Environmental Biotechnology PDF

**Environmental Biotechnology** **INTRODUCTION** The environment around us has always been a source of new products and stimulated our imagination in developing new technologies. Our species is very successful at harnessing the environment for our benefit. We are also good at destroying or harming...

**Environmental Biotechnology** **INTRODUCTION** The environment around us has always been a source of new products and stimulated our imagination in developing new technologies. Our species is very successful at harnessing the environment for our benefit. We are also good at destroying or harming the environment for immediate gains. The readily visible world has been charted and mapped, but areas under the ocean and in the deep recesses of jungles are still unknown. In fact, many parts of the visible world still harbor unknown life forms invisible to the naked eye, including bacteria and viruses. These are found in the air, water, and land. Many have unique metabolisms, and some have abilities never seen before. Many can live in extreme environments, once thought too hot or too dry for life to exist. Estimates predict that about 10^31^ to 10^32^ viral particles are present in the biosphere, an order of magnitude more than host cells. The **virosphere**, as it is sometimes known, is probably one of the biggest sources of novel genes. At the time of writing, only about 0.1% to 1% of microorganisms have been cultured. Even the majority of those found growing at moderate temperatures in soil or other normal habitats have not been cultured. In addition to DNA inside life forms, there is much free DNA in the environment that might also be a source of new genes. The field of environmental biotechnology has revolutionized the study of these previously hidden life forms and DNA. What kinds of secrets do they harbor? What kinds of new enzymes and proteins can be identified? Molecular biology techniques are now being applied directly to the environment to investigate the uncultured viruses and bacteria. PCR is routinely used to amplify random sequences from many environmental samples in the hope of identifying new genes. After PCR, the DNA is sequenced. Then bioinformatics reveals whether or not the sequence (or a close relative) has already been identified or if it is completely novel. Microarrays are also being created to compare the numbers and types of organisms present in different environments. Almost every recombinant DNA methodology discussed in the first half of this book can be applied to environmental samples. Environmental biotechnology is divided into different areas. These include direct studies of the environment, research with a focus on applications to the environment, or research that applies information from the environment to other venues. This chapter focuses on direct analyses of the environment and the natural biochemical processes that are present, whereas upcoming chapters cover research with environmental applications or results from environmental research with practical applications. Surveying different environments may identify new life forms, new metabolic pathways, or novel individual genes. Genomics techniques have revolutionized this field, and it is rapidly expanding. **IDENTIFYING NEW GENES WITH METAGENOMICS** **Metagenomics **is the study of the genomes of whole communities of microscopic life** **forms. Approaches include shotgun DNA sequencing, PCR, RT-PCR, and other genetic methods. Metagenomic research sometimes allows us to identify microorganisms, viruses, or free DNA that exist in the natural environment by identifying genes or DNA sequences from the organisms. Metagenomics applies the knowledge that all creatures contain nucleic acids that encode various protein products; therefore, organisms do not have to be cultured, but can be identified by a particular gene sequence, protein, or metabolite. The term *meta*, meaning more comprehensive, is also used in *meta-analysis*, which is the process of statistically combining separate analyses. Metagenomics is the same as genomics in its approach. The difference between genomics and metagenomics is the nature of the sample. Genomics focuses on one organism, whereas metagenomics deals with a mixture of DNA from multiple organisms, "gene creatures" (i.e., viruses, viroids, plasmids, etc.), and/or free DNA. Most microorganisms have never been cultured or previously identified. Using metagenomics, researchers investigate, catalogue, and analyze the current microbial diversity. They identify new proteins, enzymes, and biochemical pathways. They also hope to provide insight into the properties and functions of the new organisms. The knowledge garnered from metagenomics has the potential to affect how we use the environment to our benefit or harm. Metagenomics has been used to identify new beneficial genes from the environment, such as novel antibiotics, enzymes that biodegrade pollutants, and enzymes that make novel products (Table 12.1). Historically, studying microbes in the environment has identified many useful products. In the early 1900s, Selman Waksman was studying actinomycetes in soil when he discovered the antibiotic streptomycin. Similarly, metagenomics research has identified (by accident) another antibiotic, called turbomycin. The researchers were looking for hemolysin-related genes in the soil by screening a metagenomic library (see later discussion). Hemolysin is a bacterial toxin that punctures holes in susceptible cell membranes, allowing the cellular contents to leak out and the cell then dies. Hemolysin lyses red blood cells and creates a clear zone around a bacterial colony growing on blood agar plates. Some *E. coli* clones from the library had dark red or orange colors. Further investigation of these clones found two novel antibiotics, turbomycin A and B (Fig. 12.1). Novel biodegradation pathways have also been found in bacteria that inhabit contaminated sites. Enzymes that can reduce the toxic effects of oil- and petroleum-based contaminants are found in bacteria that utilize the pollutants as an energy source. Even bacteria that thrive in environments contaminated with radioactivity have been identified. **CULTURE ENRICHMENT FOR ENVIRONMENTAL SAMPLES** Various methods are used to enhance the starting material for metagenomics research, because a metagenomic library is only as good as its contents. Enrichment strategies include stable isotope probing (SIP), BrdU enrichment, and suppressive subtraction hybridization. **Stable isotope probing (SIP) **was originally developed to trace single carbon** **compounds during their metabolism by cultured methylotrophs (bacteria specialized for growth on single-carbon compounds). Labeled precursor carbons were traced into fatty acids during bacterial growth. The method was adapted to the environmental samples used to create metagenomic libraries. Here, an environmental sample of water or soil is first mixed with a precursor such as methanol, phenol, carbonate, or ammonia, that has been labeled with a stable isotope such as ^15^N, ^13^C, or ^18^O (Fig. 12.2). If the organisms in the sample metabolize the precursor substrate, the stable isotope is incorporated into their genome. Then, when the DNA from the sample is isolated and separated by centrifugation, the genomes that incorporated the labeled substrate will be "heavier" and can be separated from the other DNA in the sample. The heavier DNA will migrate further in a cesium chloride gradient during centrifugation. As described later, the DNA can either be used directly or cloned into vectors to make a metagenomic library. This technique is particularly useful to find new organisms that can degrade contaminants, such as phenol in the example given. ** Adding 5-bromo-2-deoxyuridine (BrdU)** is a related technique for enriching for the DNA of active bacteria in an environmental sample. Rather than entering only a metabolic subset of bacteria, BrdU is incorporated into the DNA and RNA of any actively growing bacteria or viruses. Note that bacteria and viruses that are dormant or dead, as well as free DNA, will not be labeled by this method. As before, the soil or water is isolated and incubated with BrdU. Any bacteria that are actively growing will take up the nucleotide analog and incorporate it into its DNA. Next, the BrdU-labeled DNA is isolated either with antibodies to BrdU or by density gradient centrifugation (see Fig. 12.2).  RNA-SIP focuses on isolating RNA from the environmental sample (rather than DNA). Small subunit ribosomal RNA (SSU rRNA), that is, the 16S rRNA of bacteria or the 18S rRNA of eukaryotes, is an excellent biomarker because it is essential to all cellular life, it is very abundant within a cell, it is variable among different species, and there is an enormous database of different SSU rRNA sequences making identification relatively easy. In RNA SIP the SSU rRNA in the environmental sample is labeled. As described earlier, 13C-labeled precursors are supplied to the environmental sample. These are incorporated into SSU rRNA independently of cell division because ribosomal RNA is produced in any cell that is making proteins, not just cells undergoing replication. This technique provides information on bacteria that are dormant as well as those that are more active. Much as before, the RNA is isolated and separated on a gradient by centrifugation. The rRNA bands tend to aggregate together during centrifugation. Therefore, each fraction must be repeatedly separated from the others. The final SSU rRNA fraction may still contain some contaminating nonlabeled rRNAs, so the fraction must be evaluated with care.  RNA-SIP can be used to identify a variety of microorganisms in environmental samples. For example, water from an aerobic industrial wastewater plant was evaluated for phenol- degrading microorganisms. The water was incubated with 13C-labeled phenol, and the SSU rRNA was isolated by centrifugation. The rRNAs were isolated and amplified with RT-PCR followed by denaturing gradient gel electrophoresis. The bands were subjected to mass spectrometry to identify which rRNA sequence was most abundant. Interestingly, an organism belonging to the genus Thauera in the β-Proteobacteria was abundant, even though this organism was most usually found in denitrifying conditions. It was previously thought that pseudomonads were degrading the phenol.  Another culture-enrichment technique, **suppressive subtraction hybridization (SSH**), takes advantage of the genetic differences between samples from two different areas. During standard subtractive hybridization, two different samples are hybridized and the mRNA that is the same is removed, leaving only mRNA that is different between the two samples. SSH works by the same principle. First, two different conditions must be established. For example, one soil sample from a polluted site could be compared with nearby soil that is not contaminated. The two soil samples will differ in their content of microorganisms, and those microorganisms enriched in the contaminated site could potentially metabolize the contaminant.  When DNA from each sample is isolated, the contaminated soil is considered the tester DNA and the "normal" soil is the driver sample. The tester sample is divided into two, and two different linkers are added to the ends of the DNA to form tester A and tester B. Tester A (with linker A), tester B (with linker B), and driver DNA are all mixed, denatured to make them single-stranded, and then rehybridized. The driver DNA is in excess to the testers, which ensures that DNA fragments from bacteria outside the contamination site outnumber those from the contaminated site. The driver DNA will anneal to all the common DNA fragments, making these double-stranded and with only one strand connected to the linker. All the tester DNA that is unique and not found in the uncontaminated soil will be free to hybridize with itself, forming A:A, B:B, or A:B hybrids. PCR primers are added to the hybridization mix; one primer recognizes linker A and the other primer is for linker B. As shown in Fig. 12.3, PCR will amplify only those hybrids that are tester:tester. Furthermore, because the A:A and B:B hybrids have inverted linkers, these hybrids will form a "panlike" structure during annealing and will not be amplified by PCR. Thus only A:B hybrids are amplified, and these represent unique sequences found only in the contaminated site. **NATURAL ATTENUATION OF POLLUTANTS** Metagenomics can also be applied to biogeochemical research. Perhaps identifying how bacteria affect the environment will help us figure out ways to maintain our species. Understanding how bacteria can live in extreme environments can reveal useful biochemical processes for biotechnology. Most important, finding out how bacteria cope with contaminated sites may provide useful enzymes for cleaning up our pollution. **Bioremediation **is one avenue in which biotechnology has made rapid advances. Many** **different humanmade compounds have contaminated the environment around us, through everyday use, accidental spillage, or intentional dumping. Many environmental biotechnologists are working on "biological" means of cleaning the environment. In fact, releasing an organism that can degrade a pollutant would provide a very easy, low-cost way of cleaning up a polluted site. Naturally occurring microorganisms often have the ability to degrade humanmade pollutants. For example, *Rhodococcus* has a highly diverse repertoire of pathways to degrade pollutants, such as short- and long-chain alkanes, aromatic molecules (both halogenated and nitro-substituted), and heterocyclic and polycyclic aromatic compounds, including quinolone, pyridine, thiocarbamate, *s*-triazine herbicides, 2-mercaptobenzothiazole (a rubber vulcanization accelerator), benzothiophene, dibenzothiphene, MTBE (see later discussion), and the related ethyl *tert*-butyl ether (ETBE). *Rhodococcus* has several features that contribute to its ability to degrade so many compounds. First, it has a range of different enzymes that degrade toxic compounds, including cytochrome P450 enzymes. These are very efficient and versatile in oxidation pathways and catalyze a variety of reactions, including epoxidation (Fig. 12.7). Other enzymes that catalyze key degradation steps include monooxygenases and dioxygenases, which help degrade aromatic compounds. Furthermore, several strains of *Rhodococcus* can survive in solvents such as ethanol, butanol, dodecane, and toluene, which would kill many other bacteria. The oil-degrading strains actually adhere to oil droplets! *Rhodococcus* species are found in all types of environments, including nuclear waste sediments, tropical soil, Arctic soil, and sites in Europe, Japan, and the United States. Genetically speaking, *Rhodococcus* also has unique attributes that are advantageous in biodegradation. The genome of *Rhodococcus* sp. strain RHA1 has 9.7 Mb of DNA, including one chromosome and three large linear plasmids. The plasmids may be critical because they are important for gene transfer and recombination events. The genes for the catabolic enzymes are often found in clusters, flanked by inverted repeats, suggesting that they are acquired and passed from one strain to another by recombination. Such horizontal gene transfer can also transfer these catabolic regions to other bacteria, including *Pseudomonas *and* Mycobacterium*., Pathway Engineering, describes some plasmids* *that encode pollutant-degrading enzymes. Different pollutants are degraded in various ways. Sometimes a single naturally occurring organism can completely degrade a pollutant. Other pollutants require more than one type of bacterium to achieve complete degradation. Some pollutants are degraded very slowly. Heavy metals cannot be chemically degraded, so they pose more of a challenge than organic molecules. Most environmental biotechnologists look for microorganisms that sequester the heavy metal in a solid phase. The conversion of such heavy metals as uranium from an aqueous phase to a solid phase can clean drinking water supplies. Thus, certain anaerobic microorganisms can reduce uranium(VI) to uranium(IV) by utilizing the metal as a terminal electron acceptor. This converts the uranium from a soluble to an insoluble form. In one uranium-contaminated site (Old Rifle, Colorado, USA), one study actually injected acetate into the ground water. Acetate is an electron donor that stimulates metal-reducing bacteria to sequester uranium into the solid phase. Within 50 days, some contaminated wells had uranium concentrations lower than the regulated level. Although these results are promising, over time the acetate dissipated and the soluble uranium levels increased again. More research is necessary to find a permanent means to keep uranium out of ground water. Another recalcitrant compound that can contaminate ground water, **methyl tert-butyl** **ether (MTBE)**, oxygenates gasoline so that it burns more efficiently. Many cases of** **MTBE contaminating groundwater have been reported. Finding a natural method to clean these sites has great applicability. The United States Environmental Protection Agency has classified MTBE as a possible human carcinogen, and drinking water must contain less than 20 to 40 µg/L. One MTBE-contaminated site in South Carolina, United States, had a large plume of MTBE-contaminated gasoline leaking from an underground storage tank at a gas station. The plume ended at a drainage ditch. The concentration of MTBE in the water was low, and in the 2-meter gap between the anaerobic and aerobic zones, the MTBE was metabolized by naturally occurring microorganisms. This led to studies that determined that MTBE could be degraded by bacteria such as *Methylobium petroleophilum *PM1 in areas that transition from anaerobic (anoxic) to* *aerobic (oxic). In fact, if anaerobic regions of the MTBE plume in South Carolina were injected with a compound that released oxygen, the concentration of MTBE decreased from 20 mg/L to 2 mg/L, suggesting that **biostimulation** may be a good approach to clean up contamination. Biostimulation is the release of nutrients, oxidants, or electron donors into the environment to stimulate naturally occurring microorganisms to degrade a contaminant. In other areas of MTBE contamination, the site may have to undergo **bioaugmentation**, that is, specific microorganisms plus their energy sources may need to be added to the site. Such microorganisms may be naturally occurring, a mixture of different organisms, or even genetically modified. **HISTORY OF PLANT BREEDING** For thousands of years humans have improved crop plants and domestic animals by selective breeding, mostly at a trial-and-error level. Over many years, animal and crop breeders have learned that improving their crops and animals has a biological basis. In fact, the father of genetics, Gregor Mendel, experimented with the common pea. He studied easily identified traits such as round versus wrinkled seeds or yellow versus green seeds. Mendel would take the pollen from one plant and put it on the stigma of another plant (Fig. 14.1), a procedure called **cross-pollination**. His experiments showed that plants have some traits that may dominate others. For example, a cross between a yellow-seeded pea plant and a green-seeded pea plant gave only seeds that were yellow. This work was published in 1865, but no one understood the importance of these findings until well after Mendel's death. Many scientists around the world still use traditional breeding techniques to enhance crop yields, increase resistance to various pests or diseases, or increase the tolerance of a particular crop to heat, cold, drought, or wet conditions. Although simply crossing a high-yielding plant with another can produce offspring with even higher yields than either parent, the process is long and tedious. Many thousands of plants must be cross-pollinated to find the one offspring with higher yield. The crosses must be done by hand, that is, pollen must be taken from one plant and manually placed on another. In addition, the possibility of finding improved traits is limited by the amount of genetic diversity already present in the plants. Consequently, if the two plants that are crossed share many of the same genes, the amount of possible improvement is limited. If a plant has no genes for disease resistance, there is no way traditional cross-pollination will develop that trait. Therefore, scientists have searched for better ways to improve plants. In the 1920s, scientists realized that mutations could be induced in seeds by using chemical mutagens or by exposure to X-rays or gamma rays. Although useful, the outcome of such treatments is even less predictable than traditional breeding methods. Nonetheless, mutation breeding has been successful, especially in the flower world. For example, new colors and more petals have been expressed in flowers such as tulips, snapdragons, roses, chrysanthemums, and many others. Mutation breeding has also been tried on vegetables, fruits, and crops. Some of the varieties of food we eat today were developed using this method. In short, a large number of seeds are exposed to the mutagen to generate various mutations in their DNA. The seeds are then planted and cultivated. However, the majority of seeds are killed by the treatment. After the viable seeds are grown, the fruit, flower, or grain of the plant is tested for improvements. If one plant is found with a desirable trait, then its progeny are tested for the trait. Novel traits are only useful if they are heritable, that is, passed from one generation to the next. Because only one original mutant plant would gain any particular desirable trait, this plant would need to be propagated a long time before any of the fruit, grain, or flower would be sold to market. It is important to realize that the actual fruits or flowers sold to the consumer were never exposed to the mutagen. Today, chemical mutagens are still used, but molecular biology techniques are being used to identify the actual gene associated with the desired phenotype (see later discussion). Recently, the emergence of molecular biology has opened the door to a much more predictable way to enhance crops. Scientists have discovered ways to move genes from foreign sources into a specific plant, resulting in a **transgenic plant**. The foreign gene, or **transgene**, may confer specific resistance to an insect, or protect the plant against a specific herbicide, or enhance the vitamin content of the crop. The major difference between transgenic technology and traditional breeding is that a plant can be transformed with a gene from any source, including animals, bacteria, or viruses as well as other plants, whereas traditional cross-breeding methods move genes only between members of a particular genus of plants. Furthermore, the transgene has a known function and has been evaluated extensively before being inserted in the plant. During traditional breeding, the identity of genes responsible for improving the crop is rarely known. **PLANT TISSUE CULTURE** One major advantage of plants is that they can often be regenerated from just a single cell, that is, each plant cell is **totipotent** and retains the ability to develop into any cell type of a mature plant (Fig. 14.2). There is no absolute separation of the germline from the somatic cells in plants (unlike animals). This unique feature of plants allows scientists to grow and manipulate plant cells in culture, then regenerate an entire plant from the cultured cells. Plant tissue culture can be done on either a solid medium in a petri dish, called **callus culture**, or in liquid, called **suspension culture**. In both cases, a mass of tissue or cells,** **known as an **explant**, must be removed from the plant of interest. In callus culture, the tissue can be an immature embryo, a piece of the apical meristem (the region where new plant shoots develop), or a root tip. For liquid culture, cells must be dissociated from one another. Liquid culture usually uses **protoplasts** (plant cells from which the cell wall is removed), microspores (immature pollen cells), or macrospores (immature egg cells). The cells are then cultured with a mixture of nutrients and specific plant hormones that induce the undifferentiated cells to grow. Different types of plants respond to different hormones. To culture wheat cells, for example, the explant is grown with 2,4-dichlorophenoxyacetic acid (2,4-D). This is an analog of the plant hormone auxin, which stimulates plant cells to dedifferentiate and grow. To culture tomato plants, the hormone cytokinin dedifferentiates the cells and induces cell division. In callus culture, undifferentiated cells form a crystalline white layer on top of the solid medium, called the **callus**. After about a month of growth, the mass of undifferentiated cells can be transferred to medium with a lower concentration of hormone or with a different hormone. Decreasing the amount of hormone allows some of the undifferentiated callus cells to develop into a plant shoot. In most cases the small shoots look like new blades of grass growing from the mass of cells. After another 30 days, the hormone is removed completely, which allows root hairs to start growing from some of the shoots. After another 30 days, small plants can be isolated and planted into soil. In liquid culture, hormones are also used to stimulate the growth of undifferentiated cells, but the shoot and root tissues grow simultaneously (Fig. 14.3). Because plant tissue culture allows many plants to be produced from one source, the technique is useful for making clones of one particular plant. If a very rare plant is identified, it can be propagated using tissue culture. Only a small cutting is needed to generate many identical progeny. Certain special plant varieties that are hard to maintain by producing seed can be maintained for the long term in culture. Plant cell culture has also been used for mutation breeding. Rather than using seeds, undifferentiated cells are exposed to the mutagen, and plants are regenerated from the mutagenized cells. Mutagens are more effective on the exposed cells of a callus rather than the protected cells within a seed. Merely growing plants by tissue culture may induce mutations. The process of regenerating a plant from a single cell may cause three different types of alterations. Temporary physiological changes can occur in the regenerated plant. For example, when blueberry plants are regenerated via tissue culture, the plants are much shorter. These changes are not permanent, and after a few years growing in the field, the regenerated blueberries are no different from any other blueberries. Another alteration that may occur is an **epigenetic** **change**. This is an alteration that persists throughout the lifetime of the regenerated plant,** **but is not passed on to the next generation. Epigenetic changes are often due to alterations in DNA methylation. Finally, true **genetic changes** affect the regenerated plant and all its progeny. These may be due to changes in ploidy level, chromosome rearrangements, point mutations, activation of transposable elements, or changes in chloroplast or mitochondrial genomes. These types of changes are relatively common, but changes in the available nutrients and hormones during tissue culture can dramatically decrease the frequency of mutation. **GENETIC ENGINEERING OF PLANTS** The first step in genetically engineering a plant is to identify a gene that will confer a specific desirable trait on the plant. In some ways, this is the hardest part of genetic engineering. The most desirable traits for a crop are increases in the amount of seed, grain, or other plant products. Increased resistance to disease or drought is also very useful. Finding the genes responsible is difficult, because multiple interacting genes usually control such traits. In addition, such genes may play other roles in plant physiology or development. So far, most successful genetic engineering of plants has relied on inserting one or a few genes that supply simple, yet useful, properties. For example, resistance to the herbicide glyphosate is due to a single gene. Making a crop such as soybean resistant to glyphosate allows the farmer to kill the weeds in the field without harming the soybeans (see later discussion). Another desirable trait often due to a single gene is the production of toxins that kill harmful insects (see later discussion). Both these cases rely on transgenes derived from bacteria. As more research into plant physiology occurs, more genes can be identified that increase the value of a crop. For example, a two-gene pathway was engineered into rice to make it more resistant to drought (see later discussion). Plants can also be engineered for novel products. Thus, golden rice expresses the biosynthetic pathway for vitamin A precursors. This rice was developed for people who rely on rice as the one main food in their diet. The addition of vitamin A precursors can prevent deficiencies that cause blindness or premature death in children in developing countries. Researchers have also engineered the human insulin gene for expression in *Arabidopsis* and safflower.  At present, insulin is produced by engineered bacteria. However, plant-produced insulin is easier to isolate and purify in bulk and should cost much less. Further research will identify new genes and useful pathways that can be engineered into plants. Perhaps engineered plants will be used to clean up oil spills or other pollutants by growing them on contaminated soil. **GETTING GENES INTO PLANTS USING THE TI PLASMID** Plants suffer from tumors, though these are quite different from the cancers of animals. The most common cause is the Ti plasmid (tumor-inducing plasmid), which is carried by soil bacteria of the *Agrobacterium* group. Specifically, the Ti plasmid of *Agrobacterium* *tumefaciens *is an important tool for plant genetic engineering. The most important aspect* *of the infection is that a specific segment of the Ti plasmid DNA is transferred from the bacteria to the plant. Scientists have exploited this genetic transfer in order to get genes with desired properties into plant cells. *Agrobacterium* is unique in the ability to transfer a segment of its DNA from one kingdom to another. Most DNA transfers occur only between closely related organisms. In nature, *Agrobacterium* is attracted to plants that have minor wounds by phenolic compounds such as acetosyringone, which are released at the wound (Fig. 14.4). These chemicals induce the bacteria to move and attach to the plant via a variety of cell surface receptors. The same inducers activate expression of the virulence genes on the Ti plasmid that are responsible for DNA transfer to the plant. This is under control of a two-component regulatory system. At the cell surface, the sensor, VirA, is autophosphorylated when it detects the plant phenolic compounds. Next, VirA transfers the phosphate to the DNA-binding protein, VirG, which activates transcription of the *vir *genes of the Ti plasmid. Two of the gene products (VirD1 and VirD2) clip the T-DNA* *borders to form a single-stranded immature T-complex. VirD2 then attaches to the 5¢ end of the T-DNA, and bacterial helicases unwind the T-DNA from the plasmid. The single-stranded gap on the plasmid is repaired, and the T-DNA is coated with VirE2 protein to give a hollow cylindrical filament with a coiled structure. This is the mature form of T-DNA and traverses into the plant. T-DNA is transferred to the plant in a process similar to bacterial conjugation. First, *Agrobacterium *forms a pilus. This rodlike structure forms a connection with the plant cell and* *opens a channel through which the T-DNA is actively transported into the plant cytoplasm. Both pilus and transport complex consist of proteins that are *vir* gene products. Once inside the plant cytoplasm, T-DNA is imported into the nucleus. Both VirE2 and VirD2 have nuclear localization signals that are recognized by plant cytosolic proteins. These proteins take the T-complex to the nucleus where it is actively transported through a nuclear pore. The single T-DNA strand is integrated directly into the plant genome and converted to a double-stranded form. The integration requires DNA ligase, polymerase, and chromatin remodeling proteins, which are all supplied by the plant. Once they are part of the plant genome, the genes in the T-DNA are expressed. These genes have eukaryote-like promoters, transcriptional enhancers, and poly(A) sites and hence are expressed in the plant nucleus rather than in the original bacterium. The proteins they encode synthesize two plant hormones, auxin and cytokinin. Auxin makes plant cells grow bigger and cytokinin makes them divide. The infected plant cells begin to grow rapidly and without control, resulting in a tumor.   T-DNA also carries genes for synthesis of opines, which are a variety of different amino acid and sugar phosphate derivatives. The type of opine differentiates the various strains of *Agrobacterium*. Opines are made by plant cells that contain T-DNA but are used by the bacteria as carbon, nitrogen, and energy sources. Notice how the bacterium tricks the plant into using its resources to supply the bacteria with food. The Ti plasmid, which is still inside the *Agrobacterium*, carries genes that allow the bacteria to take up these opines and break them down for food. Note that other bacteria, which might be present by chance, cannot use opines because they do not possess the genes for uptake and metabolism. This ensures that the plant feeds only the bacteria with the Ti plasmid. So how are Ti plasmids used to improve plants? First, the Ti plasmid is disarmed by cutting out the genes in the T-DNA for plant hormone and opine synthesis. Then, the transgene of interest, such as an insect toxin gene, is inserted into the T-DNA region of the Ti plasmid. The Ti plasmid is also streamlined by removing genes that are not involved in moving the T-DNA. These smaller plasmids are much easier to work with and can be manipulated in *Escherichia coli *rather than their original host,* Agrobacterium. *Now, when the T-DNA enters* *the plant cell and integrates into the chromosome, it will bring in the transgene instead of causing a tumor. The transferred region of the plasmid must also have other elements in order for the transgene to function properly (Fig. 14.5). Expression of the transgene requires a promoter that works efficiently in plant cells. This may be one of two types. A **constitutive promoter** will turn the gene on in all the plant cells throughout development; thus every tissue, even the fruit or seed, will express the gene. A more refined approach is to use an **inducible** **promoter **that has an on/off switch. An example of this is the** ***cab*** **promoter from the gene encoding chlorophyll *a*/*b* binding protein. This promoter is turned on only when the plant is exposed to light; therefore, root tissues and tubers such as potatoes will not express the gene. Many different promoters may be used, but ideally, the promoter should turn on only in tissues that need transgene function. Another important component for the genetically modified T-DNA region is some sort of selectable marker. Including an herbicide or antibiotic resistance gene in the T-DNA region can be used to track whether the foreign DNA has been inserted into plant cells. The selectable marker may cause problems because it must be expressed constitutively throughout the plant. Many people worry that the protein product of the selectable marker could cause allergies or reactions if expressed in fruit, grain, or vegetables. However, systems exists that can remove this gene once the transgenic plant has been isolated (see later discussion). In practice, *Agrobacterium* is used to transfer genes of interest into plants using tissue culture. Either dissociated plant cells called *protoplasts* or a piece of callus are cultured with *Agrobacterium *harboring a Ti plasmid with modified T-DNA. After coculture, the plant* *cells are harvested and incubated with the herbicide or antibiotic used as the selectable marker. This kills all the cells that were not transformed with T-DNA or failed to express the genes on the T-DNA. The transformed cells can then be induced to produce shoot and root tissue by altering the hormone conditions in the medium as described earlier (Fig. 14.6). The small transgenic plants can then be screened for transgene expression levels (see later discussion). Recently, a method for ***in planta Agrobacterium*** **transformation** was developed and has revolutionized the plant transformation world. *In planta* transformation is also known as the **floral dip** method. The method was developed using the model plant *Arabidopsis* but has been extended to other plants, such as wheat and maize. First, *Arabidopsis* plants are grown until flower buds begin to form. These buds are removed and allowed to regenerate for a few days. Once they begin to regenerate, the plants are dipped into a suspension of *Agrobacterium *containing a surfactant. The surfactant allows the* Agrobacterium *to adhere to the plant and transfer its T-DNA. Because the flower buds are just beginning to form, the T-DNA somehow becomes part of the germline through the ovarian tissue. The plant is allowed to finish growing and set seed. These seeds are harvested and grown in selective media to find those that have integrated and expressed T-DNA. Although the method gives a low percentage of transformants, so many seeds can be screened that the overall procedure works well. **PARTICLE BOMBARDMENT TECHNOLOGY** Another strategy for getting a gene of interest into plant tissue is to blast the DNA through the plant cell walls with a particle gun. Unlike Ti plasmid transfer by Agrobacterium, this technique works with all types of plants. The basic idea is that DNA is carried on microscopic metal particles. These are fired by a gun into plant tissue and penetrate the plant cell walls. This technique is rather nonspecific, yet it has been very successful in the plant world. First, either a leaf disk (just a round piece of leaf tissue) or a piece of callus is isolated from the plant, placed on a dish, and put in a vacuum chamber. The DNA to be inserted (carrying the gene of interest, with any promoter and enhancer elements, plus selectable markers) is coated on microscopic gold or tungsten beads. Gold beads are preferable because tungsten can be toxic to some plants. The beads are placed at the end of a plastic bullet. One variant of the method uses a blast of air or helium to project the bullet toward the sample. In the first gene guns, actual firearm blanks were used to accelerate the bullet. Between the bullet and plant tissue is a plastic meshwork stop. When the bullet hits the stop, the DNA-coated beads are thrown forward through the meshwork and continue on through the vacuum chamber and into the plant tissue. An alternative method is to accelerate the beads by a strong electrical discharge. The high voltage vaporizes a water droplet, and the resulting shock wave propels a thin metal sheet covered with the particles at a mesh screen. The screen blocks the metal sheet but allows the DNA-coated particles to accelerate through into the plant tissue (Fig. 14.7). One advantage to this method is that the strength of the electrical discharge can be controlled; therefore, the amount of penetration into the tissue can be changed at will. W hen the beads penetrate the tissue, some will actually enter the cytoplasm or nucleus of the leaf or callus cells. The DNA dissolves off the beads inside the cells and the DNA is free to recombine with the chromosomal DNA of the plant (Fig. 14.8). The leaf or callus tissue is then transferred to selection media where the cells that integrated the DNA carrying the selectable marker are able to grow, but other cells die. The transformed plants are regenerated using tissue culture techniques, and finally screened for the gene of interest. Particle guns have also been used on animal tissues. In addition, scientists have modified the technology in order to transform DNA into the mitochondria of yeast, as well as the chloroplasts of *Chlamydomonas*, a small green alga.   **PLANT BREEDING AND TESTING** Making a transgenic plant is a relatively small step; evaluating and testing the transformed plants is the most time consuming part of the whole process. The expression level of the transgene may vary considerably, depending on the number of integrated transgenes and their location. The term **event** refers to each independent case of transgene integration. For example, if one copy of the transgene inserted into chromosome 2 of the first transformant, this would be referred to as event 1. If, in the same experiment, a separate transformed plant received the same transgene, but integrated into chromosome 4, that would be a second event. The location of integration affects the expression of the transgene. If the transgene in event 1 integrated into a region of heterochromatin, the gene would probably be silenced and never be expressed at all, even if provided with a strong promoter. In contrast, if in event 2, the transgene integrated just downstream of a very active chromosomal gene, it would probably show high expression levels. The number of integrated transgenes can also vary. Often a single transformed plant will gain multiple copies of the same transgene. The first issue to address is whether the transgene causes any harmful side effects to the plant. Does the transgene function as expected? Does the transgene affect the crop quality? Does the transgene affect the ecosystem? The answer to these questions depends on the individual transgene being used (see later discussion for specific examples). If no harmful effects are found, then the transgene must be transferred from the experimental plant to one with a much higher yield. Most transgenic plants are made from old varieties that are good for work in laboratories, but do not make a lot of seeds per acre or are very susceptible to diseases. Furthermore, as discussed earlier, the regeneration of plants through tissue culture may itself cause mutations. In order to overcome these problems, the transgene is moved by traditional cross-breeding into high-yielding varieties that farmers are already using. First, the pollen from the plant with the transgene is harvested and put onto the corn silk or stigma of the high-yielding variety. The seeds from this cross are harvested and grown. This is the F1 generation, and the plants containing the transgene are selected. For example, if the transgene makes the plant resistant to an herbicide, the F1 generation is sprayed to kill the plants without the transgene. The pollen from the F1 plants that survive is **back-crossed** to the original high-yielding parent. The seeds are grown, plants with the transgene are selected, and the whole process is repeated about four or five times. This crossing scheme will ensure that about 98% of the genes in the final plant are from the high-yielding variety, and the remaining genes are from the original transgenic plant. Because it takes an entire summer for one generation of corn, soybeans, or cotton to grow, this backcrossing scheme can take many years to complete. Once the transgene is back-crossed into a suitable variety, field tests are performed to determine how the transgene affects the growth, yield, disease resistance, and other important traits of the plant. These field tests must be done over many different locations so that soil type, terrain, rainfall, and other factors can be allowed for. The field tests may also take many years. Different amounts of rain from one year to the next can greatly affect crop yields. The plant breeder selects only the plants that consistently have the highest yield with the best disease resistance. The other plants are never grown again. The other issue in releasing transgenic plants to the public is passing the tests of government regulatory agencies. These agencies regulate all stages of the transgenic construction process. An Institutional Committee for Biosafety regulates how the transgene is handled when making the transgenic plant, whether in *E. coli, Agrobacterium*, or the plant itself. These committees are usually associated with the university or company where the work is done, but they all follow guidelines from the National Institutes of Health (NIH). The guidelines regulate the environment in which the transgenic plant may be grown (laboratory, greenhouse, etc.). In order to test the transgenic crop in the field, the Animal and Plant Health Inspection Service of the U.S. Department of Agriculture must be notified and must approve the plan. The scientist must provide extensive data on the transgene, its potential effect on the plant, the ecosystem, and any other crops similar or related to the transgenic plant. Two other agencies must also approve the transgenic crop. If the transgenic plant gives a food product such as corn, the Food and Drug Administration (FDA) must do rigorous testing for possible allergies to the transgenic plant. The potential toxicity of the transgenic crop and whether or not the nutritional quality of the product is affected by the transgene are also tested. The Environmental Protection Agency also evaluates the transgenic crop for potential effects on the environment and on animals or insects that also inhabit the farmers' fields. These are just the beginning of the regulations since anything sold overseas must also satisfy the regulatory commissions from all the countries in which the product is sold. At this time, overseeing transgenic technology is a hot issue that is constantly changing. As transgenic technology becomes more common and more information becomes available, the regulatory issues will change and adapt to the type of crops being produced. **TRANSGENIC PLANTS WITH INSECT RESISTANCE** Although weeds are a nuisance, even worse enemies of plants, and correspondingly more expensive to farmers, are: **1.**     Insects and roundworms **2.**     Fungal diseases (molds, blights, rusts, and rots) **3.**     Viral diseases of plants It is possible to engineer plants for resistance to all of these, but we will consider just the insects here. Spraying crops with insecticides is a very costly and hazardous procedure. Insecticides are often more toxic to humans than are herbicides, because insecticides target species closer to our own. Many insect biochemical pathways are found not only in humans, but also in rodents or birds that may inhabit crop fields. Luckily, naturally occurring toxins exist that are lethal to insects but harmless to mammals. The prime example is the toxin from a soil bacterium called *Bacillus thuringiensis*. **Bt toxin**, as it is called, has been sprayed on crops to prevent insects such as the cotton bollworm and European corn borer from destroying cotton and corn, respectively. Damage from the European corn borer plus the cost of insecticides to control it cost farmers about \$1 billion annually. Damage by the corn borer also makes corn plants susceptible to infection with a toxic fungus that can harm humans if ingested. Bacteria of the genus *Bacillus* produce spores that contain a crystalline or **Cry protein**. When insects eat *Bacillus* spores, the Cry protein breaks down and releases **delta endotoxin** (i.e., the Bt toxin). This toxin binds to the intestinal lining of the insect and generates holes, which cripple the digestive system, and the insect dies (Fig. 14.14). Different species of *Bacillus* produce a family of different but related Cry proteins. These were originally classified according to insect susceptibility: CryI killed *Lepidoptera* (butterflies and moths), CryII killed both *Lepidoptera* and *Diptera* (flies), CryIII killed *Coleoptera* (beetles), and CryIV killed *Diptera* (but not *Lepidoptera*). However, as the number of known Cry variants increased, this classification became too simplistic, because sequence similarities did not always correspond to the spectrum of insecticidal activity. Cry proteins are now classified with Arabic numerals into some 20 subfamilies based on sequence relationships. Instead of spraying insects with the toxin, scientists have used transgenic technology to insert the *cry* genes directly into plants. When a cloned toxin gene was inserted into tomato plants, it partly protected against tobacco hornworm. However, the plants made only low levels of the toxin because the toxin gene is from a bacterium and is designed to express well in bacteria, not plants. Therefore the toxin gene was altered to enhance expression. The original toxin is a big protein that has 1156 amino acids. However, only the first 650 amino acids are necessary for the toxic effect; therefore, the protein was truncated by deleting the last half of the gene. The shorter, truncated protein requires less energy to produce. Next, the toxin gene was placed under the control of a promoter that gives constant high-level expression in plants. Certain promoters from plant viruses, such as cauliflower mosaic virus, satisfy these requirements and give 10-fold increases in toxin production. When genes from one organism are expressed in a very different host cell, codon usage also becomes a problem. As explained in Previews Pages, the genetic code is redundant in the sense that several different codons can encode the same amino acid. So although a protein has a fixed amino acid sequence, there is considerable choice in which codons to use. Different organisms favor different codons for the same amino acid and have different levels of the corresponding tRNAs. If a gene uses codons for a rare tRNA, the supply of this may limit the rate of protein synthesis. In practice this is relevant only to genes expressed at high levels---exactly the situation here. Therefore the insect toxin gene was altered by changing many of the bases in the third position of redundant codons. Almost 20% of its bases were altered to make it more plantlike in codon usage. This did not alter the amino acids encoded and, therefore, the toxin protein itself was not affected by this procedure. However, the rate at which plant cells made the protein greatly increased and gave another 10-fold increase in toxin production. Transgenic Bt crops such as cotton and corn have many advantages over spraying the fields with the toxin itself. One advantage is the toxin doesn't drift over other areas, which prevents cross-contamination. Using transgenic crops reduces the amount of insecticide needed. In 1998, about 450,000 kg less insecticide was used on the U.S. cotton crop alone. Yet only 45% of the cotton crop was actually transgenic. **FUNCTIONAL GENOMICS IN PLANTS** Because the complete DNA sequences of rice, poplar tree, and Arabidopsis are known, plant scientists are following functional genomics strategies. Rather than working on one specific gene, the entire genome can be screened. Most of this type of work is still done with Arabidopsis, but some has now moved into crop species such as rice, corn, and soybeans. Functional genomics in plants uses a variety of techniques, some of which have been discussed previously, but most rely on the removal or blockage of gene expression. Novel genes and metabolic pathways useful for understanding basic plant physiology are being analyzed in the hope of improving our current crops. Insertions are one method to find the function of new genes. Transposons or T-DNA insertions are two methods used to generate plant mutants. Here, instead of including a transgene, the T-DNA or transposon includes only a reporter gene. When the T-DNA or transposon integrates into the plant chromosome, the insertion may disrupt a plant gene. When the insertion knocks out the function of a plant gene, the resulting phenotype can be screened and assessed. By cloning the regions upstream and downstream of the insertion, the plant gene that corresponds to the phenotype can be identified. Gene silencing is another method to identify the function of plant genes. As described, gene silencing by RNA interference (RNAi) is a phenomenon that was originally described in plants. RNAi is triggered by double-stranded RNA, which is cut into short segments (siRNA, short-interfering RNA). The RISC enzyme complex uses siRNA to identify homologous RNA (in particular mRNA) and cut it up. This prevents mRNA from being expressed into protein. This is exploited in the laboratory by transforming a plant with small oligonucleotides that stimulate RISC to abolish the expression of a chosen gene. The plant can then be assessed for any visible phenotype associated with the gene knockdown. Another method for generating gene knockouts is **fast neutron mutagenesis**. This uses **fast neutrons **to induce DNA deletions. Fast neutrons are created by nuclear processes such** **as nuclear fission, where free neutrons with a kinetic energy close to 1 MeV are generated. These neutrons cause deletions in exposed DNA. Therefore, seeds from the plant of interest exposed to fast neutrons acquire random mutations. The dose of fast neutrons and, consequently, the number of deletions per genome can be controlled. Seeds treated with fast neutrons, known as M1 seeds, are grown into plants. Each plant has a different deletion or set of deletions and a potentially different phenotype.  The seeds from each of these plants, called the M2 seeds, are collected. Most are saved as seed stock; the remaining M2 seeds are grown into plants. The DNA from the plants is isolated and collected into pools of varying sizes. For example, if the original pool had DNA from 100 plants, successively smaller pools of the 100 plants are made, down to just one or two plants per pool. These are usually screened by PCR to find specific genes with deletions. PCR primers are made to amplify a target gene from the largest pool of DNA. If a deletion was generated within the target gene in one of the plants, the PCR primers will amplify two bands, the wild-type gene plus a shorter segment from the deleted gene. The smaller DNA pools are then screened for the deletion until a specific M2 seed can be associated with the genetic deletion (Fig. 14.16). Yet another method of creating plant mutations is called **TILLING (targeting-induced** **local lesions in genomes**; Fig. 14.17). First, point mutations are created in a collection** **of seeds by soaking them in a chemical mutagen, such as EMS (ethyl methane sulfonate), which induces G/C and A/T transitions in DNA. As before, the M1 seeds are grown into plants and the second-generation seeds (M2) are mainly saved as stocks. Some M2 seeds are grown and the DNA is harvested and pooled into a large megapool and successively smaller pools as described earlier. PCR primers are used to amplify selected regions of the DNA. The PCR primers carry fluorescent labels. Consequently, the PCR products are labeled with two different labels, one at either end.  The key to identifying point mutations (as opposed to deletions) is to create heteroduplexes of mutant and wild-type DNA. Therefore, the PCR products are denatured to single strands and then slowly cooled so that the DNA strands reanneal. During reannealing, some mutant strands will anneal with wild-type strands and the heteroduplex will have a mismatched nucleotide. The enzyme CEL-1 cleaves mismatched DNA. If the PCR product is cleaved by CEL-1, it will have only one fluorescent label, whereas uncleaved DNA (with no mismatches) will have both fluorescent labels. When the PCR products are separated by gel electrophoresis, the digested mutant strands can be identified. **FOOD SAFETY ASSESSMENT AND STARLINK CORN** Many controversies exist over the use of genetically modified crops. The first point to make is that all crops have been genetically modified in some way. Even the oldest variety of edible corn bears no resemblance to its ancestor, teosinte. These changes have occurred through cross-pollination and selective breeding. Thus, using the term *transgenic crop* instead of *genetically modified crop *is more accurate. A main concern with transgenic crops is the health issue. Does the transgenic crop include toxins or allergens, change the nutrient level of the food, or promote antibiotic resistance in humans or cattle? The spread of antibiotic resistance due to reporter genes or selective markers is no longer an issue because new transgenic varieties no longer contain these marker genes. The nutrient level of transgenic food is strictly controlled and evaluated before any transgenic crop is released to the public. The allergenic potential of transgenic crops has caused much controversy. In 2000, an unapproved transgenic corn called **Starlink** was detected in taco shells found in the grocery store. Starlink corn has two transgenes. One makes it resistant to the European corn borer by encoding the toxin from *Bacillus thuringiensis* (see earlier discussion). This Bt transgene is the Cry9C isoform. The second transgene, from *Streptomyces hygroscopicus,* makes the corn resistant to a commonly used broad-spectrum herbicide.  The Cry9C isoform of Bt toxin is much more resistant to stomach acid. Also, after cooking and processing, Starlink corn had a higher concentration of Cry9C protein than expected, suggesting that this protein is more stable than other isoforms of Bt toxin. (In contrast, cooking, processing, and digestive enzymes readily break down the Cry1A isoform of Bt toxin.) Because the findings for Cry9C protein came from only one study, the EPA demanded more tests to ensure that it would not cause an allergic reaction if consumed by the public. The companies that developed Starlink corn pushed the EPA for some sort of approval. The EPA responded by giving split approval---that is, Starlink corn could be grown, as long as it was only used to feed livestock. What the EPA failed to realize is that after corn is grown, it is hauled to the nearest grain elevator. The corn is then mixed with all the other corn in the region and shipped to processing centers. So the company and farmers were following the EPA guidelines in good faith, but the next step in the process made it impossible to keep the Starlink corn separate from all the other varieties. In September of 2000, a coalition of groups opposed to genetically modified foods announced they had detected traces of Starlink in taco shells. Further studies confirmed this and all the products were taken off the shelves. The Centers for Disease Control (CDC) examined all the people who complained of an allergic reaction to the contaminated taco shells. They first determined the type of antibody that the body would produce in response to Cry9C protein. Next they took blood samples and coded them. The blood samples were examined for the presence of Cry9C antibodies by both the CDC and an outside lab. Both concluded that none of the samples contained antibodies to Cry9C. This suggested that any allergic reactions were to some other component in the meals eaten. After this, the company offered to buy back all remaining Starlink corn, providing the farmer with a premium price, so that no more food became contaminated. In addition, all Starlink seed was pulled from the market to prevent its future growth. In all, Starlink was on the market for only 2 years, 1999 and 2000. In 1999, the amount of Starlink grown in the United States represented only 0.4% of the corn crop and in 2000, 0.5%. Because this was such a small percentage of the overall crop, the Cry9C in the taco shells was massively diluted by other varieties of corn. Starlink is no longer grown anywhere in the world, and the EPA has revoked all approval. Assaying allergic potential is critical to development of transgenic crops. In another example, soybean plants were transformed with a gene from the Brazil nut. The gene was intended to increase the methionine content of soybeans, which would improve them as cattle feed. Because many people are allergic to Brazil nuts, the FDA ordered tests for allergenicity by skin prick tests and immunoassays. This transgene was found to cause allergic reactions. The work was discontinued, and none of the transgenic plants were ever released to the public.