6 Recombinant DNA Technology

Outcomes

The transformation of cells with rDNA produces organisms called bioengineered or genetically modified organisms (GMOs). The GMOs contain new traits from another organism. The first GMOs were Escherichia coli cells that were transformed with genes from humans to produce various proteins for pharmaceutical purposes. The production of GM plants became possible after Bob Fraley and others succeeded in using Agrobacterium tumefaciens to transform plant cells with rDNA in the early 1980s (Vasil, 2008a). Since this breakthrough in plant biotechnology, GM crops are now routinely developed and grown in many parts of the globe.

The following points summarize the outcomes of rDNA technology in agriculture.

• From 1996-2007, 1.7 billion acres were cultivated with GM crops in 12 developing and 11 developed countries.
• In 2007, world acreage for GM crops was soybean 57%, maize 13%, and canola 5%. The developing countries accounted for 43% of the world’s acreage of GM crops. U.S. acreage for GM crops was soybean 90%, cotton 85%, and maize 50%.
• More than 1 billion humans have consumed foods derived from rDNA.
• More than 90% of farmers who plant GM crops are small, resource-poor farmers in developing countries.
• GM crops have contributed about $17-18 billion US dollars to the economies of developed and developing countries.

The preceding information was adapted from Vasil, (2008b). Some current statistics on the adoption of genetically engineered crops in the U.S. can be found here:

Preparation of Plant DNA

Samples must be immediately frozen to prevent degradation due to enhanced cellular metabolism. It is important to remember that for recombinant DNA procedures to work, a pure DNA sample must be obtained. The challenge is that plant cells produce numerous other compounds that often act as contaminants and may inhibit cloning or sequencing of the DNA. Also, tissues and organs from the same plant, or different plants, usually contain different compositions of metabolites, for example, proteins, lipids, and carbohydrates. These compounds must be separated from the DNA during isolation. To achieve this, scientists use different molecules’ chemical and physical properties inside the cell. For example, DNA is negatively charged, making it soluble in aqueous solution. However, the polar sugar phosphate groups of the DNA are repelled by nonpolar solutions. Therefore, the final step in many DNA purification protocols involves alcohol precipitation. Other compounds, for example, proteins, can also be easily separated from DNA by altering the salt concentration in the extraction buffer.

DNA isolation methods from plant tissues are becoming more robust and simpler as new technologies emerge. Despite the versatility of the latest DNA isolation technologies, one must understand the science behind various procedures in DNA isolation. Consider the protocol in the following article by Richards et al. (1994), describing methods of preparation of genomic DNA from plant tissue:

Article: Preparation of Genomic DNA from Plant Tissue

As you review the article, note how various compounds and enzymes are incorporated at different steps of the procedures.

Determination of DNA Concentration

The simplest way to determine the concentration of DNA is by using a UV (ultraviolet light) spectrophotometer. Each UV spectrophotometer operates slightly differently. Ensure you know which cuvettes to use and how to clean them. Quartz cuvettes are often used in a spectrophotometer if DNA is analyzed at 260 nm. The nitrogenous bases in DNA (and RNA) absorb UV light at about 260 nm wavelength (abbreviated as A260). As a rule, obtaining an A260 value of 1.0 with pure double-stranded DNA means that the concentration of the DNA is 50 μg/ml. To determine the DNA concentration in a sample, use the following simple formula.

Unknown concentration (μg/ml) = (50 μg/ml) x (measured A260 x dilution factor)

The dilution factor is obtained based on the following:

The DNA you have isolated is for important subsequent steps such as cloning and sequencing. Therefore, you do not want to waste all your valuable samples on concentration determination. Instead, scientists often combine an aliquot of their sample with a larger volume of an appropriate buffer for concentration analysis on a UV spectrophotometer.

For example, a 5 μl aliquot of the original DNA sample mixed with 995 μl of analysis buffer will result in a final volume of 1 ml of DNA-buffer solution. The dilution factor in this case is 200 (1000 μl/5 μl). Such that the following formula can determine the concentration of DNA:

Unknown concentration (μg/ml) = (50 μg/ml) x (measured A260 x 200)

The purity of the DNA can be determined by spectrophotometry because cellular compounds such as proteins and carbohydrates also absorb UV at different wavelengths. Proteins absorb UV at 280 nm, and carbohydrates at 230 nm. Thus, the ratios of the absorbance at 260 nm/ absorbance at 280 nm or 230 nm are estimates of the purity of the DNA sample. For good quality DNA, the ratios should be between 1.65 and 1.85 for A260/A280 and close to 2 for the A260/A230.

Digestion of Plant DNA with Restriction Endonucleases

Restriction endonucleases are a group of enzymes primarily derived from bacteria. Although there are several restriction enzymes, those most useful for rDNA technology recognize specific short sequences in DNA and cleave the DNA at that site to produce cohesive or blunt-ended fragments (Figure 2).

More than 500 restriction enzymes have been identified and can be purchased commercially. One may wish to digest the DNA they have isolated to test for the presence of a SNP marker (as discussed in Plant Breeding Methods) or for Southern blot analysis (below). Thus, one may ask how often a restriction enzyme cuts within a genomic sequence. It is not possible to give an exact answer to this question. However, let us assume there are four bases on any strand of DNA. This means the probability of detecting an A (adenine) at a particular location is 1/4. Since most restriction enzymes recognize specific sequences of 6 bases long, the probability of finding such a site is (1/4) ⁶ = 1 site in every 4,096 base pairs (bp). Assuming you have isolated genomic DNA from maize and want to digest it with a restriction enzyme that cuts every 4,096 (4,100) bp, how many fragments will you obtain? To answer this question, you need to know the genome size of the plant you are working with. The approximate size of the maize genome is 2,500,000,000 bp. Thus, using an enzyme that cuts every 4,100 nucleotides, one would expect to obtain 2,500,000,000 bp/4,100 bp, approximately 610,000 fragments. Note that nucleotide distribution is not always random; thus, frequency may differ for a given DNA. Also, methylation of specific bases in genomic DNA can prevent cleavage at some sites. The subject of DNA methylation was covered in Chapter 2.

Separation of Digested Genomic DNA on Agarose Gel

The only physical features of nucleic acid fragments routinely used for their characterization are their size and nucleotide sequence. The molecular weight of DNA is most conveniently evaluated by electrophoresis in agarose gels. Agarose forms a gel by hydrogen bonding when cooled from the melted state. This gel, an interwoven network of agarose chains, interferes with the movement of DNA through the gel (Figure 3). Pore size, which affects the rate of movement of DNA fragments of a given size, depends on agarose concentration. The gel is submerged in an electrolyte solution; the sample is loaded into wells on one end, and current is applied to facilitate the movement of DNA fragments (Figure 3). Since DNA is negatively charged, it will migrate in the electrical field. Fragments separate according to size. The distance of the migration each time is proportional to 1/log MW. Separation of DNA fragments of 100 to 50,000 nucleotides is routine, though not all on the same gel. Larger and chromosome-sized DNA can also be separated by a modified method called pulsed field electrophoresis:

Polyacrylamide gels can be used for high resolution and separating very small fragments (<200 bp). The principles are the same, but the sieving properties allow the separation of much smaller fragments. Polyacrylamide also enables the resolution of DNA fragments that differ by a single base, and it is used widely for DNA sequencing. Following gel electrophoresis, DNA can be visualized by staining with ethidium bromide or other DNA stains—for example, SYBR Safe DNA Gel Stain.

Detection of DNA by Southern Blotting and Hybridization

A key property of duplex DNA is the ability of strands to separate (denature) and then to reform a duplex (renature) spontaneously. This DNA feature is essential for its biological function and is integral to many of the techniques used in rDNA studies. Stable duplex formation in a solution requires a minimum number of complementary base pairs, which depends on the reaction conditions, such as the solution’s temperature and salt concentration. When the new duplex is formed between DNA strands from different sources, for example, DNA cloned on a plasmid with genomic DNA from different organisms, the reaction is called hybridization.

A more sensitive method for analyzing complex mixtures or unknown samples involves hybridizing nucleic acids. The procedure for DNA detection on a blot is called Southern blot hybridization, after the name of the individual who developed it (Figure 4). As mentioned earlier, a band on a gel represents many fragments of the same length. For a 1 kb fragment, the lower end of ethidium bromide sensitivity is 10–100 ng (0.15 pmol) or 4 x 10¹⁰ molecules. In contrast, sensitivity is 1 pg or 4 x 105 molecules for hybridization. Also, a band on a gel may represent many different sequences that are approximately the same size. Hybridization also allows the detection of a single species in a complex mixture, since DNA will only hybridize to its complement.

The stringency applied to the process is important in designing a hybridization experiment. Hybridization stringency can be thought of as the degree to which hybridization conditions allow imperfectly paired or mismatched duplexes to form. The stringency can be determined by all factors that affect duplex stability (Table 1). At very high stringency, only perfectly paired duplexes will form. As stringency is decreased, small amounts of mismatch will be allowed. A lower stringency is useful for hybridizing a related but not identical sequence in another organism.

Principles of Sequencing

Sanger’s Dideoxy DNA-Sequencing Procedure

Fred Sanger developed the Sanger DNA-sequencing procedure (dideoxy DNA sequencing) in the 1970s. The method uses enzymatic reactions to incorporate specific terminators of DNA chain elongation called 2′,3′-dideoxynucleoside triphosphates (ddNTPs). The ddNTP molecules can be incorporated into the growing DNA chain through their 5′ triphosphate groups. However, because they lack a hydroxyl (OH) group on the 3′-C of the sugar moiety (Figure 5), they cannot form a phosphodiester bond with deoxynucleotide triphosphates (dNTPs) during the sequencing reaction, resulting in termination of DNA chain elongation.

Maxam & Gilbert DNA-Sequencing Procedure

Allan Maxam and Walter Gilbert developed the Maxam and Gilbert DNA-sequencing procedure in 1977. The procedure is based on the chemical degradation of DNA chains. In this procedure (Figure 6), a segment of DNA is labeled at one end with a radioactive label (³²P ATP). A solution containing the labeled DNA is distributed into four different tubes. A chemical that destroys one or two of the four bases (G, A+G, C, C+T) in the DNA is added to each tube. The addition of a chemical called piperidine to the DNA treated with base-modifying chemicals results in cleavage of the strand at the position of the modified base. It is important to control the reaction conditions such that only a few sites are cleaved per DNA molecule. The length of the cleaved fragments depends on the distance between the modified base and the labeled end of the DNA segment. For example, if there are C residues 4, 7, and 10 bases away from the labeled end, the fragments from cleavage at the modified C residue will be 3, 6, and 9 bases in length. The cleaved products of each of the four reactions (G, A+G, C, and C+T) will be evaluated side by side on an acrylamide gel to separate them based on size. The gel is assessed by autoradiography, and the banding pattern on film is scored to determine the DNA sequence.

Next Generation Sequencing

Next-generation (nextgen) sequencing (NGS) is a high-throughput sequencing method combining parallel processes to produce thousands or millions of sequence data points at once. One main difference of next-generation sequencing from the conventional sequencing technologies described above is that no cloning of DNA fragments in E. coli is required for next-generation sequencing. The NGS is divided into (i) second-generation and (ii) third-generation sequencing. The primary difference between the two is whether a single molecule or multiple molecules are used as the template for sequencing. In the second generation, sequencing PCR products representing thousands of copies of a DNA molecule are used as a template. In the third-generation sequencing, a single DNA molecule is sequenced using a single DNA polymerase molecule. Examples of the second-generation nextgen sequencing technologies are:

Pyrosequencing or 454 Sequencing

The 454 sequencing uses a procedure known as sequencing by synthesis, or pyrosequencing. A video describing pyrosequencing can be viewed here.

Illumina sequencing

The Illumina system uses a terminator-based method to detect single bases as they are incorporated into a growing DNA strand. Watch a video describing Illumina sequencing.

SOLiD Sequencing

The SOLiD system is based on a technique of oligonucleotide ligation and detection. A video with more information about the system can be viewed here.

Ion Torrent Sequencing

Ion Torrent Sequencing, or amplicon sequencing, directly translates DNA bases into digital information on a semiconductor chip. A video with more details on the system can be viewed here.

Examples of third generation sequencing are:

Single Molecule Real Time (SMRT) Sequencing

Pacific Biosciences developed the SMRT system and uses a single DNA polymerase molecule-based approach for real-time sequencing of single DNA molecules. A video with more information about the system can be viewed here:

Oxford Nanopore

In Nanopore Technologies, protein-nanopores are applied to determine the sequence of a DNA molecule. In Oxford Nanopore Technologies, a specifically designed nanopore is inserted into a membrane created by a synthetic polymer. This membrane has very high electrical resistance. Applying a potential across the membrane generates an ionic current through the nanopore. A single DNA molecule passing through the nanopore causes characteristic disruptions in the current, detected as signals and specific to the DNA building blocks, i.e., deoxynucleotides (Figure 5). Each nucleotide produces a unique signal, which calls the bases in real-time. Thus, we gather the sequence of a molecule as it travels through a nanopore in real-time. This video describes Nanopore technology.

Helicos High Speed Gene Sequencing

Illumina described earlier that the Helicos sequencing technology also uses a terminator-based method to detect single bases incorporated into a growing DNA strand. The main difference between Helicos with the Illumina system is that in Helicos, only a single molecule is sequenced, whereas in Illumina, a cluster of copies of a DNA molecule is sequenced. There are also other differences. Watch this video and compare the differences with the Illumina sequencing technology:

Sequence Alignment and Assembling Aspects of Next-gen Sequencing

In general, next-gen second-generation technologies result in a larger number of reads that are shorter than those produced using capillary electrophoresis. Therefore, next-gen sequencing requires more robust algorithms to assemble the large quantity of data generated. Third-generation next-gen sequencing, the SMRT system, has alleviated this problem by producing more than 1,000 bases long sequences.

Several academic and private institutions provide core services for next-generation sequencing, for example, the MGH Next Generation Sequencing Core and Beckman Coulter Genomics.

Polymerase Chain Reaction

Polymerase chain reaction (PCR) is a method by which millions of copies of a DNA fragment are produced in a test tube in minutes or a few hours. The basic steps in the PCR reaction were discussed in Plant Breeding Methods. The method was invented by Kary Mullis in 1985 and exploits certain features of DNA replication. Refer to Crop Genetics and Chapter 1 for information about DNA replication .

The basic materials for conducting PCR using DNA were listed in Plant Breeding Methods. To perform a PCR reaction, small amounts of the DNA template are added to special PCR tubes (with thin walls for efficient heat exchange) that contain the following components:

1. Reaction buffer required to maintain optimum pH
2. Forward and reverse oligonucleotide primers are required to recognize the two ends of the target fragment
3. Taq polymerase that maintains activity at very high temperatures for DNA synthesis
4. dNTPs (A, C, G, T), building blocks for the new DNA strand synthesis
5. Magnesium chloride as a cofactor for the activity of Taq polymerase

The PCR reaction is performed in a thermocycler (Figure 7). A thermocycler contains a heat block for which a computer program controls the temperature. It is important to test all the variables in a PCR protocol for determining the best PCR conditions and concentrations of reagents. This is called PCR optimization. Once a PCR reaction is optimized, you may scale up your protocol to include larger volumes and number of samples to be analyzed, e.g., high-throughput PCR assays for screening a population. When hundreds of individuals are screened to search for markers of interest, technicians prepare large batches of “master mix” that contain all the correct concentrations of polymerase, buffer, dNTP, and MgCl2. A large batch of primer solution is also prepared. When thousands of samples are screened, robotics approaches are commonly used.

Reverse-transcription and PCR

Recall in Chapter 1 that you learned about RNA synthesis from DNA through transcription. Transcription is an important step in gene expression. The mRNA produced can be isolated and “copied” back to DNA by reverse transcription (Figure 8). The first step of reverse transcription mimics a strategy used by retroviruses (e.g., HIV) with RNA genomes. As part of their gene-transmission package, retroviruses also contain an enzyme called RNA-dependent DNA Polymerase, commonly referred to as reverse transcriptase. After infecting a host cell, the retrovirus uses its reverse transcriptase to copy its single-stranded RNA genome into a complementary DNA (cDNA) strand. The reverse transcriptase then synthesizes the second DNA strand from the first strand to make a double-stranded DNA copy, which integrates into the host genome.

The cDNAs produced in vitro can be used for PCR analysis, like chromosomal DNA. The combination of reverse-transcription and PCR (RT-PCR) is valuable in gene cloning and mRNA quantification.

Reverse Transcription Quantitative PCR (RT-qPCR)

Conventional PCR with end-point detection is sufficient for certain laboratory assays, such as confirming the presence or absence of a transgene. However, in many cases, a researcher may wish to quantify how strongly the transgene is expressed across different plant tissues.

Unlike standard PCR, which only reveals whether amplification has occurred after the final cycle, quantitative PCR (qPCR), or real-time PCR, monitors the accumulation of PCR products during each amplification cycle. Fluorescent dyes or probes enable real-time detection of double-stranded DNA as it is synthesized. For gene expression analysis, RNA must first be reverse transcribed into cDNA (Figure 8). Using primers specific to the sequence of interest, this cDNA is then amplified by qPCR, allowing quantification of transcript levels. The primary advantage of qPCR over conventional PCR is its ability to measure transcript abundance with high sensitivity and accuracy across a broad dynamic range, rather than simply detecting the presence or absence of a target sequence.

Use of PCR in Cloning Genes

Once a particular gene’s sequence is known, PCR makes it possible to isolate that gene from any DNA or cDNA sample. Recall in Chapter 1, you learned that at the genomic level, a gene is made up of regulatory sequences, coding, and non-noncoding sequences. Thus, if the goal is to use PCR to isolate both regulatory and non-regulatory sequences of a gene, the approach would be to use DNA as the starting material. Alternatively, isolating the gene from cDNA would be the strategy if only the coding sequence is required. The cDNA must be synthesized from tissues expressing the gene of interest. Thus, prior knowledge of where the gene is functional is important in constructing the cDNA molecules for cloning the gene of interest.

Cloning Vector Definition and Requirements

A cloning vector is a specialized DNA sequence that can enter a living cell and detect its presence to a researcher by conferring a selectable property on the host cell (e.g., resistance to antibiotics), with the means for self-replication. A vector must also possess easily distinguishable physical traits, such as size or shape, to allow purification from the host cell’s genome.

In cloning genes by PCR, restriction enzyme sites (Figure 9) are added at the 5′-end of the primers to facilitate cloning of PCR fragments. A few additional nucleotides (~6 nts) are added at the 5′-end of the restriction sites to enable restriction digestion of PCR products before cloning in a plasmid vector. Alternatively, PCR products may be cloned directly into a T-vector without restriction digestions in E. coli. A fter cloning into E. coli, the fragment is analyzed by sequencing and then sub-cloned into suitable vectors for expression studies.

Ligase Enzyme and Gene Cloning

Cutting and joining vector and DNA fragments from different origins results in rDNA. Recall that restriction endonucleases are used to cut DNA. To join DNA molecules together, an enzyme called DNA ligase is used. The enzyme DNA ligase joins restriction fragments by forming new phosphodiester bonds. The ligated vector and DNA fragment can now be transformed into a host cell for replication and expression.

The transformation of E. coli takes several steps. First, a gene of interest is inserted into a plasmid that contains a selectable marker, usually encoding for resistance to an antibiotic. Second, the plasmid construct containing the gene of interest is transformed into bacterial cells by briefly exposing the mixture of ligated plasmid-DNA fragment (rDNA molecule) and bacterial cells to cold (0 °C) and heat (37–42 °C). The next step is to grow the transformed cells on selection media containing an antibiotic. Only the cells that have been transformed with the plasmid containing the gene of interest and the marker for resistance to the antibiotic will survive. In addition to using an antibiotic, plasmid vector systems that include the lacZ gene encoding β-galactosidase allow for easier selection of positive colonies that may harbor the rDNA molecule of interest.

Types of Cloning Vectors

One vector type is a plasmid (Figure 10), an autonomously replicating extra-chromosomal circular DNA faithfully passed on to progeny. Plasmids are double-stranded circular DNA ranging from about 1 kb to 200 kb. The most useful plasmids for cloning are 2–10 kb because smaller plasmids are easier to manipulate and usually produce a higher copy number when grown in bacterial host cells.

Generally, not more than 10 kb is cloned into plasmids. A vector called bacteriophage lambda (e.g., Figure 10) is used for highly efficient cloning of large DNA fragments. Lambda is a linear duplex of about 48.5 kb with 12 bp cohesive ends, which permit the cloned DNA to circularize when introduced into the cell. Large chromosomal DNA fragments close to 23 kb are stable when introduced into a lambda phage vector.

A third vector type is called a cosmid (Figure 11). A cosmid is a plasmid with cohesive ends like lambda, which allows cloning much larger fragments (35–45 kb). A cosmid replicates like a plasmid after being introduced into the host bacterial cell. The large size of cosmids makes them a challenge to work with.

In the examples above (Figures 9 to 11), the T7, SP6, and T3 primer-binding sites can be used to validate the presence of the cloned insert (from several base pairs to about 10 kb) by PCR.

Ti Plasmid Vector and Plant Transformation

Crown galls are plant tumors that develop at the site of infection by certain species of the soil bacterium Agrobacterium. Rather than entering plant cells, Agrobacterium transfers a DNA segment known as T-DNA from its circular, extra-chromosomal tumor-inducing (Ti) plasmid into the host cell genome. Ti plasmids are retained in Agrobacterium because part of their T-DNA encodes genes for unusual amino acids that the bacterium can utilize. The T-DNA also carries genes that disrupt the host plant’s hormone balance, triggering abnormal cell growth and tumor formation. Scientists have harnessed Agrobacterium’s ability to stably integrate T-DNA into plant genomes as a tool for introducing recombinant DNA into plant cells. They removed the tumor-inducing genes from the T-DNA and engineered the plasmid to replicate in both E. coli and Agrobacterium. Replication in E. coli is advantageous because it allows verification of the cloned gene and produces large amounts of construct DNA for sequencing and later transfers into Agrobacterium. Plant transformation using Agrobacterium was introduced in Plant Breeding Methods and will be explored further in Chapter 7.

Gene Libraries in Plasmid, BAC, and YAC Vectors

There are two types of gene libraries, genomic and cDNA libraries. A genomic library is a collection of cellular clones containing sequences of an organism’s entire genome. Each clone contains only a single genomic DNA fragment. Genomic libraries are made by digesting genomic DNA with a restriction enzyme, ligating every fragment to a vector, and transforming such rDNA molecules into host cells. A cDNA library is a collection of cloned cDNA molecules generated by reverse transcription of mRNA extracted from plant tissues that are actively expressing the genes of interest (Figure 8). Unlike genomic libraries, cDNA libraries lack promoters and introns, since they are derived directly from processed mRNA rather than genomic DNA.

Now, let’s ask the following question. How many clones should be present in a library? To answer this question, one must know the genome size of the plant species they are working with. Let us take the example of maize. As mentioned earlier, the maize genome is estimated to contain 2,500 Mb (2,500,000 kb). If you were to reliably digest a 2,500,000 kb fragment from a maize cell into a series of 250 kb restriction fragments, you would generate 2,500,000/250 = 10,000 genomic fragments. To clone such large fragments, you would need to use a special cloning vector, e.g., a bacterial artificial chromosome (BAC) vector, pBeloBAC11, if you clone each of the 10,000 genomic fragments into this vector and transform them into E. coli, you will obtain a perfect genomic library. The number of clones in this “perfect” library represents what is known as a genomic equivalent. Therefore, to compute the number of clones that constitute one genomic equivalent, divide the length of the genome by the average size of the inserts carried by the cloning vector.

In addition to BAC vectors, one can use a yeast artificial chromosome (YAC) vector to clone large plant DNA fragments in individual yeast cells. YAC clones are much bigger and can accommodate large DNA inserts up to 10 Mb.

BAC and YAC vectors are instrumental in the construction of genomic libraries because they can accommodate large DNA inserts, reducing the number of clones required for a genome equivalent library. The fewer the number of clones, the less effort is placed in searching the clones for the sequence of interest. For example, you have seen above that the genomic equivalent of BAC clones with 250 kb inserts contains 10,000 clones. However, a genomic equivalent of plasmid clones with 10 kb inserts would contain 250,000 clones.

Now the main question is what the size of a library should be. As outlined above, we cannot create a perfect library due to sampling issues. The larger the library, the better the chance we can identify our DNA sequence of interest. Usually, the possibility or probability of finding a random DNA fragment in a three-genomic equivalent library is 0.95, while in the six-equivalent library it is 0.99. In general, a six-genomic equivalent library is constructed to start with. If one fails to obtain the fragment of interest, they may continue to construct a larger genomic library in the same or an alternative vector.

Protein expression in bacteria

Developing protein expression systems is an ongoing effort in both industrial and academic research settings. The development of expression vectors has allowed scientists to produce plant proteins in bacterial cells. The most popular expression systems are bacteria, for example, E. coli.

The process begins with ligating a DNA fragment containing the coding region of the gene of interest to a plasmid with promoter and other regulatory sequences recognized by the transcriptional machinery of the host cell. Most expression vectors provide options to fuse the gene of interest to a “protein tag” that can be used as a “handle” to isolate the desired protein from a pool of thousands of E. coli proteins. For example, the pGEX vector system from Sigma provides the option to link the protein of interest to glutathione-S-transferase (GST) with a short amino acid sequence that can be cleaved by thrombin. The GST-fused recombinant protein is separated from the solution using Glutathione Sepharose, which binds GST. Subsequently, the GST portion on the recombinant protein is removed by digestion with thrombin.

A concern about using bacterial cells as hosts to produce plant proteins is that some plant proteins require additional modifications that can only occur inside a plant cell. One of these modifications is the ordered insertion of sugars into some proteins that occur in specialized compartments of the plant cell. Lack of proper modifications of proteins can affect their folding and function. To overcome this problem, scientists use yeast as hosts to express plant proteins that require additional modifications related to their function in a eukaryotic organism. Alternatively, the protein may be expressed in an insect cell line through baculovirus-mediated infection. However, at times, even yeast or insect cells may fail to provide a suitable environment for expressing proteins of plant origin. In such cases, the best solution is to express the protein in plant cells.

Gene Cloning by Chromosome Walking, Chromosome Jumping

After creating a library of thousands of genomic fragments, the next step is isolating interest gene from the library. Suppose you know by genetic information that a particular trait you are interested in is in a chromosomal region. If you do not know the gene responsible for the trait, how would you identify the gene in a region that may be more than 2 Mb? The answer is, you would start by selecting genomic clones that lie closer to the region of interest and then “walk” through the region by identifying overlapping clones from your genomic library.

Recall that a genomic library is prepared from fragments obtained from digesting genomic DNA. The gene responsible for your trait will be in your library’s thousands of DNA inserts. To identify clones close to the gene of interest, you will first need to find DNA markers linked to the trait you are interested in. This is done by searching for markers that co-segregate with the trait in a plant population. In other words, recombination between your gene (trait) and markers is not seen.

The next step is to screen the genomic library with the co-segregating markers (probes) in isolating clones that hybridize to those markers. The ends of the clones that hybridize to co-segregating markers are used as probes to identify overlapping clones. The procedure is repeated to systematically “walk” toward the gene of interest, one clone at a time. You might think the procedure is lengthy, and you are right. For example, it may take more than a month to walk >100 kb. For that reason, chromosome jumping (Figure 12) is used to cover a region quickly. The chromosome jumping method is described in this article.

It is worth noting that with the advent of next-generation (nextgen) sequencing (see below) technologies and genome sequences for many crop plants, one may avoid arduous traditional map-based cloning or chromosomal jumping experiments in cloning genes based on their map positions. A new approach, based on next-generation sequencing of the bulk of recessive mutants following bulk-segregant analyses, has been demonstrated for identifying genes in Arabidopsis.

Gene Silencing as a Tool to Uncover Gene Function in Biochemical Pathways

Recall from Chapter 2 that RNA interference (RNAi) can be accomplished more efficiently by expressing a portion of the target gene, usually around 400 bp, that has been engineered as an inverted repeat in transgenic crop plants. Following transcription of this engineered gene, the RNA molecules form a hairpin structure that is then cleaved into small fragments of double-stranded RNA, which interfere with the accumulation and function of the endogenous mRNA molecules of the target gene. Below is an example of how scientists applied RNAi to investigate gene function in the jasmonic acid (JA) pathway (Figure 13).

JA is a key plant hormone that regulates development and mediates responses to stresses such as leaf injury. Research by Paul Staswick and his team at the University of Nebraska–Lincoln revealed that amino acid conjugates of JA, particularly jasmonoyl-L-isoleucine (JA-Ile), serve as crucial signaling molecules (Staswick et al., 2002; Staswick & Tiryaki, 2004; Suza & Staswick, 2008). In Arabidopsis, the JASMONATE RESISTANT 1 (JAR1) gene encodes an enzyme that conjugates JA with isoleucine (Figure 13A). Building on this knowledge, Suza et al. (2010) applied RNAi (Figure 13B) to identify and characterize a tomato gene predicted to encode an enzyme that catalyzes JA-Ile formation in response to leaf wounding (Figure 14), analogous to the role of JAR1 in Arabidopsis.

Gene Silencing as a Tool for Trait Improvement

Consider mutation breeding described in Plant Breeding Methods. One can create a mutant population for selecting desirable phenotypes. Most mutations lead to loss of gene function. If the gene is a negative regulator of a particular trait, loss of that “switch” due to a mutation will lead to expression of a specific trait. One good example is the mlo locus in barley. Mlo is a negative regulator of non-race-specific powdery mildew resistance in barley. A mutation in the Mlo gene led to the mlo allele, which now provides broad-spectrum resistance to all races of the powdery mildew pathogen. This is one of the success stories of mutation breeding in barley. However, the entire process of mutation breeding is labor-intensive. We can accomplish the same objective by manipulating targeted genes in transgenic crop plants.

Decreased expression of an existing gene, such as Mlo in barley, could be accomplished quite effectively by introducing a copy of the gene in the antisense (reversed) orientation. A normal transcribed region, or a portion of it, is reversed so that the RNA molecules transcribed from the engineered gene will hybridize to the RNA molecules of the endogenous native gene, leading to the degradation of the mRNA encoded by the endogenous gene. This process is highly gene-specific and leads to downregulation of the targeted gene, even though the original gene is not mutated in any way. The downregulation of the Mlo gene is a result of RNAi.

RNAi provides a powerful tool for genetic study and for biotechnological crop improvement. The following are examples of the application of RNAi for Crop Improvement.

Example 1: Enhancing Carotenoid and Flavonoid Content in Tomatoes

Tomato contains lycopene and β-carotene in disproportionate amounts. Lycopene is an important antioxidant that prevents heart disease and cancer in humans. β-carotene is a precursor to vitamin A. Vitamin A is essential to humans, and failure to consume adequate vitamin A is a leading cause of health problems and infant mortality in developing countries. Efforts to genetically enhance both lycopene and β-carotene had previously been unsuccessful. However, Davuluri et al. (2005) took an approach to shut down the expression of the DE–ETIOLATED1 (DET1) gene in tomato fruits. The DET1 gene is a regulatory gene that represses several signaling pathways modulated by light. A mutation in this has been shown to increase the pigment concentration in tomato. Mutants also contain increased levels of flavonoids and carotenoids. Davuluri and co-workers fused a section of the DET1 coding sequence in both the reverse and forward orientations to a promoter (Figure 15) that only directs the expression of the “inverted repeats” in the fruit. The inverted repeats of DET1 resulted in the formation of dsRNA. The DET1 dsRNA was a substrate for dicer to produce siRNAs (about 21-25 nucleotides in length). These DET1 siRNAs are complementary to the region of DET1 that was used to produce the inverted repeats. The binding of DET1 siRNAs to the DET1 mRNA transcribed from the DET1 gene in fruits resulted in patches of DET1 siRNA-DET1 RNA duplex. The double-stranded patches trigger the degradation of the entire DET1 mRNA and suppression of the DET1 gene in fruits. The result of DET1 silencing was enhanced production of both lycopene and β-carotene in transgenic fruits.

Fruit-specific RNAi-mediated suppression of DET1 enhances carotenoid and flavonoid content in tomatoes. To read about this, see http://www.nature.com/nbt/journal/v23/n7/pdf/nbt1108.pdf

Example 2: Caffeine-Free Coffee

Scientists and plant breeders have sought to remove caffeine from coffee for over a century. Unfortunately, the chemical processes used currently to produce decaf coffee also destroy the flavor of the highly consumed beverage.

Caffeine is produced in coffee plant cells from a natural plant chemical called xanthosine. Caffeine biosynthesis involves three subsequent additions of methyl groups to xanthosine (Figure 16). The methylation reactions are controlled by three distinct methyltransferases, xanthosine methyltransferase (XMT), 7-N-methylxanthine methyltransferase (MXMT; theobromine synthase), and 3,7-dimethylxanthine methyltransferase (DXMT; caffeine synthase). Ogita and colleagues from Nara Institute of Science and Technology in Japan used RNAi to downregulate the expression of MXMT in coffee plants. The result was transgenic RNAi plants with 70% less caffeine than controls (Ogita et al., 2003, 2004). The commercial application of this strategy awaits further improvement to target RNAi only to coffee beans and further to reduce caffeine content beyond the current 70% level. Other reports claim that the transgenic RNAi coffee plants show reproductive abnormalities, making it challenging to produce sufficient seed for breeding purposes (Borrell, 2012).

Read: Plant biotechnology: Make it a decaf

Read: Producing decaffeinated coffee plants

Read: Application of RNAi to confirm theobromine as the major intermediate for caffeine biosynthesis in coffee plants with potential for construction of decaffeinated varieties.

Example 4: Increased Resistance to Nematodes

Plant parasitic nematodes cause significant crop losses worldwide. In the early 2000s, the cost of agriculture due to plant parasitic nematodes was estimated to be around US$125 billion (Chitwood, 2003). Control of nematodes relied heavily on costly and toxic chemicals. Cultural systems such as crop rotation were less effective because the nematodes can remain dormant (e.g., cyst nematode) for many years. The discovery that when short RNAs are ingested by the model worm Caenorhabditis elegans result in gene

silencing, inspired researchers to apply this technology to target nematodes (Figure 17). One of the targets of economic importance is the soybean cyst nematode (Heterodera glycines), a major pest of soybean responsible for billions of dollars each year in the US. Transgenic soybean expressing dsRNA for a gene involved in reproduction in H. glycines showed a significant reduction in the number of H. glycines eggs in root tissue (Steeves et al., 2006).

Read: RNAi in plant parasitic nematodes

Read: Controlling soybean cyst nematodes by RNAi

Example 5: Engineering Male Sterility

Male sterility is an important trait in hybrid seed production. To demonstrate the usefulness of RNAi in plant breeding, let us consider the induction of male sterility by reducing the expression of key genes involved in floral development. The maize Zea Apetala1 (ZAP1) encodes a transcription factor that controls inflorescence architecture. The expression of ZAP1 is restricted to the sterile organs of the male floret (Mena et al., 1995)—Consequently, RNAi silencing of ZAP1 results in male sterility (Figure 18).

Overexpression of Transgenes for Crop Improvement

Example 1: Improving Cotton Tolerance to Drought and Salt Stress

Abiotic stresses, including drought and salinity, seriously threaten agriculture worldwide. Genes controlling many steps in plant response to abiotic stress and their associated metabolic and signaling networks have been identified and characterized. Researchers are using this knowledge and lessons learned from plant species adapted to hostile environments to figure out ways to engineer abiotic stress tolerance.

A gene from Arabidopsis encoding a membrane-localized proton pump called AVP1 was overexpressed in several plant species, including cotton (Pasapula et al., 2011). The result was enhanced tolerance to drought and salinity stress. Also, transgenic cotton overexpressing AVP1 had 20% higher fiber yield than controls under dry conditions in the field, suggesting potential for application of this approach in other crop species.

Example 2: Improving Rice Tolerance to Salinity Stress

Sodium ion (Na⁺) is toxic to plant cells when present in excessive levels. Plant cells have a mechanism to regulate their Na⁺ concentration through the action of Na⁺ transporters. However, the degree of tolerance to salt varies markedly among plant species. An Arabidopsis gene encoding a membrane-localized sodium (Na⁺) transporter was overexpressed in rice’s root cortical and epidermal cells (Plett et al., 2010). The resulting transgenic rice grown under high salt had reduced Na⁺ in the shoot due to decreased root-to-shoot flux of Na⁺. This genetic modification enabled the plants to gain fresh weight under salinity stress.

Other Crop Improvement Approaches

Example: Transgenic Virus-Resistant Papaya

The advent of recombinant DNA technology enabled scientists to study cross-protection by transforming plants with viral genes. Cross-protection refers to the phenomenon in which infection by a mild strain of a virus protects a plant from subsequent infection by more severe strains. This strategy has been successfully applied to control certain plant virus diseases.

Plant virus infections can devastate agriculture, as seen in papaya production in Hawaii during the early 1990s. The papaya ringspot virus (PRSV) caused systemic rotting, wilting, chlorosis, and the appearance of yellow spots. By 1997, PRSV had nearly destroyed Hawaii’s papaya industry, forcing many farmers out of business. To combat PRSV, researchers built upon earlier work from Roger Beachy’s laboratory (Nelson et al., 1988), which showed that transgenic plants overexpressing the tobacco mosaic virus (TMV) coat protein (CP) exhibited resistance to infection (Figure 19). Using a similar approach, Fitch et al. (1992) developed transgenic papaya plants resistant to PRSV by overexpressing the PRSV coat protein gene. This intervention ultimately rescued the papaya industry in Hawaii and demonstrated that transgenic virus resistance can be a powerful strategy for managing viral diseases in horticultural crops.

Read: Virus Tolerance, Plant Performance of Transgenic Tomato Plants Expressing Coat Protein from Tobacco Mosaic Virus

Read: Virus-Resistant Papaya Plants Derived from Tissues Bombarded with the Coat Protein Gene of Papaya Ringspot Virus

Read: Transgenic Virus-Resistant Papaya: From Hope to Reality for Controlling Papaya Ringspot Virus in Hawaii

Transient Gene Expression

All examples in the previous sections involved stable transformation of plant cells. Stable transformation means that a plant cell has received and has stably integrated a transgene into its genome. In some cases, such as promoter analysis or to verify that a construct works properly, it isn’t necessary to regenerate plants after transformation. Genes may be expressed in cell nuclei immediately or shortly after they have been treated with the appropriate DNA construct, even though new plants have not been regenerated. In this case, it is not necessary to use a selectable marker.

With transient expression, DNA may or may not be integrated into the genome. A reporter gene is useful for this purpose. To define the elements of a promoter required for expression of a gene, it can be dissected by creating deletions of the promoter and by mutating certain candidate nucleotides that may have a regulatory function. The modified promoters derived from either deletion or mutation are investigated for their activity by fusing them to a reported gene, the expression of which can easily be monitored. The advantage of transient expression studies is obviously that they are much more rapid, avoiding the many months needed for regeneration, recovery, and analysis of progeny. A disadvantage is that it may not be as accurate, since it is difficult to know how much DNA is in the nuclei of cells that are assayed. However, there are ways to standardize this by co-transforming with a second gene whose expression level does not change under the conditions being investigated. The other pitfall of transient expression study could be artifact in expression levels and patterns, significantly different from the expression patterns of the same gene in stably transformed plants. For these reasons, transient expression is often a preliminary approach, and more precise results should be obtained from stably transformed plants.

An example of a transient method of gene silencing is called viral-induced gene silencing, or VIGS for short. In the VIGS method, rather than transforming a plant, the silencing construct is incorporated into a viral genome, and the plant is then infected with the virus. The silencing DNA spreads from cell to cell in the plant along with the viral DNA or RNAs, and then the silencing portion interferes with the accumulation of the plant mRNAs corresponding to the gene construct.

Application of Next-Generation Sequencing in Crop Genetics and Breeding

Next-generation sequencing (NGS) technologies have transformed crop genetics and breeding by enabling a wide range of applications. These include the development of genomic resources, molecular marker discovery, QTL mapping, wide crosses and alien gene introgression, expression analysis, association genetics, and population biology. A more detailed discussion of this subject is provided in Molecular Plant Breeding.

Traditionally, map-based cloning was a lengthy and resource-intensive process, often requiring years of high-density and high-resolution mapping. With the advent of NGS, this process has been greatly simplified. Instead of constructing extensive genetic maps, researchers can now create two bulks by pooling DNA from homozygous families carrying different alleles at the target locus. This bulked segregant analysis approach, combined with NGS, dramatically reduces time and effort.

The rapid decline in sequencing costs has further accelerated genomics research. Entire genomes can now be sequenced quickly, leading to breakthroughs in functional genomics (studying gene expression patterns), metagenomics (sequencing microbial communities from specific environments), comparative genomics, molecular marker development, genome-wide association studies (GWAS), and cloning of genes underlying traits of interest.

Sequencing genomic DNA, including bacterial artificial chromosomes (BACs), reduced representation genomes (RRGs), or cDNA from reference genotypes, can generate essential genomic resources such as expressed sequence tags (ESTs), gene catalogs, and draft genome assemblies. These resources provide critical insights into genome architecture and serve as the foundation for crop genetics research.

Another powerful application of NGS lies in parental genotyping of mapping populations or wild relatives. This accelerates the development of molecular markers such as simple sequence repeats (SSRs) and single-nucleotide polymorphisms (SNPs). These markers enable constructing genetic maps, identifying QTLs, and tracking alien genome introgression during wide crosses. Once QTL-associated markers are identified for a trait of interest, they can be applied in marker-assisted selection (MAS) to efficiently select progeny carrying favorable alleles.

For developing functional or gene-based markers, NGS of cDNA from contrasting genotypes can identify candidate genes associated with specific traits. Expression mapping of these candidate genes and phenotyping of segregating populations can reveal expression QTLs (eQTLs). Markers associated with eQTLs serve as highly reliable tools for MAS in crop breeding programs.

NGS also plays a major role in association genetics and population biology. Genomes or pooled PCR products from thousands of candidate genes can be sequenced across hundreds of individuals using barcoding strategies. The resulting sequence data enables the identification of SNPs and haplotypes across genes or genomes, which can then be used for association studies and population-level analyses.

Chapter Summary

Recombinant DNA technology has contributed significantly to the development of agricultural biotechnology. The transformation of cells with rDNA produces organisms called bioengineered or genetically modified organisms (GMOs) with improved traits such as pest and disease resistance, herbicide tolerance, and enhanced nutritional content. Sequencing is the determination of the order of the nucleotides on a DNA molecule. The more recent sequencing technologies are referred to as next-generation and third-generation (NGS and long-read sequencing technologies, respectively). Nextgen technologies produce many short reads compared to the earlier sequencing approaches developed by Sanger and Maxam-Gilbert. Therefore, more robust algorithms are required to assemble and analyze next-gen data (and to identify genetic variations, annotate genes, and study complex genomes). The tools for plant rDNA technology include vectors, restriction enzymes, ligation enzymes, bacterial hosts, methods to isolate and multiply nucleic acids, methods to quantify nucleic acids, and Agrobacterium as a vector to insert foreign DNA into plants, which enables stable transformation and functional analysis of genes.