Open access peer-reviewed chapter

Genetic Resistance to the Reniform Nematode in Cotton

Written By

Seloame T. Nyaku, Yonathan Tilahun, Kathy Lawrence, Venkateswara R. Sripathi, Abreeotta J. Williams and Govind C. Sharma

Submitted: December 8th, 2015 Reviewed: May 24th, 2016 Published: November 9th, 2016

DOI: 10.5772/64389

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Abstract

Among major nematode pests of Upland, cotton production is the reniform nematode, which is a serious threat in various cotton-producing regions. The availability of germplasm lines with tolerance or resistance to this menacing pest is a valued asset. To date, various laboratories and research institutions have collaborated to transfer the reniform nematode resistance from wild gene pools of cotton into widely cultivated Upland cotton, which have led to positive results. This chapter focuses on the current status of these introgressions and resistance mechanisms in cotton. In this overview, four major themes are being pursed: (1) tolerance mechanisms in cotton to the reniform nematode, (2) genotype evaluations, (3) introgression of reniform resistance into Upland cotton, and (4) functional analysis of reniform infection in Upland cotton. Genetic resistance in Upland cotton to the reniform nematode is the only practical solution because conventional control measures are the most cost-effective and environmentally sustainable and therefore have been and will be actively pursued. Resistance genes, if successfully introgressed into crop plants from wild relatives, should complement management of the reniform nematode with traditional methods.

Keywords

  • functional analysis
  • introgression
  • reniform nematode
  • resistance mechanisms
  • tolerance

1. Introduction

Cotton (Gossypium hirsutumL.) is one of the most important fiber crops of the world. However, in selected regions, yields are being reduced drastically by the reniform nematode, Rotylenchulus reniformis. This pest was reported in 1940 [1] and to date has become a significant pest of cotton in the southeastern and south-central United States [2, 3]. The estimated cotton production loss that occurred in the United States due to reniform nematode in 2005 was 115 × 106 kg (526,000 bales), conservatively worth US$1 kg–1 [4, 5].

The reniform nematode is a semi-endoparasite and derives its name from having a kidney-shaped body, usually of the adult female, and the males are, however, vermiform. The reniform life cycle is in four stages beginning from the egg. The first juvenile stage (J1) molts to become the second juvenile (J2) which also occurs within the egg. The J2 then hatches 1–2 weeks after eggs are laid especially when conditions are favorable [5]. Cuticles may sometimes overlay the other and are seen usually in the third- and fourth-stage juveniles, J3 and J4 [6]. Generally, after molting, there is a reduction in the body size of the reniform nematode [5]. The young adult reniform nematode females move towards the germinating host seedlings and to certain organic substances secreted by the roots [7]. Penetration by the nematode is in the elongation zone of the host plant root, proximal to the meristem. The epidermis and cortex are pierced by the vermiform young female intracellularly with the pericycle being the permanent feeding site [8, 9]. Half to two-thirds of the nematodes’ body usually remains outside the root, and the nematode swells with kidney-like shape morphology.

Among the cellular changes observed after penetration of the cells by the reniform nematode include the formation of dense granular cytoplasm, hardening of cell walls through accumulation of polysaccharides, disintegration of cell walls, and enlargement of cells [1012]. The adult female reniform nematode oviposits into a gelatinous matrix, produced by the vaginal glands, and the number of eggs within an egg mass varies, from 60 to 200 by a single adult female nematode [13]. The reniform nematode has the capability to undergo anhydrobiosis, a survival mechanism in the absence of water, making it thrive for at least 2 years in the absence of a host in dry soil [14]. The state of anhydrobiosis for the reniform nematode can last for more than 20 years in the absence of host plants [15].

Root-knot nematode (Meloidogynespp.), another damaging nematode of cotton, are sedentary endoparasites that feed within the roots of host plants, resulting in drastic yield losses [16, 17]. The major observable signs on plants are the root knots that indicate colonization of roots of infected plants by these organisms. Symptoms include low yields, stunted growth, wilting, and predisposition to other pathogens. The most damaging stage is the second-stage juveniles (J2), which penetrate and invade their hosts near root tips and then migrate intercellularly toward the vascular cylinder. These nematodes puncture the cell wall continuously, and there is secretion release from the stylet into the cytoplasm, and the cytoplasm contents are ingested. The cells enlarged to form giant cells [18], and these have multiple nuclei and are very active metabolically in their functions [19]. The life cycle of Meloidogynelasts for about 3 weeks, and eggs are then released into a gelatinous matrix by the adult female. During this period of feeding within the plant roots, physiological changes occur which alter gene expression mechanisms of the host [20].

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2. Tolerance and resistance mechanisms of cotton to reniform nematode

Studies have shown changes in host gene expression during infection of roots by nematodes especially within the syncytium [2123]. During the formation of syncytium, a number of changes occur within plant cells, among these are changes in cell cycle mechanisms, hormone regulation events, and cell wall architecture [24, 25]. Establishment of nematode feeding cells is through specific processes which are controlled and directed by encoded products of nematode parasitism genes found within the esophageal glands. These products are then delivered into the feeding cell through the nematode stylet [26]. Processes and events occurring at the nematode feeding sites play roles related to various degrees of susceptibility of cotton plants to the reniform nematode. Among these events are early degradation of syncytia [10], formation of wall deposits, and absence of hypertrophy in pericycle cells. These are some known mechanisms proposed for cotton resistance to reniform nematode.

Various Upland cotton varieties were planted in Unites States (US) in 2015 [27]. Among the varieties grown in United States, the largest and least acreages of cultivation were ‘Deltapine’ (31.2%) and 'Seed Source Genetics' (0.02%), respectively. ‘Deltapine’ cultivar covered about 50.1% of the acreage planted to Upland cotton in the southeastern states (Alabama, Florida, Georgia, North Carolina, South Carolina, and Virginia). Bayer CropScience ('FiberMax' and 'Stoneville') also covered significantly higher acreages (21.61 and 16.93%), respectively. An early study was conducted on susceptible (DP50-HR: 'Deltapine 50' with higher reproduction) and resistant (DP50-LR: 'Deltapine 50' with lower reproduction) genotypes, which were inoculated with 3000 reniform nematodes [28]. Roots collected 3, 6, 9, 12, and 15 days after inoculation (dai) revealed reduced reproduction in susceptible genotypes with degeneration of the syncytial cells and absence of hypertrophy within the pericycle cells.

Reniform nematodes show significant variations in populations [2934]. Cotton genotypes will, therefore, have varying reproductive and pathogenicity responses to reniform nematode variants if present in populations. The ‘LONREN ’genotypes (‘LONREN-1’ and 'LONREN-2' both of which are resistant to the reniform nematode) were used in association with susceptible genotypes ('FiberMax 966' and 'Deltapine 555BR') to six levels of inocula (0, 500, 1000, 5000, 10,000, or 50,000), consisting of eggs and vermiform life stages of reniform nematodes [35]. High inoculum levels (10,000 and 50,000) significantly reduced the root dry mass of 'LONREN' genotypes; however, higher levels in the susceptible genotypes rather stimulated root mass. Thus, the 'LONREN' genotypes may be involved in hypersensitive responses to nematode parasitism.

Another study was conducted on five cotton genotypes with various resistance/tolerance levels to reniform nematode isolates obtained from cotton field in Louisiana [36]. High reproduction of isolates was observed from Evan (33,793 juveniles/250 g soil) and Avoyelles (27,800 juveniles/250 g soil) genotypes. Data revealed that the G. arboreum('A2-190') and 'LONREN-2' were the most resistant genotypes among the nematode isolates. However, 'TX-110' and 'BARBREN-713' both had high levels of resistance to the reniform nematodes in terms of pathogenicity. The 'LONREN-1', 'LONREN-2', and 'A2-190' genotypes showed hypersensitivity to reniform nematode invasion of roots as seen in stunted plant growth. This study therefore confirms the presence of variation in both pathogenicity and reproduction of reniform nematode isolates in cotton fields. Isolates obtained from various infested cotton fields may have various adaptive mechanisms in the soils in which they occur [37, 38]. Further, genetic variation within specific genes, for example, rRNA and ITS1 and microsatellite loci of the reniform nematode [2931, 34, 39, 40] might influence pathogenicity and reproductive ability of nematodes. Simple sequence repeat (SSR) markers have been used to reveal the polymorphism in reniform nematode sampled from infested fields in Mississippi [40].

Exploring of host resistance to nematode parasitism is the most effective and environmentally friendly method of nematode management. Resistant cultivars pose an incompatible interaction with nematodes. A hypersensitive reaction is produced in a response to the feeding or invading nematode leading to lignification of the cells in close proximity to the reniform nematode's head, or the cells either collapse or become necrotic [41, 42]. However, in a compatible interaction, there is a formation of active syncytia with cells becoming hypertrophic, dense cytoplasm, enlarged nuclei, in most instances a partial disintegration of cell walls [29]. Application of nematicides to the resistant genotypes of cotton, for example, 'LONREN' have a positive impact on plant response and yield [43]. This hypersensitivity which is produced can be reduced in seedlings after nematicide applications. Four-resistant breeding lines from ('LONREN-1' × 'FM966') cross, a susceptible line from the 'LONREN' × 'FM966' cross, 'LONREN-1', 'BARBREN-713', and the 'DP393' (susceptible cultivar) were explored in a nematicide trial [43]. There was an increase in plant height and yield among plants to which nematicides were applied in greenhouse experiments. Furthermore, the number of reniform nematodes isolated was about 50% lower in resistant genotypes 45 days after planting (DAP). Differences were also observed among 'BARBREN-713' and 'LONREN'-derived lines in relation to reniform nematode egg number counts, with lower counts noted on 'BARBREN-713'. There was also a reduction in stunting of genotypes that received aldicarb treatment.

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3. Evaluation of G. hirsutumgenotypes to reniform nematode

Within the Gossypiumgenus, about 50 diploid and allotetraploid species are known. Two widely cultivated diploids include Gossypium arboreumL. and G. herbaceumL., and the two cultivated allotetraploids are G. hirsutumL. and G. barbadenseL. [44]. There are eight diploid genome groups (A, B, C, D, E, F, G, and K) with the same chromosome number (2n = 26) [45]. Among the wild diploid species, three geographical groups are known (Australian, American, and Afro-Arabian).

Towards the end of the 1980s, some germplasm lines ('La. RN 4-4', 'La. RN 909', 'La. RN 910', and 'La. RN 1032') were developed in Louisiana State University in Baton Rouge with low-to-moderate levels of resistance to the reniform nematode [46]. These lines were from Upland cotton and were the first with some levels of resistance to the reniform nematode. Towards the latter part of 1997, the Unites States Department of Agriculture (USDA) Agricultural Research Service (ARS) scientists in Texas released four germplasm lines ('N220-1-91', 'N222-1-91', 'N320-2-91', and 'N419-1-91') with slightly higher levels resistance to the reniform nematode compared to earlier released lines [47].

Further screening of G. hirsutumgenotypes has been conducted [4649]. However, most of these lines have varying tolerance levels to the reniform nematode [47]. Several genotypes (G. hirsutum(110), G. herbaceum(7), G. arboreum(14), G. barbadense(6),wild Gossypiumspp. (33), Hibiscusspp. (22), and other Malvaceae (7) were evaluated for their resistance against the reniform nematode, through their ability to reproduce on roots [50]. The genotypes (G. stocksii, G. somalense, and G. barbadenseand 'Texas110' possessed high resistance to the reniform nematode. Gossypium hirsutum893, G. herbaceumPI 408775, G. arboreumPI 41895, PI 417891, and CB 3839 also have low nematode egg productions (20%).

Resistance has also been explored in wild accessions of G. hirsutumand G. barbadenseto M. incognitarace 3 and reniform nematodes [51]. The G. barbadenseaccessions (‘TX-1347’ and ‘TX-1348’) had lower reproduction on them compared to the G. hirsutumaccessions although were susceptible to M. incognitarace 3. A greenhouse study was conducted on 52 cultivars of cotton to identify those with some resistance to the reniform nematode; however, all the cultivars were susceptible to this organism [48]. Among the various cultivars, those with the lowest reproductive factors (RF) were ‘SG 105’, 'DP 543 BGII'/'RR', 'DP 445BG'/'RR', and 'FM 989 R'. Other G. hirsutumlines ('MT2468 Ren1', 'MT2468 Ren2', and 'MT2468 Ren3') with resistance to the reniform nematode were released [52]. This effort was from Scientists at the USDA ARS and the Mississippi Agricultural and Forestry Experiment Station.

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4. Introgression of reniform nematode resistance into Upland cotton

Researchers over the years have screened genotypes and believed to have resistance to the reniform nematode. They have proved that these genotypes are not 100% resistant to the reniform nematode [4850]. This, therefore, has resulted in multi-institutional collaborations on the introgression of resistance into Upland cotton from close and distant relatives of Gossypium[53, 54]. Monosomic addition lines were developed with the aim of transferring resistance into Upland cotton from G. longicalyx[55]. However, after screening 12 lines that were segregating, resistance to this pest was minimal. However, there have been successful introgressions into the reniform nematode into G. hirsutumfrom G. arboreum[56], G. aridum[57], G. barbadense[5860] making the introgressed lines resistant to the reniform nematode.

The introgression of the reniform nematode resistance trait of G. longicalyxinto Upland cotton was achieved through the development of two 52-chromosome trispecies hybrids [(G. hirsutum × G. longicalyx) chromosome-doubled × G. armourianum] (abbreviated as HLA) and [(G. hirsutum × G. herbaceum) chromosome-doubled × G. longicalyx] (abbreviated as HHL) [61]. The G. longicalyx(F1) had the resistance trait. In this approach, there was a need to include a genotype that could reduce ploidy characteristics existing between the 'donor' and 'recipient' genotypes. Therefore, either G. armourianumKearney (D2–1) or G. herbaceumL. (A1) were used in genetic crosses. A number of recurrent backcrossings were undertaken that produced 28 lines with reniform nematode resistance, and segregation (1:1) was observed in all the backcrosses for resistant and susceptible traits [61, 62]. This demonstrated that the trait responsible for reniform nematode resistance is single gene controlled and heritable. This effort led to the release of two G. hirsutumgermplasm lines, 'LONREN-1' (Registration # GP-977, PI 669509) and 'LONREN-2' (Registration # GP-978, PI 669510). The release was by both USDA and Cotton Incorporated to enable cotton farmers’ access to the reniform nematode-resistant germplasm with improved yields. The introgressed chromosome segments in both lines differ slightly in terms of size, with that of 'LONREN-2' much smaller than that of 'LONREN-1'. Experimental assays revealed a 95% reduction in nematode populations in growth chambers and 50–90% reductions in field trials. A disadvantage with these lines, however, is the reduced plant growth and stunting in fields with very high populations of reniform nematodes, reflecting in poor yields [63, 64]. This gene conferring high resistance to the reniform nematode has also been introgressed from the African species G. longicalyx(Hutch. & Lee, 2n = 2x = 26; 2F1) [65]. The allele conferring this resistance (Renlon) was linked to chromosome 11. In that study [65], two trispecies hybrids made up of G. hirsutum(AD)1, the recipient species, G. longicalyx(F1) the donor parent, and two wild diploids, G. armourianum(D2–1) and G. herbaceum(A1), were used as 'bridges'.

Various screens were performed on G. hirsutum, G. longicalyx, G. armourianum, and HLA trispecies hybrids to identify markers that could be linked to resistance in the reniform nematode [65]. Sixty-two simple sequence repeat (SSR) markers were used in amplifying pooled DNA from G. hirsutumcultivars ['Auburn (Aub)-623', 'AcalaNemX', and 'Deltapine (DP)-458'] for the identification of polymorphisms on the A-sub-genome. Some primer pairs revealed polymorphisms among various groups of resistant (F1 and HLA) and susceptible (AD)1 and (D)2–1 genotypes. These primer pairs were selected and used for screening genotypes in BC2F1 populations to identify markers associated with the reniform resistance locus Ren. A phenotypic marker in G. longicalyxcould be linked with green seed fuzz (Fzglon) which was closely associated with the reniform nematode. This association was confirmed by screening 984 resistant and susceptible genotypes in various backcrosses.

Transfer of reniform nematode resistance into Upland cotton has also been successfully achieved through crossing the resistant G. arboreum(A-genome) with a D-genome species [66]. Backcrossing with Upland cotton (AD)1, and screenings of the BC2F1 and the BC2F2 for resistance to reniform nematode showed plants with resistance to this pest. Another successful reniform nematode resistance introgression into G. hirsutumwas from G. aridum[57]. This study involved mapping of traits from various genotypes that have been backcrossed. This included a trispecies hybrid, [G. arboreum × (G. hirsutum × G. aridum)], crossed with ’MD51ne’ (G. hirsutum). One hundred and four simple sequence repeat markers were then used to identify markers linked with resistance in the reniform nematode in 50 and 26 resistant and susceptible progenies, respectively, obtained from the above crosses. Among the various markers utilized, 25 of them were resistance specific to the reniform nematode and were all localized to chromosome 21 of cotton. The locus of resistance was Renariand two of these markers identified were BNL3279_132 and BNL2662_090.

Reniform nematode resistance introgression into G. hirsutumfrom G. arboreumaccession (’A2-190’) crossed with a hexaploid (AD1) × D4 bridging line (G 371) is another success story [56]. The progenies were tetraploid triple-species hybrids. They were progressively backcrossed into G. hirsutumproducing 277 BC1 new genotypes. Growth chamber experiments revealed that the ‘G371’, a hexaploid bridging line, poses resistance to the reniform nematode. Therefore, resistance to the reniform nematode was controlled by dominant genes.

Trispecies hybrids with G. thurberihave been utilized in transferring resistant traits into G. hirsutum[67]. This hybrid [(G. hirsutum × G. thurberi) a hexaploid [2n = 6× = 78, (AADD)1 × D1] × G. longicalyx(F)] was used in cytogenetic analysis. Fifteen simple sequence repeat (SSR) markers were also utilized in accessing the introgressions in the various hybrid plants through specific bands, which confirmed successful introgressions. Cytogenetic analysis revealed a chromosome configuration 2n = 52 = 14.13 I + 15.10 II + 1.03III + 0.9 IV + 0.03 V + 0.13 VI (I, II, III, IV, V, and VI refer to univalents, bivalents, trivalents, tetravalents, pentavalents, and hexavalents, respectively). Mitotic chromosome analysis provided evidence on the number of chromosome within the genomes of the species and hybrids. The chromosomes varied in number for G. hirsutum(52), G. thurberi(26), G. longicalyx(26), and (G. hirsutum × G. thurberi) (78), and three-species hybrid [(G. hirsutum × G. herbaceum) × G. longicalyx] (HTL) (52).

Large germplasm lines (‘1866’ and ‘907’) of G. hirsutumand G. barbadense,respectively, have been effectively screened in efforts to identify resistance to the reniform nematode [68]. Introgressions from G. barbadenseinto G. hirsutumagainst the reniform nematode, and markers linked to resistance in this pest have been a success [5860, 68, 69].

Greenhouse assays also complement field screening of germplasm lines for nematode resistance in these lines [70]. In one of these assays, a single test plant was screened against six susceptible and resistant plants of G. hirsutum, 'Deltaple-16' and moderately resistant G. barbadense'TX-1348', respectively [51]. Screening assays revealed about 5 and 12% of G. hirsutumand G. barbadenseaccessions having lower numbers of reniform nematodes compared to as susceptible check 'TX-1348', respectively. Moderate resistance was observed in the G. barbadensecultivar 'TX-110' (PI 163608) and G. barbadenseaccession 'GB 713' (PI 608139). Among the various G. barbadenseaccessions screened with moderate resistance to M. incognita, none of these was resistant to the reniform nematode.

Texas AgriLife Research released two breeding lines 'TAM RKRNR-9' (Reg. No. GP-941; PI 662039) and 'TAM RKRNR-12' (Reg. No. CP-942; PI 662040) of Upland cotton in 2010 [58]. Both of these lines poses resistance to M. incognitaand the reniform nematode. These germplasm lines were developed through the crosses between 'M-315 RNR', an M. incognitaresistant G. hirsutumline, and a reniform nematode-resistant G. barbadenseline ‘TX-110'. Both germplasm lines are commercially available and can be utilized in cotton breeding research. In other collaborative efforts, Research Scientists at the USDA and Mississippi Agricultural and Forestry Experiment Station released three germplasm lines, 'M713 Ren1' (Reg. No. GP-958, PI 665928), 'M713 Ren2' (Reg. No. GP-959, PI 665929), and 'M713 Ren5' (Reg. No. GP-960, PI 665930) in 2012 [59]. The reniform nematode resistance within these lines was from the G. barbadense accession'GB 713'.

SSR markers associated with reniform nematode quantitative trait loci (QTLs) involved in resistance are known [69]. This was achieved through genotyping of 300 F2 populations of 'GB713' × 'AcalaNem-X' crosses. QTLs were localized to chromosomes 21 and 18, respectively. The QTLs on chromosome 21 were on map positions 168 (LOD 28.0) and 182.7 (LOD 24.6), with the specific SSR markers BNL 1551_162 and GH 132_199 on position 154.2 and 177.3 and BNL 4011_155 and BNL 3279_106 on positions 180.6 and 184.5 associated with these loci, respectively. However, the only single QTL on chromosome 18 was on the map position 39.6 (LOD 4.0) with the specific SSR markers BNL 1721_178 and BNL 569_131 on positions 27.6 and 42.9, respectively. The authors, therefore, suggested the following designations Renbarb1and Renbarb2for QTLs located on chromosome 21 and Renbarb3for those on chromosome 18. Further experiments in controlled environments on 'GB-713' showed a reduction in reniform and M. incognitanumbers by 90% [60].

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5. Functional analysis of reniform nematode infection in cotton

5.1. Parasitism genes

Sequencing of the nematode parasitome usually involves gene products that the nematode secretes during its cycle; critical study of genes will enhance the understanding of the nature of damage caused by nematodes to host plants [22]. Majority of these genes encode cell wall-modifying proteins such as galactosidases, xylanases, pectinases, and expansins [71]. Isolation of the parasitome from plant–parasitic nematodes, for example, β-1-4-endoglucanases (cellulases) was from the subventral glands of Heterodera glycinesand Globodera rostochiensis[72, 73]. These proteins were the first isolated molecules in plant–parasitic nematodes. Cellulases are needed by nematodes to degrade cell walls for easy penetration to the roots [74].

Cellulase genes have been identified within reniform nematode [7577], H. glycines[78], G. tabacum[79], Pratylenchus penetrans[80], M. incognita[81], and H. schachtii[82]. Within the cyst nematode, cellulase activity has been observed in J2 juveniles, but rarely in J3 juveniles [78]. These cellulases are associated with the glycosyl hydrolase family 5, with two domain structures (i.e., a catalytic domain with and without a cellulose-binding domain). The cellulase gene (RR-ENG-1) of 1341-bp length has also been molecularly characterized in the reniform nematode. This gene had a 19-bp 5′-untranslated region (UTR), a 1245-bp open-reading frame (ORF), and an 80-bp 3′-UTR region [75]. Multiple sequence alignment of the cDNA and genomic sequences revealed seven introns and eight exons for RR-ENG-1gene.

Further BLAST analysis gave hits to HG-ENG-6mRNA in H. glycines. Semiquantitative RT-PCR used in studying gene expression revealed RR-ENG-1highly expressed in the second-stage juvenile (J2) and adult vermiform life stages. However, expression levels in the adult female were much lower. Recent characterization of the reniform nematode genome revealed hits to HG-ENG-6and ENG-1genes in H. glycines(Accession # AA025506) and Radophilus similis(Accession# ACB38289), respectively [77]. Similarly, transcriptome sequencing of the reniform nematode revealed hits to ENG-2(Accession # AAK21881.1 and AAK21883.2), ENG-1(Accession # AAD45868.1), and ENG-7(Accession # AAK21887.1) genes in Meloidogynespp. [76].

Two cellulase cDNAs (HG-ENG-1and GR-ENG-1), that code for a secretion of signal peptide, cellulase catalytic domain, cellulose binding domain (CBD), and a small peptide linker, are within the cyst nematode genome. Secretion of these cellulases was within the subventral esophageal gland cells, and their presence was confirmed through mRNA in situ hybridization and immunolocalization. The cellulase activity is usually enhanced by the presence of the CBD. In G. rostochiensis, characterization of its cellulase cDNA (GR-ENG-2) showed absence of a CBD, which inhibits synthesis of crystalline cellulose [72, 83]. Horizontal gene transfer from prokaryotic microbes such as bacteria to nematodes is known [84]. Cyst nematode cellulases are similar to those found in bacteria, and theories of horizontal gene transfer to ancestor cyst nematodes have been proposed [26, 85]. This phenomenon has also been observed in Meloidogynespecies EST data and the existence of associations to parasitism [86]. Two Meloidogynegenes of rhizobial origin encode l-threonine aldolase (mi01644) and a protein of unknown function (mi00109). Sympatric organisms share the same soil ecological niche and both Meloidogyneand rhizobia fall into this class [87]. Another group of parasitism genes characterized in the reniform nematode genome are the C-type lectins (CTLs) [76, 77, 88]. The C-type lectins (CTLs) are a family of Ca2+-dependent carbohydrate-binding proteins with roles in innate immune response. A 5′- and 3′-RACE analysis was used in the identification of 11 reniform nematode CTL transcripts (RR-CTL-1RR-CTL-11), and these ranged from 1083 to 1194 bp with 93–99% sequence identity with the other [88]. Multiple sequence alignment of cDNA and genomic sequences showed three intronic regions. Specific BLAST hits were to Heligmosomoides polygyrusand H. glycines. The genes RR-CTL-1, RR-CTL-2, and RR-CTL-3 expressions were constant in the life cycle of the reniform nematode. The Rr-ctl transcripts were not constant in the various juvenile stages and were 839-fold higher in sedentary female nematodes compared to any other juvenile stage. A previous expresses sequence tag (EST) study [89] revealed C-type lectin domain peptides. These groups of peptides are carbohydrate-binding proteins, and therefore, calcium is a requirement for their effective functioning [90] and it is found in metazoans [91]. The CTLgenes have been characterized in some animal–parasitic nematodes, Heligmosomoides polygyrus, Toxocara canis, and Nippostrongylus brasiliensis[92, 93]. These CTLs have also been identified in the subventral glands of M. graminicola[94]. Characterization of the reniform nematode genome and transcriptome through 454 sequencing showed hits to BM1_02750(Accession # XP_001892052) and CLEC-180(Accession #NP_501229.2) in Brugia malayiand Caenorhabditis elegans,respectively [77].

5.2. Parasitism proteins

Parasitism proteins released from the cell of nematodes induce changes in host cell physiology through specific signals in the nucleus within the host cells [95]. The regulation of the host’s cells by nematodes occurs during transcription and direct interaction between nematode-secreted protein and plant-protein target has been observed. In the root knot nematode (RKN), a 13-aa secretory peptide 16D10 has interactions with two SCARECROW-like transcription factors [96].

The root knot and cyst nematodes both secrete chorismate mutase (CM) which affects the cellular shikimic acid pathway [71]. An overexpression of this protein in M. javanicain plant roots influences indole acetic acid (IAA) secretion which may result in improper tissue development [97]. Other proteins (14-3-3, sxp-ral-2, and ranbpm-like family proteins) influence cell-cycle, calcium binding, and defense regulation mechanisms in plants [71]. The 14-3-3 proteins have also been identified in reniform ESTs and these had homologies to 14-3-3 sequences of C. briggsae(Accession # XP_002643936.1) and C. elegans(Accession# NP_509939.1) [76]. Within the reniform, nematode genomic sequence hits to 14-3-3 sequences were observed in Bursaphelenchus xylophilus(Accession# ACZ13351), Ancylostoma caninum(Accession # ACO59962), and M. incognita(Accession# AAR85527) [77].

Expansin-like proteins in the potato cyst nematode (G. rostochiensis) can imitate some of their host genes in their function. An example is the hg-syv46 parasitism gene with a C-terminus having a similar function to CLAVATA3/ESR (CLE), a conserved domain in Arabidopsis thaliana[98, 99]. Three reniform nematode genes that code for putative CLE motifs (rr-cle-1, rr-cle-2, and rr-cle-3) have been isolated [100]. These peptides have an amino-terminal signal peptide with specific roles in secretion and pose a C-terminal CLE motif which can be associated with that of Heteroderaspp. The parasitism gene (16D10) is a conserved gene found in root knot nematode species controlling signaling events in RKN and its host associations. A double-stranded RNA (dsRNA) within the genomes of nematodes produces RNA interference (RNAi) of the targeted transcript. The technique of RNAi was first studied in C. elegans[101] and has been applied in gene silencing of variety organisms [102]. There are in vitroassays that facilitate stimulation of parasitic J2 nematodes to intake dsRNA from solutions through the nematodes stylet for RNAi induction [96, 103]. Specific nematode parasitism genes that are targeted could be easily knocked out and their functions effectively studied at the molecular level through RNAi approach [103].

5.3. Host-pathogen interactions

Plant host-pathogen interactions have fascinated plant pathologists over hundreds of years [104]. Contemporary studies on interactions began with seminal work on the gene-for-gene concept in plant host-pathogen interactions [105]. The gene-for-gene concept states that for every host R (resistance) gene, a corresponding Avr (avirulence) gene exists; thus, a successful host-defensive response requires a successful interaction [105]. The ranbpm-like family proteins and chorismate mutase (CM) interact with plant R genes, indicating nematode parasitism proteins act as avirulence genes [106, 107].

Proteins made by animal parasitic nematodes have been found in some plant parasitic nematode genomes and they are conserved [108]. These proteins function to challenge the host’s immune system invoking specific responses [109]. The last two decades have advanced the global approach to studying gene expression. We now have advanced recombinant DNA technologies to study gene expression at the mRNA or total RNA (transcriptome) and protein (proteome) levels. Approaches employed in the study of plant response to nematode parasitism include differential display [110, 111], promoter-reporter gene fusions [112114], RNA blotting, protein immunolocalization, in situhybridization [79, 115], and differential library screening [116]. Identification of genes playing useful roles in parasitism have been achieved through development of cDNA libraries from esophageal gland cells of the soybean cyst nematode [117, 118]. Other techniques used in the study of gene expression in plants at a single time point during early stages of infection of nematodes include oligonucleotide and cDNA microarrays [119, 120]; for example, in soybean cyst nematode, 1358 cDNAs from the esophageal glands were identified in expression analysis [121].

Recently, RNA-Seq analysis has been used in transcriptome sequencing of cotton (G. hirsutumL.) genotypes to measure comparative transcript abundance in reniform nematode susceptible (’DP90’ & ’SG747’), resistant (’BARBREN-713’), and hypersensitive (’LONREN-1’) genotypes of cotton (G. hirsutumL.) with and without reniform nematode infestation [122]. Several resistance genes that encode proteins known to be tightly linked to pathogen perception and resistance, for example, LRR-like and NBS-LRR domain-containing proteins were identified.

Most gene expression is a regulated process with genes being active in some situations and inactive in others. Gene expression relates to the physical signals from the environment and developmental cues of the organism in question. The rate of protein synthesis was once thought to be proportional to the concentration of mRNA; therefore, gene expression regulation depends on the regulation of the steady-state concentration of mRNAs [123]. However, mRNA levels and protein concentrations only partially correlate, a finding based on thoroughly composed reference datasets accounting for factors where ribosome occupancy and density and open-reading frame (ORF)-specific translation elongation rates were considered [124]. Therefore, many regulatory mechanisms involved with gene expression operate at many levels. The mechanisms influence on alterations in DNA structure, modification of transcription, stability, or translation of mRNA, or alterations in protein activity through post-translational modification.

Plants display varying levels of resistance to most pathogens in their environment, often being able to recognize pathogens through specifically distinct methods of detection [125]. A series of mechanisms of defense have been developed by plants, some of which can be constitutive or inducible. Resistance in plants is defined as the inability of a pathogen to propagate and spread on a host plant, usually involving a response referred to as the hypersensitive response (HR) [125]. The lines between a plant and its pathogens are truly battlegrounds where there are deployments of defense. The most current description of the action that takes place applies also to cotton-reniform interactions. Inducible defense responses in this type of interaction follow the ‘zigzag’ model [126].

According to the model suppression of immune-associated macroscopic programmed cell death (PCD) triggered by MAPK cascades or by the ETI cognate elicitors R3a/Avr3a occurs in susceptible hosts [126, 127]. The study showed that nematodes injected into the plant, thereby suppress PAMP-triggered immunity (PTI) or effector-triggered immunity (ETI) that are associated with the activation of PCD. Therefore, resistance is the ability of the host plant to evade the suppression of PCD. In other words, the host is resistant when its cells are able to undergo PCD and susceptible when the cells cannot undergo PCD.

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6. Conclusion

There has been much progress and success made by researchers in introgression of reniform nematode resistance into Upland cotton from distant and close relatives. However, introgressed genotypes, solely developed to withstand reniform nematode parasitism, may have low crop yields in fields where mixed-populations of nematodes occur. The challenge now is to develop germplasm lines that may be able to withstand more than two nematode types in the same field. The rapid advances in genomics, transcriptomics, and proteomic analysis provide huge datasets, from which several resistance genes to the reniform nematode and other nematodes have been identified. These genes could be further explored and transferred into Upland cotton with various trials initiated in specific cotton-producing localities, for this overarching goal to be achieved.

References

  1. 1. Smith AL. Distribution and relation of meadow nematode,Pratylenchus praunsis, toFusariumwilt of cotton in Georgia. Phytopathology. 1940; 30:710. doi: 10.12691/wjar-1-4-1.
  2. 2. Lawrence GW. Managing the reniform nematodes with nematicides. In: Proceedings of the Beltwide Cotton Conference, 6–10 January 1999; Orlando, FL. p. 100–101.
  3. 3. Lawrence GW, McLean KS. The reniform nematode. In: Kirkpatrick TL, Rothrock CS, editors. Compendium of Cotton Diseases. APS Press, St. Paul, MN; 2001. p. 42–44.
  4. 4. Blasingame D. Cotton disease loss estimate committee report. In: Proceedings of the Beltwide Cotton Conference, 3–6 January 2006; San Antonio, TX. p. 155–157.
  5. 5. Robinson AF, Percival, AE. Resistance toMeloidogyne incognitarace 3 andRotylenchulus reniformisin wild accessions ofGossypium hirsutum and G. barbadensefrom Mexico. Journal of Nematology. 1997; 29:746–755.
  6. 6. Linford MB, Oliveira JM.Rotylenchulus reniformisnov. gen., n. sp., a nematode parasite of roots. In: Proceedings of the Helminthological Society of Washington; 1940. p. 35–42.
  7. 7. Riddle DL, Bird AF. Responses of the plant parasitic nematodesRotylenchulus reniformis,Anguina agrostis, andMeloidogyne javanicato chemical attractants. Parasitology. 1985; 91:185–195. DOI: 10.1017/S0031182000056626.
  8. 8. Yik CP, Birchfield W.Gossypiumgermplasm resistant to the reniform nematode. Journal of Nematology. 1981; 13:465. (Abstr.).
  9. 9. Khan RM, Khan MW. Histopathology of cauliflower roots infected with reniform nematode. Pakistan Journal of Nematology. 1985; 3:83–85. (Abstr.).
  10. 10. Rebois RV, Madden PA, Eldridge BJ. Some ultrastructural changes induced in resistant and susceptible soybean roots following infection byRotylenchulus reniformis. Journal of Nematology. 1975; 7:122–139.
  11. 11. Vovlas N, Ekanayake HMRK. Histological alterations induced byRotylenchulus reniformisalone or simultaneously withMeloidogyne incognitaon banana roots. Nematropica. 1985; 15:9–17.
  12. 12. Gaur HS, Perry RN. The biology of the plant parasitic nematodeRotylenchulus reniformis. Agricultural Zoology Reviews.1991; 4:177–212.
  13. 13. Sivakumar CV, Seshadri AR. Life history of the reniform nematodeRotylenchulus reniformisLinford and Oliveira, 1940. Indian journal of Nematology. 1971; 1:7–20.
  14. 14. Radewald JD, Takeshita G. Desiccation studies on five species of plant-parasitic nematodes of Hawaii. Phytopathology. 1964; 54: 903–904.
  15. 15. Birchfield W. Host-parasite relations ofRotylenchulus reniformisonreniformisonGossypium hirsutum, Phytopathology. 1962; 52:862–865.
  16. 16. Mai WF. Plant-parasitic nematodes: their threat to agriculture. In: Sasser J, Carter CC, editors. An Advanced Treatise onMeloidogyne. North Carolina State University Graphics, Raleigh, USA;1985. 1:11–17.
  17. 17. Sasser JN. Root-knot nematode: a global menace to crop production. Plant Disease. 1980; 64:36–41.
  18. 18. Hussey RS. Host-parasite relationships and associated physiological changes. In: Sasser J, Carter CC, editors. An Advanced Treatise onMeloidogyne. 1985; 1:143–153.
  19. 19. Jones MGK. Early stages of nematode-induced giant-cell formation in roots ofImpatiens balsamina. Journal of Nematology. 1978; 10:70–84.
  20. 20. Sijmons PC, Atkinson HJ, Wyss U. Parasitic strategies of root nematodes and associated host cell responses. Annual Review of Phytopathology. 1994; 32:235–259. doi: 10.1146/annurev.py.32.090194.001315.
  21. 21. Hussey RS, Davis EL, Baum TJ. Secrets in secretions: genes that control nematode parasitism in plants. Brazilian Journal of Plant Physiology. 2002; 14:183–194. doi: 10.1590/S1677-04202002000300002.
  22. 22. Gheysen G, Fenoll C. Gene expression in nematode feeding sites. Annual Review of Phytopathology. 2002; 40: 191–219. doi: 10.1146/annurev.phyto.40.121201.093719.
  23. 23. Williamson VM, Gleason, CA. Plant-nematode interactions. Current Opinion Plant Biology. 2003; 6:327–333. doi: 10.1016/S1369-5266(03)00059-1.
  24. 24. Bird DM. Manipulation of host gene expression by root-knot nematodes. Journal of Parasitology. 1996; 82:881–888. doi: 10.2307/3284193.
  25. 25. Davis EL, Mitchum MG. Nematodes: sophisticated parasites of legumes. Plant Physiology. 2005; 137:1182–1188. doi: 10.1104/pp.104.054973.
  26. 26. Davis EL, Hussey RS, Baum TJ. Getting to the roots of parasitism by nematodes. Trends in Parasitology. 2004; 20:134–141. doi: 10.1016/j.pt.2004.01.005.
  27. 27. Cnavar. Cotton varieties planted in 2015 [{internet]. 2016. Available from: https://www.ams.usda.gov/mnreports/cnavar [Accessed: 2016-05-20].
  28. 28. Agudelo PR, Robbins T, Kim KS, Stewart J McD. Histological changes inGossypium hirsutumassociated with reduced reproduction ofRotylenchulus reniformis. Journal of Nematology.2005; 37:185–189.
  29. 29. Agudelo P, Robbins RT, Stewart JM, Szalanski AL. Intra-specific variability ofRotylenchulus reniformisfrom cotton-growing regions in the United States. Journal of Nematology. 2005; 37(1):105–114.
  30. 30. Tilahun Y, Soliman K, Lawerence KS, Cseke LJ, Ochieng JW. Nuclear ribosomal DNA diversity of a cotton pest (Rotylenchulus reniformis) in the United States. African Journal of Biotechnology. 2008; 7(18):3217–3224.
  31. 31. Nyaku ST, Sripathi VR, Kantety RV, Gu YQ, Lawrence KS, Sharma GC. Characterization of the two intra-individual sequence variants in the 18S rRNA gene in the plant parasitic nematode,Rotylenchulus reniformis. PLoS ONE. 2013; 8(4): e60891. doi: 10.1371/journal.pone.0060891.
  32. 32. Nyaku ST, Kantety RV, Tilahun Y, Lawrence KS, Soliman KM, Cebert E, Sharma GC. 18S and ITS1 genomic sequence variations inRotylenchulus reniformisisolates in Alabama. Journal of Cotton Science. 2013; 17:184–194.
  33. 33. Deng D, Zipf A, Tilahun Y, Sharma GC, Jenkins J, Lawrence K. An improved method for the extraction of nematodes using iodixanol (OptiPrep™). African Journal of Microbiology Research. 2008; 2:167–170.
  34. 34. van Den Berg E, Palomares-Rius JE, Vovlas N, Tiedt LR, Castillo P, Subbotin SA. Morphological and molecular characterization of one new and several known species of the reniform nematode,RotylenchulusLinford & Oliveira, 1940 (Hoplolaimidae: Rotylenchulinae), and a phylogeny of the genus. Nematology. 2016; 18:67–107. doi: 10.1163/15685411-00002945.
  35. 35. Sikkens, RB, Weaver DB, Lawrence KS, Moore SR, van Santen E. LONREN upland cotton germplasm response toRotylenchulus reniformisinoculum level. Nematropica. 2011; 41:68–74.
  36. 36. Bhandari B, Myers GO, Indest MO, Overstreet C. Response of five resistant cotton genotypes to isolates ofRotylenchulus reniformiscollected from reniform nematode infested fields of Louisiana. Nematropica. 2015; 45:252–262.
  37. 37. Koenning SR, Walters SA, Barker KR. Impact of soil texture on the reproductive and damage potentials ofRotylenchulus reniformisandMeloidogyne incognitaon cotton. Journal of Nematology. 1996; 28:527–536.
  38. 38. Sturhan D. Biological races. In: Zuckerman BM, Mai WF, Rohde RA, editors. Plant parasitic nematodes volume II. Academic Press, New York, NY; 1971. p. 51–71.
  39. 39. Leach M, Agudelo P. Genetic variability ofRotylenchulus reniformis. Plant Disease. 2012; 96:30–36. doi: 10.1094/PDIS-02-11-0132.
  40. 40. Arias RS, Stetina SR, Tonos JL, Scheffler JA, Scheffler BE. Microsatellites reveal genetic diversity inRotylenchulus reniformispopulations. Journal of Nematology. 2009; 41:146–156.
  41. 41. Barker KR. Resistance/tolerance and related concepts/terminology in plant nematology. Plant Disease. 1993; 77:111–113.
  42. 42. Das S, De Mason DA, Ehlers JD, Close TJ, Roberts PA. Histological characterization of root-knot nematode resistance in cowpea and its relation to reactive oxygen species modulation. Journal of Experimental Botany. 2008; 59:1305–1313.doi: 10.1093/jxb/ern036.
  43. 43. Schrimsher DW, Lawrence KS, Sikkens RB, Weaver DB. Nematicides enhance growth and yield ofRotylenchulus reniformisresistant cotton genotypes. Journal of Nematology. 2014; 46(4):365–75.
  44. 44. Kohel RJ, Lewis CF, editors. Cotton. Agronomy 24. American Society of Agronomy, Madison, WI; 1984. 605 p.
  45. 45. Percival AE, Wendel JF, Stewart JM. Taxonomy and germplasm resources. In: Smith CW, Cothren JT, editors. Cotton: Origin, History, Technology, and Production. John Wiley & Sons, Inc., New York, NY; 1999. p. 33–63.
  46. 46. Jones JE, Beasley JP, Dickson JL, Caldwell WD. Registration of four cotton germplasm lines with resistance to reniform and root-knot nematodes. Crop Science. 1988; 28:199–200.doi: 10.2135/cropsci1988.199b.rgp.
  47. 47. Cook, CG, Robinson AF, Namken, LN. Tolerance toRotylenchulus reniformisand resistance toMeloidogyne incognitarace 3 in high yielding breeding lines of upland cotton. Journal of Nematology. 1997; 29:322–328.
  48. 48. Usery SR, Lawrence KS, Lawrence GW, Burmester CH. Evaluation of cotton cultivars and tolerance toRotylenchulusreniformis. Nematropica. 2005; 35: 121–133.
  49. 49. Weaver DB, Lawrence K, van Santen E. Reniform nematode resistance in upland cotton germplasm. Crop Science. 2007; 47:19–24.doi:10.2135/cropsci2006.02.0130.
  50. 50. Yik CP, Birchfield W. Resistant germplasm inGossypiumspecies and related plants toRotylenchulus reniformis. Journal of Nematology. 1984; 16:146–153.
  51. 51. Robinson AF, Percival AE. Resistance toMeloidogyne incognitarace 3 andRotylenchulus reniformisin wild accessions ofGossypium hirsutumandG. barbadensefrom Mexico. Journal of Nematology. 1997; 29:746-755.
  52. 52. McCarty JC, Jenkins JN, Wubben MJ, Hayes RW, LaFoe II. Registration of three germplasm lines of cotton derived from L. accession T2468 with moderate resistance to the reniform nematode. Journal of Plant Registrations. 2012; 6:85–87. doi: 10.3198/jpr2011.07.0381crg.
  53. 53. Robinson AF. Reniform in U.S. cotton: when, where, why, and some remedies. Annual Review of Phytopathology. 2007; 45:263–288. doi: 10.1146/annurev.phyto.45.011107.143949.
  54. 54. Starr JL, Koenning SR, Kirkpatrick TL, Robinson AF, Roberts PA, Nichols RL. The future of nematode management in cotton. Journal of Nematology. 2007; 39:283–294.
  55. 55. Ehou MK. Attempt at utilization of the wild diploid speciesG. longicalyxfor the improvement of the cultivated tetraploid speciesG. hirsutumby the method of addition and substitution lines. [thesis] (In French.) Univ. Pierre et Mari Curie, Paris; 1983.
  56. 56. Sacks EJ, Robinson AF. Introgression of resistance to reniform nematode (Rotylenchulus reniformis) into upland cotton (Gossypium hirsutum) fromG. Arboreumand aG. hirsutum/G. aridumbridging line. Field Crops Res. 2009; 112:1–6. doi: 10.1016/j.fcr.2009.01.006.
  57. 57. Romano GB, Sacks EJ, Stetina SR, Robinson FA, Fang DD, Gutiérrez OA, Scheffler JA. Identification and genomic location of a reniform nematode (Rotylenchulus reniformis) resistance locus (Renari) introgressed fromGossypium ariduminto upland cotton (G. hirsutum). Theoretical Applied Genetics. 2009; 120:139–150. doi: 10.1007/s00122-009-1165-4.
  58. 58. Starr JL, Smith CW, Ripple K, Zhou E, Nichols RL, Faske TR. Registration of TAM RKRNR-9 and TAM RKRNR-12 germplasm lines of upland cotton resistant to reniform and root-knot nematodes. Journal of Plant Registrations. 2011; 5:393–396.doi: 10.3198/jpr2011.01.0048crg.
  59. 59. McCarty JC, Jenkins JN, Wubben MJ, Gutiérrez OA, Hayes RW, Callahan FE, Deng D. Registration of three germplasm lines of cotton derived fromGossypium barbadenseL. accession GB713 with resistance to the reniform nematode. Journal of Plant Registrations. 2013; 7:220–223. doi: 10.3198/jpr2012.08.0024crg.
  60. 60. Bell AA, Robinson FA, Quintana J, Duke SE, Starr JL, Stelly DM, Zheng X, Prom S, Saladino V, Gutiérrez OA, Stetina RS, Nichols RL. Registration of BARBREN-713 germplasm line of upland cotton resistant to reniform and root-knot nematodes. Journal of Plant Registrations. 2015; 9:89–93. doi: 10.3198/jpr2014.04.0021crg.
  61. 61. Robinson AF, Bell AA, Dighe ND, Menz MA, Nichols RL, Stelly DM. Introgression of resistance to nematodeRotylenchulus reniformisinto upland cotton (Gossypium hirsutum) fromGossypium longicalyx. Crop Science. 2007; 47:1865–1877. doi: 10.2135/cropsci2006.12.0776.
  62. 62. Bell AA, Robinson AF, Quintana J, Dighe ND, Menz MA, Stelly DM, Zheng X, Jones JE, Overstreet C, Burris E, Cantrell R, Nichols RL. Registration of LONREN-1 and LONREN-2 germplasm lines of upland cotton resistant to reniform nematode. Journal of Plant Registrations. 2014; 8:187–190. doi: 10.3198/jpr2013.11.0069crg.
  63. 63. Sikkens RB, Weaver DB, Lawrence KS, Moore SR, van Santen E. LONREN upland cotton germplasm response toRotylenchulus reniformisinoculum level. Nematropica. 2011;41:68–74.
  64. 64. Weaver DB, Sikkens RB, Lawrence KS, Sürmelioğlu Ç, van Santen E, Nichols RL.RENlonand its effects on agronomic and fiber quality traits in upland cotton. Crop Science. 2013; 53:913–920. doi: 10.2135/cropsci2012.08.0504.
  65. 65. Dighe N, Robinson AF, Bell A, Menz M, Cantrell R, Stelly D. Linkage mapping of resistance to reniform nematode in cotton (Gossypium hirsutumL.) following introgression fromG. longicalyx(Hutch. & Lee). Crop Science. 2009; 49:1151–1164. doi: 10.2135/cropsci2008.03.0129.
  66. 66. Avila CA, Stewart J McD, Robbins RT. Introgression of reniform nematode resistance into upland cotton. In: Proceedings of the Beltwide Cotton Conference; January 3–6, 2006; San Antonio, TX; p. 154.
  67. 67. Konan NO, Baudoin JP, D–Hont A, Mergeai G. Creating a tri-species hybrid cotton resistant to reniform nematode. Cashiers Agriculture. 2007; 16(1), 37–39.
  68. 68. Robinson AF, Bridges AC, Percival AE. New sources of resistance to the reniform (Rotylenchulus reniformis) and root-knot (Meloidogyne incognita) nematode in upland (Gossypium hirsutumL.) and sea island (G. barbadenseL.) cotton. Journal of Cotton Science. 2004; 8:191–197.
  69. 69. Gutiérrez OA, Robinson AF, Jenkins JN, McCarty JC, Wubben MJ, Callahan FE, Nichols RL. Identification of QTL regions and SSR markers associated with resistance to reniform nematode inGossypium barbadenseL. accession GB713. Theoretical and Applied Genetics. 2011; 122:271–280. doi: 10.1007/s00122-010-1442-2.
  70. 70. Luzzi BM, Boerma HR, Hussy RS. Resistance to three species of root-knot nematode in soybean. Crop Science. 1987; 27:258–262. doi: 10.2135/cropsci1987.0011183X002700020027x.
  71. 71. Mitchum MG, Hussey RS, Davis EL, Baum TJ. Application of biotechnology to understand pathogenesis of nematode plant pathogens. In: Punja ZK, De Boer S, Sanfacon H, editors. Biotechnology & Plant Disease Management. CABI International; Nosworthy Way, Walliford, Oxfordshire, UK; 2007. p. 58–86. doi: 10.1079/9781845932886.0058.
  72. 72. Smant G, Yan Y,Stokkermans JWPG, de Boer JM, Davis EL, Pomp H, Zilverentant JF, Overmars HA, Helder J, Hussey RS, Schots A, Bakker J. Endogenous cellulases in animals: isolation of β-1,4-endoglucanase genes from two species of plant-parasitic cyst nematodes. Proceedings of the National Academy of Sciences. 1998; 95:4906–4911. doi: 10.1073/pnas.1118560109.
  73. 73. Yan Y, Smant G, Stokkermans J, Qin L, Helder J, Baum TJ, Schots A, Davis EL. Genomic organization of four β-1,4-endoglucanase genes in plant-parasitic cyst nematodes and its evolutionary implications. Gene. 1998; 220:61–70. doi: 10.1016/S0378-1119(98)00413-2.
  74. 74. Wang X, Meyers D, Yan Y, Baum TJ, Smant G, Hussey RS, Davis EL. In planta localization of a β-1,4-endoglucanase secreted byHeterodera glycines. Molecular Plant-Microbe Interactions. 1999; 12:64–67.doi: 10.1094/MPMI.1999.12.1.64.
  75. 75. Wubben MJ, Ganji S, Callahan FE. Identification and molecular characterization of a β-1,4-endoglucanase gene (Rr-eng-1) fromRotylenchulus reniformis. Journal of Nematology. 2010; 42:342–351.
  76. 76. Nyaku ST, Sripathi VR, Wiley G, Najar FZ, Cseke LJ, Sharma GC, Roe BA, Cseke SB, Moss E, Kantety RV. The expressed parasitism genes in the reniform nematode (Rotylenchulus reniformis). American Journal of Plant Science. 2013; 4:780–791. doi: 10.4236/ajps.2013.44097.
  77. 77. Nyaku ST, Sripathi VR, Kantety RV, Cseke SB, Buyarappu R, Gu YQ, Lawrence K, Senwo Z, Sripathi P, McEwan R, George P, Sharma GC. Characterization of the reniform nematode genome through shotgun Sequencing. Genome. 2014; 57(4):209–21. doi: 10.1139/gen-2014-0019.
  78. 78. Gao B, Allen R, Maier T, Davis EL, Baum TJ, Hussey RS. Identification of a new β-1,4-endoglucanase gene expressed in the subventral gland cells ofHeterodera glycines. Journal of Nematology. 2002; 33:12–15.
  79. 79. Goellner M, Wang XH, Davis EL. Endo-beta-1,4-glucanase expression in compatible plant-nematode interactions. Plant Cell. 2001; 13:2241–2255. doi: 10.1105/tpc.010219.
  80. 80. Uehara T, Kushida A, Momota Y. PCR-based cloning of two β-1,4-endoglucanases from the root-lesion nematodePratylenchus penetrans. Nematology. 2001; 3:335–341.
  81. 81. Rosso MN, Favery B, Piotte C, Arthaud L, De Boer JM, Hussey RS, Bakker J, Baum TJ, Abad P. Isolation of a cDNA encoding a beta-1,4- endoglucanase in the root-knot nematodeMeloidogyne incognitaand expression analysis during plant parasitism. Molecular Plant-Microbe Interactions. 1999; 12:585–591. doi: 10.1094/MPMI.1999.12.7.585.
  82. 82. de Meutter J, Vanhomle B, Bauw G, Tytgat T, Gheysen G. Preparation and sequencing of secreted proteins from the pharyngeal glands of the plant parasitic nematodeHeterodera schachtii. Molecular Plant Pathology. 2001; 2:297–301. doi: 10.1046/j.1464-6722.2001.00078.x.
  83. 83. de Boer JM, Yan Y, Bakker J, Davis EL, Baum TJ. In situ hybridization to messenger RNA ofHeterodera glycines. Journal of Nematology. 1998; 30:309–312.
  84. 84. Hotopp JCD, Clark ME, Oliveira DCSG, Foster JM, Fischer P, Torres MCM, Giebel JD, Kumar N, Ishmael N, Wang S, Ingram J, Nene RV, Shepard J, Tomkins J, Richards S, Spiro DJ, Ghedin E, Slatko BE, Tettelin H, Werren JH. Widespread lateral gene transfer from intracellular bacteria to multicellular eukaryotes. Science. 2007; 317:1753–1756. doi: 10.1126/science.1142490.
  85. 85. Keen NT, Roberts PA. Plant parasitic nematodes: digesting a page from the microbe book. Proceedings of the National Academy of Sciences. 1998; 95:4789–4790. doi: 10.1073/pnas.95.9.4789.
  86. 86. Scholl EH, Thorne JL, McCarter JP, Mck Bird D. Horizontally transferred genes in plant-parasitic nematodes: a high-throughput genomic approach. Genome Biology. 2003; 4:R39.1–R39.11. doi: 10.1186/gb-2003-4-6-r39).
  87. 87. Bird DM, Koltai H. Plant parasitic nematodes: habitats, hormones, and horizontally-acquired genes. Journal of Plant Growth Regulation. 2000; 19:183–194. doi: 10.1007/s003440000022.
  88. 88. Ganji S, Jenkins JN, Wubben MJ. Molecular characterization of the reniform nematode C-type lectin gene family reveals a likely role in mitigating environmental stresses during plant parasitism. Gene. 2014; 537:269–278. doi: 10.1016/j.gene.
  89. 89. Wubben MJ, Callahan FE, Scheffler BS. Transcript analysis of parasitic females of the sedentary semi-endoparasitic nematodeRotylenchulus reniformis. Molecular and Biochemical Parasitology. 2010; 172:31–40. doi: 10.1016/j.molbiopara. 2010.03.011.
  90. 90. Zelensky AN, Gready JE. The C-type lectin-like domain superfamily. FEBS Journal. 2005; 272(24):6179–217. doi: 10.1111/j.1742-4658.2005.05031.x.
  91. 91. Drickamer K. C-type lectin domains. Current Opinion in Structural Biology. 1999; 9:585–590. doi: 10.1016/S0959-440X(99)00009-3.
  92. 92. Harcus Y, Nicoll G, Murray J, Filbey K, Gomez-Escobar N, Maizels RM. C-type lectins from the nematode parasitesHeligmosomoides polygyrusandNippostrongylus brasiliensis. Parasitology International. 2009; 58:461–470. doi: 10.1016/j.parint. 2009.08.011.
  93. 93. Loukas AC, Mullin NP, Tetteh KKA, Moens L, Maizels RM. A novel C-type lectin secreted by a tissue-dwelling parasitic nematode. Current Biology. 1999; 9: 825–828. doi:10.1016/S0960-9822(99)80366-2.
  94. 94. Haegeman A, Bauters L, Kyndt T, Rahman MM, Gheysen G. Identification of candidate effector genes in the transcriptome of the rice root knot nematodeMeloidogyne graminicola. Molecular Plant Pathology. 2013; 14(4):379–390. doi: 10.1111/mpp.12014.
  95. 95. Elling AA, Davis EL, Hussey RS, Baum TJ. Active uptake of cyst nematode parasitism proteins into the plant cell nucleus. International Journal of Parasitology. 2007; 37:1269–1279. doi:10.1016/j.ijpara.2007.03.012.
  96. 96. Huang G, Dong R, Allen R, Davis EL, Baum TJ, Hussey RS. A root-knot nematode secretory peptide functions as a ligand for a plant transcription factor. Molecular Plant–Microbe Interactions. 2006; 19:463–470. doi: 10.1094/MPMI-19-0463.
  97. 97. Doyle EA, Lambert KN. 3.Meloidogyne javanicachorismate mutase 1 alters plant cell development. Molecular Plant Microbe–Interactions. 2003; 6:123–131. doi: 10.1094/MPMI.2003.16.2.123.
  98. 98. Olsen AN, Skriver K. Ligand mimicry? A plant-parasitic nematode polypeptide with similarity to CLAVATA3. Trends Plant Science. 2003; 8:55–57. doi: 10.1016/S1360-1385(03)00003-7.
  99. 99. Wang X, Mitchum MG, Gao BL, Li C, Diab H, Baum TJ, Hussey RS, Davis EL. A parasitism gene from a plant-parasitic nematode with function similar to CLAVATA3/ESR (CLE) ofArabidopsis thaliana. Molecular Plant Pathology, 2005; 6:187–191. doi: 10.1111/J.1364-3703.2005.00270.x.
  100. 100. Wubben MJ, Gavilano L, Baum TJ, Davis EL. Sequence and spatiotemporal expression analysis of CLE-Motif containing genes from the reniform nematode (Rotylenchulus reniformisLinford& Oliveira). Journal of Nematology. 2015; 47(2):159–165.
  101. 101. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. Potent and specific genetic interference by double-stranded RNA inCaenorhabditis elegans. Nature. 1998; 391:806–811. doi: 10.1038/35888.
  102. 102. Hannon GJ. RNA interference. Nature. 2002; 418:244-251. doi: 10.1038/418244a.
  103. 103. Lilley CJ, Bakhetia M, Charlton WL, Urwin PE. Recent progress in the development of RNA interference for plant parasitic nematodes. Molecular Plant Pathology. 2007; 8:701–711. doi: 10.1111/j.1364-3703.2007.00422.x.
  104. 104. Ward HM. On the relations between host and parasite in the bromes and their brown rust,Puccinia dispersa(Erikss). Annals of Botany. 1902; 16:233–316.
  105. 105. Flor HH. Current status of the gene-for-gene concept. Annual Review of Phytopathology. 1971; 9:275–296. doi: 10.1146/annurev.py.09.090171.001423.
  106. 106. Moffett P, Sacco M. A nematode avirulence determinant and a co-factor for a disease resistance protein converge on the Ran signaling pathway. Phytopathology. 2006; 96:S80. doi: 10.1094/PHYTO.2006.96.6.S1.
  107. 107. Lambert KN, Bekal S, Domier LL, Niblack TL, Noel GR, Smyth CA. Selection ofHeterodera glycineschorismate mutase-1 alleles on nematode-resistant soybean. Molecular Plant-Microbe Interactions. 2005; 18:593–601. doi: 10.1094/MPMI-18-0593.
  108. 108. Baum TJ, Hussey RS, Davis EL. 2006. In: Setlow JK, editor. Root-knot and cyst nematode parasitism genes: the molecular basis of plant parasitism. Genetic Engineering, volume 28. p. 17–43. doi: 10.1007/978-0-387-34504-8_2.
  109. 109. Jasmer DP, Goverse A, Smant G. Parasitic nematode interactions with mammals and plants. Annual Review of Phytopathology. 2003; 41:245–270. doi: 10.1146/annurev.phyto.41.052102.104023.
  110. 110. Hermsmeier D, Hart JK, Byzova M, Rodermel SR, Baum TJ. Changes in mRNA abundance withinHeterodera schachtii-infected roots ofArabidopsis thaliana. Molecular Plant-Microbe Interactions. 2000; 13:309–315. doi: 10.1094/MPMI.2000.13.3.309.
  111. 111. Vercauteren I, Van Der Schueren E, Van Montagu M, Gheysen G. Arabidopsis thaliana genes expressed in the early compatible interaction with root-knot nematodes. Molecular Plant-Microbe Interactions. 2001; 14:288–299. doi: 10.1094/MPMI.2001.14.3.288.
  112. 112. Goverse A, Biesheuvel J, Wijers GJ, Gommers FJ, Bakker J, Schots A, Helder J.In plantamonitoring of the activity of two constitutive promoters, CaMV 35S and in developing feeding cells induced byGlobodera rostochiensisusing green fluorescent protein in combination with confocal laser scanning microscopy. Physiological and Molecular Plant Pathology. 1998; 52:275–284. doi: 10.1006/pmpp.1998.0154.
  113. 113. Mazarei M, Lennon KA, Puthoff DP, Rodermel SR, Baum TJ. Expression of anArabidopsisphosphoglycerate mutase homologue is localized to apical meristems, regulated by hormones, and induced by sedentary plant-parasitic nematodes. Plant Molecular Biology. 2003; 53:513–530. doi: 10.1023/B:PLAN.0000019062.80459.80.
  114. 114. Mazarei M, Lennon KA, Puthoff DP, Rodermel SR, Baum TJ. Homologous soybean and Arabidopsis genes share responsiveness to cyst nematode infection. Molecular Plant Pathology. 2004; 5:409–423. doi: 10.1111/j.1364-3703.2004.00241.x.
  115. 115. Niebel A, Engler JD, Tire C, Engler G, Van Montagu M, Gheysen G. Induction patterns of an extension gene in tobacco upon nematode infection. Plant Cell. 1993; 5:1697–1710. doi: 10.1105/tpc.5.12.1697.
  116. 116. Gurr SJ, McPherson MJ, Scollan C, Atkinson HJ, Bowles DJ. Gene expression in nematode-infected plant roots. Molecular and General Genetics. 1991; 226:361–366.
  117. 117. Gao B, Allen R, Maier T, Davis EL, Baum TJ, Hussey RS. The parasitome of the phytonematodeHeterodera glycines. Molecular Plant Microbe Interactions. 2003; 16:720–726. doi: 10.1094/MPMI.2003.16.8.720.
  118. 118. Wang X, Allen R, Ding X, Goellner M, Maier T, de Boer JM, Baum TJ, Hussey RS, Davis EL. Signal peptide-selection of cDNA cloned directly from the esophageal gland cells of the soybean cyst nematodeHeterodera glycines. Molecular Plant Microbe Interaction. 2001; 14:536–544. doi: 10.1094/MPMI.2001.14.4.536.
  119. 119. Puthoff DP, Nettleton D, Rodermel SR, Baum TJ. Arabidopsis gene expression changes during cyst nematode parasitism revealed by statistical analyses of microarray expression profiles. The Plant Journal. 2003; 33:911–921. doi: 10.1046/j.1365-313X.2003.01677.x.
  120. 120. Khan R, Alkharouf N, Beard H, MacDonald M, Chouikha I, Meyer S, Grefenstette J, Knap H, Matthews B. Microarray analysis of gene expression in soybean roots susceptible to the soybean cyst nematode two days post invasion. Journal of Nematology. 2004; 36:241–248.
  121. 121. De Boer JM, McDermott JP, Wang X, Maier T, Qui F, Hussey RS, Davis EL, Baum TJ. The use of DNA microarrays for the developmental expression analysis of cDNAs from the esophageal gland cell region ofHeterodera glycines. Molecular Plant Pathology. 2002; 3:261–270. doi: 10.1046/j.1364-3703.2002.00122.x.
  122. 122. Li R, Rashotte AM, Singh NK, Lawrence KS, Weaver DB, Locy RD. Transcriptome analysis of cotton (Gossypium hirsutumL.) genotypes that are susceptible, resistant, and hypersensitive to reniform nematode (Rotylenchulus reniformis). PLoS One. 2015; 10:1–27. doi: 10.1371/journal.pone.0143261.
  123. 123. D’haeseleer P, Liang S, Somogyi R. Genetic network inference: from co-expression clustering to reverse engineering. Bioinformatics. 2000; 16:707–726.
  124. 124. Brockmann R, Beyer A, Heinisch JJ, Wilhelm T. Posttranscriptional expression regulation: what determines translation rates? PLoS Computational Biology. 2007; 3:0531–0539. doi: 10.1371/journal.pcbi.0030057.
  125. 125. Odjakova M, Hadjiivanova C. The complexity of pathogen defense in plants. Bulgarian Journal of Plant Physiology. 2001; 27:101–109.
  126. 126. Jones JDG, Dangl JL. The plant immune system. Nature. 2006; 444:323–329. doi: 10.1038/nature05286.
  127. 127. Niu J, Liu P, Liu Q, Chen C, Guo Q, Yin J, Yang G, Jian H. Msp40 effector of root-knot nematode manipulates plant immunity to facilitate parasitism. Scientific Reports. 2016; 6:1–13. doi: 10.1038/srep19443.

Written By

Seloame T. Nyaku, Yonathan Tilahun, Kathy Lawrence, Venkateswara R. Sripathi, Abreeotta J. Williams and Govind C. Sharma

Submitted: December 8th, 2015 Reviewed: May 24th, 2016 Published: November 9th, 2016