Species-specific primers for diagnosis of selected plant parasitic nematodes.
\r\n\tThis book aims to provide in-depth study and analysis of various fabrication and processing techniques of nanoplates, and their characterization and applications in recent technology. The shape dependent properties of nanoplates have opened novel research areas in modern Nanoscience and Nanotechnology that will be discussed in this book.
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He has completed post-doctorates from South Korea and China. His research interests include the fabrication of nanomaterials and their structural, magnetic, optical, and electrical characterizations. He has authored more than 90 articles and served as the editor of eight books. Presently he is serving as the Editor-In-Chief of 'Journal of Materials, Processing and Design” (USA) and Executive Editor of 'International Journal of Nano Studies & Technology” (Canada), and also as the Editorial Board Member of few more journals of materials science.",institutionString:"Pakistan Institute of Nuclear Science and Technology",position:null,outsideEditionCount:0,totalCites:0,totalAuthoredChapters:"0",totalChapterViews:"0",totalEditedBooks:"8",institution:{name:"Pakistan Institute of Nuclear Science and Technology",institutionURL:null,country:{name:"Pakistan"}}}],coeditorOne:null,coeditorTwo:null,coeditorThree:null,coeditorFour:null,coeditorFive:null,topics:[{id:"17",title:"Nanotechnology and Nanomaterials",slug:"nanotechnology-and-nanomaterials"}],chapters:null,productType:{id:"1",title:"Edited Volume",chapterContentType:"chapter",authoredCaption:"Edited by"},personalPublishingAssistant:{id:"205697",firstName:"Kristina",lastName:"Kardum Cvitan",middleName:null,title:"Ms.",imageUrl:"https://mts.intechopen.com/storage/users/205697/images/5186_n.jpg",email:"kristina.k@intechopen.com",biography:"As an Author Service Manager my responsibilities include monitoring and facilitating all publishing activities for authors and editors. From chapter submission and review, to approval and revision, copyediting and design, until final publication, I work closely with authors and editors to ensure a simple and easy publishing process. I maintain constant and effective communication with authors, editors and reviewers, which allows for a level of personal support that enables contributors to fully commit and concentrate on the chapters they are writing, editing, or reviewing. I assist authors in the preparation of their full chapter submissions and track important deadlines and ensure they are met. I help to coordinate internal processes such as linguistic review, and monitor the technical aspects of the process. As an ASM I am also involved in the acquisition of editors. 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Sarria",coverURL:"https://cdn.intechopen.com/books/images_new/5884.jpg",editedByType:"Edited by",editors:[{id:"146466",title:"Prof.",name:"Andreia",surname:"Ferreira de Castro Gomes",slug:"andreia-ferreira-de-castro-gomes",fullName:"Andreia Ferreira de Castro Gomes"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}},{type:"book",id:"7325",title:"Nanostructures in Energy Generation, Transmission and Storage",subtitle:null,isOpenForSubmission:!1,hash:"8e49924dd2c3e28c82fdc115ce04f925",slug:"nanostructures-in-energy-generation-transmission-and-storage",bookSignature:"Yanina Fedorenko",coverURL:"https://cdn.intechopen.com/books/images_new/7325.jpg",editedByType:"Edited by",editors:[{id:"199149",title:"Dr.",name:"Yanina",surname:"Fedorenko",slug:"yanina-fedorenko",fullName:"Yanina Fedorenko"}],productType:{id:"1",chapterContentType:"chapter",authoredCaption:"Edited by"}}]},chapter:{item:{type:"chapter",id:"55761",title:"Methods and Tools Currently Used for the Identification of Plant Parasitic Nematodes",doi:"10.5772/intechopen.69403",slug:"methods-and-tools-currently-used-for-the-identification-of-plant-parasitic-nematodes",body:'Nematodes are diverse, microscopic multicellular animals comprising free living to plant parasitic species. They parasitize a wide range of plant species, including monocots and dicots and are one of the most limiting factors for major crops, causing an estimated annual crop loss of $78 billion worldwide and an average crop yield loss of 10–15% [1–3].
Reliable, fast, and proper nematode diagnosis and specimen identification are mandatory for choosing adequate management control strategies and for avoiding spreading of exotic nematodes in quarantine materials [4–7]. Nonetheless, nematodes are one of the most difficult organisms to be identified, either due to their small, microscopic sizes or due to the difficulties in observing key diagnostic characters/features under conventional light microscope [5, 7–10]. In addition, the differences of some of these morphological and morphometric characters are subtle, subjective, and have overlapping characters or show intraspecific variation which compromise proper identification or may lead to erroneous identity of the species [5, 10, 11]. Furthermore, nematode identification using classical morphology requires well trained and experienced nematode taxonomists which are in decline these days due to lack of interest of young scientists in classical taxonomy [10].
Currently, new methods and tools using biochemical and molecular approaches have been successfully used as diagnostic for plant parasitic nematodes [4, 6, 7, 10, 12–16]. Despite the feasibility and accuracy in using biochemical and molecular-based detection tools and methods these days, diagnoses based on morphology are still sufficient or are required in some specific cases. Thus, when possible, the use of integrative diagnostic/taxonomical approaches using morphological, biochemical, and molecular data may be more time consuming but overall may lead to a more accurate diagnosis of nematode species, especially for those cryptic or newly described species.
Diagnostic laboratory that provides testing for plant parasitic nematodes has been increasing in recent years due to increased occurrence, damage, and dissemination of plant parasitic nematodes, lack of proper control management strategies, and high population density of key nematode pests in agricultural systems [17]. The current withdrawal of most chemical nematicides from the market is direct consequence of their toxicity and side effects to environment and human health. Alternative means in controlling plant parasitic nematodes for a sustainable cropping system include the use of resistant cultivars, the use of non and poor hosts, crop rotation, crop succession, and biological control [5]. However, accurate and fast identification of nematodes to species and subspecies levels is mandatory not only to be successful in choosing a proper management strategy but also for studying their genetic and biological variability or to avoid global spread of exotic and quarantine pathogens [4, 6, 7, 18, 19].
The goal of this chapter is to report a literature review of methods and tools to identify the most common genera of plant parasitic nematodes and its use to other nematode species as well.
Nematode diagnosis and taxonomy have traditionally relied on morphological and anatomical characters using light microscopy. Lately, despite the increased interest in molecular diagnosis, due to its feasibility which allow quick and easy identification of specimens and allow researchers and extensions folks to use these techniques for routine use [10], classical taxonomy using reliable and nonoverlapping morphological characters is still an important tool for the identification of nematodes mainly for the following reasons: (i) it allows a clear link between function and morphological aspects of the specimen analyzed; (ii) it is still a method that provides fast results; (iii) it is suitable for quantitative evaluations; (iv) it is cheaper; and (v) it is used for population surveys of plant parasitic nematodes with the objective to recommend management control strategies [10].
Other disadvantage in using classical taxonomy, besides the drawback of obscure morphological characters afore mentioned, the preparation of nematode specimens for classical identification, may result in modifications of the nematode, which may be difficult in its proper identification. For instance, much of the nematode body composition is proteins and fats, which undergo immediate coagulation or other alteration during the processing of the specimens, due to the use of substances such as sucrose, formaldehyde, glycerin, and the heating to which they are normally subjected during fixation [20]. In this way, specimens after being extracted, killed, fixed, and mounted on microscopic preparations, frequently present artifacts that make it difficult to locate external or internal structures of diagnostic value, or even produce characteristics that are not natural [20].
According to Inserra et al. [20], other disadvantages of light microscopy, in relation to other methods (electron microscopy, isozyme electrophoresis, and molecular methods), are as follows: the need for specimens in excellent preservation conditions; some characteristics of diagnostic value show high intraspecific variation, reason for the need for more than safe examination, need for abundant and updated scientific literature, the need for a taxonomist to be deepened in taxonomical studies, who probably would be a specialist in only a few groups of nematodes; several morphological and morphometric characteristics of diagnostic value are modified by environmental factors, such as geographic location, host plant species, host plant mineral nutrition, and light. However, the main disadvantage is that microscopic examination is not sufficient for identifying morphological characters that are extremely difficult to observe [20]. Thus, using integrative diagnostic approaches with more than one diagnostic method is less prone to errors.
Root-knot nematodes (RKNs), Meloidogyne spp., are the most aggressive, damaging, and economically important group of plant parasitic nematodes infecting important crops worldwide. Currently, about 97 Meloidogyne spp. have been described [5], of which M. arenaria, M. incognita, M. javanica, and M. hapla represent up to 95% of RKN in cultivated soils.
This group of nematodes is highly diverse, showing a continuum of diversity in terms of cytogenetics (variable chromosomes numbers with aneuploidy and polyploidy states), mode of reproduction (ranging from amphimixis to obligatory-mitotic parthenogenesis), specialization in parasitism, species complexes, cryptic species, interspecific hybridization, and broad host ranges [4, 6, 7, 18, 21]. Overall, this high level of diversity contributes to an extremely complex relationship with their hosts that lead to highly successful parasitism. For example, the three major Meloidogyne spp. (i.e., M. incognita, M. javanica, and M. arenaria) are highly polyphagous, infecting more than 3000 plant species [5].
Diagnosis of Meloidogyne spp. has traditionally relied on the characterization of female perineal patterns and morphometrics. However, since these morphological characters overlap in some RKN species (e.g., in M. paranaensis, M. konaensis, and M. enterolobii), misidentification of species using morphology as the only criteria is often frequent [22, 23].
The morphology of female perineal patterns has been a character most frequently used in several laboratories for the identification of Meloidogyne species, a character located in the posterior body region of adult females. This area comprises the vulva-anus area (perineum), tail terminus, phasmids, lateral lines, and surrounding cuticular striae. Preparation of perineal patterns for the observation and identification of Meloidogyne spp. has been covered by different authors. A more detailed account on root-knot nematode perineal pattern development was given by Karssen [23]. Figure 1 summarizes the perineal patterning for 12 major Meloidogyne species that are considered important to major crops [5].
Comparison of perineal patterns for 12 major Meloidogyne spp. A, B: M. arenaria; C, D: M. hapla; E, F: M. incognita; G, H: M. javanica; I: M. acronea; J: M. chitwoodi; K, L: M. enterolobii; M: M. ethiopica; N, O: M. exigua; P: M. fallax; Q, R: M. graminicola; S, T: M. paranaensis. Drawings not to scale [5].
For many years, the identification of Meloidogyne spp. has been relied upon the characterization of adult female perineal pattern and the use of several morphometric and morphological features of juveniles. To these characters were added features of male (although they are rarely seen), such as the form of the labial region, including annulation, and the form of stylet and basal knobs. However, with increasing numbers of described species, the value of many of these characters, themselves showing often large intraspecific variation, was eroded almost to the point where robust identification tended to involve a fair measure of serendipity. As an example, what may be termed the incognita-type of perineal pattern is now known to occur in a substantial number of species, some of which were commonly misidentified as M. incognita.
As an alternative to morphological identification of Meloidogyne spp., esterase patterning has been used for diagnosing Meloidogyne spp. from a wide range of samples and has been proved to be species-specific for a number of species [13, 24, 25]. Meloidogyne spp., isozyme electrophoresis patterning has discriminated all of these otherwise cryptic species, however, this technique is restricted to females [24]. Examples of esterase patterning for major Meloidogyne spp. are shown in Figure 2.
Esterase phenotypes (Est) of major Meloidogyne spp. associated with coffee. Rm = ratio of migration in relation to the fast band of M. javanica. Dotted lines indicate weak bands [24].
One of the earliest examples of the use of isozyme phenotypes to distinguish Meloidogyne spp. was given by Esbenshade and Triantaphyllou [25], who reported esterase patterns for 16 Meloidogyne species, with the most common phenotypes being A2 and A3 (M. arenaria), H1 (M. hapla), I1 (M. incognita), and J3 (M. javanica). In landmark surveys for Meloidogyne spp. using isozyme [12, 25] study, approximately 300 populations originate from 65 countries and several continents. In later surveys, Carneiro et al. [22] found 18 esterase phenotypes among 111 populations of Meloidogyne spp. from Brazil and other South American countries. Isozymes continue to be widely used for diagnosis of Meloidogyne spp. despite some limitations. Nonetheless, isozyme phenotyping has been used for a large number of species [6]. Schematic diagrams of isozyme patterns based on surveys, including those conducted in the international Meloidogyne project have been published [8, 12, 22, 25] and provide important references.
Several isozyme systems have been used, nonetheless, carboxylesterase/esterase EST proved to be the most useful in discriminating Meloidogyne species. Others, such as malate dehydrogenase (MDH), are also often included to confirm species identification [25]. Enzyme phenotypes patterning are designated, indicating the Meloidogyne species that each specifies and the number of bands detected. Phenotypes with the same number of bands are differentiated by small letters [12, 25]. Enzyme patterns are usually compared with a known standard, with M. javanica being frequently used to determine migration distances among bands. Isozymes are used primarily with female egg-laying stage, using single individuals. Miniaturization and automation of the electrophoresis systems and the use of precast polyacrylamide gels (i.e., PhastSytem, Pharmacia Ltd, Uppsala, Sweden) have made isozyme phenotyping a widely used technique in most labs [22, 23, 25]. Classical electrophoresis methods using vertical and horizontal systems were also described in details in Refs. [13, 25], respectively.
Aside from the initial equipment cost, the consumables required are relatively inexpensive and isozymes have been often used for field surveys, diagnosis, as well as with routine screening of glasshouse cultures to assure species stability and pure cultures. The relative stability of isozyme phenotypes within Meloidogyne species makes them an attractive system, although there are some drawbacks. For instance, the occurrence of intraspecific variants and the difficulty in resolving the same esterase phenotype between species (e.g., M. exigua vs. M. naasi) have required the use of an additional enzyme system (e.g., MDH) to confirm species identity. In addition, weak bands on the polyacrylamide gel may need to use a larger number of females per well (e.g., M. exigua) [22]. For some species, there are more than one esterase phenotype for a same species—e.g., M. javanica (J3, J2, and J2a), M. incognita (I1, I2, and S2), M. arenaria (A2 and A1), M. exigua (E1, E2, and E3), and M. paranaensis (P1 and P2) [8, 9, 26].
In surveys with the objective to study Meloidogyne biodiversity and nature conservancy, isozymes are a convenient first stage in species identification and have enabled the study of species diversity and frequency of a particular species, as well as their abundance. Females recovered after allowing multiplication of field samples on a generally susceptible host such as tomato (Solanum lycopersicum) can be tested for their isozyme phenotypes and the associated egg mass reserved for further characterization, if necessary [22]. Thus, novel isozyme phenotypes have been frequently found in these surveys in conserved areas, overall adding to the understanding of species ecology and biogeography of Meloidogyne spp. The Esbenshade and Triantaphyllou [25] listed Est F1 as an undescribed phenotype from Brazil; Later, M. paranaensis was described showing this phenotype [27]. In addition, Carneiro et al. [22] listed the patterns Est K3, Est Y3, and Est L3 as atypical esterase phenotypes; later, M. ethiopica and M. inornata were identified showing these new esterase patterns [8–9]. Recently, M. luci (Est L3) was described as a new species [28]. The phenotype Est Sa4 (Rm 73.5, 78.0, 53.0, 59.0), a new esterase phenotype from coffee in Central America, was later described as M. izalcoensis [29].
Isozyme electrophoretic profiles, often using esterase (EST) and malate dehydrogenase (MDH), have been established for a number of species [6] and can provide a useful routine diagnostic test particularly for morphologically variable species, such as M. arenaria. This species showed different profiles and high intraspecific variability, it may be an indication of the existence of species swam. Recently, the Est phenotype A3 of M. arenaria was identified as M. morocciensis [9].
Although isozyme electrophoresis is currently one of the best methods for Meloidogyne spp. diagnosis, it seems likely that DNA-based methods and tools will soon usurp this method for many applications where finer resolution, particularly of intraspecific variation, is paramount [6]. Nonetheless, the use of an integrative diagnosis, combining more than one approach, such as morphology, morphometrics, biochemical, and molecular data is less prone to error and could be used when possible.
Since the development of polymerase chain reaction (PCR) and the vast amount of genetic data generated with DNA sequencing, molecular-based detection tools have been widely developed and successfully used for the diagnosis of plant parasitic nematodes. Molecular-based detection tools have the following advantages as compared with other methods, (i) can be used in a high throughput manner, (ii) DNA information can be acquired easily with the vast amount of databases and sequencing information, (iii) are cheap, fast, and accurate, (iv) DNA markers are independent of phenotypic variation and developmental stage of the nematode [14].
DNA-based markers have been proved reliable and have allowed diagnosis and description of new species for several groups of nematodes, including key genera such as Meloidogyne, Pratylenchus, Globodera, and Heterodera [4, 6, 7, 10, 18, 30–32]. DNA-based detection tools make excellent methods of nematode diagnosis since they are simple, accurate, and fast [6, 7] and can be used with a wide range of sample types, including host tissue, eggs, egg masses, soil extracts, and fixed samples [16].
Nowadays, most labs worldwide are commonly using molecular methods to diagnose nematodes since cost associated with reagents and equipment are affordable and there has been a crescent interest in molecular taxonomy by young scientists [10, 16]. These methods have been used ordinary and are sensitive enough to detect individual nematodes from complex types of samples, including soil samples and species mixtures in the field [21, 33–35]. Some limitations of molecular-based detection tools include problems associated with optimization and validation of tools and methods, DNA extraction protocols, conditions of samples (i.e., quarantine specimens), amount of target DNA in a sample, cross contamination, false positive and negative results, which overall should be used carefully as to not compromise the ultimate result of diagnosis [16].
A vast amount of examples of nematode diagnosis has mostly been based on amplification of target DNA by PCR using species-specific primers. PCR-based detection methods have revolutionized the area of diagnostics of nematodes and have been used due to improved sensitivity, specificity, speed, relatively ease to perform, and cost effectiveness compared with other diagnostic procedures [4, 6, 7, 10, 18]. One of the approaches to design DNA-based markers that can aided diagnosis of nematodes has been based usually on conserved regions in the ribosomal DNA (rDNA) cistron, i.e., the external transcribed spacer (ETS), internal transcribed spacers 1 and 2 (ITS1 and ITS2), and the intergenic spacer regions 1 and 2 (IGS1 and IGS2) [7]. Schematic representation of these genetic regions is shown in Figure 3.
Schematic representation of nuclear rRNA genes in eukaryotic cells. SSU = 18S—small subunit; LSU = 28S—large subunit; ETS—external transcribed spacer region; ITS1 and ITS2—internal transcribed spacers; IGS1 and IGS2—intergenic spacer regions; arrows indicate possible starting point for primer amplification. Open box indicates the D2-D3 expansion segments for the 28S rRNA.
In this way, sequences that are divergent among nematode species and conserved within several isolates of a same species make ideal target for designing species-specific primers [7]. Ribosomal DNA regions have been very suitable for choosing a target marker since they are multicopy genes and provide sequences with enough variation that can be used for diagnosis and phylogenetic relationships among species [7].
Diagnostics of nematodes have also been based on other genomic target regions such as mitochondrial DNA (mtDNA). Mitochondrial DNA genomes are relatively small circular molecules ranging from 12 to 20 kilobases [7, 36]. Divergences in mtDNA sequences due to insertions, deletions, and accelerated ratio of mutations compared with nuclear DNA [7] have provided target markers suitable for discriminating nematode species [37–39].
Satellite DNAs (satDNAs) are highly repeated tandem arrays of short sequences ranging from 70 to 2000 bp. It has different signature sequences, copy numbers, length, and polymorphic regions that can be explored to find species-specific markers [6, 7]. Such PCR-based detection using satDNA markers in nematode diagnosis has been reported by several labs [7, 40, 41] and represents a target option for designing diagnostic primers.
One of the first methods used to differentiate nematode species was restriction fragment length polymorphism (RFLP), a method that uses restriction enzymes to digest whole genomic DNA or an amplified segment of it to generate DNA banding patterns according to divergences in sequences among isolates [7, 42]. This technique can also be coupled with DNA hybridization with radioactive or nonradioactive labeled probes [7]. Although being effective in differentiating nematode isolates, this method is less used nowadays due to technical complexity and the need for a large amount of target DNA, which usually requires preculturing of nematode populations [6, 7].
Alternatively, species-specific primers have been designed from sequences randomly scattered throughout nematode genomes, e.g., DNA band obtained from random amplified polymorphic DNA (RAPD) or amplified fragment length polymorphism (AFLP) gels, with posterior cloning and sequencing of bands differential across related species and their conversion into species-specific sequence characterized amplified region (SCAR) markers [21, 43–46]. SCAR-based markers and rDNA-based specific primers have been used to diagnose nematodes with either conventional or real-time PCR (q-PCR) [6, 7, 10].
Numerous primers and approaches used for diagnosis of nematodes using conventional and quantitative PCR were designed based on several target regions in the nematode genome (e.g., SCAR, rDNA, ITS, D2-D3 segment, IGS, among others). Table 1 summarizes some of these strategies used in some main studies.
Nematode species | Target region | Method | References |
---|---|---|---|
Meloidogyne spp. | |||
M. arabicida and M. izalcoensis | SCAR* | PCR | [45] |
M. arenaria | SCAR | PCR | [43] |
M. chitwoodi | IGS | PCR | [47] |
SCAR | PCR | [48] | |
M. exigua | SCAR | PCR | [21] |
M. enterolobii | mtDNA | PCR | [49] |
SCAR | PCR | [44] | |
M. ethiopica | SCAR | PCR | [46] |
M. fallax | IGS | PCR | [47] |
SCAR | PCR | [48] | |
M. graminis | ITS | PCR | [11] |
M. hapla | satDNA | PCR | [50] |
M. hapla | SCAR | PCR | [48] |
IGS | PCR | [51] | |
M. incognita | SCAR | PCR | [43] |
SCAR | PCR | [21] | |
M. javanica | SCAR | PCR | [43] |
SCAR | PCR | [52] | |
M. marylandi | 28S D2-D3 | PCR | [11] |
M. naasi | ITS | PCR | [52] |
M. naasi | 28S D2-D3 | PCR | [11] |
M. paranaensis | SCAR | PCR | [21] |
Other parasitic nematodes | |||
Bursaphelencus xylophilus | satDNA | PCR | [53] |
B. xylophilus | satDNA | qPCR | [54] |
B. xylophilus | heat shock protein | qPCR | [55] |
Ditylenchus destructor D. dipsaci | rDNA | PCR/qPCR | [56] |
H. glycines | rDNA | qPCR | [57] |
H. schachtii | ITS | PCR | [58] |
H. glycines | SCAR | qPCR | [59] |
Pratylenchus penetrans | rDNA | qPCR | [60] |
Species-specific primers for diagnosis of selected plant parasitic nematodes.
*SCAR—sequence characterized amplified region; IGS—intergenic spacer region; ITS—internal transcribed spacer; mtDNA—mitochondrial DNA; satDNA—satellite DNA; PCR—polymerase chain reaction; qPCR—quantitative real-time PCR.
In particular, successful SCAR markers have been developed for diagnosing some of the major tropical Meloidogyne spp. associated with important crops such as coffee, guava, and grapevine, including M. arenaria [43], M. incognita [21], M. paranaensis, M. exigua [21], M. enterolobii [44], M. arabicida, M. izalcoensis [45], and M. ethiopica [46] (see a complete list of references for species-specific primers in Table 1). These primers were validated in several population studies, using DNA from a single juvenile (J2), or in multiplex PCR reactions containing mixtures of species, and have become an excellent practical diagnostic kit for certain crops-associated Meloidogyne spp. [8, 21, 45, 46]. Interestingly, [61] established a diagnostic key for the identification of seven RKN species, i.e., M. incognita, M. javanica, M. arenaria, M. enterolobii, M. hapla, M. chitwoodi, and M. fallax using a combination of IGS PCR, SCAR markers, and RAPD profiling.
Quantitative PCR (q-PCR) is a technique that amplifies and quantifies nucleic acids simultaneously. Its advantage over conventional PCR is that it is fast, sensitive and does not need postamplification processing of samples normally seen in conventional PCR, which can lead to false results in some cases, reviewed in Refs. [7, 10]. As new genomic sequences become available for plant parasitic nematodes, there have been a vast number of examples of this approach to detect and quantify nematodes from a wide range of samples, including greenhouse, field experiments, ecological studies, experiments with plant-nematode interactions, and virus load in nematode vector and are being used as the advantage to be fast, accurate and to be applicable in high throughput processing systems of large number of samples, reviewed in Ref. [10]. Application of q-PCR in nematode diagnosis using rDNA target or other marker has been showed for major nematode species, including M. incognita, M. chitwoodi, M. fallax, M. javanica, Bursaphelenchus xylophilus, Globodera rostochiensis, and G. pallida. For a complete list of nematodes, see Ref. [10].
The concept of DNA barcode for nematode taxonomy has been employed in Ref. [62] and is related to a DNA sequence of a particular region in the genome as a mean to give unique signature (barcode) for the identification of nematode species [7]. Although theoretically sounds, this approach has not been widely accepted since there has not been an unique DNA locus that can define the limits of species boundary and be used as universal identification of nematodes, besides the missing link of DNA barcode approach with classical taxonomy which has shown clear methods for species-level resolution [7, 14].
Lately, there has been increased interest of labs to perform molecular diagnosis of nematodes directly from soil samples without the need to extract the target nematode species, a strategy commonly used for communities of bacteria and fungi [7]. There are available commercial kits for the direct extraction of nematode DNA from soil extracts and has been successfully employed in some labs. Alternatively, nematodes can be extracted from soil samples using conventional methods such as Baerman funnel, Whitehead tray or other method can be pooled for DNA extraction using commercial kits or other ordinary DNA extraction method. This strategy has been used by several authors and has been proved reliable and time saving [33, 34]. Nonetheless, there are drawbacks associated with this strategy, including underestimation of nematodes due to their uneven distribution in the soil, the limited amount of soil sample used for DNA extraction in commercial kits and the cost associated with sample processing [7].
Alternatively, a method to enrich nematode from soil extract using antibody-based capture was proposed by Chen et al. [63]; however, its use as routine diagnosis in labs needs to be analyzed.
The accurate identification of nematodes to species and subspecies levels is essential for their control and is a prerequisite to meaningful research. Many nematode species are easily identified based on distinct morphological characters and restricted host ranges. Several species are difficult to identify due to their similarity to other species or poor taxonomic descriptions. The difficulty in identifying nematodes species may result from morphological variations within and between populations from a same species.
Problems in the morphological identification of nematodes species, such as large number of described species within a specific group, e.g., as to compare several observed characters seen by light or scanning microscopy, and lack of apparent differences in a certain feature between species, e.g., perineal patterns, have encouraged much interest in the use of biochemical and molecular techniques as routine methods for the identification of nematodes. Biochemical and molecular methods used for the identification of nematodes are now widely used and are essential for diagnosis of a wide range of plant parasitic nematodes.
A clear understanding of species boundaries and adequate sampling of known species across their geographic areas are lacking in several regions and important crops. The future prospects in nematode taxonomy and diagnostics are dependent on molecular-based methods and tools that will discriminate not only at the species level but also at the level of host races, thereby opening up opportunities for more focused management strategies. Such techniques offer the possibility of rapid, unequivocal diagnostics and should help resolve the present problems associated with relatively morphologically conserved organisms that reproduce, for the most part, parthenogenetically, as is the case of Meloidogyne spp.
As for the case of taxonomy for Meloidogyne spp., once such molecular techniques are widely employed, no doubt the number of current nominal species will be shown to be junior synonyms, while others, conversely, will be shown to be species complexes, possibly of sibling species. It seems likely that molecular methods will replace isozymes as the preferred diagnostic tool for Meloidogyne spp. because of their inherently higher resolution and the opportunity to develop DNA chips for rapid and reliable field identification.
Molecular tools will also enhance our understanding of phylogenetic relatedness of nematodes and its relationship with other plant parasitic nematodes. Although nucleic acids-based detection techniques have been useful in diagnosing nematode species due to their high throughput characteristics, fast, sensitivity and cost effectiveness, the use of integrative diagnose approaches for nematode identification, combining morphology, biochemical and molecular data are more appropriate when necessary to strength nematode identification, define species boundaries, and to have a more suitable molecular database for nematode species.
A vast amount of genetic data are becoming available with nematode genome sequencing, which provides tools to perform comparative genomes and finds target DNA regions that can be used as diagnostic marker.
Molecular-based detection tools and methods are aimed to aid nematode taxonomy and should not totally exclude classical taxonomy approaches since in some cases, they must be complementary for accurate diagnosis.
Antimicrobial resistance is a global public health crisis. According to Public Health England [1], each year approximately 25,000 people die across Europe due to hospital-acquired infections caused by antibiotic-resistant and MDR bacteria such as Mycobacterium tuberculosis, Methicillin-resistant Staphylococcus aureus and multiresistant Gram-negative bacteria. Gram-negative infections include those caused by Escherichia coli, Klebsiella pneumoniae and Pseudomonas aeruginosa [2]. Nevertheless, it is estimated that by 2050, the global yearly death toll will increase to 10 million. Accelerating emerge of antimicrobial resistance seriously threatens the effectiveness of treatments for pneumonia, meningitis and tuberculosis, in addition to diminishing prevention of infections acquired during surgeries and chemotherapies. The crisis of the antibiotic resistance requires urgent, coordinated action. Misuse and overuse of antibiotics must be controlled, implementation of new policies regarding prescriptions has to be internationally addressed; and development of new therapeutics is urgently required [1].
Félix d’Herelle, known as the father of bacteriophage (or phage) therapy [3], brought an evolutionary discovery of phages as therapeutics for various infections and conditions. Phage therapy was widely enforced in the 1920s and 1930s to combat the bacterial infections. However, in the 1940s, the newly discovered antibiotics replaced the phage therapy (except Russia, Georgia and Poland) [4].
The emergence of MDR bacteria prompted a renewal of the interest to the phage therapy as an alternative treatment to overcome a broad spectrum of resistant bacterial infections. Phage therapy and phage cocktails that contain a mixture of different bacteria-specific phages, drawn interest within molecular biology and modern medical research as potential antimicrobials that could tackle the crisis of antimicrobial resistance. Nonetheless, the phage therapy remains controversial due to its disadvantages such as bacteriophage resistance: bacteria-phage evolutionary arms race that could put a burden on a long-time application of phage therapy as an anti-infectious agent [5].
Phage therapy has many advantages, primary because phages are very specific (generally limited to one species) and easy to obtain as they are widely distributed in locations populated by bacterial hosts including soil and seawater, and they do not have any known chemical side effects like antimicrobials [6].
Understanding host-phage interactions and ‘the war between bacteria and phages’ are steps towards designing engineering ‘broad-spectrum phage’ that can overcome the limitations of phage therapy and potentially overcome a wide range of resistant bacterial infections [6].
Phages are obligate intracellular parasites that distinctively infect bacterial cells. Although phages are very specific to their host, generally limited to one species, they pose an enormous threat to bacteria as in some habitats they outnumber their hosts by nearly 10-fold number [7]. Phages are the most abundant, ubiquitous and diversified organisms in the biosphere [8, 9]. Phage-host interaction and fight for the survival led to the evolution of bacterial and viral genomes and, therefore, to the evolution of resistance mechanisms. Bacteria, continuously, evolve many molecular mechanisms, driven by gene expression to prevent phage infection. These evolving phage-resistance mechanisms in bacteria induce the parallel co-evolution of phage diversity and adaptability [10, 11]. The co-evolving genetic variations and counteradaptations, in bacteria and phages, drive the evolutionary phage-host arm race [11, 12].
Leigh Van Valen, an evolutionary biologist, metaphorised the co-evolutionary arm race and proposed the Red Queen hypothesis [13].
‘It takes all the running you can do, to stay in the same place’ the Red Queen says to Alice in Through the Looking-Glass.
The Red Queen hypothesis proposes that to survive, microorganisms must constantly adapt, evolve and thrive against ever-evolving antagonistic microorganisms within the same ecological niche [14].
Bacteria have developed various anti-phage mechanisms including non-adaptive defences (non-specific) and adaptive defences associated with Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) along with CRISPR-associated (Cas) proteins [7, 15, 16, 17, 18].
The non-specific adaptations (analogues to innate immunity in multicellular organisms) act as primary mechanisms to evade viral infection, and they include mechanisms that inhibit phage adsorption and prevent nucleic acid entry, superinfection exclusion systems, restriction-modification systems and abortive infection [7, 19].
On the other hand, the adaptive resistance (analogues to the acquired immunity in multicellular organisms) serves as a second line of defence, which is very efficient and phage-specific.
Interestingly, it was observed that the bacterial anti-phage mechanisms are generally present in a genomic array, known as ‘defence islands’ [20]. The ‘defence islands’ are enriched in putative operons and contain numerous overrepresented genes encoding diverged variants of antiviral defence systems. Moreover, scientific evidence and characteristic operonic organisation of ‘defence islands’ show that many more anti-phage mechanisms are yet to be discovered [21, 22, 23, 24].
Although bacteria have developed several resistance mechanisms against phages, phages can circumvent bacterial anti-phage mechanisms on the grounds of their genomic plasticity and rapid replication rates. These counterstrategies include point mutations in specific genes and genome rearrangements that allow phages to evade bacterial antiviral systems such as CRISPR/Cas arrays by using anti-CRISPR proteins and abortive infection by hijacking bacterial antitoxins, as well as escaping from adsorption inhibition and restriction-modification mechanisms [15, 16, 17, 18].
This chapter will comment on the genetic basis of bacterial resistance to phages and different strategies used by phages to evade bacterial resistance mechanisms.
Phage adsorption to host-specific receptors on the cell surface is the initial step of the infection and host-phage interaction. Depending on the nature of bacteria, whether it is Gram-positive or Gram-negative proteins, lipopolysaccharides, teichoic acids and other cell surface structures can serve as irreversible phage-binding receptors [19]. These receptors might be present in the cell wall, bacterial capsules, slime layers, pili or flagella [25].
Bacteria have acquired various barriers to inhibit phage adsorption, such as blocking of phage receptors, production of extracellular matrix (e.g. capsule, slime layers) and production of competitive inhibitors [26, 27, 28, 29, 30, 31]. The diversity of phage receptors in the host is influenced by co-evolutionary adaptations of phages to overcome these barriers [32]. This includes diversity-generating retroelements (DGRs) and phase variation mechanisms causing phenotypical differences within the bacterial colony [7, 33, 34].
Phase variation is a heritable, yet reversible process regulating gene expression in bacteria; genes can switch between a functional (expression) and a non-functional state leading to phenotypical variations within the bacterial population even when strains have identical genotype. Sørensen et al. [35] investigated the underlying resistance mechanism of Campylobacter jejuni (NCTC11168) to phage F336. They have discovered that phage F336 relies on the hypervariable O-methyl phosphoramidate (MeOPN) modification of capsular polysaccharides (CPS) for successful adsorption to the bacterial surface. Nevertheless, loss of MeOPN receptor on the bacterial cell surface due to phase variation in the cj1421 gene encoding the MeOPN-GalfNAc transferase (MeOPN transferase attaches MeOPN to GalfNAc and Hep side chains of CPS) results in phage resistance [35, 36].
DGRs are genetic elements diversifying DNA sequences and the proteins they encode ultimately mediating the evolution of ligand-receptor interactions. Error-prone DGRs and random mutations in the bacterial genes encoding cell surface receptors lead to the alternation and change in the structural composition of the phage receptors, making them non-complementary to the phage’s anti-receptors, known as receptor-binding proteins (RBP) [34] (Figure 1(1)).
Bacterial defence mechanisms preventing phage adsorption and phage’s counteradaptations. (1) Phage adsorption to a host-specific receptor site on a host cell surface. Bacterium evolves phage resistance by the modification of these cell surface receptors; phage is incapable of binding to the altered receptor. (2) Phage’s adaptation to these modifications through mutations in receptor-binding protein gene that leads to the co-evolution of bacterial genetic variation. Bacteria are also capable of producing proteins that mask the phage recognition site receptors (3 and 4), thus making the receptor inaccessible for phage adsorption [28, 29, 30, 31]. Image courtesy of springer nature: https://www.ncbi.nlm.nih.gov/pubmed/20348932.
Yet, phage’s replication is exceedingly error-prone, therefore causing many random mutations in the genes encoding the RBP or tail fibres. Phages also possess DGRs that mediate phage’s tropism by accelerating the variability in the receptor-coding genes through reverse transcription process [37]. The changes in the nucleotide sequence in the RBP-coding gene may ultimately lead to the adaptation to the modified receptor (Figure 1(2)), thus the ability to adsorb and infect the bacterial cell.
Unsurprisingly, bacteria also exhibit different strategies to block their receptors [28, 29, 30, 31].
Figure 1(4) demonstrates the findings from studies conducted on Staphylococcus aureus by Nordstrom and Forsgren [38]. Mutants of Staphylococcus aureus producing higher anticomplementary protein A were found to adsorb fewer phages than Staphylococcus aureus mutants with scarce of protein A, which had an apparent increased ability to adsorb phages [38]. These findings indicate that some bacteria, including Staphylococcus aureus, are capable of production of surface proteins that mask the phage receptors making them inaccessible for phage recognition and attachment (Figure 1(3)).
Receptors located on bacterial cell surface serve a vital role in bacterial metabolism; they may function as membrane porins, adhesions or chemical receptors [19]. Therefore, mutation or complete loss of the receptor might be lethal for bacteria. To inhibit phage adsorption, bacteria can produce surface molecules, such as exopolysaccharides.
Exopolysaccharides are extracellular polysaccharides acting as a physical barrier, composing slime or capsules surrounding bacterial cells that lead to inaccessible host receptors for efficient phage adsorption [39] (Figure 2). Studies conducted by Looijesteijn et al. [40] shown that exopolysaccharides produced by Lactococcus lactis function as external protection from phages and the cell wall destructing lysozyme, due to masked cell surface receptors [40].
Bacterial strategies to inhibit phage adsorption and phage strategies to access host receptors. Some bacteria are capable of the production of exopolysaccharides, which act as an outer shield, protecting a cell from the phage infection [28, 29, 30, 31]. If the phage does not possess any polysaccharide-degrading enzymes, it cannot access the host cell membrane receptor. However, some phages evolved mechanisms allowing them to recognise these extracellular matrixes and degrade them by the means of hydrolases and lyases [15, 16, 17, 18]. Image courtesy of Springer Nature: https://www.ncbi.nlm.nih.gov/pubmed/20348932.
Nevertheless, some phages evolved mechanisms allowing them to recognise these extracellular matrixes and degrade them by utilising hydrolases and lyases (Figure 2) [15, 16, 17, 18]. The polysaccharide-degrading enzymes allow phages to gain access to the receptor that may lead to the viral propagation. They are commonly present bound to the RBPs or exist as free soluble enzymes from previously lysed bacterial cells [41].
If phage bypasses primary antiviral strategies, it is now able to initiate infection by adsorption to a specific receptor site on a host cell surface through phage RBP [42, 43]. Upon interaction with the cell receptors, the phage injects its genetic material (single or double-stranded DNA or RNA) into the cytoplasm of the host. Depending on the nature of the phage and growth conditions of the host cell, it follows one of the two life cycles: lytic or lysogenic (Figure 3).
Lytic and lysogenic life cycles of a temperate coliphage λ that infects Escherichia coli [44, 45]. cos—cohesive sites: the joining ends that circularise the linear phage λ DNA. Image courtesy of Springer Nature: https://www.nature.com/articles/nrg1089.
In the lytic cycle, virulent phages degrade host’s genome leading to the biosynthesis of viral proteins and nucleic acids for the assembly of phage progeny. Eventually, the bacterial cell lysis, releasing a multitude of newly assembled phages, is ready to infect a new host cell [46].
In contrast, temperate phages might enter the lytic or lysogenic cycle, if the host cell exists in adverse environmental conditions that could potentially limit the number of produced progeny (Figure 3 demonstrates typical lifecycle of temperate phage using coliphage λ as an example) [44, 45]. In the lysogenic phase, repressed phage genome integrates into the bacterial chromosome as a prophage. This process causes the proliferation of prophage during replication and binary fission of bacterial DNA.
Prophage only expresses a repressor protein-coding gene. The repressor protein binds to the operator sites of the other genes and ultimately inhibits synthesis of phage enzymes and proteins required for the lytic cycle.
When the synthesis of the repressor protein stops or if it becomes inactivated, a prophage may excise from the bacterial chromosome, initiating a lytic cycle (induction) which leads to the multiplication and release of virulent phages and lysis of a host cell [44, 45].
If the phage remains in the nearly dormant state (prophage), the lysogenic bacterium is immune to subsequent infection by other phages that are the same or closely analogous to the integrated prophage by means of Superinfection exclusion (Sie) systems [47].
Sie systems are membrane-associated proteins, generally, phage or prophage encoded, that prevent phage genome entry into a host cell [47]. Figure 4 shows the role of Sie system (proteins Imm and Sp) in blocking phage T4 DNA entry into Gram-negative Escherichia coli. Despite successful attachment to the phage-specific receptor, phage DNA is directly blocked by Imm protein from translocating into the cytoplasm of the cell. Sp system, on the other hand, prevents the degradation of the peptidoglycan layer by inhibiting the activity of T4 lysozyme [26, 27, 28, 29, 30, 31, 48].
Superinfection exclusion systems preventing phage DNA entry in Gram-negative Escherichia coli. (a). Standard T4 phage: upon attachment to phage-receptor on the surface of the host cell, an inner-membrane protein aids the translocation of phage DNA into the cell’s cytoplasm. (b) Imm encoding phage T4: Imm protein directly blocks the translocation of the phage DNA into the cytoplasm of the cell. (c) Imm and Sp encoding phage T4: phage DNA is prevented from entering the cell’s cytoplasm by Imm; and Sp protein prevents degradation of the peptidoglycan layer by inhibiting the activity of T4 lysozyme [28, 29, 30, 31]. Image courtesy of Springer Nature: https://www.ncbi.nlm.nih.gov/pubmed/20348932.
The evolution of bacterial genomes allowed bacteria to acquire vast mechanisms interfering with every step of phage infection. In a case where a phage succeeded to inject its viral nucleic acid into a host cell, bacteria possess a variety of nucleic acid degrading systems such as restriction-modification (R-M) systems and CRISPR/Cas that protect bacteria from the phage invasion.
It has been reported that R-M systems can significantly contribute to bacterial resistance to phages [49].
R-M systems incorporate activities of methyltransferases (MTases) that catalyse the transfer of a methyl group to DNA to protect self-genome from a restriction endonuclease (REase) cleavage and REases, which recognise and cut foreign unmethylated double-stranded DNA at specific recognition sites, commonly palindromic. To protect self-DNA from the degradation, methylases tag sequences recognised by the endonucleases with the methyl groups, whereas unmethylated phage (nonself) DNA is cleaved and degraded (Figure 5) [26, 27, 50, 51, 52].
General representation of the bacterial restriction-modification (R-M) systems providing a defence against invading phage genomes. R-M systems consist of two contrasting enzymatic activities: a restriction endonuclease (REase) and a methyltransferase. REase recognises and cuts nonself unmethylated double-stranded DNA at specific recognition sites, whereas MTase adds methyl groups to the same genomic recognition sites on the bacterial DNA to protect self-genome from REase cleavage [50, 51]. Image courtesy of: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3591985/.
R-M systems are diverse and ubiquitous among bacteria. There are four known types of R-M within bacterial genomes (Figure 6). Their classification is mainly based on R-M system subunit composition, sequence recognition, cleavage position, cofactor requirements and substrate specificity [26, 27, 50, 51].
Four distinct types of restriction-modification (R-M) systems. (a) Type I R-M system is composed of three subunits forming a complex: hsdR (restriction), hsdM (modification) and hsdS (specificity subunit that binds to an asymmetrical DNA sequence and determines the specificity of restriction and methylation). Two hsdM subunits and one hsdS subunit are involved in methylation of self-DNA. On the other hand, two complexes of hsdR, hsdM and hsdS (where each complex consists of two hsdR, two hsdM and one hsdS subunit) bind to the unmethylated recognition sites on phage DNA and cleave the DNA at random, far from their recognition sequences. Both reactions—methylation and cleavage—require ATP. (b) Type II R-M system is composed of two distinct enzymes: palindromic sequence methylating methyltransferase (mod) and endonuclease (res) that cleave unmethylated palindromic sequences close to or within the recognition sequence. (c) Type III R-M system is formed of methyltransferase (mod) and endonuclease (res) that form a complex. Methyltransferase transfers methyl group to one strand on the DNA, whereas two methyltransferases (endonuclease complexes) act together to bind to the complementary unmethylated recognition sites to cleave the DNA 24–26 bp away from the recognition site. (d) Type IV R-M system contains only endonuclease (res) that recognises methylated or modified DNA. Cleavage occurs within or away from the recognition sequences [26, 27, 50, 51]. Image courtesy of: https://www.annualreviews.org/doi/abs/10.1146/annurev-virology-031413-085500?journalCode=virology.
Due to the diversity of R-M systems, phages acquired several active and passive strategies to bypass cleavage by REases. Passive mechanisms include reduction in restriction sites, modification and change of the orientation of restriction sites, whereas more specific, active mechanisms include masking of restriction sites, stimulation of MTase activity on phage genome or degradation of an R-M system cofactor (Figure 7) [15, 16, 17, 18].
Phage’s passive and active strategies to bypass restriction-modification (R-M) systems. (a) Phages that possess fewer restriction sites in their genome are less prone to DNA cleavage by the host restriction endonuclease (REase). (b) Occasionally phage DNA might be modified by bacterial methyltransferase (MTase) upon successful injection into a host cell. Methylated recognition sites on viral DNA are, therefore, being protected from the cleavage and degradation by REase, leading to the initiation of the phage’s lytic cycle. In addition, some phages encode their own MTase that is cooperative with the host REase; thus viral DNA cannot be recognised as nonself. (c) Some phages, for example, coliphage P1, while injecting its DNA into a host cell, it also co-injects host-genome-binding proteins (DarA and DarB) that mask R-M recognition sites. (d) Phages such as Coliphage T7 possess proteins that can mimic the DNA backbone. Ocr, a protein expressed by Coliphage T7, mimics the DNA phosphate backbone and has a high affinity for the EcoKI REase component, thereby interfering with R-M system. (e) In addition, some phages (e.g. Ral protein of Coliphage λ) can also stimulate activity of the bacterial modification enzyme in order to protect own DNA from the recognition by the bacterial REase as nonself. The peptide Stp encoded by Coliphage T4 can as well disrupt the structural conformation of the REase-MTase complex [15, 16, 17, 18]. Image courtesy of: https://www.nature.com/articles/nrmicro3096.
Fewer restriction sites in the evading genome lead to the selective advantage of this phage as its DNA is less prone to cleavage and degradation by the host REase (Figure 7a). Also, some phages incorporate modified bases in their genomes that may lead to successful infection of the host cell as REase may not recognise the new sequences in the restriction sites. A decrease in the effective number of palindromic sites in DNA or change in the orientation of restriction-recognition sites can affect R-M targeting. Alternatively, the recognition sites within the viral genome can be too distant from each other to be recognised and cleaved by the REase [15, 16, 17, 18, 53].
Interestingly, phage genome might be methylated by bacterial MTase upon successful injection into a host cell. Methylated recognition sites on viral genomes are therefore being protected from the cleavage and degradation by REase, leading to the initiation of the phage’s lytic cycle. Viral progeny remains insensitive to this specific bacterial REase until it infects a bacterium that possesses a different type of REase, in which case the new progeny will become unmethylated again and will, therefore, be sensitive to the R-M system of the cognate bacterium [28, 29, 30, 31].
The fate of the host cell chiefly confides in the levels of R-M gene expression and ultimate proportion of the R-M enzymes and their competition for the sites in the invading phage genome [52].
Furthermore, some phages encode their own MTase that is cooperative with the host REase, and thereby viral DNA cannot be recognised as nonself. Phages can also stimulate the activity of host modification enzymes that can rapidly methylate viral DNA, thus protecting it from the activity of REase.
Alternatively, phages can bypass R-M systems by masking restriction sites. For example (Figure 7c), coliphage P1, while injecting its DNA into a host cell, it also co-injects host-genome-binding proteins (DarA and DarB) that mask R-M recognition sites [53, 54].
As shown on an example of a Coliphage T7 (Figure 7d), some phages code for proteins that directly inhibit REase. Coliphage T7 possesses proteins that can mimic the DNA backbone. Ocr, a protein expressed by Coliphage T7, directly blocks the active site of some REases by mimicking 24 bp of bent B-form DNA, and it has a high affinity for the EcoKI REase component, thereby interfering with R-M system [53].
Lastly, phage-bacteria arm race allowed phages to gain capabilities of degrading necessary cofactors of R-M systems. For instance, coliphage T3 encodes S-adenosyl-l-methionine hydrolase that destroys an essential host R-M cofactor (the S-adenosyl-l-methionine). The removal of this necessary co-factor will lead to the inhibition of the REase, thereby successfully infecting the host cell [15, 16, 17, 18].
CRISPR along with CRISPR-associated (Cas) proteins is the type of adaptive heritable ‘immunity’ of bacteria, thus very specific and effective; and it is prevalent within the bacterial domain [55]. The CRISPR are DNA loci consisting of short palindromic repeats (identical in length and sequence), interspaced by segments of DNA sequences (spacer DNA) derived from previous exposures to phages. The spacer DNA sequences act as a ‘memory’, allowing bacteria to recognise and destroy specific phages in a subsequent infection. Genes encoding Cas proteins are adjacent to CRISPR loci [56].
Although some studies have suggested that CRISPRs can be used for pathogen subtyping [57], it has been found that CRISPR typing is not useful for the epidemiological surveillance and outbreak investigation of Salmonella typhimurium [58].
The CRISPR/Cas phage resistance is mediated in three-step stages: adaptation (acquisition), where spacer phage-derived DNA sequences are incorporated into the CRISPR/Cas system; expression, where cas gene expression and CRISPR transcription lead to pre-CRISPR RNA (pre-crRNA) that is then processed into CRISPR RNA (crRNA); and interference, during which the crRNA guides Cas proteins to the target (subsequently invading DNA) for the degradation. The cleavage of the target (proto-spacer) depends on the recognition of complementary sequences in spacer and protospacer [59, 60].
CRISPR/Cas systems have been classified into three major types: Types I, II and III, which are further divided into subtypes that require different types of Cas proteins. Although the CRISPR/Cas array is diverse among the bacteria and it is continuously co-evolving in response to the host-phage interactions, the defence activity in all three types of the CRISPR is comparable [21, 22, 23] Figure 8 illustrates the defence mechanisms in three distinct CRISPR/Cas arrays.
Image showing mechanisms of adaptation, expression and interference in three different types of CRISPR/Cas arrays. Type I and Type II CRISPR/Cas arrays rely on the protospacer adjacent motif (PAM), contained within phage nucleic acid, to ‘select’ the phage-derived protospacer. Next steps in the adaptation stage are similar in all three types; protospacer is incorporated by Cas 1 and Cas2 proteins into the bacterial genome at the leader end of the CRISPR loci to form a new spacer. In expression step, CRISPR loci are transcribed into pre-crRNA. The crRNA processing and interference stage is distinct in each type of the CRISPR/Cas system. In Type I, the multisubunit CRISPR-associated complex for antiviral defence (CASCADE) binds crRNA to locate the target, and with the presence of Cas3 protein, the invading target genome is degraded whereas in Type II, Cas9 protein is essential in the processing of the crRNA. TracrRNA recognises and attaches to the complementary sequences on the repeat region that is then cut by RNase III in the presence of Cas9. Lastly, in Type III, processing of pre-crRNA into crRNA is dependent upon the activity of Cas6. Mature crRNA associated with Csm/Cmr complex targets foreign DNA or RNA for the degradation [21, 22, 23]. Image courtesy of: https://www.nature.com/articles/nrmicro2577.
The Type II, CRISPR/Cas9, which was first identified in Streptococcus pyogenes, gained considerable interest within scientific studies as a precise genome editing tool. CRISPR/Cas9 system is unique; a single Cas 9 protein (in addition to prevalent Cas 1 and Cas 2) is involved in the processing of crRNA and destruction of the target viral DNA [56, 61].
In the adaptation stage, phage-derived protospacer (snippet of DNA from the invading phage) is incorporated into the bacterial genome at the leader end of the CRISPR loci. In expression phase, the Cas9 gene expresses Cas9 protein possessing DNA cleaving HNH and RuvC-like nuclease domains; CRISPR locus is then transcribed and processed into mature crRNA. Finally, in interference step, the complex consisting of Cas9, crRNA and separate trans-activating crRNA (tracrRNA) cleave 20 base pairs crRNA-complementary target sequence that is adjacent to the protospacer adjacent motif (PAM) [62].
To bypass CRISPR/Cas that has an incredibly dynamic rate of evolution, phages acquired array of strategies to succeed in propagation; this includes mutations in the protospacers or in the PAM sequences and expression of anti-CRISPR proteins, and even some phages encode their own functional CRISPR/Cas systems [15, 16, 17, 18, 63].
Phages can evade interference step of Type I and Type II CRISPR/Cas system by a single point mutation or deletion in their protospacer region or in the PAM sequence (Figure 9). Phages with single-nucleotide substitutions or deletions positioned close to PAM sequence can bypass the CRISPR/Cas activity and complete their lytic cycles; in contrast, phages with multiple mutations at PAM-distal protospacer positions do not [15, 16, 17, 18, 28, 29, 30, 31].
Evasion by mutation. Mutations in the phage protospacers or in the PAM sequences allow the phage to escape interference step of the CRISPR/Cas system that would lead to the degradation of the phage genome [15, 16, 17, 18]. Adapted image courtesy of: https://www.nature.com/articles/nrmicro3096.
In some circumstances, however, although the phage successfully evades CRISPR/Cas interference, the host cell may survive by the acquisition of new spacer sequences (derived from invading phage) into their own CRISPR/Cas system. This new spacer provides the bacterium with an accelerated spectrum of phage resistance [15, 16, 17, 18].
Prophages integrated within Pseudomonas aeruginosa possess genes that encode anti-CRISPR proteins directly suppressing CRISPR/Cas-mediated degradation of the phage genome (Figure 10). According to Wiedenheft [64], these proteins might interrupt CRISPR RNA processing by preventing mature crRNA from binging to the crRNA-guide complex or by preventing the assembled crRNA-guided complex from interacting with target substrates through binding to it [64].
Anti-CRISPR proteins expressed against CRISPR subtype I-F systems. Temperate phages such as Pseudomonas aeruginosa possess genes encoding anti-CRISPR proteins that directly interfere with the bacterial CRISPR/Cas system [15, 16, 17, 18]. Adapted image courtesy of: https://www.nature.com/articles/nrmicro3096.
Prophages do not only contribute to bacterial resistance to invading phages, they can also encode proteins that contribute to bacterial virulence and antimicrobial resistance [58, 66].
Bacteria can also resist phages by possessing phage-inducible chromosomal islands (PICI) which prevent phage replication. Nevertheless, phages evolved their genomes to overcome this very specific antiviral strategy. For example, Vibrio cholerae ICP1 phages possess their own CRISPR/Cas systems that inactivate PICI-like elements (PLE) in Vibrio cholerae (Figure 11). Studies conducted by Naser et al. [67] have shown that phage CRISPR arrays have evolved by the acquisition of new spacers targeting diverse regions of PLEs carried by Vibrio cholerae strains. Furthermore, the addition of the new spacers within phage CRISPR/Cas loci enables the phages to expand their ability to counter PLE-mediated phage defence of diverse Vibrio cholerae strains [67].
Phage-encoded CRISPR/Cas systems in Vibrio cholerae ICP1 phages. Upon adsorption and injection of viral genome into a host cell, phage crRNAs and CRISPR/Cas complexes are expressed and target phage-inducible chromosomal island (PICI) in the host genome; in the Vibrio cholerae, they are termed as PICI-like elements (PLE). If the spacers within phage CRISPR locus are complementary to the bacterial PLE, the CRISPR machinery is then able to specifically target this genetic element and inactivate it, leading to the viral propagation. However, in the absence of such targeting, phage CRISPR/Cas system can acquire new spacers to evolve rapidly and ensure effective targeting of the PLE to restore phage replication [15, 16, 17, 18, 65]. Adapted image courtesy of: https://www.nature.com/articles/nrmicro3096.
Abortive infection (Abi) systems promote cell death of the phage-infected bacteria, inhibiting phage replication and providing protection for bacterial populations [68].
Abi systems require both toxins and antagonistic antitoxins. Antitoxins are proteins or RNAs that protect bacterial cell from the activity of toxins in a typical cell life cycle, whereas toxins are the proteins encoded in toxin-antitoxin locus that disrupt cellular metabolism (translation, replication and cell wall formation), causing cell death. During an infection, the expression of the antitoxin encoding gene is suppressed, leading to the lethal activation of the toxin [69]. Figure 12 illustrates the mechanism of Abi systems in Escherichia coli [70].
Abortive infection (Abi) systems in Escherichia coli. The Rex system is a two-component Abi system. A phage protein-DNA complex (formed during phage replication) activates the sensor protein RexA, which in turn activates RexB. RexB is an ion channel that causes depolarisation of the bacterial membrane leading to cell death [28, 29, 30, 31]. Image courtesy of Springer Nature: https://www.ncbi.nlm.nih.gov/pubmed/20348932.
Interestingly, phages evolved an array of tactics to circumvent Abi systems. This includes mutations in specific phage genes and encoding own antitoxin molecules that suppresses bacterial toxin [15, 16, 17, 18]. Figure 13 provides a broad overview of the strategies employed by the phages to by-pass Abi systems.
Escaping abortive infection mechanisms. (a) In a typical cell life cycle, antitoxins protect bacterial cell from the activity of toxins. (b) During phage infection, the expression of antitoxin encoding gene is suppressed, leading to the lethal activation of the toxin. (c) Mutations in certain phage genes can lead to escaping Abi systems activity, thereby a successful viral propagation without killing the host cell. (d) Some phages encode molecules that functionally replace the bacterial antitoxins, thus suppressing toxin activity and avoiding host cell death [15, 16, 17, 18]. Image courtesy of: https://www.nature.com/articles/nrmicro3096.
Bacteria-phage interaction is therefore very complex, and it is crucial to understand the molecular basis of this interaction and how bacteria and phages ‘fight’ each other. It has been reported that Anderson Phage Typing System of Salmonella Typhimurium can provide a valuable model system for study of phage-host interaction [71].
The rapid emergence and dissemination of MDR bacteria seriously threaten global public health, as, without effective antibiotics, prevention and treatment of both community- and hospital-acquired infections may become unsuccessful and lead to widespread outbreaks.
Carbapenems and colistin are antibiotics of last resort, generally reserved to treat bacteria which are resistant to all other antibiotics. Until not long ago, colistin resistance was only described as chromosomal, however, in 2016 Liu et al. reported the emergence of the first plasmid-mediated colistin resistance mechanism, MCR-1, in Enterobacteriaceae [72]. Furthermore, the increasing occurrence of colistin resistance among carbapenem-resistant Enterobacteriaceae has also been reported [73]. This is of significant concern as infections caused by colistin and carbapenem-resistant bacteria are very challenging to treat and control, as the treatment options are greatly limited or non-existent. Thus, the discovery and development of alternative antimicrobial therapeutics are the highest priorities of modern medicine and biotechnology.
Phages should be considered as great potential tools in MDR pathogens as they are species-specific (specificity prevents damage of normal microbiota), thus harmless to human; they have fast replication rate at the site of infection, and their short genomes can allow to further understand various molecular mechanisms implied to ‘fight’ bacteria. In addition, this understanding can enable scientists to ‘manipulate’ viral genomes and engineer a synthetic phage that combines the antibacterial characteristics of multiple phages into a single genome.
The escalating need for new antimicrobial agents attracted new attention in modern medicine, proposing several potential applications of phages as antibacterial therapeutics including phage therapy, phage lysins and genetically-engineered phages.
Phage therapy utilises strictly lytic phages that have bactericidal effect. As phages are host-specific, ‘phage cocktails’ containing multiple phages can broaden range of target cells. Nevertheless, selection of suitable phages is at the paramount to the successful elimination of clinically important pathogens, and it includes avoidance of adverse effects, such as anaphylaxis (adverse immune reaction) [74].
In order to hydrolyse and degrade the bacterial cell wall, phages possess lysins.
The spectrum of efficiency of natural lysins (derived from naturally occurring phages) is generally limited to Gram-positive bacteria; however, recombinant lysins have shown an ability to destabilise the outer membrane of Gram-negative bacteria and ultimately lead to rapid death of the target bacteria [74].
Bioengineered phages have the potential to solve inherent limitations of natural phages such as narrow host range and evolution of resistance. Various genetic engineering methods have been proposed to design phages with extended antimicrobial properties such as homologous recombination, phage recombineering of electroporated DNA, yeast-based platform, Gibson assembly and CRISPR/Cas genome editing [75].
Engineering of synthetic phages could be tailored to enhance the antibiotic activity, to reverse antibiotic resistance or to create sequence-specific antimicrobials [74].
The antagonistic host-phage relationship has led to the evolution of exceptionally disperse phage-resistance mechanisms in the bacterial domain, including inhibition of phage adsorption, prevention of nucleic acid entry, Superinfection exclusion, cutting phage nucleic acids via restriction-modification systems and CRISPR, as well as abortive infection.
Evolvement of these mechanisms has been induced by constant parallel co-evolution of phages as they attempt to coexist. To survive, phages acquired diverse counterstrategies to circumvent bacterial anti-phage mechanisms such as adaptations to new receptors, digging for receptors and masking and modification of restriction sites and point mutations in specific genes and genome rearrangements that allow phages to evade bacterial antiviral systems such as CRISPR/Cas arrays, as well as mutations in specific genes to bypass abortive infection system. Conclusively, the co-evolving genetic variations and counteradaptations, in both bacteria and phages, drive the evolutionary bacteria-host arm race.
Besides, accumulating evidence shows that phages contribute to the antimicrobial resistance through horizontal gene transfer mechanisms. Indeed, many bacterial strains have become insensitive to the conventional antibiotics, posing a growing threat to human; and although in the past, western counties withdrew phage therapy in response to the discovery of therapeutic antibiotics, now, phage therapy regains an interest within the research community. There are apparent advantages of phage therapy, such as specificity, meaning only target bacteria would encounter lysis, but not healthy microbiota inhabiting human’s system. Additionally, ‘phage cocktails’, containing multiple bacteria-specific phages, could overcome the issue of phage-resistance as phages do adapt to these resistance mechanisms. However, ‘phage cocktails’ would require large numbers of phages that would have to be grown inside pathogenic bacteria in the laboratory, putting laboratory staff and the environment at risk.
Alternatively, building up the understanding of host-phage interactions and ‘the war between bacteria and phages’ could potentially lead to defeating antimicrobial resistance by designing synthetic phages that can overcome the limitations of phage therapy.
Dr Manal Mohammed is funded by a Quinton Hogg start-up award, University of Westminster.
abortive infection capsular polysaccharides clustered regularly interspaced short palindromic repeats crispr RNA diversity-generating retroelement deoxyribonucleic acid multidrug-resistant O-methyl phosphoramidate methyltransferase protospacer adjacent motif phage-inducible chromosomal island PICI-like element receptor-binding protein restriction endonuclease restriction-modification ribonucleic acid superinfection exclusion trans-activating crRNA
IntechOpen implements a robust policy to minimize and deal with instances of fraud or misconduct. As part of our general commitment to transparency and openness, and in order to maintain high scientific standards, we have a well-defined editorial policy regarding Retractions and Corrections.
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\\n\\n1. RETRACTIONS
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\\n\\n1.2. REMOVALS AND CANCELLATIONS
\\n\\n2. STATEMENTS OF CONCERN
\\n\\nA Statement of Concern detailing alleged misconduct will be issued by the Academic Editor or publisher following a 3rd party report of scientific misconduct when:
\\n\\nIntechOpen believes that the number of occasions on which a Statement of Concern is issued will be very few in number. In all cases when such a decision has been taken by the Academic Editor the decision will be reviewed by another editor to whom the author can make representations.
\\n\\n3. CORRECTIONS
\\n\\nA Correction will be issued by the Academic Editor when:
\\n\\n3.1. ERRATUM
\\n\\nAn Erratum will be issued by the Academic Editor when it is determined that a mistake in a Chapter originates from the production process handled by the publisher.
\\n\\nA published Erratum will adhere to the Retraction Notice publishing guidelines outlined above.
\\n\\n3.2. CORRIGENDUM
\\n\\nA Corrigendum will be issued by the Academic Editor when it is determined that a mistake in a Chapter is a result of an Author’s miscalculation or oversight. A published Corrigendum will adhere to the Retraction Notice publishing guidelines outlined above.
\\n\\n4. FINAL REMARKS
\\n\\nIntechOpen wishes to emphasize that the final decision on whether a Retraction, Statement of Concern, or a Correction will be issued rests with the Academic Editor. The publisher is obliged to act upon any reports of scientific misconduct in its publications and to make a reasonable effort to facilitate any subsequent investigation of such claims.
\\n\\nIn the case of Retraction or removal of the Work, the publisher will be under no obligation to refund the APC.
\\n\\nThe general principles set out above apply to Retractions and Corrections issued in all IntechOpen publications.
\\n\\nAny suggestions or comments on this Policy are welcome and may be sent to permissions@intechopen.com.
\\n\\nPolicy last updated: 2017-09-11
\\n"}]'},components:[{type:"htmlEditorComponent",content:'IntechOpen’s Retraction and Correction Policy has been developed in accordance with the Committee on Publication Ethics (COPE) publication guidelines relating to scientific misconduct and research ethics:
\n\n1. RETRACTIONS
\n\nA Retraction of a Chapter will be issued by the Academic Editor, either following an Author’s request to do so or when there is a 3rd party report of scientific misconduct. Upon receipt of a report by a 3rd party, the Academic Editor will investigate any allegations of scientific misconduct, working in cooperation with the Author(s) and their institution(s).
\n\nA formal Retraction will be issued when there is clear and conclusive evidence of any of the following:
\n\nPublishing of a Retraction Notice will adhere to the following guidelines:
\n\n1.2. REMOVALS AND CANCELLATIONS
\n\n2. STATEMENTS OF CONCERN
\n\nA Statement of Concern detailing alleged misconduct will be issued by the Academic Editor or publisher following a 3rd party report of scientific misconduct when:
\n\nIntechOpen believes that the number of occasions on which a Statement of Concern is issued will be very few in number. In all cases when such a decision has been taken by the Academic Editor the decision will be reviewed by another editor to whom the author can make representations.
\n\n3. CORRECTIONS
\n\nA Correction will be issued by the Academic Editor when:
\n\n3.1. ERRATUM
\n\nAn Erratum will be issued by the Academic Editor when it is determined that a mistake in a Chapter originates from the production process handled by the publisher.
\n\nA published Erratum will adhere to the Retraction Notice publishing guidelines outlined above.
\n\n3.2. CORRIGENDUM
\n\nA Corrigendum will be issued by the Academic Editor when it is determined that a mistake in a Chapter is a result of an Author’s miscalculation or oversight. A published Corrigendum will adhere to the Retraction Notice publishing guidelines outlined above.
\n\n4. FINAL REMARKS
\n\nIntechOpen wishes to emphasize that the final decision on whether a Retraction, Statement of Concern, or a Correction will be issued rests with the Academic Editor. The publisher is obliged to act upon any reports of scientific misconduct in its publications and to make a reasonable effort to facilitate any subsequent investigation of such claims.
\n\nIn the case of Retraction or removal of the Work, the publisher will be under no obligation to refund the APC.
\n\nThe general principles set out above apply to Retractions and Corrections issued in all IntechOpen publications.
\n\nAny suggestions or comments on this Policy are welcome and may be sent to permissions@intechopen.com.
\n\nPolicy last updated: 2017-09-11
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