Sunday, 27 October 2019

Plastid Transformation

Plastid Transformation

Introduction to Plastid Transformation

The genome of eukaryotic cells is unevenly distributed and kept in different subcellular compartments. While the vast majority of genetic information is sheltered in the nucleus, small portions of DNA reside in organelles, namely the mitochondria and in the case of plants in the plastids. These unique organelles which in their most prominent manifestation are called chloroplasts developed from cyanobacterial ancestors in a process described by the well-accepted endosymbiosis theory (Gould et al. 2008). In brief, a pre-eukaryotic cell must have engulfed and taken up an ancestor of today’s cyanobacteria and subsequently formed a close endosymbiotic relationship with the newly developing organelle. Residues of this evolutionary ancestry are still apparent today in certain prokaryotic characteristics retained by the plastids.

However, chloroplast transformation makes high demands on vector design, transformation method as well as plant regeneration. This is exemplified by the so far not solved problem to generate fertile transplastomic lines of the model plant Arabidopsis thaliana (Sikdar et al. 1998), an example depicting the obstacles of this technique which need to be overcome to gain broad applicability. On a routine basis, so far only tobacco chloroplasts are transformed, and therefore most examples given in this article refer to tobacco chloroplast transformation. This great progress has been made in expanding the range of this technique to other plant species by Nevertheless.

Technique For Transforming DNA Transfer to the Chloroplast

Technique For Transforming DNA Transfer to the Chloroplast
Schematic drawing of the plastid transformation and regeneration process. Usually, a leaf, an explant,  is bombarded with DNA-coated tungsten or gold particles. When the DNA is delivered to one chloroplast, integration of the transgene takes place, generating a heteroplasmic cell in which a small number of plastid genomes are transgenic (open circles). Subsequent differentiation and shoot regeneration from this cell result in a heteroplasmic plantlet. To obtain a homoplastic transgenic plant requires several cycles of regeneration under selection.

Delivery of foreign DNA to the chloroplast requires its transport through several physical barriers: the cell wall, the cytoplasmic membrane, and the chloroplast double-membrane system. Since no bacterial or viral pathogen is known which could be utilized for DNA delivery, transgene transmission needs to employ rather rigid physical methods. The most effective method microprojectile bombardment. It utilizes microprojectile bombardment with plasmid-coated gold or tungsten particles, the so-called biolistic method, which was first used to transiently transform onion epidermal cells (Klein et al. 1987). 

Subsequent refinement of the technique eventually enabled the transformation of smaller cell types as well as subcellular targets, like plastids in the unicellular algae Chlamydomonas (Boynton et al. 1988) or in tobacco (Svab et al. 1990). Other sophisticated methods have also been developed, like polyethylene glycol (PEG)-mediated transformation of protoplasts (Golds et al. 1993) or even the direct injection of DNA into the organelle via a femtoliter syringe (Knoblauch et al. 1999). 

Although plastid transformation with PEG requires some experience in the enzymatic digestion of the cell wall and the treatment of protoplasts as well as the regeneration of plants, it can be basically performed with standard laboratory equipment. Micromanipulation of cells, on the other hand, requires specialized equipment, which is limiting for its use; and so far no reports have shown the successful regeneration of transplastomic plants from this particular gene transfer method. The most successful and widely used method is the biolistic transfer of DNA, depicted by the successful transformation of the plastids of numerous plant species.

Once the transgene DNA has been delivered to the chloroplast, stable integration via homologous recombination has to take place (see below) to generate a stable transgenic trait which will be passed on after plastid division to its descendants. Every chloroplast harbors up to a hundred copies of its genome, grouped in nucleoids representing aggregates of 7–10 copies. Since cells can contain up to 100 chloroplasts, a single integration event creates a transplastomic cell in which only a minority of genomes is altered, the so-called heteroplasmic state. 

For the generation of a stable transplastomic plant, wild-type plastid genomes have to be winnowed. As the sorting of plastid DNA during cell division in shoot-regeneration is a stochastic process a small percentage of altered homoplastic plants can be generated in the absence of selection pressure (5.6%; Lutz and Maliga 2008). To increase the efficiency of transformation routinely, a homoplasmic state of the engineered plants is reached by successive regeneration under strong selective pressure with an appropriate antibiotic. It is estimated that it takes between 20 and 30 cell divisions to deplete the undesired wild-type chloroplast genomes (Maliga 2004; Verma and Daniell 2007). 

Since this number could not be reached in a single plant regeneration process, explants have to go through several cycles of regeneration under selection. So far a given plant's ability to regenerate from fully differentiated tissue is the biggest obstacle for applying the plastid transformation to a large number of plant species. Tobacco is by far the best-analyzed system regarding plastid transformation, and therefore most experiments referred to in this section are made in tobacco.

Vector Design

Nicotina tabacum
Graphic map of the Nicotiana tabacum plastid genome (GeneBank accession number NC001879), made with the web-based program OGDRAW (Lohse et al. 2007). Genes on the outside of the circle are transcribed counter-clockwise, those on the inside clockwise. IRB inverted repeat B, IRA Inverted repeat A,  LSC large single-copy region, SSC single copy region. Numbered arrows Transgene integration sites. Site of integration into the inverted repeat region; Dashed arrows (numbers 8–13) therefore integration sites are in duplicate.

  • Flanking Regions
  • Promoters and UTRs
  • Transgene Stacking and Control of Gene Expression

Flanking Regions

Agrobacterium sp. mediated transformation utilizes universal vector systems in which the transgene expression cassette is flanked by two rather short sequence stretches, termed left border (LB) and right border (RB). These sequences facilitate almost random insertion of the transgene cassette into the host genome, resulting in multiple individual lines differing in site and numbers of transgene integration. 

In absolute contrast, the insertion of foreign DNA into the chloroplast genome relies on targeted integration of transgenes by homologous recombination, facilitated by a bacterial recombination system inherited from the plastids cyanobacterial ancestors (Cerutti et al. 1992). Hence, a transgene could be targeted to virtually any site in the chloroplast genome by designing the flanking regions according to the desired location. This is not only a big advantage for the positioning of expression cassettes to defined locations but also enables the targeted inactivation of plastid genes for functional studies and gene knock-outs. 

For the former, preference is naturally given to intergenic regions to circumvent deleterious effects and interference with endogenous gene expression. The targeted sequence is mutated in vitro and reinserted into the plastome for gene knock out. In the case of tobacco, numerous studies describe the targeted knock-out of plastidial genes for functional studies (reviewed by Maliga 2004). Additionally, a total of 13 sites on the plastome has been utilized for the integration of an expression cassette, demonstrating that modification and integration of heterologous genes can be performed at any given site in the circular plastid genome.

Promoters and UTRs

For the expression of heterologous genes, the choice of promoters and regulatory sequences is highly important. In general, chloroplasts mainly utilize a prokaryotic transcription and translation machinery, a heritage from their cyanobacterial ancestors. Gene organization in operons, a eubacterium-type RNA polymerase as well as sigma-like factors, and a specific codon-usage in the open reading frames are highly similar to those found in bacteria. However, during evolution novel mechanisms of gene organization and regulation of expression also evolved in plastids. Processes like intron splicing and RNA editing can be found in chloroplasts which are absent in bacteria. 

Aiming for the high-level expression of a given gene, regulatory sequences are required which provide efficient transcription, translation, and RNA stability. The strongest promoter described so far is the s 70 -type promoter of the ribosomal RNA operon (Prrn; Svab and Maliga 1993; Kuroda and Maliga 2001a, b). In the majority of vectors used to date, this particular promoter is used to drive the expression of the marker gene aadA (see below) to provide sufficient expression for the selection of transformed cells. Another promoter in use is the endogenous psbA promoter (driving expression of the abundant D1-protein; Zoubenko et al. 1994).
In plastids, there is also a high degree of translational control of gene expression which is in clear contrast to prokaryotes. Therefore, the 5 0 -UTRs and 5 0 -regulatory sequences have proven indispensable for RNA stability and efficient translation initiation. While in Eubacteria virtually all mRNAs contain a Shine–Dalgarno sequence for accurate translation initiation, only 40% of chloroplast mRNAs contain such sequences, indicating that alternative pathways of regulation exist (Hager and Bock 2000). Numerous studies have investigated in detail the effect of swapping different regulatory sequences and determine the consequence of protein accumulation. For example, Eibl et al. (1999) could show that such variations in 5’-UTRs result in up to 100-fold differences in protein accumulation in the case of the reporter gene uidA (encoding beta-glucuronidase). 

Not only the 5 0 -UTR but also the first codons of the open reading frame (the so-called “downstream box”) seems to contribute to translation efficiency (Kuroda and Maliga 2001a, b), demonstrating that high variability of expression levels could be expected for any given sequence and expression cassette. The highest expression level reported so far was over 70% of the total soluble protein (TSP) of phage derived lytic protein driven by the rrn promoter fused to the Escherichia coli phage T7 gene 10 (T7g10) 5 0 -untranslated region (Melanie Oey 2008). This extraordinarily high content of recombinant protein almost exhausted the protein biosynthesis capacity of the chloroplasts and resulted in plants with impaired growth. Although extraordinarily high amounts of protein aspired, several proteins have been reported to be toxic to the plastid at elevated levels (Hennig et al. 2007), and therefore certain threshold levels might exist which should be taken into account to preserve the plants’ viability.

Transgene Stacking and Control of Gene Expression

A demanding task in the generation of transgenic plants is the option of simultaneously introducing two or more genes into an organism. Conventional approaches require the combination of separate expression cassettes each containing a promoter and terminator region framing the gene of interest. In contrast, plastids are thought to offer a unique option of combining multiple ORFs under the control of one promoter, yielding a polycistronic transcript from which translation can be initiated independently (Staub and Maliga 1995). 

However, except for a few examples like the cry2Aa2 operon from Bacillus during genesis (De Cosa et al. 2001; QuesadaVargas et al. 2005), this technique has not been utilized for the simultaneous production of two or more recombinant proteins so far. This is probably due to the hitherto unpredictable secondary structure interactions in polycistronic transcripts, which determine the translatability and processing into monocistronic mRNAs and subsequently result in poor protein accumulation. 

Although the similarities between transcription and translation in bacteria and in plastids are striking, one cannot generally extrapolate results obtained in bacteria to plastids. For instance, an operon encoding for hemoglobin a- and b-subunits, which worked well in E. coli, did not lead to detectable expression when integrated into plastids (Magee et al. 2004). A recent study identified a so-called inter cistronic expression element (IEE), a short sequence that mediates the cleavage of a polycistronic precursor into stable monocistronic transcripts (Zhou et al. 2007). It will be interesting to see whether this novel element leads to concerted high-level expression of recombinant proteins.


  • Antibiotic Resistance Markers
  • Other Selection Markers
  • Marker Gene Excision
  • Analysis

Antibiotic Resistance Markers

For the selection of plastid transformants, aminoglycoside antibiotics have proven highly useful, and especially spectinomycin has become indispensable as a selective marker. Its mode of action is plastid-specific as it binds to the prokaryotic-type plastid ribosomes and inhibits protein synthesis. The 16S rRNA Mutations, one of the target sites of spectinomycin, confer resistance to the antibiotic and early transformation vectors contained such mutant genes. 

An endogenous gene of the mutant is recessive until the homoplasmic stage is reached, transformation efficiency with such marker genes is low. The use of a gene encoding an antibiotic detoxifying protein, namely the aminoglycoside-3 00 -adenyl-transferase (AAD, encoded by the aadA gene) as a marker greatly expedites the development of transplastomic plants.

 Applicability testing for resistance markers in plastid transformation, especially as it was observed that spontaneous spectinomycin resistant lines were formed under strong selection. No spontaneous resistance in higher plants was reported for kanamycin, and thus this particular marker was also adapted for selection of plastid transformants. First successful attempts using the neo gene (encoding for neomycin phosphotransferase II, NPTII) generated transplastomic tobacco, but at a lower efficiency than with aadA. 

Koop and co-workers used the aminoglycoside phosphotransferase gene (APhA-6) gene from Acinetobacter baumannii, which yielded a much higher transformation efficiency (Huang et al. 2002). Both selection markers are in use, even in combination but have not displaced the aadA gene as the most frequently used antibiotic resistance marker.

Other Selection Markers

Another approach is to insert a gene that confers resistance to a toxic compound, for example, herbicides. This approach is widely used for nuclear transformation, and it is thought to be also functional in plastid transformation. Resistance against glyphosate could be obtained via 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS; Ye et al. 2001), and integration of the bar gene made tobacco plants resistant to phosphinothricin (Lutz et al. 2001). However, all expression cassettes also contained the aadA gene for initial selection on spectinomycin and herbicide resistance was only observed when nearly homoplastic plants were obtained.

Marker gene Excision

In principle, any marker gene becomes dispensable as soon as the homoplastic stage is achieved. Sophisticated protocols to subsequently remove the marker flank the sequence to be removed by two directly oriented recombinase target sites. Examples include the 34-bp loxP sites, which are recognized by the Cre recombinase derived from the P1 bacteriophage (Corneille et al. 2001; Hajdukiewicz et al. 32 H. Warzecha and A. Hennig 2001), and the attP (215 bp) and attB (54 bp) sites, recognized by the phiC31 phage integrase Int (Lutz et al. 2004). As long as no integrase is present, the genomes harboring those sites are stable. 

To initiate excision, a second, nuclear transformation step has to be performed. Using a construct in which the recombinase is genetically fused to a chloroplast transit peptide will target the protein to all chloroplasts and initiate site-specific excision of the unwanted genes. Further backcrosses are required where also the nuclear transgene is removed to obtain a marker-free plant line. Several more methods have been described for marker gene removal, and an excellent overview is given by Lutz and Maliga (2007).


Initial plastid transformation and subsequent regeneration of transplastomic plants need to be carefully monitored by different techniques. While the presence of the transgene can be easily checked by PCR, no precise statement about the integration site can be made with gene-specific primers. To rule out an accidental insertion of the expression cassette into the nuclear genome, a PCR with a primer combination bridging the transgene/genome border is advisable. 

Differentiation between the homoplastic and heteroplasmic states can be made by restriction fragment length polymorphism (RFLP) analysis. Restriction digestion of plastid genomes eventually generates fragments of variable length, regarding the integration of the transgene, and therefore resulting in a specific pattern on a subsequent DNA blot hybridized with a specific probe. The gradual disappearance of the wild-type signal and attainment of the homoplastic condition can be monitored with this technique. 

However, the occurrence of promiscuous plastid DNA in the nucleus sometimes feigns a heteroplasmic state and generates a need for isolating chloroplast DNA prior to analysis (Ruf et al. 2000). An unambiguous test for plastid transformation is testing for maternal inheritance of the resistance trait. Transgenic plants pollinated with wild-type pollen generate uniformly green seedlings on a selective medium only if they are homoplastic.


During the past years, tremendous progress has been made in developing and improving plastid transformation techniques and in expanding the methodology to various plant species. Although the method is time-consuming and tedious, it can be established without great effort. Especially for the production of recombinant proteins, plastid transformation has become a valuable tool, notably due to the enormous rate of protein accumulation reported.

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