​Introduction

Forward Genetics vs. Reverse Genetics

Variants help us understand the 'normal.' Variation can be measured at many scales – from macro (body size, morphology) to different levels of micro variation (crude protein profiles to DNA sequence variation). Forward genetics refers to a process where studies are initiated to determine the genetic underpinnings of observable phenotypic variation. In many cases the observable variation has been induced using a DNA damaging agent (mutagen) but also may be naturally occurring. The investigator eventually ends up sequencing the gene or genes thought to be involved (Figure 1). 

Figure 1

With the advent of whole genome sequencing many researchers are now in a very different position. They have access to all of the gene sequences within a given organism and would like to know their function. So, instead of going from phenotype to sequence as in forward genetics, reverse genetics works in the opposite direction – a gene sequence is known, but its exact function is uncertain. In reverse genetics, a specific gene or gene product is disrupted or modified and then the phenotype is measured (Figure 1). Here we will overview some of the techniques for reverse genetics with a special emphasis on the TILLING (Targeting Induced Local Lesions IN Genomes) technique which is being utilized on plant pathogens in the genus Phytophthora.

Specific Reverse Genetic approaches

The goal in reverse genetics is to investigate the impact of induced variation within a specific gene and to infer gene function. The process of disruption or alteration can either be targeted specifically as in the case of gene silencing or homologous recombination or can rely on non-targeted random disruptions (e.g., chemical mutagenesis, transposon mediated mutagenesis) followed by screening a library of individuals for lesions at a specific location. Following is a brief overview of some commonly used targeted and non-targeted approaches.

Gene Silencing

RNA interference (RNAi) is the process by which expression of a target gene is inhibited by antisense and sense RNAs. It works based on the ability of double-stranded sequences to recognize and degrade sequences that are complementary to them (Lewin 2004). RNAi was first discovered in Caenorhabditis elegans when the introduction of double-stranded RNA was observed to be an efficient method for silencing gene expression (Fire et al. 1998; Kuttenkeuler and Boutros 2004). RNAi- based silencing is an exciting strategy for reverse genetics (Waterhouse, Graham, and Wang 1998). RNA interference has recently become a powerful tool to silence the expression of genes and analyze their loss-of-function phenotype, allowing analysis of gene function when mutant alleles are not available. Having been shown to work in a similar manner in all metazoans, RNAi has proven to be applicable to many organisms and has been used to generate a wide variety of loss-of-function phenotypes (Kuttenkeuler and Boutros 2004). The phenomenon of post-transcriptional gene silencing observed in plants may also be due to a related RNAi mechanism (Waterhouse, Graham, and Wang 1998). RNAi based silencing utilizes the endonuclease Dicer to cleave single stranded RNAs, abbreviated siRNAs, from double stranded RNA; the RISC complex then destroys specific target mRNAs based on sequence complementarity with the siRNA (Pattanayak et al. 2005).

RNAi has been used for a systematic analysis of gene function in C. elegans by generating loss of function phenotypes, creating a library of worms expressing dsDNA corresponding to different genes (Lewin 2004). Genome-wide RNAi screens against these libraries of predicted genes have allowed study of a variety of biological processes in C. elegans. A genome-wide library of double-stranded RNAs that target every gene in the Drosophila genome has also been published that is suitable for high throughput cell-based assays (Kuttenkeuler and Boutros 2004). One difficulty in using RNAi as a reverse genetic technique is that throughput is limited by the ability to deliver siRNAs to target loci (Henikoff, Till, and Comai 2004). It is also labor intensive, can give ambiguous results, and can be unsuitable for isolating mutants that have lethal or sterile phenotypes (Gilchrist and Haughn 2005).

Targeted gene disruption by homologous recombination

Homologous recombination is a reciprocal exchange of DNA sequences, as in between two chromosomes that carry the same genetic loci (Lewin 2004). Just as homologous recombination has been found to be mainly initiated with a double-strand break, gene targeting by homologous recombination is associated with the repair of double strand breaks. The double-strand break repair and synthesis-dependent strand-annealing models are the most generally accepted models to explain gene targeting (Iida and Terada 2004).

Homologous recombination has been widely used in embryonic stem (ES) cells in mice, and has allowed construction of precise mutations in nearly every gene. A reverse-genetic system using homologous recombination has recently been developed for Drosophila. It is promising, but is a lengthy procedure and requires generation of specific transgenic flies (Stemple 2004). Reproducible gene targeting by homologous recombination is now also feasible in rice. With the combination of site-specific recombination systems (such as Cre-lox), the future of gene targeting by homologous recombination as a routine procedure for engineering the genome of rice and presumably other plants is bright (Iida and Terada 2004).

Insertional mutagenesis/transposon mediated mutagenesis

Transposons are mobile genetic elements that can relocate from one genomic location to another (Hayes 2003). They are DNA sequences that can insert themselves at a new location in the genome without having any sequence relationship with the target locus (Lewin 2004). Transposon-based signature-tagged mutagenesis has been successful in identifying essential genes as well as genes involved in infectivity of a variety of pathogens. Strategies for insertional mutagenesis using transposons have been developed for a number of animal and plant models (Hayes 2003). Reverse-genetics is currently being done in Drosophila and C. elegans by utilizing libraries of individuals who carry transposable element insertions, many of which have been mapped, and some of which will disrupt the expression of nearby genes. In Drosophila P-elements, imprecise excision can be driven to generate a mutation in the nearest gene. Transposon-based methods are also being used in Arabidopsis, maize and other plants (Stemple 2004). One drawback of insertional mutagenesis is the low frequency of mutations, necessitating the screening of large numbers of individuals to find mutations in any given gene (Gilchrist and Haughn 2005). Also, insertions in essential genes will usually cause lethality, and less severe mutations must be generated in these genes in order to understand gene function (Till et al. 2003).

The segment of the Ti plasmid of Agrobacterium tumefaciens known as T-DNA that carries genes to transform the plant cell has also been utilized for insertional mutagenesis. T-DNA insertional mutagenesis has been used to obtain gene knockouts for greater than 70% of Arabidopsis genes (Alonso et al 2003), but no comparable resources exist for rice or maize even as high-coverage genomic sequence is becoming increasingly available (Henikoff, Till, and Comai 2004). Unlike other successful gene targeting systems (namely mouse, Physcomitrella, and Drosophila), the precise mechanism of T-DNA integration into the plant genome remains largely unknown (Iida and Terada 2004). Like RNA suppression techniques, insertional mutagenesis is limited by its host range and by its limited range of allele types (McCallum et al. 2000).

Gene replacement via transformation is a commonly used tool for many filamentous fungi (Fang, Hanau, and Vaillancourt 2002; Lalucque and Silar 2004; Takano et al. 2000) . The transformation can be mediated in many ways, including Agrobacterium (Zhang et al. 2003) and various other transformation vectors (Scott-Craig et al. 1998; Takano et al. 2000).

Chemical mutagenesis/TILLING

Two of the most widely used mutagens for chemical mutagenesis experiments are EMS (ethylmethanesulfonate) and ENU (ethylnitrosourea). EMS is a chemical mutagen that alkylates guanine bases. The alkylated guanine will then pair with thymine instead of the preferred cytosine base, ultimately resulting in a G/C to A/T transition. EMS is the most commonly used mutangen in plants. In Arabidopsis, five percent of EMS-induced mutations in targeted coding regions result in premature termination of the gene product, while fifty percent result in missense mutations that alter the amino-acid sequence of the encoded protein (Gilchrist and Haughn 2005). This high level of missense mutations relative to terminated gene products is very useful in analyzing gene function. ENU also induces point mutations, and is a more potent mutagen than EMS. It is also an alkylating agent, mutagenizing by transferring an ethyl group to oxygen or nitrogen radicals in the DNA molecule, which leads to mispairing and ultimately results in base pair substitutions, and sometimes base pair losses if not repaired (Guénet 2004).

Chemical mutagenesis is attractive for reverse genetics because it results in induced point mutations, which create a diverse range of alleles for genetic analysis. It induces a large number of recessive mutations per genome that are randomly distributed (Gilchrist and Haughn 2005) . Because chemical mutagenesis is already widely used in many organisms for forward genetic screens, it promises to be generally applicable for reverse genetics (Henikoff, Till, and Comai 2004). Until recently, chemical mutagenesis has not been widely used as a tool for reverse genetics because of the lack of high-throughput techniques for detecting point mutations (Gilchrist and Haughn 2005). The TILLING strategy provides a high throughput strategy to detect single base changes within genetic targets and can be applied to a wide variety of organisms.

TILLING Phytophthora

Whole genome sequences for the soybean pathogen Phytophthora sojae and the sudden oak death pathogen P. ramorum were made public near the end of 2004. Large-scale genomic sequencing projects are underway for the potato late blight pathogen P. infestans and the vegetable pathogen P. capsici. The compilation of these genomic sequences allows the application of many sophisticated research tools and promises a better understanding of this devastating group of pathogens. In an effort to better understand gene function within Phytophthora a project to develop a reverse genetic TILLING resource for Phytophthora has been initiated. Figure 2 provides an overview of the process as it is being applied to Phytophthora. Zoospores present an ideal life stage for mutagenesis as they are uni-nucleate and single mutant individuals can readily be isolated following mutagenesis. The mutation rate for EMS or ENU is not known for Phytophthora and lethality is being used as an indicator of dose response. Genomic DNA is extracted from the mutant individuals and pooled 2 to 4-fold. The genomic DNA library can then be repeatedly screened. Specific genes are amplified from the pools of genomic DNA, and the PCR products are heated up and allowed to cool slowly to form heteroduplexes between wild type and mutant strands of DNA. The heteroduplexes are treated with the single strand specific endonuclease CEL1 which cuts 3' of single base mismatches producing novel fragments of DNA. CEL1 treated PCR products are then resolved on a polyacrylamide gel and screened for the presence of novel fragments. Pools containing novel fragments are then analyzed to determine exactly which mutant is carrying an induced point mutation, and the PCR product from this mutant is sequenced to determine if the induced change is predicted to be silent, missense, or a knockout mutation.

Figure 2

Since Phytophthora is diploid through most of its life cycle (including the zoospore stage) all of the induced point mutations exist in the heterozygous state. Once a non-silent point mutation has been identified the mutant isolate is taken through the sexual stage and the sexual progeny are screened to identify individuals homozygous for the mutation under investigation. Homozygous mutants are then tested to determine if there is an altered phenotype.

Limitations/roadblocks to completing reverse genetics

Completing reverse genetics is not without its pitfalls, and not all techniques can be applied to all organisms. To be successful, there are several aspects that must be checked. For organisms that do not have efficient transformation systems available techniques such as TILLING that can be applied without transformation may be the only practical choice. In these cases, the rate of mutagenesis is an important factor that can be difficult to determine. The load of mutations must be balanced with the recovery of mutants (Till et al. 2003) – in other words, the genome can't be so riddled with mutations that it is impossible to see a mutant phenotype. Also to be considered is the fertility of the mutagenized organism, especially in the first generation, but also in subsequent generations (Perry et al. 2003) , both before and after mutagenesis. This is especially true for diploid organisms, because if the sexual machinery is not intact and working properly, then it is impossible to obtain a homozygous mutant. The mutagenized organisms must also be kept alive long enough to screen a mutant population for a specific target. For some organisms, like Arabidopsis or Phytophthora, this is not a problem as the seeds or cultures are relatively easy to store. It presents a challenge for other organisms, such as zebrafish or rats, because they must be stored and kept alive through the mutant screening stage.

Supplementary information: examples of TILLING in Action

*Arabidopsis

A public TILLING resource was set up in 2001 for the Arabidopsis community. This effort was called the Arabidopsis TILLING Project (ATP) and is now called the Seattle Tilling Project (STP) and is a joint effort between the Comai Laboratory at the University of Washington and the Henikoff Laboratory at the Fred Hutchinson Cancer Research Center in Seattle, Washington. Users are charged a fee that covers partial costs of the services provided to request mutations in genes of interest (Gilchrist and Haughn 2005; Till et al. 2003). Training sessions, workshops, and on-going support to researchers interested in developing TILLING in other organisms has been made available through the ATP and has served to make the TILLING technique more wide-spread. This group has also developed and made publicly available web-based software programs for PCR primer design and visualization of polymorphisms (Gilchrist and Haughn 2005).

*Zebrafish

Target selected mutagenesis using TILLING is also being done in zebrafish. The Hubrecht Laboratory at The Netherlands Institute for Developmental Biology has generated a library of 4608 ENU-mutagenized F1 animals and has kept a living stock. The DNA from these animals has been screened for mutations in 16 genes using TILLING followed by re-sequencing. This resulted in 255 mutations being identified, 14 of which resulted in a premature stop codon, 7 in a splice donor/acceptor site mutation, and 119 in an amino acid change. They were able to knock out 13 different genes in only a few months time through reverse genetics (Wienholds et al. 2003) .Thus far, mutant phenotypes have been characterized for two of the zebrafish genes modified via TILLING - the gene for Dicer1 (Stemple 2004; Wienholds et al. 2003) and the gene for adenomatous polyposis, a tumor suppressor, which through was found to have a previously unknown function for signaling in cardiac-valve formation (Hurlstone et al. 2003; Stemple 2004).

Acknowledgements

This work was supported by a National Science Foundation, Faculty Early Career (CAREER) Development award (#0347624) to Kurt Lamour.

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