18.9 Manipulating genomes: targeted gene disruption, transformation and vectors
Analysing natural genomes rapidly leads to ideas about modifying genomes. Of course, since the dawn of agriculture, the practical man has been involved in modifying the genomes of his/her cultivated plants and animals by a process of artificial selection. Indeed, although we were unaware of it at the time, by selecting particular brews or ferments that produced the most satisfactory end products in brewing, baking and other food fermentations (cheese, salami, soy, miso) we have also been unconsciously applying selection pressure to the fungi and bacteria involved in those processes for a very long time.
During the course of the twentieth century increasing knowledge of genetics enabled applied genetics to be much more formalised and very considerable advances were made in breeding improved varieties. Classical genetics of this sort puts the emphasis on the phenotype. What matters is the phenotypic characteristic of the new strain; those features that make it more useful or advantageous. In time, deeper analysis might establish the genetic basis of a phenotype or trait and enable genetic manipulation (mutation, controlled breeding) to further enhance the trait and/or combine it with others.
Automated DNA sequencing generates large volumes of genomic sequence data quite quickly. The consequence is that many genetic sequences are discovered well in advance of information about their function in the life of the organism. Molecular analysis enables us to start from the other end of this line of activity; we can seek to find the possible phenotypes that may obtain from a specific genetic sequence obtained by DNA sequencing. So, if classical, 20th century, genetics is considered to be forward genetics, proceeding from phenotype to genetic sequence; then the molecular genetics of the 21st century has come to be called reverse genetics. Reverse genetics attempts to link a specific genetic sequence with precise effects on the organism.
In practice this proceeds from functional analysis by experimental design and can eventually lead to functional design. The essential flow of activity is: gene sequence → change or disrupt the DNA (deletion, inactivation by insertion, point mutation) → mutant phenotype → function → alter function → change sequence → new (improved?) phenotype.
This has come to be called functional genomics; it is the study of gene function on the genomic scale. In filamentous fungi it is a field of research that has made great advances in very recent years and which continues to advance at rapid pace. Transformation and gene manipulation systems have been developed and applied to many economically important filamentous fungi and oomycetes; further advances in functional genomics will undoubtedly reveal a mass of new information in the near future (Weld et al., 2006).
Several different techniques are commonly used in reverse genetics and functional genomics:
- Random deletions, insertions and point mutations; generated by creating large populations of mutagen treated organisms (=large libraries of mutants) using chemical mutagens (point mutations), gamma radiation (deletions) or DNA insertions (insertional knockouts). These mutant libraries are screened for specific changes in the sequence of interest using PCR.
- Directed deletions and point mutations; site-directed mutagenesis is a more refined technique that can modify chosen parts of the sequence of interest, such as regulatory regions in the promoter of a gene or codon changes in the ORF to identify/modify specific amino acids to affect directly the protein function. The technique can also be used to create ‘gene knockouts’ by deleting a gene function (forming what is known as a null allele) (Fig. 20). Directed deletions have been created in every non-essential gene in the yeast genome (Winzeler et al., 1999) and methods are available for efficient gene targetting in filamentous fungi (Krappmann, 2007). As you might now suspect, there is an international collaborative effort to generate systematically specific deletion mutants of individual Saccharomyces cerevisiae genes called the European Functional Analysis Project (EUROFAN; see http://mips.gsf.de/proj/eurofan/).
|Fig. 20. Replacing the target gene wild type chromosomal sequence with mutant derivatives using homologous recombination.|
- Knockouts are gene deletions; an alternative approach is to substitute genes at specific times and in specific cells with experimental sequences and this is called ‘gene knockin’. The method involves insertion of a protein coding cDNA ‘signal’ or ‘reporter’ sequence at a particular site, and is particularly applicable to study the function of the regulatory sites (promoters, for example) controlling expression of the gene being replaced (Fig. 21). This is accomplished by observing how the easily-observed reporter phenotype responds to regulation.
|Fig. 21. Gene disruption/deletion by homologous integration of a targeted marker sequence. Top panel shows the preparation of short regions which are homologous to the target gene sequences (SFH, or short flanking homology region). Bottom panel: one-step gene disruption by homologous recombination uses a readily-identified marker flanked by the pre-prepared SFH.|
- Gene knockouts and knockins are permanent sequence alterations. A number of gene silencing techniques target the expression machinery and are generally temporary. This approach is often called gene knockdown since the effect is usually to grossly reduce expression of the gene. Gene silencing may use double stranded RNA, also known as RNA interference (RNAi) or Morpholino oligos.
- RNA interference relies on a specific cellular pathway (called the RNAi pathway) interacting with the introduced double-stranded RNAs (dsRNAs, typically over 200 nucleotides long), which are made to be complementary to some target messenger RNA (mRNA). An RNase-like enzyme called Dicer in this pathway generates small interfering RNAs (siRNAs) about 20-25 nucleotides long. The siRNAs assemble into complexes containing ribonuclease (known as RISCs, or RNA-induced silencing complexes). The siRNA strands guide the RISCs to their complementary target RNA molecules, which they cut and destroy; thereby systematically interfering with expression of the target gene, so that the effect of the absence of that gene activity can be catalogued.
- Morpholino antisense oligos block access to the target mRNA without the need for mRNA degradation. Morpholinos contain standard nucleic acid bases, but instead of the bases being linked to ribose rings connected by phosphate groups, those bases are bound to morpholine rings linked through phosphorodiamidate groups. The latter are uncharged and therefore not ionised in the usual physiological pH range; this and the other structural differences mean that Morpholinos are not sensitive to the same chemistry or enzymes as natural polynucleotides, but they still bind to complementary sequences of RNA by standard nucleic acid base-pairing. Morpholinos (usually 25 bases in length) base pair with regions of the natural RNA and this binding blocks splicing and translation, and therefore expression of the target gene.
A significant advantage of site-directed (or insertional) mutagenesis over random chemical or radiation mutagenesis is that the genes mutated by insertion are tagged (= physically identified) by the transforming DNA (T-DNA), which is used to disrupt the genes. This means that the molecules are readily identifiable in vitro, and, if the inserted sequence carries an expressed phenotype distinct from the recipient (such as an antibiotic resistance, ability to use an exotic substrate or render a toxin harmless) then the successfully-transformed cells can be identified, ideally selected, in vivo.
All of these approaches require that a recipient cell is transformed by uptake of the constructed DNA so that the latter can at least form a partial heterozygote that ideally undergoes homologous recombination and integrates the constructed DNA into the resident chromosome . The first barrier to successful transformation is the fungal cell wall, and most transformation techniques depend on three main ways of breaching the wall (which can be combined to improve efficiency) (Weld et al., 2006):
- enzymic removal of the wall to create protoplasts or sphaeroplasts,
- use of electroporation by applying electric shocks,
- or by ‘shooting’ micrometre-sized particles (usually of dense, relatively inert, metals like tungsten or gold) coated with DNA or RNA into the cells; a process called biolistic transformation.
Protoplasts are fungal or plant cells that have had their entire cell wall removed. The word sphaeroplast is used when some wall material remains, even if this is a functionless remnant. Protoplasting has been used for a very long time (e.g. Peberdy & Ferenczy, 1985) and is the most widely used aid to transformation although it is the most laborious and time-consuming method, and the homologous recombination frequency can be relatively low. However, sphaeroplasts are generally readily prepared by enzymatic treatment of mycelial suspensions with a commercially available lytic enzyme preparation (with commercial brand names such as: Driselase, Novozyme, Glucanex).
The most prominent enzyme activities of Driselase are endo-β(1,4)-glucanase, β-glucosidase and β(1,3)-glucanase. Chitinase activity does not seem to be required for most Ascomycota (Wiebe et al., 1997; Koukaki et al., 2003) though this enzyme produced good yields of protoplasts from germinated spores of the basidiomycete Coprinopsis (Moore, 1975). Young, actively growing hyphae, such as germinating spores, generally give the best results; the enzyme treatment removes the wall at the hyphal apex and sphaeroplasts are released from the apical tips of hyphae after about 90 minutes exposure to the enzyme and, with longer treatments, from other, older, regions of the hyphae. Yields of sphaeroplasts depend on the mycelial concentration treated and seem to be best when this is in the range 1-10 mg dry weight of mycelium ml-1. Protoplasts and sphaeroplasts are osmotically fragile and a variety of solutes (at concentrations in the range 0.6 to 1.5 M) have been used, including glucose, sucrose, sorbitol, mannitol, ammonium sulfate, or magnesium sulfate to provide osmotic support during purification and treatment. These osmotic stabilisers also have to be used during the transformation process and after it, when the sphaeroplasts are ‘germinating’ and regenerating their walls.
Electroporation is a technique for the introduction of nucleic acids and other macromolecules into cells. A brief electric pulse (lasting in the region of 1 to 20 ms) at a potential gradient of about 0.5 to 10 kV cm-1 is applied to temporarily permeabilise cell membranes so as to enable entry of large, charged molecules across the hydrophilic membrane. It is an efficient system for transformation of protoplasts of filamentous fungi (Fariña, Molina & Figueroa, 2004), germinating conidia (Gangavarama et al., 2009), or whole yeast cells (Chen et al., 2008). The frequency of transformation by electroporation is enhanced markedly by pre-treatment of whole cells with dimethyl sulfoxide (dissolves charged and uncharged molecules and easily penetrates cell membranes), dithiothreitol (prevents disulfide bonds forming between cysteine residues of proteins) or lithium acetate (which permeabilises the cell wall).
Biolistic transformation is an unusual process in which microparticles coated with DNA or RNA are introduced into cells by being accelerated to velocities of approximately 500 m s-1 by the forces generated by explosion of gunpowder or by explosive expansion of cold helium gas at pressures in the region of 1,300 pounds per square inch (1300 psi; = 8963 kPa). Developed first for transforming plant cells, it has been used successfully with bacteria, mammalian cells and fungi (Armaleo et al., 1990; Sanford, Smith & Russell, 1993; Aly et al., 2001), as well as mitochondria and chloroplasts. The constructed DNA is precipitated onto the 0.5-0.65 µm diameter particles of (usually) tungsten with CaC12, the effect of which is enhanced by spermidine. As with sphaeroplasts, the biolistically treated cells need osmotic stabilisation with 1.5 M mannitol + sorbitol in the medium, presumably because of the physical damage done to the cell wall by the particle bombardment. The great majority of transformants result from penetration of single particles, delivering on average 10-30 biologically active plasmids into the cell. The end-products of biolistic and sphaeroplast transformation are the same: recipient cells containing either the introduced DNA integrated into the chromosome at its homologous site, or free in the cytoplasm replicating like an independent plasmid. However, biolistic transformation works best on stationary phase cells, while spheroplasting is most effective with log-phase cells.
Agrobacterium tumefaciens-mediated transformation (AMT). As an alternative to the above methods, a fairly recent trend has been towards approaches based on Agrobacterium tumefaciens-mediated transformation (usually abbreviated to AMT). A. tumefaciens is a gram-negative bacterium and a common plant pathogen that causes crown gall tumours on plants. This tumorous growth of the plant tissue is induced when the bacterium transfers to the host plant some bacterial DNA (called Ti-DNA) which is located on a 200 kbp plasmid (the tumour-inducing or Ti plasmid). The Ti-DNA integrates into the plant genome, then Ti-DNA genes that encode enzymes for the production of plant growth regulators are expressed, and their expression results in uncontrolled growth of the plant cells. However, for use as a cloning vector, the Ti-region of the plasmid can de deleted and replaced by other DNA sequences because plasmid virulence, transfer and integration are controlled by genes elsewhere on the plasmid.
What is significant for the present discussion is that A. tumefaciens is able to transfer its Ti-DNA to a very wide range of fungi and fungal tissues and produces a significantly higher frequency of more stable transformants than biolistic transformation (Michielse et al., 2005). AMT is a relatively simple system to work with, does not require the production of protoplasts or sphaeroplasts. Indeed a major attraction of AMT is the variety of starting materials can be used: protoplasts, spores, mycelium, and fruit body tissues have all produced successful transformation. Even fungi that have not been transformed by other systems have been successfully transformed by co-cultivation with Agrobacterium. The approach is applicable to 'Zygomycota', Ascomycota and Basidiomycota and shows great potential for fungal biotechnology and medicine (Michielse et al., 2005; Sugui, Chang & Kwon-Chung, 2005).
Despite the confident descriptions given above and the use of phrases like ‘relatively simple system’, applying a transformation system to an organism for the first time is often not as straightforward as might be suggested. There are many variables that have to be optimised and even after reliable transformation systems have been developed, there may still be difficulties to overcome before it is possible to analyse gene function. A major potential problem for genetic analysis of any filamentous fungus is the multinucleate nature of the hyphae. Multiple nuclei can confuse results because gene replacement and insertional mutagenesis rely on the isolation of homokaryotic transformants derived from a single transformation event to study loss of function mutants (Weld et al., 2006). The consequence is that methods have to be carefully refreshed and optimised every time they are applied to a new organism.
DNA cloning requires vectors. Cloning involves inserting DNA molecules of interest into specialised carriers called vectors that enable replication within a host cell, producing many copies of the inserted piece of DNA carried by the vector. Cloning vectors are ‘engineered’ to contain one or several recognition sites for restriction enzymes. Digesting both the vector and the DNA to be cloned with the same restriction enzyme produces complementary ‘sticky ends’ in both molecules, allowing the foreign (or heterologous) DNA fragment to be inserted into the vector. A vector carrying an inserted fragment of DNA is known as a recombinant plasmid. The replicated molecules are called clones because all the copies made in the host cell are identical. After harvesting from the host cell, the cloned DNA can be purified for further analysis.
There are several types of cloning vector, which differ in origin, nature of host cell, and in their capacity for the size of inserted DNA they can carry. The simplest vectors are bacterial plasmids, which are circular, double‑stranded, DNA molecules that replicate in the host independently of the main bacterial chromosome. Commonly used plasmids can carry up to 15 kb of foreign DNA.
DNA fragments up to 25 kb in length can be accommodated in vectors derived from the bacteriophage (‘phage’) lambda (λ), which is a double stranded DNA virus that infects the bacterium Escherichia coli. The λ phage replicates in its host as a circular molecule, but has a linear DNA molecule 50 kb long in mature (infective) virus particles. The virus chromosome circularises after infection because it has complementary single stranded overlaps at each end known as cos (for cohesive end) sites.
A completely artificial, larger capacity vector has been engineered by inserting cos sites into a plasmid. These are called cosmids. They can carry up to 45 kb of inserted DNA and have the additional advantages that they use a virus coat to infect host bacteria (a very efficient way of entering the host), but replicate like a plasmid and can be constructed to use plasmid‑derived markers for recombinant selection. We have briefly described cosmids, bacterial artificial chromosomes (BACs), and yeast artificial chromosomes (YACs) above, in relation to their use in DNA sequencing (see the section entitled Sequencing fungal genomes CLICK HERE to view now.
The largest capacity vectors currently available are yeast artificial chromosomes (YACs) which can carry DNA inserts of up to 1 million base pairs (1 megabase = 1 Mb) in length. YAC vectors are plasmids that contain yeast centromere DNA, two yeast telomeres separated by a restriction site, and yeast replication origins (autonomous replication sequences, or ARS) as well as two selectable markers. Restriction enzyme digestion produces two fragments, one a telomere + selectable marker + cloning site, the other a telomere + selectable marker + replication origin + centromere + cloning site, which are mixed with the DNA to be cloned. Among the constructs which result will be some which behave like yeast chromosomes during mitosis. Any that are constructed with two centromeres, without a centromere, or lacking a telomere will fail to segregate. Consequently, the presence of both selectable markers coupled with proper mitotic segregation is sufficient to identify the desired constructs.
Interestingly, at meiosis a diploid cell containing two homologous YACs will show 1:1 segregations into progeny spores just like a regular pair of homologous chromosomes. Currently, YACs offer the highest capacity of any cloning vector, but recombination can affect insert stability in some YACs. Choice of cloning vector, and consequently insert size, depends upon the purpose in mind when the cloning is initiated, as well as considerations about stability, reliability and practicality. The capacity of cloning vectors is important because the genetically functional regions of filamentous fungi are larger than those in yeast because the genes contain more introns.
All yeast vectors are shuttle vectors, meaning that they can be propagated (that is, grown) in cell cultures of both yeast and the bacterium Escherichia coli. These vectors contain a bacterial plasmid backbone that contains all the functions required for maintenance and selection in E. coli. They also contain yeast chromosomal elements that determine their characteristics and behaviour within yeast cells. The main types of yeast vectors are:
- Yeast Integrative plasmids (YIp); which are maintained as a single copy providing they have integrated successfully into the genome.
- Yeast Replicative plasmids (YRp); which contain a chromosomal origin of replication (ARS), and because of this origin of replication are maintained autonomously at high copy number (which means 20 to 200 copies of the plasmid per yeast cell).
- Yeast Centromeric plasmids (YCp); which contain both ARS and centromere sequences and are consequently maintained in the cell as a single copy autonomously replicating supplementary chromosome.
- Yeast Episomal plasmids (YEp); which is also an autonomously replicating plasmid (contains ARS) but contains the origin of replication from yeast’s own 2µ plasmid so it is maintained at a copy number of about 20 to 50 copies per cell. This type of vector is used for gene over-expression purposes (as are YRps). Gene over-expression creates a gain of function mutation and requires the use of multicopy vectors and strong promoters.
Vectors must carry selectable markers so that successful insertion can be selected, and shuttle vectors require selectable markers that are effective in both hosts. The preferred marker for yeast shuttle vectors is called KanMX. This is based on the kanamycin resistance gene from Escherichia coli which is an aminoglycoside antibiotic that works by affecting the 30S ribosomal subunit and causing mistranslation. Aminoglycosides are also toxic to eukaryotes because they block translation by inhibiting polypeptide synthesis. The E. coli kanr gene (which encodes an enzyme that destroys the antibiotic) can also be expressed in yeast to make the yeast cell resistant to the aminoglycoside antibiotic known as G418 (also called Geneticin). KanMX is an expression cassette in which the kanr gene is fused to promoter and terminator sequences from the Ashbya (=Eremothecium) gossypii (Ascomycota), which is a filamentous fungus belonging to the same family as Saccharomyces cerevisiae (Güldener et al., 1996).
Updated December 17, 2016