1	Microprojectile Bombardment
When the difficulties in transforming some plant species first became apparent, a number of procedures that are alternatives to transformation by A. tumefaciens were developed. However, with both the improvement of Ti plasmid-based transformation protocols and the success of microprojectile bombardment, the procedures currently used by nearly all researchers, other methods for transforming plants are now only rarely used. At the present time, essentially any plant can be transformed using one of these two procedures.
Microprojectile bombardment, also called biolistics, is by far the most important alternative to Ti plasmid DNA delivery systems for plants (Table 10.4). Spherical gold or tungsten particles (approximately 0.4 to 1.2 μm in diameter, or about the size of some bacterial cells) are coated with DNA that has been precipitated with CaCl2 , spermidine, or polyethylene glycol (Fig. 10.6). The coated particles are accelerated to high speed (300 to 600 meters/second) with a special apparatus called a particle gun (or gene gun). The original version of the gene gun used a small amount of gunpowder to provide the propelling force. The device that is currently used employs high-pressure helium as the source of particle propulsion (Fig. 10.7). The DNA-coated metal projectiles can penetrate plant cell walls and membranes; however, at the density used, the particles do not significantly damage the cells.
Once inside a cell, the DNA is slowly released from the particles and, in some cells, integrates into the plant genomic DNA. Microprojectile bombardment can be used to introduce foreign DNA into plant cell suspensions, callus cultures, meristematic tissues, immature embryos, protocorms, coleoptiles, and pollen in a wide range of plants, including monocots and conifers, plants that are less susceptible to Agrobacterium-mediated DNA transfer. Furthermore, this method has also been used to deliver genes into chloroplasts and mitochondria, thereby opening up the possibility of introducing exogenous (foreign) genes into these organelles.
Typically, plasmid DNA (here it is not necessary to utilize the Ti plasmid) dissolved in buffer is precipitated onto the surfaces of the microprojectiles. Using this procedure, it is possible to increase the transformation frequency by increasing the amount of plasmid DNA; however, too much plasmid DNA can be inhibitory. It is estimated that, on average, there are approximately 10,000 transformed cells formed per bombardment. With this technique, cells that appear to be transformed, based on the expression of a marker gene, often only transiently express the introduced DNA. Unless the DNA becomes incorporated into the genome of the plant, the foreign DNA will be degraded eventually.
The configuration of the vector that is used for biolistic delivery of foreign genes to plants influences both the integration and expression of those genes. For example, transformation using microprojectile bombardment is more efficient when linear rather than circular DNA is used. Moreover, large plasmids (>10 kb), in contrast to small ones, may become fragmented during microprojectile bombardment and therefore produce lower levels of foreign-gene expression. However, large segments of DNA may be introduced into plants using yeast artificial chromosomes (YACs). The YACs are engineered to contain plant selectable markers, as well as yeast selectable markers, which are already present on the YAC vector (chapter 3). YACs up to 150 kb in total size have a good chance of being transferred to plant cells, and the transferred DNA can be stably integrated into the plant cell. Thus, the production of transgenic plants that contain several foreign genes is feasible; eventually, entire biosynthetic pathways may be introduced into plant cells.
To predictably increase plant fruit yield, it is essential to alter a variety of plant processes. These processes include increasing the ability of the plant to assimilate carbon and nitrogen, transporting the newly synthesized sugars and amino acids to plant tissues, and converting the newly synthesized sugars and amino acids to storage compounds in the sink tissues. This is not an easy task. However, one group of researchers recently reported making multiple genetic modifications to tomato plants to increase the yield of tomato fruit. For a start, researchers needed to monitor the concentrations of a number of plant metabolites from plants grown under greenhouse conditions. This information was used to develop a computer model of the genes that were involved in fruit yield. Based on this model, these researchers introduced 20 separate genes into tomato plants by biolistic cotransformation. Of these genes, 6 were involved in metabolite assimilation, 3 in transport, and 11 in sink metabolism. Moreover, 16 genes were overexpressed and 4 were silenced. Although this procedure only increased the fruit yield by 23%, it was nevertheless quite impressive that so many foreign genes could be successfully introduced by microprojectile bombardment.
Although microprojectile bombardment enables scientists to transform monocots and numerous other plants, some plant varieties remain recalcitrant to this procedure. To overcome this hurdle, scientists overexpressed two plant growth-stimulating genes, Bbm (baby boom) and Wus2 (Wuschel2). The products of both of these genes are considered to be regulators of plant embryogenesis, so their presence makes plants more likely to take up input foreign DNA. A number of different protocols have employed these genes to increase the frequency of transformation of several different monocots. One of those protocols is shown in Fig. 10.8. Some similar procedures employ only the Wus2 gene, and some procedures simultaneously introduce the target gene and the Wus2 (and Bbm) gene(s). With these changes, transformation frequencies of other recalcitrant plants are greatly improved. To date, this approach has been shown to be effective in corn, sorghum, sugarcane, and rice.
2	Chloroplast Engineering
Most higher plants have approximately 50 to 100 chloroplasts per leaf cell, and each chloroplast has about 10 to 100 copies of the chloroplast DNA genome (Fig. 10.9). Thus, while a single copy of a foreign gene inserted into plant chromosomal DNA is present in one copy per plant cell, a single copy of the same gene inserted into chloroplast DNA will be present in 500 to 10,000 copies per plant cell. The many copies of chloroplast DNA in each plant cell means that much higher levels of foreign gene expression should be attainable when the introduced gene is present as part of the chloroplast DNA. Researchers using this approach have reported that a recombinant protein may be expressed to a level of around 1 to 7% of the soluble plant protein. For example, active human somatotropin accumulated to 7% of the total soluble protein in tobacco chloroplasts. This is 10- to 100-fold higher than protein expression levels that are typically obtained when foreign DNA is inserted into the plant genomic DNA.
While the vast majority of plant genes are found as part of the nuclear DNA, both the chloroplast and mitochondrion contain genes that encode a number of important and unique functions. Moreover, not all of the proteins that are present in these organelles are encoded by organellar DNA. For chloroplasts, in particular, some proteins are encoded in the nuclear DNA, synthesized in the cell’s cytoplasm, and then imported into the chloroplast by a specialized transport mechanism. Accordingly, there are two ways that a specific foreign protein can be introduced into the chloroplast. In one approach, a fusion gene encoding the foreign protein and additional amino acids that direct the transport of the protein to the organelle can be inserted into the nuclear chromosomal DNA using the A. tumefaciens– Ti plasmid system and a plant selectable marker (Fig. 10.10A). After the recombinant protein is synthesized, it can be transported into the chloroplast by the chloroplast targeting/transit peptide. This peptide is removed upon insertion of the target protein through the chloroplast membrane. This approach does not necessarily result in very high levels of expression of the target protein.
In the other approach, the gene for the foreign protein can be inserted directly into the chloroplast DNA (Fig. 10.10B). In this case, foreign DNA is typically introduced into the chloroplast genome by microprojectile bombardment of the genetic construct on a plasmid vector. The plasmid generally contains both the foreign DNA and a chloroplast selectable marker flanked by specific chloroplast DNA sequences. Chloroplast selectable markers are designed to be expressed only within chloroplasts and not within the cytoplasm. Homologous recombination is the normal mode of DNA integration into the chloroplast genome; in this case, integration is targeted to a region of the DNA that does not encode proteins (an intergenic region).
The efficient expression of foreign genes in the chloroplast requires an appropriate (chloroplast-specific) promoter sequence and the presence of translational signals in the 5′ and 3′ untranslated regions of the mRNA. Translational control sequences that function only within the chloroplast and not within the cytoplasm are included on the cloned DNA together with the gene of interest and a 3′ chloroplast untranslated region containing a stem-and-loop structure, which is expected to act as a transcription termination signal (Fig. 10.10B). An additional advantage of inserting a target gene into the chloroplast genome is that since chloroplast transcription and translation are prokaryote-like, several genes may be introduced and expressed at the same time in a single operon.
Chloroplast DNA is inherited in a non-Mendelian fashion. In most angiosperms (flowering plants), it is maintained in egg cells but not sperm (pollen) cells and is therefore transmitted uniparentally by the female. Thus, pollen cannot transmit the contents of the chloroplast genome to the zygote. This has practical importance, since this trait can prevent the spread of foreign genes, localized in the chloroplast, through pollen to neighboring plants, thereby addressing one of the concerns of some critics about the cultivation of genetically engineered plants.
Most chloroplast transformation protocols include selectable markers that confer resistance to antibiotics that inhibit protein synthesis on prokaryote-type ribosomes (Table 10.5). These antibiotics inhibit greening (chlorophyll production), the rate of plant proliferation, and shoot formation. Plant cells with transformed chloroplasts that express the antibiotic resistance genes are readily identified in the presence of (otherwise) inhibitory antibiotics by their greening, faster proliferation, and shoot formation. Since only one, or at most a few, of the chloroplasts actually incorporates the foreign DNA, repeated rounds of growth of cultured plant cells on agar plates containing selective antibiotics are often required so that the chloroplast population becomes enriched and eventually dominated by the transformants. Eventually, all chloroplast genomes that do not carry the transgenic DNA with their selectable marker gene are lost.
Chloroplasts belong to a group of plant organelles called plastids, which contain approximately 120 to 180 kb of circular double-stranded DNA bounded by a double membrane. Plastids include nonpigmented organelles called amyloplasts, which synthesize and store starch grains, chloroplasts, which contain chlorophyll, elaioplasts, which contain oil, and chromoplasts, which contain other pigments. While photosynthetic tissues, such as green leaves, contain chloroplasts, other tissues contain other types of plastids. For example, tomato fruit contains a large number of chromoplasts. Thus, in much the same way that foreign DNA may be expressed as a part of the chloroplast genome, it can also be targeted for expression in the chromoplast.
2.1	Very-High-Level Protein Expression
Facilitating the move toward greater use of plants to produce foreign proteins is the promise of very high expression levels following the expression of foreign genes in the chloroplast. With this is in mind, it is worth examining in some detail how one group of researchers was able to obtain an exceptionally high level of synthesis of a therapeutic protein in transgenic tobacco plants.
The target protein for experiments to optimize the expression of therapeutic proteins in plants was a bacteriophage lytic protein that acts as an antibiotic against pathogenic bacteria (group A and group B streptococci). The PlyGBS lysin gene was isolated from a bacteriophage that infects Streptococcus agalactiae, a group B streptococcus. Infections caused by group A and B streptococci are the leading causes of neonatal sepsis (the presence in a newborn baby of a bacterial bloodstream infection) and meningitis (inflammation of the protective membranes covering the brain and spinal cord), and as persistent colonizers of the human genital and gastrointestinal tracts, pathogenic group B streptococci can sometimes be transmitted to a fetus during pregnancy. The PlyGBS lysin kills all known group B streptococcus strains and reduces colonization by these bacteria in both the vagina and the oropharynx (the part of the pharynx between the soft palate and the upper edge of the epiglottis). PlyGBS lysin is also active against group A streptococci such as Streptococcus pyogenes, a human pathogen that colonizes the skin and the mucous membranes of the upper respiratory tract, causing a variety of infections.
To maximize the production of the PlyGBS lysin in transgenic plants, several high-expression strategies were combined (Fig. 10.11). Following puri -cation of the bacteriophage DNA, the plyGBS gene was isolated and its sequence was determined. To ensure that a high level of the PlyGBS lysin could be produced in chloroplasts, the plyGBS gene was completely resynthesized so that its codon usage corresponded to the codon usage found in the AT-rich tobacco chloroplast genome. The synthetic gene was then fused to the strongest known promoter in chloroplasts, i.e., the constitutive rRNA operon promoter. The promoter was followed by the gene 10 leader RNA from phage T7, which provides a strong ribosome-binding site. Next, the plyGBS gene expression cassette was inserted into a chloroplast transformation vector between chloroplast DNA sequences to target the transgene to the intergenic spacer between the trnfM and trnG genes by homologous recombination. Chloroplasts were transformed by microprojectile bombardment of tobacco leaves followed by selection on spectinomycin-containing plant regeneration medium. A spectinomycin resistance gene was used as the chloroplast selectable marker gene (Fig. 10.11).
Transgenic tobacco plants regenerated from the cells of the transformed tobacco leaves expressed an extraordinarily high level of the PlyGBS lysin. Not surprisingly, plants expressing the plyGBS gene cassette were somewhat debilitated in their growth compared to wild-type tobacco plants. This is a reflection of the enormous metabolic load imposed on these plants by the overproduction of the lysin protein. Notwithstanding the slower growth, the plants developed normally and produced viable seeds. When this technology is commercialized, these transgenic plants will likely be confined to a greenhouse environment to avoid release of lysin into the environment that might lead to the development of lysinresistant bacteria.
It has been suggested that for bacteriophage lysins to be effective at killing their bacterial hosts, the lysins must be highly resistant to degradation by bacterial proteases. Thus, at least in part, the extraordinary stability of the lysin protein itself may facilitate its stable accumulation in chloroplasts. Since most other proteins are not as stable as the PlyGBS lysin, it may be difficult to express other foreign proteins in chloroplasts to the extent observed for the PlyGBS lysin. Nevertheless, the strategy of (i) optimizing the codon usage of the foreign gene, (ii) fusing the foreign gene to the rRNA operon promoter and the T7 gene 10 leader peptide, and (iii) expressing the construct in plant chloroplasts should provide an effective means of obtaining relatively high levels of many different foreign proteins.
3	Use of Reporter Genes in Transformed Plant Cells
It is essential to be able to detect the foreign DNA that has been integrated into plant genomic DNA so that those cells that have been transformed can be identified; this requires the use of a selectable marker. Dominant-marker selection provides a direct means of obtaining only transformed cells in culture. For example, in the presence of the antibiotic kanamycin, only plant cells that express a neomycin phosphotransferase gene can grow. Furthermore, in studies of plant transcriptional regulatory signals and the functioning of these signals in specific plant tissues (such as leaves, roots, and flowers), it is often important to be able to quantify the level of expression of a gene with a readily identified product. Quantification and other applications require the use of reporter genes that encode an activity that can be assayed.
A number of different genes have been tested as reporters for transformation, including genes that can be used as dominant selectable markers and genes whose proteins produce a detectable response to a specific assay (Table 10.6). Many of these reporter genes are from bacteria and have been equipped with plant-specific regulatory sequences for expression in plant cells. The desired outcome of a particular experiment often dictates which reporter gene will be used. Clearly, when the expression of a reporter gene interferes with normal plant functions, it cannot be used. Moreover, the presence of some reporter genes and their products may taint a commercial product. In this context, it is best to remove the reporter gene once transformants with the desired traits have been selected, especially in crop plants.
Some reporter gene products, e.g., β-d-glucuronidase (GUS), both -re- +y and bacterial luciferases, and green fluorescent protein (GFP), can be detected in situ in intact plant tissues. One of the most popular of these systems is the E. coli uidA gene, which encodes the protein GUS, a stable enzyme that is not normally present in plants and that catalyzes the cleavage of a range of β-d-glucuronides. The GUS activity in transformed plant tissues can be localized by the presence of a blue color that is formed after the hydrolysis of the uncolored synthetic substrate 5-bromo-4-chloro-3-indolyl β-d-glucuronic acid. Alternatively, GUS activity in plant extracts can be quantitatively and sensitively assayed by a fluorometric analysis that involves the hydrolysis of the substrate 4-methylumbelliferyl β-d-glucuronide to form a fluorescent product.
GFP is an ideal in vivo marker for monitoring transgenic plants because it fluoresces green when excited with either ultraviolet or blue light and does not require the addition of any substrates or cofactors. Normally, wild-type plants (which do not contain the gene for GFP) fluoresce reddish purple when they are excited by ultraviolet or blue light. Moreover, derivatives of the gene that encodes the GFP protein that have different spectral properties and increased levels of fluorescence have been constructed (see chapter 4). Importantly, the expression of GFP has no effect on the survival and growth of transformed plants under field conditions.
4	Manipulation of Gene Expression in Plants
4.1	Transient Gene Expression
The development of transgenic plants that can act as factories for the production of large amounts of therapeutic proteins is a time-consuming, expensive, and inefficient process. Nevertheless, it is possible to achieve high yields of recombinant proteins in plants. Unfortunately, all of the strategies currently available have some limitations. For example, high expression has been reported for nuclear transgenes in seeds; however, because the amount of seed harvested per hectare is much lower than the green biomass harvest (i.e., only a small percentage), the yield of the recombinant protein per hectare is relatively low. Similarly, recombinant proteins can be expressed in high yield in chloroplasts; however, proteins synthesized in chloroplasts are not glycosylated, a situation that is problematic for many therapeutic proteins.
To get around the above-mentioned constraints to the high-level expression of foreign proteins from genes stably integrated into the plant genome, it is possible to transiently express foreign genes in plants in large quantities. Foreign genes that are transiently expressed in plants do not integrate into the plant nuclear or chloroplast DNA. Rather, they exist stably for a limited period (often weeks to months) in the plant cell cytoplasm, typically under the control of strong expression signals such as those found in plant viruses. In this way, the protein may be produced rapidly (transgenic plants do not need to be produced, selected, or propagated) and in high yield. Since the foreign gene is not incorporated into the plant genome, it is not inherited.
With “first-generation viral vectors,” the virus expresses all of its own genes in addition to the target gene, which is spliced into its genome, either under the control of a strong viral promoter or fused to a gene encoding the viral coat protein, which is highly expressed and is present on the surface of assembled viruses (Fig. 10.12). The foreign gene may be delivered to plant cells either using infectious (naked) nucleic acid versions of the vector that are not assembled into a whole virus or as mature viral particles. When mature viral particles are used, the entire process can be performed on a large scale by spraying plants in the field with a mixture of viral particles and an abrasive compound such as carborundum (silicon carbide). Depending on how efficiently the viral particles can move through the plant, it may take about 2 to 3 weeks for most of the tissues of the transformed plants to become infected. This approach has been somewhat successful in that there are numerous reports of plant expression of various therapeutic proteins as fusions to viral coat proteins (Fig. 10.12).
After purification of recombinant vaccine antigens from plants and injection into animals, some of these proteins have been found to be immunogenic and, in some instances, to demonstrate some level of protection against pathogenic agents in animal models, i.e., they have been used as vaccines. Unfortunately, only small epitopes (i.e., fewer than 25 amino acids) have been successfully expressed as coat protein fusions. This is probably because larger epitopes can prevent the viral particle from assembling. To overcome this limitation, scientists constructed “second-generation viral vectors.” In this approach, the recombinant viral DNA is delivered to the plant by A. tumefaciens infection. This avoids the high level of metabolic resources that are devoted to the synthesis of large amounts of viral coat protein and instead facilitates a high level of expression of the target protein. The viral genome is modified so that only the viral elements required for efficient expression of the target gene are maintained (Fig. 10.13). Since the modified viral DNA is delivered to the plant by A. tumefaciens, viral genes responsible for infectivity, amplification/replication, cell-to-cell movement, assembly of viral particles, shutoff of the synthesis of plant cell components, and systemic spread of the virus are dispensable and therefore removed. In addition, by selectively modifying the vector, e.g., (i) introducing silent mutations (mutations that do not alter the encoded amino acids) to remove potential cryptic splice sites, (ii) changing the codon usage, and (iii) adding multiple plant introns, it has been possible to construct more efficient viral delivery systems. The plant introns are thought to make the viral RNA more plant mRNA-like and therefore more efficiently transcribed, processed, and translated. With the improved vectors, there is typically one successful infection event per 10 to 20 copies of A. tumefaciens that are introduced by vacuum infiltration, a process in which the aerial part of the plant is immersed in an A. tumefaciens suspension and then a weak vacuum is then applied for 10 to 30 seconds. Vacuum infiltration replaces the viral functions of infection and systemic movement. Amplification of the viral genome within a plant cell and movement from one plant cell to another are performed by the viral proteins (i.e., replicase and movement protein) that remain encoded in the modified viral DNA. Starting with vacuum infiltration, the entire process takes 4 to 10 days before recombinant protein can be isolated from plant leaves. Moreover, the approach is effective with a range of plants. It is estimated that in a 1-hectare plant growth facility, this system may be able to produce up to 400 kilograms of recombinant protein per year. Since this system does not require a large amount of land, growth of transformed plants could be limited to a contained greenhouse facility.
One group of workers demonstrated that with a transient expression system, it was possible to dip a head of lettuce, obtained from a grocery store, into an A. tumefaciens cell suspension, vacuum infiltrate the bacterium (carrying the engineered viral DNA), and within about 1 week’s time obtain around 20 to 100 milligrams of functional target protein per kilogram of fresh lettuce leaf tissue (Fig. 10.14). With this system, plant growth facilities are not required; the system is very efficient and inexpensive and is easily scaled up. This transient-expression system, based on expression from viral RNA replicons (where an RNA replicon is RNA that replicates from a single origin of replication) delivered into plant cells by A. tumefaciens, allows production of recombinant proteins at yields of up to 5 grams per kilogram of fresh leaf biomass. Such high yields are possible because plant biomass accumulation takes place prior to infection.
Notwithstanding the high yields of foreign protein that can be obtained, this approach requires special equipment for vacuum infiltration and may be expensive to perform on a large scale. In addition, containment of the genetically engineered agrobacteria producing modified viral replicons that are used for transfection is required. To avoid these problems, researchers have developed an ethanol-inducible system that provides efficient release of viral RNA replicons from proreplicons (which lack a replication gene) contained in a stably transformed cassette (Fig. 10.15A). The cassette carrying the viral vector has two components, the viral replicon and a gene encoding the movement protein, and each of these components is placed separately under the transcriptional control of an ethanol-inducible promoter. In this system, the viral replicon and the movement protein are under the control of the alcohol dehydrogenase gene (alc) promoter from the fungus Aspergillus nidulans. The alc promoter is activated only when it is bound by the AlcR transcriptional activator, encoded on a separate construct, in the presence of ethanol (Fig. 10.15B). In practice, to activate the production of a target protein, transgenic plants carrying the two constructs shown in Fig. 10.15 are sprayed twice a day for around 5 days with a solution of 4% ethanol. Alternatively, expression of the target gene was strongly induced when the roots were soaked and then sprayed once with 4% ethanol. Ethanol was chosen as the inducer in this system because it is relatively inexpensive and is nontoxic and because the system exhibits a very low background level of expression in the absence of the inducer. This system should be readily amenable to scale-up. What remains is to determine whether this system is as effective and reliable in practice as it seems.
4.1.1	Transient Expression of Antibodies
The production of human therapeutic agents, especially monoclonal antibodies, is very expensive, requiring large investments in specialized equipment with skilled personnel to run it and significant operating costs. However, with the enormous health benefits of these therapeutic agents, there is widespread demand for more inexpensive versions of these highly efficacious therapeutic molecules which can be achieved in relatively inexpensive plant-based systems. For example, to produce humanized monoclonal antibodies in the tobacco-like plant Nicotiana benthamiana, the following steps were undertaken (Fig. 10.16). Starting with isolated cDNA, the nucleotides encoding the amino acid sequence of a mouse monoclonal antibody directed against Zika virus were altered to (i) humanize the monoclonal antibody (removing all mouse DNA sequences) and (ii) optimize its codon expression in plants. The resynthesized cDNAs were spliced into geminivirus expression vectors (second-generation transient expression vectors), which were delivered to plant leaves using Agrobacterium. The strain of N. benthamiana that was used was previously engineered so that plant-specific β1,2-linked xylose and α1,3-linked fucose molecules were no longer added to the glycans on the surface of the foreign monoclonal antibody. This monoclonal antibody molecule was produced at a level of ~1.5 grams per kilogram of leaf tissue and was correctly assembled so that it recognized the Zika virus envelope protein and neutralized the Zika virus. Using affinity chromatography, this monoclonal antibody could be purified to near homogeneity in a single step (see chapter 3). This is an important demonstration of the ability of plant systems to be an effective and relatively inexpensive tool for the production of multiprotein human therapeutic proteins.
4.2	Plant Promoters
Many early experiments to express foreign proteins utilized promoters that were constitutively active in a wide range of plant cells. However, in order to minimize any deleterious effects on plant growth from the expression of foreign genes, it is necessary to regulate the expression of introduced genes, spatially and temporally, and the amount of the foreign protein that is produced. More recently, many additional plant promoters have been isolated and characterized and used to express foreign proteins in specific cells at certain times during the growth and development of the plant. For example, instead of the strong constitutive 35S promoter from cauliflower mosaic virus, which is expressed in all dicot plant tissues (and to a lesser extent in monocot plant tissues) and throughout the life of the plant, researchers have used the strong promoter for the gene encoding the small subunit of the photosynthetic enzyme ribulose bisphosphate carboxylase, which is active only in photosynthetic (green) tissues, such as leaves. Similarly, plant promoters active only in specific tissues, such as roots, seeds, or #owers, or only during periods of environmental stress—e.g., the pathogenesisrelated promoters—have been used to control the expression of some foreign genes.
While a major factor in the initiation of transcription in plants is the binding of RNA polymerase II to the promoter sequence, other factors also affect this process. Much of the specificity of transcription in plants is controlled by sequence-specific transcription factors and/or enhancerbinding proteins (Fig. 10.17). Enhancer elements are regions on the DNA that bind to enhancer-binding proteins, which interact with transcription factors and RNA polymerase to increase the level of transcription from a particular promoter. Enhancer sequences are generally considered to be important determinants of the tissue and temporal specificity of gene expression. While the importance of promoter and enhancer sequences has been recognized, a thorough understanding of how these and other elements regulate plant gene expression is still not completely available. Nevertheless, using a variety of different promoters, scientists have been able to engineer both tissue-specific gene expression and high levels of gene expression.
4.3	Manipulation of Genes in Plants
The targeted editing of a plant genome could offer tremendous advantages over traditional methods of plant breeding to create new cultivars with advantageous combinations of alleles, especially when several traits are needed for crop improvement. Traditional methods of plant breeding are designed to combine desirable alleles of different genes; they involve extremely tedious and time-consuming genetic crosses and selection for the sought-after phenotype, and often, as with closely linked loci, traditional methods cannot be used to produce the desired result. While the ability to create transgenic plants has solved some of these problems, consumers and regulators in a number of countries, especially in Europe, have expressed serious concerns and reservations about their commercialization and widespread use. On the other hand, in situ modification of an existing plant gene is currently not considered to be genetic engineering in many countries and therefore should allow some specifically altered plants to more rapidly reach the marketplace in those countries. With the targeted stable modification of plant genomes, no foreign DNA is introduced into the plant. Some recently developed and perfected techniques, including the use of zinc finger nucleases, transcription activator-like effector nucleases (TALENs), and the clustered regularly interspaced short palindromic repeat CRISPR/Cas9 system have dramatically increased the ease with which it is possible to target and modify specific plant genes.
The most promising of the three above-mentioned gene-targeting systems (and also the newest) exploits the bacterial CRISPR/Cas system (see chapter 2). The simplest form of this technology consists of a single guide RNA molecule (sgRNA) where part of the molecule matches the target DNA sequence and an endonuclease from the bacterium Streptococcus pyogenes (Cas9). The sgRNA and the Cas9 endonuclease form a complex in which the sgRNA guides the complex to its target DNA and the nuclease activity of Cas9 then cleaves the target DNA in both strands. With the CRISPR/Cas9 system, only the 20 nucleotides in the sgRNA need to be changed to confer different target specificities, and therefore it is possible to alter several genes at the same time, knocking out redundant or parallel pathways.
The first reports of the use of the CRISPR/Cas9 system to alter plant genomic DNA were published in mid-2013. Since that time, there have been hundreds of reports of the use of this technology to alter the genomes of a variety of plants (Table 10.7). The CRISPR/Cas9 system is used to create single-base-pair deletions and whole or partial gene knockouts and to introduce gene modifications, gene fragment deletions, and gene insertions (Fig. 10.18). Given the very rapid pace of the development of this technology, one can envision a wide range of potentially commercializable CRISPR/Cas9-modified plants being developed within the next 5 to 10 years. Initially, CRISPR/Cas9-modified plants had fewer regulatory hurdles to overcome in Europe, as they were viewed as not being genetically modified organisms. However, on 25 July 2018, the Court of Justice of the European Union ruled that crops developed using this technology must be treated the same way as other genetically modified organisms. In a different venue, in 2014, the New Zealand Environmental Protection Agency ruled that plants produced by gene-editing methods, where no foreign DNA was introduced into the edited plant, were not to be regulated as genetically modified organisms; however, a subsequent court challenge overturned this ruling.
For the most part, sgRNA and Cas9 coding sequences have been introduced into plant genomes by either Agrobacterium-mediated transformation or by microprojectile bombardment of callus cultures followed by plant regeneration. This limits this technology to those cultivars that are amenable to tissue culture, significantly restricting the use of this approach. To circumvent this limitation, one group of researchers developed a microprojectile bombardment-based delivery system that enabled the components of the CRISPR/Cas9 system to be transiently expressed to edit genomes in wheat embryos (Fig. 10.19), thereby completely avoiding tissue culture and plant regeneration. In this case, gold particles coated with CRISPR/Cas9 components (and GFP as a marker gene) were delivered to wheat seed embryos. The bombarded embryos that exhibited transient GFP expression were subsequently grown into adult plants. Subsequent analysis of these plants by PCR for the target gene (i.e., targeted by the sgRNA) indicated that 5.2% of the selected embryos carried mutant (changed) alleles, with one-third of those plants inheriting the mutation in the next generation. Moreover, in no instances was the gene encoding the Cas9 enzyme integrated into the genomic DNA. It is suggested that this method should be useful in editing the genomes of plant species and cultivars that are otherwise recalcitrant to this procedure because they cannot be grown in culture and subsequently regenerated.
4.3.1	Clonal Seeds from Hybrid Rice
Hybrid plants are typically constructed from genetically different plant varieties in order to increase their vigor relative to their homozygous parents. Commercial hybrid plants are constructed to stably reproduce a valued attribute of each initial variety into the offspring. Typically, commercial hybrids are developed to improve traits such as disease resistance, plant size, #ower color, fruit size, or taste. Unfortunately, hybrid seed production is a relatively slow process and is quite expensive for many crops. Generally, hybrids are produced by transferring the pollen from one plant to the pistil of another plant. To prevent plants from self-pollinating, all of the stamens (the pollen-producing organs) must be removed from the plants that are going to be pollinated. Finding the desired hybrid may take years of constructing and testing genetic hybrids. In a recent effort to avoid the painstaking process of creating genetic hybrids with the desired traits, scientists, using the CRISPR/Cas9 system, simultaneously created mutants of several specific rice genes, including REC8, PAIR1, OSD1, and MTL. In this case, the genetically modified rice strains behaved identically to the initial wild-type strain. However, following the self-propagation of F1 hybrids it was observed that the resultant plants could propagate clonally through seeds, a trait not usually associated with hybrid plants. Thus, the introduced genetic modifications had turned meiosis into a mitosislike cell division (Fig. 10.20). Importantly, the genes that were targeted for modification in rice are highly conserved in numerous other plants so that this method of producing stable hybrid plants should be applicable to a wide range of crops.
4.3.2	Conferring Male Sterility
Hybrid plant seed is typically produced from a genetic cross between two genetically different plant lines. The hybrid seeds usually result in larger, more vigorous plants (hence the term hybrid vigor). To facilitate the production of hybrid seeds, scientists often use plants that are cytoplasmically male sterile. Male sterile plants do not produce any functional pollen, although they do produce viable eggs. In addition, in jurisdictions where there is significant concern regarding the possible unintentional widespread dissemination of transgenic plants, the employment of male sterile transgenic plants that do not produce functional pollen (and hence do not spread their DNA) clearly obviates those concerns. Recently, a group of scientists reported using TALEN-mediated genome editing to knock out six (nearly identical) CmDMC1 genes in chrysanthemum; this plant is a hexaploid species that contains six copies of the entire plant genome. The CmDMC1 gene is involved in meiotic homologous recombination. Interestingly, the transgenic plants in which all six of these genes were knocked out exhibited both male and female sterility. Under the Cartagena Protocol on Biosafety, ornamentals must be unable to outcross with wild relatives prior to their approval for practical use. Thus, this manipulation obviates the difficulty of producing male sterile plants when the plants (such as chrysanthemum) are polyploid and may become a widely used approach to prevent the #ow of transgenes from genetically engineered plants to their wild relatives.
4.4	Facilitating Protein Purification
Transgenic plants have some advantages over bacteria and animal cells as expression systems for foreign proteins. For example, transgenic bacteria and animal cells are typically grown in expensive bioreactors under precisely defined conditions and require highly skilled personnel to oversee the entire growth process. On the other hand, plants may be grown in the field or in greenhouses by somewhat less highly skilled individuals at a much lower cost and in more or less unlimited quantities. Thus, one of the main advantages of expressing a foreign protein in a transgenic plant is that it is relatively inexpensive to grow plants on a large scale. For commercial production of a recombinant protein in any host organism, following growth, the organisms must be harvested and processed, and then the recombinant protein is purified. Protocols for the purification of proteins from transgenic bacteria, yeast, and animal cells are well established, and many proteins produced by bacteria and animal cells are currently in the marketplace. However, transgenic plants generally produce a much lower level of foreign proteins than bacteria and yeast; therefore, for commercial production of a range of protein products, the purification of target proteins produced by transgenic plants may require special strategies and approaches.
4.4.1	Oleosins
A novel way to facilitate recombinant protein purification from plants is to fuse the foreign protein to plant oleosins. Oleosins, or oil body proteins, are found in the seeds of a wide range of plants. These proteins are quite hydrophobic and are mostly found embedded within small oil droplets (0.6 to 2 mm in diameter) called plant oil bodies, thus stabilizing the oil bodies as discrete intracellular organelles for storing neutral lipids. However, the N- and C-terminal regions of oleosins are more hydrophilic than the rest of the protein and are exposed to the aqueous environment. It is therefore possible to engineer fusions between oleosins and watersoluble proteins at the DNA level (Fig. 10.21), with the expectation that the fusion protein will be targeted to plant oil bodies, making it relatively easy to purify. In this case, the water-soluble target protein will not be embedded in the oil body but, rather, will be exposed to the aqueous environment. Since it is relatively straightforward to purify oil bodies from plant seeds, the purification of the recombinant protein is simplified. A cleavable linker is included between the oleosin and the target protein so that the recombinant protein can be recovered by cleavage of the fusion protein. Expression in seeds is particularly attractive, as proteins accumulate stably and seeds can be stored without deterioration prior to being processed. This system has the potential to significantly lower the costs of purifying target proteins produced in plant seeds.
4.4.2	Rhizosecretion
Harvesting the variety of protein products that can be produced in transgenic plants can be difficult, since they are generally localized within plant cells that must be disrupted before their purification. Moreover, since the cost of purifying a recombinant protein can be as much as 90% of the total cost of producing that protein, if plants are going to be used as costeffective bioreactors for protein production, it is essential that purification costs be kept to a minimum. One way around this problem is to engineer plants to secrete foreign proteins through the roots in a process that has been called “rhizosecretion.” If a plant engineered for rhizosecretion is grown hydroponically, the protein will be secreted directly into the culture medium (Fig. 10.22).
Normally, roots secrete large amounts of small molecules, such as sugars and amino acids; however, they secrete only low levels of relatively few proteins. The small organic molecules are first secreted to the root intercellular space (apoplast) before they are exuded/secreted by the roots. In one series of experiments, three different proteins—xylanase from the thermophilic anaerobic Gram-positive bacterium Clostridium thermocellum, GFP from the jellyfish Aequorea victoria, and human placental secreted alkaline phosphatase—were tested to determine whether they could be engineered for secretion through the roots of transgenic plants. The three proteins were directed to the root apoplast using three different secretion signals (Fig. 10.23). Each protein was efficiently exuded by the roots of transgenic tobacco plants, as long as the genetic construct contained a DNA sequence encoding a signal peptide, even one that was not of plant origin, placed upstream (at the 5′ end) of the gene whose protein was targeted for secretion. Both the 35S promoter, which is expressed in all plant cell types, and the mas2′ promoter, which is preferentially expressed in roots, directed the synthesis of a significant amount of the target protein in root tissue. With the 35S promoter, the foreign protein could also be recovered from the guttation #uid (i.e., the leaf exudate). However, given the ease of collecting root as opposed to leaf exudate, this approach appears to have the most promise at the present time.
4.5	Protein Glycosylation
A large number of mammalian proteins, including many potentially therapeutic molecules, are glycosylated (i.e., they contain specific sugars attached to the hydroxyl group of either serine or threonine or the amide group of asparagine) (see chapter 3). While the addition of glycans (polysaccharides) containing high levels of mannose at specific asparagine residues on proteins is initially identical in mammalian and plant cells, subsequent trimming of the sugar residues generates complex N-glycans with very different structures and properties in the two different types of organisms. Although differences in glycosylation may not directly alter the activity of a protein, other properties, such as protein folding, stability, solubility, susceptibility to proteases, blood clearance rate, and antigenicity, can be affected profoundly. Thus, especially for monoclonal antibodies, glycosylation of the protein is considered to be a critical attribute of the final product.
To avoid some of the problems that result from the incorrect glycosylation of mammalian proteins that are produced in plant cells, it is possible to modify the plant so that it does not add “problematic” carbohydrate residues. For example, when the aquatic plant Lemna minor was engineered to produce light and heavy chains that are part of a human monoclonal antibody, the plant was simultaneously transformed with an RNA interference construct (see chapter 6) that specifically inhibited the expression of the plant glycan modification enzymes α-1,3-fucosyltransferase and β-1,2-xylosyltransferase. The antibodies that were produced by these transgenic plants contained a single major N-glycan species without any plant-specific N-glycan residues (Fig. 10.24A). While these truncated glycan side chains are not identical to the glycans that are produced when a human monoclonal antibody is synthesized in Chinese hamster ovary (CHO) cells in culture, they share the same core molecules. In fact, when human monoclonal antibodies produced in genetically modified plants were tested, they displayed better antibodydependent cell-mediated cytotoxicity and effector cell receptor-binding activities than antibodies that were produced in CHO cells. Thus, with this system, it may be possible to produce a wide range of mammalian proteins in which the glycan side chains no longer limit the use of the final protein. In addition, it should be much less expensive to produce mammalian proteins in transgenic plants than in mammalian cell culture.
To further improve this system, the gene encoding the enzyme endoN-acetyl-β-d-glucosaminidase (endo T) from the fungus Hypocrea jecorina was inserted into the genome of the plant Arabidopsis thaliana under the transcriptional control of a seed-specific promoter from Phaseolus vulgaris (common bean), and the enzyme was localized to the Golgi (of seed cells), where many proteins are glycosylated. The A. thaliana strain that was used was a mutant that lacked the enzyme N-acetylglucosaminyltransferase-I; this resulted in the accumulation of proteins with high mannose glycan side chains that are able to act as substrates for the enzyme endo T. Endo T hydrolyzed the bonds between adjacent N-acetylglucosamine residues in the glycans of some proteins, leaving only the innermost Nacetylglucosamine residue (Fig. 10.24B). This system, which has been called GlycoDelete, yields proteins with only a single N-acetylglucosamine residue attached to the amide group of asparagine. This reduces any difficulties in purifying the target protein that might otherwise arise as a consequence of heterogeneous glycosylation (with differently glycosylated proteins having somewhat different physical and biochemical properties) and does not interfere with the functioning of the target proteins. Moreover, the developers of this technology argue that plant-produced pharmaceuticals that are synthesized in seeds could be stored in a stable manner as part of the seed until the protein product is needed.
The technology for producing uniform Fc portions of monoclonal antibodies (which are all glycosylated) currently exists for a number of different plants. This enables enhanced Fc effector functions, greater consistency between batches, decreased risks related to the immunogenicity of the therapeutic monoclonal antibody, and the possibility for economically manufacturing monoclonal antibodies in plants. This approach should facilitate and significantly reduce the cost of monoclonal antibody production. In fact, glycoengineered monoclonal antibodies produced in plants have recently begun clinical trials.
4.6	Gene Stacking
In order to develop transgenic plants that have increased commercial value, it is often necessary to introduce more than a single foreign gene into plants. Currently, the two most commonly introduced traits in commercially available transgenic plants are insect resistance and herbicide tolerance, each of which may be encoded in a single gene (see chapter 11). To produce transgenic plants that are both insect resistant and herbicide tolerant requires plants to contain at least two transgenes. This stacked transgenic plant enables farmers to have a crop that is simultaneously insect resistant and herbicide tolerant. In fact, there are numerous commercial examples of transgenic plants that have stacked genes. In the future, crops that contain multiple transgenes are expected to become quite common. There are several methods that can be used to produce stacked transgenic plants (Table 10.8). These methods include hybrid stacking, cotransformation, multigene cassette transformation, and retransformation.
With expanding populations worldwide, there has been a substantial loss of arable land necessary to feed this increasing population. One possible solution to this problem is to grow more food in urban environments. However, this approach is fraught with problems. Nevertheless, by judicious modification of the genomes of some plants, it may be possible to grow them to a smaller size than is normally the case on much smaller plots of land without decreasing the final crop yield. Recently, a group of scientists reported using a gene stacking strategy to generate and combine mutations in cherry tomatoes for condensed shoots, rapid #owering, and precocious growth termination. These mutations were all produced using CRISPR/Cas9 genome editing and produced tomatoes plants that were compact and early-yielding plants that were deemed suitable for urban agriculture. The engineered plants were then grown in hydroponic vertical farms and compared to wild-type plants grown in the same way. Although the yield of fruits per plant on the smaller genetically manipulated plants was slightly lower than with larger wild-type plants, this reduction in yield was readily compensated for by the fact that more, smaller genetically manipulated plants could be grown in the same space than the larger wild-type plants. Moreover, editing the same set of three genes in groundcherry led to a similar result, suggesting that this approach may be effective with a number of different crops.
4.7	CRISPR-Based Directed Evolution
The process of directed evolution involves introducing random mutations into a target gene in a host organism and then screening for proteins with potentially superior performance. In prokaryotes, directed evolution is often achieved by error-prone PCR of a target gene followed by extensive screening and testing (see chapter 3). However, this technique, which is highly effective in prokaryotes, is impractical in plants. Therefore, some researchers have developed the ability to produce random mutations in targeted genes in plants using CRISPR/Cas9. In this instance, the Cas9 endonuclease generates double-strand breaks within a region of the target gene defined by the sgRNA, and these breaks are then repaired by nonhomologous end joining, a procedure that generates small insertions, small deletions, or base substitutions. This procedure may also be performed using a nickase Cas9 enzyme that cuts only one of the two target DNA strands. In addition to generating random mutations in one portion of a gene, it is possible to simultaneously target two or more regions of the same gene using two or more sgRNAs. CRISPR-based directed evolution is a technique that is still in its infancy; however, it is currently being used in attempts to alter a wide variety of traits in several different plants in a rapid and simple manner.
4.8	Polycistronic Gene Expression
Plants and other eukaryotes are monocistronic, where each gene is under the transcriptional control of a separate promoter. This makes it extremely difficult for scientists to genetically transform plants with several genes that are part of the same pathway and ensure that these genes are transcribed and translated to the same extent (e.g., as part of a metabolic pathway). For example, multiple rounds of genetic transformation to introduce several genes (even if those genes contain the same transcriptional promoter) generally result in those genes being inserted in different portions of the plant chromosomal DNA and therefore expressed at differing levels due to the influence of the epigenetic environment. However, scientists have recently developed systems that can get around this roadblock. Using synthetic biology, where the desired genes are synthesized and then logically assembled (see chapter 2), researchers were able to assemble multiple transcription units in a single DNA fragment, each with its own promoter, transcription terminator, and signal peptide-coding sequence for secretion of the produced protein (Fig. 10.25A). Those transcription units are separated by short linker sequences of nucleotides encoding 18 to 22 amino acids, known as 2A peptides, which are self-cleaving peptide sequences (Fig. 10.25B). These 2A sequences (peptides) were first discovered in the eukaryotic footand-mouth disease virus and function by inducing ribosomal skipping during protein translation between Pro and Gly in the C-terminal end of the peptide, where the amino acid sequence of the 2A peptide from foot-and mouth disease is Val-Lys-Gln-Thr-Leu-Asn-Phe-Asp-Leu-Leu-Lys-LeuAla-Gly-Asp-Val-Glu-Ser-Asp-Pro-Gly-Pro. Similar self-cleaving sequences have been found in a wide range of viral families, with different 2A peptides having different efficiencies of self-cleavage. With this system, comparable levels of transgene expression can be expected regardless of the gene positions in the polycistronic expression cassette. It now remains to be determined whether this system is a useful and effective means of introducing entire functional metabolic pathways into plants.
5	Production of Marker-Free Transgenic Plants
Usually, at the same time that a foreign gene is introduced into plants, a selectable marker gene is also introduced. Although none of these marker genes or their products have to date been shown to have an adverse effect on human, animal, or environmental safety, their inclusion in transgenic plants has raised some concerns. For example, it is possible that the products of some marker genes might be either toxic or allergenic. Also, the antibiotic resistance genes that are often used as selectable markers might be transferred to pathogenic soil microorganisms. While gene transfer from plants to environmental or medically related bacteria, or from plant products consumed as food to intestinal microorganisms or human cells, is generally considered to be highly unlikely (see chapter 13), there are nevertheless some inherent risks that have not been totally addressed, and therefore both regulatory and public concerns remain in many places in the world. Moreover, the presence of a selectable marker makes it technically more difficult to transform a transgenic plant with additional foreign genes, since the same selectable marker cannot be used more than once. To allay these concerns, strategies for the production of transgenic plants without any marker genes have been developed.
5.1	Removing Marker Genes from Nuclear DNA
One experimental approach that has been used to produce markerless transgenic plants includes cotransformation of plants with two separate DNAs, one carrying the marker gene and the other carrying the target foreign gene. With cotransformation, approximately 30 to 80% of the segregation during a few rounds of matings. In this way, the selectable marker is removed from the plant genome during the segregation and recombination that occur during sexual reproduction. At present, this strategy is the most widely used approach to produce marker-free transgenic plants.
As an alternative to cotransformation with two separate DNAs, a selectable marker gene may be cloned between two plant transposable elements (Ds elements) and inserted into the T-DNA along with the target gene and a transposase gene that, after plant transformation, excises the DNA between the Ds elements and integrates it at another chromosomal site (Fig. 10.27A). Any sequence that is between two Ds elements can be mobilized to a new location in the genome—excising the gene from the original location—provided that the appropriate transposase enzyme is available. Following insertion of the T-DNA into the host plant DNA, about 90% of the time, the selectable marker that is between the two Ds elements will be moved to another site on the chromosomal DNA. About half of the time, the new location for the selectable marker will be far away from the original location. Thus, a selectable marker gene can be used to identify transformed plant cells, and subsequently, it can be removed by breeding.
Any procedure that uses sexual crossing to segregate the selectable marker from the gene of interest cannot be applied to woody plants (because of their very long generation times), vegetatively propagated plants (where new plants grow from parts of the parent plant rather than from seeds), or sterile plants. One simple way around this problem is to #ank the selectable marker with specific DNA sequences that mark it for eventual removal from the genome (Fig. 10.27B). In this way, the selectable marker and #anking sequences are only transiently present as a component of the transgenic plant. For example, if the selectable marker gene is fused to a recombinase gene and #anked by sequences that are recognized by the recombinase, both from the yeast Zygosaccharomyces rouxii, the selectable marker gene will be initially expressed. However, the recombinase is also expressed and mediates excision of the region between its recognition sites, thus removing both the selectable marker and recombinase genes after growth in culture for several months. In this case, transformed plant cells are selected immediately following transformation, characterized to ascertain whether they contain and express the target gene, and then grown in culture until the selectable marker has been spontaneously excised. In addition to the system outlined here, several similar excision systems that result in the removal of the selectable marker have been developed. Importantly, regardless of the details, it is no longer necessary to produce transgenic plants that retain antibiotic resistance genes or other undesirable selectable markers.
In the selection of genetically modified plants following genome editing by the CRISPR/Cas9 methodology, it is necessary to distinguish modified from unmodified plants. This is a straightforward procedure if the modified plants carry a selectable marker. Procedures similar to those described above may be employed to remove the selectable markers from both transgenic plants and CRISPR/Cas-edited plants. Unfortunately, many of the procedures that have been developed to eliminate the selectable markers (typically antibiotic- or herbicideresistant genes) in transgenic plants require growing the plants for two or more generations, where a generation is typically 4 to 12 months, depending on the plant. To shorten the time that is required to select marker-free plants following genome modification by CRISPR/Cas, one group of researchers used a gene encoding Discosoma (a genus of anemone) red fluorescent protein, i.e., DsRED, as a selectable marker (Fig. 10.28). Following CRISPR/Cas modification of rice, tomato, and Arabidopsis plants, DsRED fluorescence was readily and rapidly detected in plant seeds. With this fluorescent marker, transgene-free homozygous genome-edited crops could be detected within a single generation following in vitro transformation.
Any system used to remove the selectable marker generally requires growing the transformed plants for at least one and up to several generations and then testing to ensure that the selectable marker has been removed. To speed up this somewhat tedious process, one group of researchers did not include a selectable marker as part of their T-DNA construct but instead included a bacterial β-glucuronidase reporter gene, a gene that is not found in plants. The enzyme β -glucuronidase (often called GUS) can cleave the colorless substrate 5-bromo-4-chloro-3-indolyl glucuronide (X-Gluc; Fig. 10.29), a compound that turns blue at the site of GUS activity in the plant. Leaves or other tissues from putative transgenic plants may be easily tested to see whether they contain GUS activity, since upon cleavage, the colorless substrate turns an insoluble blue color that results from the synthesis of 5,5 ′ -dibromo-4,4 ′ -dichloro-indigo. Thus, putative transgenic cotton plants were identified by an in situ GUS histochemical assay of pollen grains, immature embryos, and tissues from putative firstgeneration transgenic plants, with 0.2 to 0.7% of the plants tested being transgenic.
5.2	Removing Marker Genes from Chloroplast DNA
The possibility of high levels of foreign-gene expression, as well as expression of several related genes under the control of the same promoter (e.g., encoding a biochemical pathway), has resulted in an increasing number of foreign genes being introduced into plant chloroplast DNA. The foreign genes are typically introduced as part of a genetic construct that includes a selectable bacterial gene, e.g., aadA, which confers resistance to the antibiotics spectinomycin and streptomycin. For removal of the marker gene, it is #anked by directly repeating DNA sequences (174 bp). Following growth of the transformed cells in the absence of selective pressure, the selectable gene will be excised by homologous recombination between the direct repeat sequences (Fig. 10.30).
An alternative approach is to develop selectable markers that do not encode antibiotic resistance or any other undesirable trait. One possible selectable marker of this type is the spinach gene for the enzyme betaine aldehyde dehydrogenase. This enzyme, which is present in the chloroplasts of a limited number of plants, converts the toxic betaine aldehyde to the nontoxic glycine betaine. In addition, glycine betaine acts as an osmoprotectant, conferring some measure of salt or drought resistance. When resistance to betaine aldehyde was used to select transgenic plants, the transformation efficiency was approximately 25-fold higher than when transformants were selected with spectinomycin. The use of this approach should facilitate the chloroplast transformation of many important crops, including cereals that are naturally resistant to spectinomycin.

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