterial plasmid (bacteriophage P1). On the other hand, some plasmids are capable of being transferred to another cell (F-plasmids).

Plasmids provide bacteria their needed extra flexibility to cope with the changes in environment. In fact, some of bacterial characteristics are not encoded in chromosome, but determined by plasmids. They will be described in details below.

a) Antibiotic resistance: Many bacteria carry their antibiotic-resistance genes in plasmids. The ability of plasmids to be exchanged between cells is one of the major reasons for the widespread dissemination of antibiotic-resistance genes.

b) Colicins and bacteriocins: are names for the phenomenon that some bacterial strains have the ability to produce antimicrobial proteins which kill their closely-related microorganisms. The genes for these protein, the colicin genes, are carried on plasmids, together with a second gene for antidotes to protect the bacteria from their own toxins. One of such plasmids, the ColE1, is of special importance, since most of the commonly used E.Coli vectors nowadays are derived from it.

c) Virulence determinants: many pathogenic bateria have their virulence thanks to the genes carried on plasmids.

d) Ti-plasmids are plasmids from Agrobacterium, the bacteria which cause the crown gall disease on plants. Upon injecting the Ti-plasmids into the host cell, the bacterial DNA is incorporated into host genome and changes its metabolism. As the result, turmor-like nodules are formed on the roots of infected plants (hence the name: Turmor inducing plasmids). Ti-plasmid is one of very rare cases, in which bacterial DNA is inserted into an eukaryotic genome. It is therefore of special importance, being used as vector to transform plant cells.

Another bacterium that affects plants is the Rhizobium. In contrast to the case of Agrobacterium, this relationship is rather symbiotic than pathogenic. Rhibozium infects leguminous (bean-family) plants, causing the formation of nodules on the roots (the process called nodulation). These nodules are the sites for nitrogen fixation, facilitated by bacteria. In return, plants share their organic products of the photosynthesis. All genes for nodulation and nitrogen fixation are carried on the bacterial plasmids.

e) Metabolic activities Genes for various metabolic activities, such as the fermentation of lactose, are found on plasmids. They can sometimes lead to confusion in bacterial identification tests, which often use metabolic traits.

Another type of plasmids that carry genes for metabolic activities are the ones responsible for the biodegradation / bioremediation of toxic compounds. Such genes are of great potential to degrade environmentally harmful substances.

2. Molecular properties of plasmids

Plasmids are circular DNA molecules with compact conformation. Plasmid size varies from a few to hundreds kb. Generally, plasmids are divided into two groups:

The first one consists of relatively small plasmids (<10kb), which are present in large number in the cell (hence the name high copy number). Their replication is not independent of chromosome replication and cell division, though there are some mechanisms to control it. Therefore, it is possible under certains circumstances to specifically inhibit the chromosome replication, while keep the plasmids duplicating. This phenomenon, termed plasmid amplification, is of great importance in plasmid isolation.

The second type of plasmids is termed F-plasmid. They are large plasmids, only present in only one or two copies per cell. Their replication is tightly controlled in the same manner as that of chromosome, and there are also several extra mechanisms to control the number of plasmid per cell. F-plasmids are also called conjugative plasmids, since they play crucial role in the conjugation (described later).

Two types of plasmids depict different strategy to distribute plasmids between two daughter cells after cell division. The high copy number plasmids rely on their sheer number to ensure that each daughter cell will receive  at least one copy of plasmid. The high number imposes a burden on the cell, therefore the plasmid size must be kept low. On the other hand, low copy number plasmids carry large genes for complicated process such as cell conjugation, so neither high number nor size reducing is an option. Instead, various regulatory mechanisms are applied to tightly control the number of plasmid per cell.

a) Plasmid replication and control:

Generally, the replication is initiated from a known point termed oriV (the vegetative origin, to distinguish it from the starting point of conjugation transfer, oriT). The mechanisms of high- and low copy number plasmids differ significantly.

– Example of a high copy number plasmid: Replication of ColE1

ColE1 is a small (6.4kb) plasmid, carrying a colcinogenic gene colE1, an immunity conferring gene imm, mob gene encoding a nuclease required for mobilization, copy number controlling gene rom, an oriT and an oriV.

The replication begins with the production of a RNA primer (termed RNA II) upstream of oriV.  The transcripton goes pass through oriV,  but the RNA transcript is cut specifically at oriV site by enzyme RNase H. DNA polymerase uses this RNA fragment as primer and facilite the replication.

Another RNA, termed RNA I,  is synthesized upstream in opposite direction. RNA I has a 108-base long sequence that is complementary to RNA II. The interaction of RNA I and II changes its secondary structure and inhibits the cleavage by RNase H. Another regulatory enzyme, mob, contributes to this by facilitating the interaction of RNA I and II. As the result, EcoE1 plasmid is many times replicated, but the replication is hindered to some extend.

– Example of a high copy number plasmid: Replication of R100

R100 is a large (89kb), conjugative plasmid, containing resistance genes for 4 antibiotics: tetracyline (tet), chloramphenicol (cat – chloramphenicol acetyltransferase), streptomycin (str) and sulfoamides (sul), as well as resistance gene for mercury salts (mer). Genes required for conjugation is arranged in cluster.

The replication begins from oriV, facilitated by protein RepA, which is expressed from the repA gene adjacent to oriV. Replication is inhibited by two genes which are also coded in plasmid. One of them copB codes for the enzyme CopB, which represses the repA-transcription. The other one, copA, is within the region of repA but trancribed in opposite direction. The product, practically an antisense-RNA binds to repA transcript and represses the translation. Together, they ensure that only a limited numbers of plasmid copies are made before the replication is repressed.

R100 is found unable to co-exist with some other plasmids such as R1, the phenomenon termed plasmid imcompatibility. The reason is: both plasmids produce very similar copA antisense-RNA, so the products are interchangeable. The concurence between two plasmids results that only one remains on the daughter cells, while other is lost. It should be noted that there are still many different causes for incompatibility.

copA and copB work in trans, but R100 also contains a sequence termed par sequence that works in cis. Par is responsible for the correct partioning of plasmid copies at cell divison.

– Replication control by DNA repeats (iterons)

Some plasmids, such as F-plasmids, exist as a sole copy in the cell. It cannot be sufficiently explained by above mentioned mechanisms. Another control mechanism is provided by the existence of many repeated short DNA sequences (17-22 bp long), termed iterons. RepA can bind to both iteron sequences, effectively lock two plasmid together into a handcuff-like structure and prevent the replication.

Iterons are also a possible cause of plasmid incompability. If two plasmids have the same iterons, they will be locked together by RecA and cannot be replicated.

– Replication of linear plasmids

Most baterial plasmids are circular, but not all of them. Linear plasmids have been characterized from several bacterial genera, including Borrelia and Streptomyces. The linear DNA poses problem for the replication that the 3′-end of the lagging strand cannot be completely replicated because there is not enough place to the Okazaki fragment to form. Bacteria adopt different strategies to overcome this problem.

In Borrelia, the ends of both strands are covalently joined together into a hairpin-loop structure. The replication takes place in conventional manner. After the replication, the ciruclar product is cut into two new linear plasmids.

In Streptomyces, a terminal protein (TP) binds to both 5′-ends and facilitates the patching of the unreplicated region at lagging strand’s 3′-end.

3. Plasmid instability

– Plasmid integrity: Plasmids often contain many recombination hot-spots: transposons and insertion sequences. They are therefore susceptible to recombination. The consequence can be either gene deletion or gene rearrangement.

– Partitioning: plasmids adopt many strategies to ensure that each daughter cell receives at least one plasmid copy.

In the case of high copy number plasmids, recombination can lead to the formation of multimeric structures such as dimers, trimers… This phenomenon reduces the number of plasmids for partitioning and hence increases the risk that one daughter cell will receive no plasmid. To counter this, new mechanisms are developed to diassociate any newly formed multimer. For example, EcoE1 utilizes host site-specific recombination enzymes, XerC and XerD, to recognize its own sequence cer. In a dimer, there are two cer sequences, which will be crossed over and rejoined by recombinase, leading to the disassociation of dimer.

Low copy number plasmids adopt another strategy: any cell without plasmid will be killed. For example, F-plasmid contains an operon ccd consisting of two genes: ccdA and ccdB. CcdB is a toxic protein which inhibits the DNA replication through its interaction with gyrase. CcdA is an antidote to CcdB, but it is short-lived and susceptible to proteolytic degradation. Only cells with plasmids can produce CcdA to protect them from the toxic CcdB, while cells without plasmid die.

Similar strategy is adopted by plasmid R1. It contains a gene hok (host killing) encoding for a toxic polypeptide , and also produce the antisense-RNA sok (supression of killing) to eliminate the hok mRNA. sok antisense-RNA is short-lived and only cells containing plasmid is able to produce it continuously and survive.

– Differential growth rate plasmid replication puts much burden on the host cell. In case of wild-type plasmids, the difference in growth rate between cells containing and not containing plasmids is small. However, in case of genetically engineered plasmids, the burden on host cell can be considerable, and differential growth rate becomes a factor to count with.

Bacteriophages

Main source: J.W. Dale, Molecular genetics of bacteria, 4th edition, Wiley Verlag, 2004

1. Introduction

Bacteriophages (or in short, phages) are viruses that infect bacteria. The infection mechanism begins with the attachment of phage on specific receptors on the surface of the bacterium, followed by injection of phage genetic material (DNA or RNA) into the host cell. The so-called early genes are first expressed by pre-existing host’s transcription and translation machinery, most of them are responsible for replication of phage nucleic acids. After that, the late genes are switched on, resulting in the assembly of phage particles, cell lyses and releasing of new phages, which in turn infect another bacterial cells.

Some phages don’t always enter the lytic cycle described above. Instead, they establish a more or less stable relationship with the host cell, in which phage genetic material is incorporated into host genome and replicated by host replication machinery, a state called lysogeny. Such phages are termed temperate phages. Lysogeny plays an important role in biotechnology, since it is one of the first methods to transfer foreign genes into bacterial genome. Under certain conditions, lysogenic phages can enter lytic cylce.

A simple experiment to identify phages is the infection of a bacterial culture on agar plate. A suspension of bacterial culture in soft agar is mixed with phage culture and poured onto an agar plate. Phages released from infected cells can only infect other cells in neighborhood. As the result, clear or turbid dots will appear on the agar plate, indicating that the growth of bacteria in this regions is heavily inhibited. Such dots are termed plaques.(Note: it is still possible to infect the bacterial culture on agar plate simply by pouring phage culture onto it, but the formed plaques is 2D and thus much more difficult to see. )

The structure of phages is very simple: it consists of genetic material, either DNA or RNA, contained by a protein coat. The size, structure and assembling mechanism, however, vary significantly and will be covered in details later.

Phages were one of the first vectors for cloning, but nowadays their use is more or less replaced by plasmids.

2.Single-stranded DNA containing phages

a) ΦX174, one of the smallest phages

ΦX174 is an icosahedral (20-face) phage that contains a circular single-stranded DNA of 5386 nucleotides, which encodes for 11 proteins. The genes are tightly packed, with very little non-coding sequence and extensive use of overlapping genes. It greatly reduces the genome size, which may offer some economic advantages, but also limits the gene choice and puts constraints on the gene evolution. That explains why overlapping genes are rarely used in higher virus and bacteria.

The mechanism of replication of phage DNA is as follows: upon entering the host cell, the phage single-stranded DNA (which is a “plus” strand) serves as template for the synthesis of the complementary “minus” strand by host machinery, forming a double-stranded DNA (termed replicative form of RF). The “minus” strand is now in turn used as template for production of a new “plus” strand. As opposed to chromosome replication, the synthesis of “plus” and “minus” strands is in fact separate. After late genes are expressed, the protein coat is assembled and the “plus” strand is packaged into it.  The “minus” strand degrades. This mechanism resembles replication of some plasmids.

b) M13 phage is  filamentous. It is termed “male-specific” phage, since it attaches specifically onto the tips of F-pili formed by F-plasmid.

M13 is somewhat unusual among phages. First, it doesn’t cause cell lysis upon releasing. Plaque-like regions can still be seen on agar plate, but they are rather areas of reduced growth. Secondly, the protein coat is not pre-synthesized. Instead, it is formed by means of protein polymerization around the single-stranded DNA molecule as it is excluded out of the cell.

The mechanism of replication resembles that of ΦX174, through the double-stranded RF. Large numbers of RF are found inside the infected cells and can be isolated easily using normal plasmid isolation techniques.

3. MS2, RNA-containing phage

MS2 is an icosahedral, RNA-containing, male-specific phage. It attaches specifically on the sides of F-pili. MS2 genome is extremely simple: it contains 3600 nucleotids, and codes for only 3 protein: a coat protein, a maturation protein and a replicase. All other needed proteins are from host. MS2 is a positive-sense virus: its RNA serves both as replication template as well as mRNA.

The replication shares the same motif as in ΦX174 and M13, utilizing the plus strand as template for production of minus strand and vice versa. The replicase, a RNA-directed RNA polymerase, is however not from bacterial cell, but translated from MS2’s RNA.

4. Double-stranded DNA containing phages – Introduction

T4 and lambda phages represent large, double-stranded DNA containing phages. They both have linear genome, which presents a serious problem in replication: the 5′-end of the lagging strand cannot be properly replicated, since there is not enough place for the Okazaki fragment. As the result, the DNA molecule gets shorter and shorter after each cycle of replication. T4 and lambda phages develop different strategies to circumvent this issue, which will be described later.

5. Bacteriophage T4 is a virulent phage of E.Coli. Its genome is linear, double-stranded and about 165kb large.Repeated sequences of 1,6kb, in the same orientation, are found on both ends of phage DNA. It reveals the replication mechanism in T4, involving the production of a long, linear precursor which is subsequently cut into many phage DNA molecules.

The mechanism of replication is as follows: First, the linear phage DNA is duplicated by DNA polymerase. The ends of newly formed DNA molecules are joined together by means of recombination between terminal repeats.  As the result, a long linear DNA molecule is formed. Now DNA is tightly coiled and packaged inside the pre-formed phage head until it is full. Then DNA molecule is cut and the remaining rest is again packaged into other phage heads. As the result, phage DNA is successfully replicated and packaged, albeit with different terminal repeats.

Control of phage development The early genes are solely transcribed by host RNA polymerase. Second group of genes, termed quasi-late or middle genes, is expressed a little later. They still utilize the host RNA polymerase, but add two other phage-encoded proteins which assist the binding of RNA polymerase to DNA.

One possible activation of late genes is via the production of a new sigma-factor, which alters the specificity of RNA polymerase. Other regulatory mechanisms may also be involved.

6. Bacteriophage lambda also contains linear double-stranded DNA. As opposed to T4, no terminal redundancy nor variation is found in lambda. Instead, short (12 bases), complementary to each other overhangs are found on both 5′-ends of lambda’s DNA, which highly suggest the replication mechanism involving circularization. These sequences are thus termed cos site, in which cos stands for cohesive.

a) The mechanism of replication is suggested as follows: at first, both ends are joined together into a non-covalently linked circle by means of interaction between two complementary overhangs. The enzyme ligase seals the nicks, forming a covalently closed circular DNA molecule. In early stage of infection, this molecule is replicated in theta-mode, the standard replication mechanism of bacterial circular DNA and plasmids. Later, it is turned into rolling circle mechanisms, resulting in a multiple length linear DNA molecule. In the last step, the sequences between two cos sites are cut and packaged into phage particles.

As opposed to T4, deletion or insertion during replication results in shorter or longer DNA packaged inside phage particles, both cases lead to phage’s destabilization. Phage lambda has therefore packing limits: the distance between two cos sites must be between 75 and 107 percent of the wild-type sequence (about 37 – 52kb). This limit is of practical importance in use of lambda as cloning vectors.

b) Lysogeny: Lambda is a temperate phage, which means it doesn’t enter the lytic cycle. Instead, it establishes a more or less stable relationship with the host by integrating its genome into the host genome. The integration is generally a site-specific recombination. Integrase, which is encoded by the phage gene int, specifically recognizes two sequences on phage (attP) and bacterial chromosome (attB) and facilitates the recombination. The result is the integration of phage DNA (termed prophage) into the bacterial chromosome. Prophage is flanked by two sequences: attL and attR, neither resembles attP nor attB, thus can be recognized by Int. The integration is therefore irreversible.

The reversal process (excision of prophage out of bacterial genome) requires another protein, Xis. Xis interacts with Int and modifies it, so Int can recognize the attL and attR sites. The following recombination facilitated by Int+Xis results in the excision of prophage and induction of the lytic cycle.

It should be noted that chromosomal integration is not an essential feature of lysogeny. In some lambda mutants as well as other bacteriophages, prophage is maintained in host cell as extramolecular circular DNA molecules, practically plasmids.

c) Lytic and lysogenic regulation of bacteriophage lambda:

– Overview of lambda genome: In lambda genome, genes are arranged in functional groups. The early genes are in the vicinity of the control region and responsible for the replication and activation of late genes. The late genes are expressed later in infection and responsible for the synthesis of the protein coat and inhibition of early genes which are now not needed anymore.

There are two major promoters leftwards and rightwards, termed PL and PR. Associated with them are correspondingly the terminator tL leftwards and tR1, tR2 rightwards (tR1 is a weak terminator and bypassed in many cases).

– Control of the lytic cycle: 

Early in the infection, only a small numbers of genes are expressed, among them the genes for replication and two genes: N and cro.

N protein is an anti-terminator. It binds to specific sites on the DNA upstream of the terminators, interacts with RNA polymerase and thus allows the transcription to proceed through these terminator sites (the transcription stops at other strong terminators, which are not affected by protein N).  The result is the expression of a new set of genes, termed delayed early genes. One product is the protein Q, which is also an anti-terminator. Q triggers the transcription of late genes, which are responsible for synthesis of phage particle.

Cro protein acts as repressor of the early genes’ promoters. The accumulation of Cro leads to the repression of early genes. Cro is also found playing important role in regulating the lytic/lysogenic switch, which is described in details below.

– Control of the lysonenic cycle:

Lysogeny is an alternative of the normal lytic pathway. When the phages enter lysogeny, all genes needed for the lytic cycle are repressed. The key repressor is coded in gene cI (c stands for clear, since mutants are not able to form lysogeny and clear rather than turbid plaques are observed on agar plate as the result). cI transcription is driven by the promoter PE (E stands for Establishment of repression), which requires two other proteins: cII and cIII, in order to be functional. cII and cIII are both delayed early genes and expressed from promoters PR and PL, correspondingly. Both cII and cIII are highly susceptible to proteolytic degradation and thus very short-lived.

cI binds to the operator region of PR and PL, termed OR and OL, and switches them off. This prevents the expression of many key proteins of the lytic cycle, most notably N and cro. However, the transcription of cII and cIII is also repressed in effect, which inhibits the further production of cI. To circumvent this problem, lambda utilizes another weaker promoter, PM (M stands for Maintenance), to continue producing cII and cIII. PM is adjacent to OR/OL regions, and the binding of cI to this region not only represses PR and PL, but also activates PM. cI has thus both negative and positive regulatory functions.

What is the function of Cro protein in this process? Cro inhibits the production of cI, either directly by binding to the PE promoter, or indirectly by switching off the PM promoter and thus inhibiting the production of cII and cIII, which are essential for cI expression.

– Completition between Cro and cI is found extremely important for the decision between lytic and lysogenic pathways. This competition is illustrated by the interaction of Cro and cI with the operator CR (The interaction with CL happends in the similar manner).

OR consists of three similar adjacent DNA regions, termed OR1, OR2 and OR3. OR1 overlapps with PR, while OR3 overlapps with PM, thus binding of repressor to OR1 switches off PR, to OR3 swichts off PM.

The affinity of cI is greatest for OR1, though the binding is cooperative (which means the binding on the first site facilitates the binding on the second one ). In present of cI, PR is repressed, while OR3 remains unoccupied, cI only binds to OR3 when its concentration is high enough and further cI production is just wasteful. This control mechanism keeps cI on a low but stable level, which represses the lytic pathway while no putting too much constraints on the host cell.

The binding of Cro is somewhat different. Cro prefers binding to OR3 first, which swicht off cI synthesis for good. PR is only repressed only at high concentration of Cro.

– Choice between lysogeny and lysis is largely determined by the state of host cells. A healthy host with full nutrients will trigger the lytic pathway. On the other hand, ill host will have lambda DNA incorporated into his genome, where it waits for a “better time” to come. Such a cunning parasitic strategy.

7. The discovery of restriction enzymes

Following experiment shows how restriction enzymes are discovered. Two different strains of E.Coli, E.Coli C and E.Coli K are infested with bacteriophage lambda. It is found out that the number of plaques in E.Coli C culture is 10 000 times higher than in E.Coli K culture. If the phages that are capable of forming plaques in E.Coli K culture, termed lambda-K, are picked out by a toothpick, resuspended in buffer and tested again with both strains of E.Coli, the number of plaques formed in both cultures are now comparable.

It is explained that E.Coli K is able to produce enzymes which inhibits the growth of bacteriophage lambda. Such enzymes, termed restriction enzymes, recognize foreign DNA and degrade it. To avoid being cut by its own restriction enzymes, bacterial DNA is heavily methylated. If a mutated phage DNA can somehow escape degradation by host restriction enzyme, it is replicated and also properly methylated by host replication machinery. In the second round, when used against E.Coli K, the methylation pattern allows it to escape the recognition by host restriction enzymes, and the growth of phage is not inhibited any more.

8. Applications of bacteriophages

– Phage typing: is a method to distinguish two different strains of the same bacterium, which have different sensitivity. Same results on two different samples might suggest that they are the same strain and originated from the same source.

– Phage therapy: Using bacteriophages to fight against pathogenic bacteria might be a new promising therapy. There are still problems with it, such as phages are often killed by human immune system, or pathogenic bacteria can develop resistance against phages. Another more practical approach is use lytic enzyme from phages to kill bacteria.

– Phage display: is an elegant method to find a protein that binds specifically to the ligands of interest. The process begins with cloning many random DNA fragments that encode for a large set of proteins into a phage genome, preferably the filamentous M13. The phages are next amplified by infecting a suitable E.Coli strain. Meanwhile, the ligand of interest is immobilized on a microtitier plate. The phage suspension is now poured onto the plate. Phages with correct proteins will bind to the ligand and stay on plate, so they can be recovered later. Other phages are just washed away.

Plastid transformation

Sources:

http://www.aloj.us.es/bioqplantas/tema4-5/plastid%20annurev.pdf

http://onlinelibrary.wiley.com/doi/10.1111/j.1467-7652.2011.00615.x/pdf

http://www.springerlink.com/content/km042p85455747l8/fulltext.pdf

1. Plastids are cellular organelles found in algae and plants, which supply energy to the cell via photosynthesis. They are also major sites for pigment, starch, amino acids, fat biosynthesis and storage. Plastids have their own genome and transcription / translation machinery. Plastid genome, termed plastome or ptDNA, is highly polyploid, circular double-stranded DNA, about 120-180kb in site and encoding for ca. 120 proteins.Plastid genome retains many features of prokaryotic genome, which suggest their endosymbiotic origin.

Plastids have been successfully transformed in about 16 plants, most particularly in tobacco. The first transformation was achieved on isolated plastids, which were then fused into protoplasts and regenerated into transgenic plants. Most widely used method nowadays is biolistics, using gene gun to shoot DNA-coated gold particles into plastids. Cheaper methods, such as PEG-mediated transformation or microinjection, are also in development.

Plastid transformation has many distinct advantages over the classic nuclear transformation:

– Much higher transgene copy number can be obtained, since there are up to 100 plastids per cell, each containing up to 100 copies of ptDNA. Higher transgene copy also means higher level of gene expression, which increases the yield and makes purification much easier. It is even possible to exhaust the protein biosynthesis capacity of chloroplasts, impairing the plant growth.

– In contrast to the case in nuclear, transgenic DNA can be incorporated into plastid genome by means of homologous recombination. It eliminates the random insertion problem (the so-called position effect) and results in uniform level of transgene expression. Furthermore, so far no gene silencing has been reported.

– Plastid compartmentalization allows the formation of disulfide bonds as well as the correct protein folding, which have significant meaning in production of edible vaccines, pharmaceuticals and antibodies. Furthermore, compartmentalization also reduces the toxic effects of foreign protein accumulation.

– Since only maternal plastid genes are inherited, there is no risk of spread of transgene into the environment via pollen.

2. Methods of DNA delievery

Several physical obstacles stand between the transgenic DNA and target plastid genome: the cell wall, cell membrane and the double-layer plastid membrane.  Unfortunately, no known viral or bacterial system is found capable of delivering foreign DNA into plastids like Agrobacterium to nuclear, direct methods of DNA delivery have to be used.  The most effective and widely used one is biolistics, using a gas-driven gene gun to deliver plasmid-coated gold or tungsten particles into plastids, followed by regeneration of callus into plants.

Another method utilizes polyethylene glycol (PEG) to destabilize the membranes, allowing plasmid to get into plastids. This method requires the cell wall to be first enzymatically removed, and the newly formed vulnerable protoplasts to be treated carefully in proper osmotic conditions. Otherwise, no specialized equipment is required.

Due to the great number of plastids per cell, only about 10% of plastids are successfully transformed. The transplastomic cells are now in the so-called heteroplastomic state. The dividing of plastids between 2 daughter cells during cell division is basically a random process. Under constant selective pressure (usually in present of antibiotics), cells containing transformed plastids (which have antibiotic-resitance genes) are at advantage. As the result, the percentage of transformed plastids will increase after each cell division cycle. It is estimated that about 20-30 cycles are needed to achieve the homoplastomy. That means explants have to go through several cycles of regeneration under constant selective pressure. Plant regeneration ability is proved to be one of the most significant obstacles hindering the use of plastid transformation in broad range of species.

3. Vector design – flanking sequences

Transgenic DNA is incorporated into plastid genome by means of homologous recombination. The plastid transformation vectors are usually E.Coli plasmid derivatives with clones ptDNA sequences flanked by flanking regions, which determine the recombination sites. If foreign DNA is to insert into plastid genome, it is advisable to have recombination sites in intergenic regions to avoid gene deletion and inactivation. 13 such sites have been identified and utilized in tobacco. If gene knock-out is the aim, target sequence is mutated and reinserted into plastome.

4. Vector design – Promoters and UTRs

In general, plastid utilize prokaryotic transcription and translation machinery, though processes like intron splicing or RNA editing, which are absent in bacteria, are also found.

The most widely used and strongest promoter so far is sigma-70 type RNA promoter of rRNA operon (Prrn), which is usually used to drive the expression of the marker gene aadA. Another promoter in used is the endogenous pbsA promoter.

In clear contrast to prokaryotes, there is a high degree of translational regulation in plastids. The 5′-UTR, which is often a truncated or mutated form of the native sequence,  is found playing important regulatory role. Changing the 5′-UTR sequence can result in significant change in level of gene expression.

5. Genetic markers

The most widely used are genes conferring resistance against antibiotics or toxic compounds:

– Aminoglycosides antibiotics, in particular Spectinomycin, has become indispensable for plastid transformation. They bind to prokaryotic-like plastid ribosomes, inhibit protein synthesis and thus inhibit greening, cell division and shoot formation. The aadA gene, encoding for the detoxifying enzyme aminoglycoside-3”-adenyl transferase, is widely used as selectable marker. Other antibiotic-resistance genes in use are: neo (encoding for neomycin phosphotransferase II ) against neomycin,  and aphA6 (encoding for aminoglycoside phosphotransferase) against kanamycin and amikamycin.

– Another gene in use is the gene encoding for betaine aldehyde dehydrogenase (BADH) from spinach, which detoxifies the toxic compound betaine aldehyde. Though this marker still requires many further researches, it is highly promising, since it circumvents the issues associated with the use of antibiotics and antibiotic-resistance genes.

As far as clean-gene technology is concerned, selectable markers should be removed after homoplastomic state is achieved. Many sophisticated methods are developed for this purpose, for example using Cre recombinase from P1 bacteriophage for marker excision.

Reporter genes, such as gus or gfp, can also be used in tandem with selectable markers to enhance the screening efficiency.

6. Analysis

Though plasmids for plastid transformation are developed for sole expression in plastids, false-positive is still possible. It can be the result of integration into nucleus, or spontaneous mutation to antibiotic resistance (which happends quite a lot in all tested species sofar). Therefore, further analyses are needed. They range from test-crossing to PCR analysis of purified ptDNA.

Vectors for plant transformation

Main source: Slater, Plant biotechnology, 2nd Edition, Oxford 2008

Vectors are the main tool to transfer transgenes into the plant genome. Vectors for plant transformation is created using the same concept as any other plasmid-derived cloning vector. In this review, most important features of a standard vector will be described, as well as insight into its development and application.

1. Common features of plasmid-derived cloning vector

– Small size: which makes the plasmids more difficult to damage, as well as enhances the yield. Small size also means there is less chance of having restriction sites outside the multiple cloning site (MCS), which makes design and integration much easier.

– Containing a selectable marker gene which offers the host a selection advantage. It will be useful for the next step where successfully transformed cells need to be isolated from others. Most popular ones are gene encoding for antibiotic resistance, very often ampicilin.

– Containing a MCS, which has many cleavage sites for a large range of restriction enzymes.

– Contaning a reporter gene which enable the easy detection of colonies containing recombinant plasmids. This is usually archieved by putting the small MCS inside the reporter gene’s encoding sequence, much too often lacZ, gene for ß-galactosidase protein. The successful insertion of the gene of interest into the plasmid results in the insertion mutagenesis of lacZ, effectively disabling it. Cells without working lacZ is unable to degrade X-gal (a galactosid analog) and remain white, while cells with working lacZ can degrade X-gal and release a blue substance, which turns the colony in blue. Reporter gene is not 100% reliable, but it does dramatically reduce the number of colonies which need to be screened.

– An origin of replication (oriC) for replication of plasmids inside E.coli cells

– The gene(s) of interest

Plasmids used for plant transformation via Agrobacterium need additionally:

– The ability to replicate not only in E.Coli but also in Agrobacterium

– The border sequences needed for integration into host’s genome.

– Additional selectable markers for further screening of successfully transformed plant cells

– Appropriate promoters and terminators allowing (particularly prokariotic ) genes to be expressed by host’s eukaryotic transcription machinery.

2. Promoters and terminators

Many plant transgenes, reporter genes, marker genes are derived from bacteria, which need their own promoters in order to be expressed. Furthermore, suitable terminators are also required to ensure that transcription is stopped at the correct positon. Promoters  and terminators are often also incorporated into the plasmid.

The genes from Ti-plasmid of Agrobacterium for opines synthesis, especially the nopaline synthase (nos) gene are used as suppyling source for both promoters and terminators used in vectors. Since they are in the T-DNA, they work without problems in plants.

The most widely used promoter is however the promoter of cauliflower mosaic virus 35S RNA gene (the 35S promoter),  a very strong constitutive one. This promoter can be used to drive expression in all tissues of transgenic plants, particularly in dicotylendons. The activity of 35S promoter can even more enhanced by the inclusion of one or more copies of enhancer region. Alternatively,maize ubiquitin I or rice actin promoters are used in monocotyledons for high-level expression

Tissue-specific promoters are used to drive the gene expression only in certains tissues. Much effort has been put into isolation of promoters with this desired characteristic, with some success.

Inducible promoters are another actively researched field in plant biotechnology. These promoters only drive gene expression under certain conditions, allowing the timing of transgene expression to be controlled closely. There are three main categories: non-plant-derived systems, plant-derived systems that respond to environmental signals, and plant-derived systems based on developmental control of gene expression.

a) Non-plant-derived systems response to the application of an exogenous chemical.  The common features of inducers are:

– Specificity: no gene expression takes place without inducer or with inducers from other types.

– Gene expression should be turned on and off rapidly with application/withdrawl of the inducer.

– Inducer should be non-toxic and cause no change in gene expression.

Some of typical inducers of this class are:

– Tetracyline: works in tandem with bacterial tet operator (from tet-Operon coding for tetracyline resistance). Without tetracyline, the repressor protein TetR binds closely to the operator and prevents the gene expression. Upon tetracyline binding, TetR is modified and releashed from the operator, allowing the transcription to begin. The regulatory machinery used in plants is a chimeric promoter consists of a 35S promoter and several copies of tet operator, while TetR is overexpressed from (another???) 35S promoter.

Tetracyline can also be used as an inhibitor. This time, it’s not TetR but another protein, tTA acting as an activator.

The drawbacks of this system are the issues with high level of TetR inside the plant cells, and the short half-life of tetracyline which means it must be applied constantly.

– Alcohol-inducible promoters: consists of a core 35S promoter and binding sites for the constitutively expressed AlcR transcription factor (from the fungus Aspergillus nidulans).  Ethanol is applied as drench or vapour. This system is not very widespread due to the activation by other metabolites, as well as the side-effects of ethanol on plants.

– Steroid-inducible promoter: are again chimeric from 35S promoter and a modified transcription factor capable of binding a steroid hormone or its analogue.

– Copper-inducible promoters: are based on the yeast metallothionein regulatory system.

Non-plant-derived systems require the application of specific substances, some of them are quite expensive, and thus are unsuitable for any agricultural scale.

b) Plant-derived systems share the same concept as non-plant-derived systems, using chimeric promoter consists of a core promoter (usually 35S promoter) and binding sites for transactivating factors. These factors can either respond to  environmental signals or be only expressed in specific stages of plant development.

– Systems based on response to environmental signals: include wound-inducible, pathogen-inducible and heat-shock-inducible.

– Systems base on developmental control of gene expression: involve senescence (aging) – specific gene expression, abscisic acid-inducible gene expression, auxin-inducible gene expression…

Plant-derived systems depends heavily on normal plant processes, but they are much simpler to use in agriculture. It’s worth noting that no system is suitable for all situations.

3. Selectable markers are used to allow successfully transformed tissues to be effectively isolated, usually by conferring resistance to a toxic substance. Since transformation is a very-low-frequency event, selectable markers help reduce the number or samples for screening considerably. The most common selectable markers are antibiotics resistance genes, especially the nptII gene (conferring resistance to kanamycin). Though plants are eukaryotic, antibiotics still inhibit the protein synthesis in organelles, particularly in chloroplasts. Antibiotics other than kanamycin are used for plant with high degree of resistance to kanamycin (cereals), or on the other end, too sensitive to it (some soft fruits). Some prominent examples are: bleomycin, spectinomycin, hygromycin. Other than antibiotics, herbicides can also be used for selection. Genes conferring resistance to herbicides such as chlorsulphuron, bialaphos… are already in use.

The use of any selectable markers needs to be controlled carefully. Too high concentration of toxic substance can lead to inability to regenerate, too low concentration can lead to high number of false-positive results. Furthermore, the use of antibiotics- and hercibide-resistance genes as selectable markers raises the concern about the spreading of antibiotics-resistance gene as well as the creation of the so-called “super weed”. Much effort has been put to discover new types of selectable markers. Some of them involve the abitity to use mannose and xylose as carbon-source other than sucrose. ability to express green fluroescent protein (GFP), the ability to grow in absense of cytokinin… to name but a few.

4. Reporter genes are used to asses the expression by promoter analysis, and as indicators of transformation. The reporter genes confer an easily detectable phenotype on transformed plant tissues, allow them to be sorted out by simple non-destructive assays. Only a small numbers of reporter genes are widely used in plant transformation vectors. Each will be described below.

a) ß-glucuronidase (GUS) is perhaps the most widely used reporter gene, because of its many advantages: GUS can be assayed sensitively, quickly and easily, both quantitative and qualitative data can be obtained, and little or no endogenous activity in most plant tissues has been reported.

4-MUG                                                                   X-gluc

GUS-assay with rice embryo

The substrate for GUS enzyme is 4-Methylumbelliferyl-b-D-glucuronide or 4-MUG. 4-MUG is hydrolysed to the flourescent 4-Methylumbelliferone (4-MU). The fluorimetrical measurement of 4-MU allows quantitative data to be obtained. Qualitative data is obtained from assay using 5-bromo-4-chloro-3-indolyl glucuronide (X-gluc). Hydrolysis with GUS results in an insoluble blue precipitation, effectively revealing the location of expression.

The gus gene in transformation vector is designed with an intron, allowing it only to be expressed in plants, in order to avoiding problem associated with GUS-expression in Agrobacterium.

b) Green fluorescent protein (GFP) has some advantages over GUS. It is easier to assayed, the assay is non-destructive, allowing some types of screening which are impossible with GUS, such as time-course experiments.

The gene is from the jellyfish Aequorea victoria, with some significantly modifications such as alternative splicing (results in shorter protein), more “plant-like” codon (???) and prevention of accumulation in nucleoplasm.

c) Luciferases are firefly’s emzymes catalyzing the ATP-dependent light emmiting oxidation of D-luciferin. Assays are highly sensitve allowing detection of low-level or highly localized gene expression, but the expression of luciferases in plants is difficult.

Other than firefly’s luciferases, bacterial luciferases (LuxA and LuxB from Vibrio harveyi) can also be used. They catalyze the oxidation of long-chain fatty aldehydes, resulting in light emission.

d) Chloramphenicol acetyltransferase (CAT) was the first bacterial gene to be expressed in plants. It is also a widely used reporter gene in mamalian, but in plants newer reporter systems such as GUS or GFP are proved to be better.

5. Origin of replication: since both E.coli and Agrobacterium are Gram-negative bacteria, a single origin of replication with broad host range (e.g. the RK2 origin) can be used for replication in both. Unfortunately, this kind of origin often result in low plasmid copy number in E.coli, which reduces the yield and makes the DNA manipulation more difficult and time-consuming. Alternatively, two orginins are used: one for replication in Agrobacterium and one is from a high-copy-number cloning vector (e.g. pUC ori). The downside is the increase in plasmid size and with it associated problems.

6. Vector Initially, deletion derivatives of Ti plasmids, termed co-integrative vectors, were used for transformation. The DNA was first cloned into an intermediate vector, which was small and easy to manipulate in vitro, and then introduced into the Agrobacterium vector by means of recombination within the bacterium. Normally, the vir genes and T-DNA were on the same Ti plasmid.

Today, binary vectors are used instead. vir genes and T-DNA are located on separate plasmids, which allows to reduce the size of plasmids, making them easier to manipulate in vitro.

Gateway cloning: even modern binary vectors (e.g. pGreen, 4.6kb ) are still too large, making cloning quite difficult and time-consuming. A new approach, named Gateway conversion technology,  is developed by Invitrogen, an US company, using DNA recombination rather than classic cut-and-paste methods using restriction enzymes and ligases. DNA fragment that needs to be cloned is suitably flanked by recombination sites, which, in the presence of recombinase, recognize the recombination sites in the vector and insert the DNA fragment into it. The recombinase is provided by bacteriophage lambda.

7. Transgenes

Arrangement of genes in the vector: analysis of the efficiency of transgene expression allows some rules to be drawn up: 1) When there are more than one genes in the vector, each should have its own promoter and terminator, since using the same promoter / terminator for many genes can lead to gene silencing. 2) There should be enough space between genes, and genes should be located in the same orientation, to avoid adjacent inverted repeats which lead to plasmid instability and gene silencing.

Integration sites into the plant genome: the integration is basically a random process, since higher plants don’t have an endogenous system for homologous recombination. Therefore, large differences in the level of transgene expression are expected. It has been tried using bacteriophage or yeast site-specific recombinases, with some success. Another approach is flanking the transgene with matrix-attachment regions (MAR), which keep the chromatin in open structure allowing gene expression.

Transgene modification differences between non-plant derived transgenes and plant genes can lead to poor level of transgene expression. Some transgene modifications are therefore needed:

– changing the transgene’s GC content

– Removal/ replacement of the so-called cryptic introns ( regions recognized by plants as introns and therefore spliced out, leading to deleterious mutation )

– Changing the upstream sequence of star codon to more plant-like

– Adding export signal to avoid the accumulation of transgene’s products in nucleoplasm

8. Clean-gene technology

Selectable markers are very useful in plant biotechnology, but together with them come several concerns about the health- and environmental issues such as the spreading of antibiotic- and herbicide resistance genes. It is therefore desirable to develop crops that contain no selectable markers. Some approaches are available as follows:

– Using other screening methods using no selectable markers, such as PCR. Proved to be quite laborious and expensive.

– Use selectable markers on separate T-DNA molecules

– Excise the selectable markers out of plant genome by means of recombination.