Identification of a stylet-secreted effector protein family as a core component of root-knot nematode feeding tubes

Significance

Root-knot nematodes represent the most damaging group of plant-parasitic nematodes impacting global crop production. These obligate, sedentary endoparasites produce an essential structure, called a feeding tube, inside the giant feeding cells they induce in host roots to take up nutrients required to complete their life cycle. The feeding tube remains one of the best described, yet the least understood feature of sedentary endoparasitism. We developed a protocol to isolate feeding tubes for proteome analysis and identified a stylet-secreted effector protein family as a core component of feeding tubes that promises to shed light on their structure and function in nematode parasitism of plants.

Abstract

Proteins secreted from a mouth stylet of sedentary plant-parasitic root-knot nematodes self-polymerize to form a unique feeding tube structure within host cells modified into giant feeding cells by the nematode. Feeding tubes have essential functions as they complex with the host endomembrane system for nutrient uptake to sustain parasitism. Despite their significance, they remain one of the least understood aspects of nematode parasitism of plants. Their small size and location within giant-cells deeply embedded within galls encasing adult females has prohibited studies to isolate and discern their molecular composition. Here, we developed a protocol for the isolation and semipurification of root-knot nematode feeding tubes from giant-cell cytoplasm of several host plant species to provide a unique view of these structures at the light and scanning electron microscopy level revealing previously undescribed features of their structure. Our methods allowed for the isolation and solubilization of sufficient quantities of enriched feeding tubes enabling a comparative proteome analysis across host species that identified proteins with an increased likelihood to function in feeding tube formation. A comparison across root-knot nematode species further narrowed candidates to a conserved class of secretory proteins that specifically localized within secretory granules of the dorsal gland of adult females and in feeding tubes formed within host cell cytoplasm to unequivocally demonstrate these proteins as core components of feeding tubes. Our finding gives scientists a look into the protein composition of feeding tubes opening the door to a better understanding of their structure and function in nematode parasitism.

Root-knot nematodes (RKN, Meloidogyne species) are one of nature’s most successful parasites or pathogens, representing a global threat to food and fiber crop production (1). These obligate sedentary endoparasitic nematodes, which have an extensive host range including most agricultural crops grown resulting in billions of dollars in economic losses annually (2), have evolved a complex feeding relationship with their host plants. The RKN life cycle involves five developmental stages separated by four molts with the first molt occurring within an egg. Second-stage juveniles (J2) hatch and infect roots of host plants to initiate feeding while growing to a saccate shape and becoming sedentary. After an initial feeding period of several days the saccate J2 rapidly molt to non-feeding third (J3)- and fourth (J4)-stage juveniles without feeding stylets and then undergo a final molt to an adult female during which the stylet reforms and feeding resumes for development into mature pyriform egg-laying adult females (3).

The successful completion of the life cycle is dependent on the RKN’s ability to dramatically modify cells within the central vascular cylinder of host roots into permanent, specialized feeding cells called giant-cells, for nutrients essential for their development and reproduction. Indeed, giant-cell formation is one of the most sophisticated responses elicited in plant tissue by any parasite or pathogen. The giant-cell cluster typically consists of 5 to 8 parasitized root cells that have been transformed into hypertrophied multinucleate, metabolically hyperactive cells filled with dense granular cytoplasm, proliferated subcellular organelles and have elaborately modified cell walls to sustain the feeding of the adult female RKN (4–6).

RKN secrete through the hollow protrusible needle-like feeding stylet a diverse arsenal of effector proteins that are produced in three large specialized transcriptionally active secretory esophageal gland cells (two subventral and one dorsal) to infect roots and direct the formation and maintenance of the giant-cells, including modulating complex changes in cell morphology, function, and gene expression (7–9). While the majority of the RKN repertoire of effector proteins currently have unknown functions, known functions of a few effector proteins include suppressing host immunity, modifying plant cell walls, and reprogramming host cells by modulating gene transcription (10, 11).

In addition to secreting the effector proteins that dramatically modify recipient host root cells into giant-cells, RKN adult females also secrete one or more proteins that form a specialized tube-like structure called a feeding tube (FT) within the cytoplasm of the giant-cells (12, 13). These unique intracellular structures are used by the nematode for efficiently withdrawing nutrients from the cytoplasm of the giant-cells, and represent one of the most fascinating features of nematode parasitism of plants. A detailed ultrastructural study showed that the RKN FTs are approximately 1 µm wide and can extend at least 75 µm in length into the giant-cell cytoplasm (12). Electron micrographs of cross-sections of the FT revealed a crystalline-like composition of the FT wall that is 190 to 290 nm wide and surrounds a lumen with a diameter of 340 to 510 nm. The distal end of the FT is sealed with wall material, which prevents the FT lumen from having direct contact with the cytoplasm. RKN adult females insert their stylets only 2 to 3 µm through the wall of the giant-cell without piercing the plasma membrane and inject secretions into the cytoplasm through a pore in the plasma membrane at the stylet orifice. Therefore, the production of a FT, which functions as an extension of the nematode’s stylet, enables the adult female to have direct access to soluble assimilates throughout the metabolically hyperactive giant-cell cytoplasm over the entire length of the FT and transport them to the stylet orifice for ingestion. While the FT provides the nematode access to soluble assimilates in the cytoplasm, the rapid maximum dilation of the pump chamber in the metacorpus of the esophagus during the ingestion phase of the feeding cycle provides the suction for FT uptake of the soluble assimilates that are ingested by the nematode (14, 15).

Although the composition of the FT wall is unknown, previous evidence indicates that FTs are probably formed from stylet secretions originating in the dorsal esophageal gland that produces proteins for secretion from the stylet of the adult female RKN (9, 16). Remarkably, the wall of a newly formed FT immediately becomes enveloped in a compact uniform membrane system formed from rearrangement of part of the giant-cell endomembrane system (12). This membrane system can extend 450 to 500 nm outward from the FT wall into the cytoplasm of the giant-cell, with smooth endoplasmic reticulum contiguous to the FT wall while ribosomes are bound to tubular membrane elements at the periphery of the system. The intimate association of the membrane system with a newly formed FT indicates that it probably is required for the FT to function by synthesizing and/or serving as a channel in transporting nutrients in the cytoplasm of the giant-cell to the FT for ingestion by the adult female (12, 17). The adult female feeds in cycles from the different giant-cells radially arranged around its head, forming a new FT each time the female reinserts its stylet into a giant-cell to initiate a new feeding cycle, resulting in the accumulation of multiple FTs in the cytoplasm of each giant-cell (12, 13). Although the membrane system associated with the FT is degraded on the discarded FT, the crystalline FT walls persist free in the giant-cell cytoplasm for some time (12).

We reasoned that microisolation procedures enabling FT enrichment and solubilization could facilitate mass spectrometry (MS)-based proteomic analysis of these structures. We developed a method for semipurification of FTs from giant-cell cytoplasm of multiple host plant species enabling the identification of nematode proteins in FT-enriched samples. We further demonstrated the feasibility of our approach by identifying a stylet-secreted effector protein family abundantly produced in the dorsal gland of actively feeding adult RKN females and localized these proteins to FTs formed in giant-cell cytoplasm to significantly advance our knowledge of the composition of these unique structures.

Results

Feeding Tube Isolation.

FTs, being stable and insoluble, were isolated by expelling onto a cavity slide the dense cytoplasm from the giant-cells in stripped galled root sections sliced in half. The golden-tinted giant-cell cytoplasm of galled roots approximately 4 to 5 weeks postinoculation (wpi) (SI Appendix, Fig. S1A) had a syrup-like consistency that yielded the most FTs and contained granular cytoplasm clumps with large numbers of nuclei and FTs (SI Appendix, Fig. S1B). The number of FTs isolated varied but ranged from 2 to 26 per dissected gall. While over 1,100 FTs were isolated and their length measured, FTs < 50 µm, which appeared to be sections of FTs perhaps broken during the isolation process or in various stages of degradation, were not included in the analysis. The remaining FTs had 74% 50 to 99 µm in length and 26% ≥100 µm in length (Table 1, Fig. 1, and SI Appendix, Fig. S2). The longest FT for eggplant, tomato, tobacco was 148, 179, and 224 µm in length, respectively. Galls from tobacco roots consistently had the largest giant-cell clusters and yielded the greatest number of FTs ≥100 µm in length (39%). The walls of FTs considered intact and full-length (~35%) were always tapered toward a sealed distal end, which frequently was bent, hooked, or coiled (Fig. 1 and SI Appendix, Fig. S2). Numerous FTs were frequently embedded in clusters of nuclei of various sizes in the granular cytoplasm (Fig. 1D and SI Appendix, Fig. S2 C and D). The isolated nuclei were round and typically with multiple small nucleoli rather than being large highly lobed nuclei with a single large nucleolus as observed in intact giant-cells (Fig. 1D, SI Appendix, Fig. S3, and 12). FTs embedded in the clumps of granular cytoplasm and nuclear clusters could be exposed by treating with 0.1% sodium deoxycholate, which dissolved the cytoplasm and nuclear membranes but not the FTs (SI Appendix, Fig. S1C). FTs treated with the sodium deoxycholate, which can expand the FT wall, were not included in the FT measurements. A few FTs were coiled (Fig. 1A and SI Appendix, Fig. S2F). A FT lumen was sometimes discernible and a useful feature in identifying FTs when viewed with light microcopy. However, SEM provided a more detailed view of the FT surface and showed that the lumen had collapsed during FT isolation (Fig. 1 E and F). What appears to be the lumen with light microscopy was the indentation in the FT wall where the lumen had collapsed. The membrane system that envelopes newly formed FTs was not discernible with either light microscopy or SEM.

Table 1.

Summary of feeding tube lengths isolated from giant-cells induced by M. incognita on three host plants

Feeding Tubes	Eggplant	Tomato	Tobacco
50 to 99 μm (74%)	167 (91%)	278 (77%)	226 (61%)
≥100 μm (26%)	16 (9%)	81 (23%)	143 (39%)
Total	183	359	369
Full Length (35%)	76 (42%)	118 (32%)	129 (35%)
Longest FT	148 μm	179 μm	224 μm

Fig. 1.

M. incognita feeding tubes of varying lengths shown as microns (µm) isolated from giant-cells in galls on roots of tobacco (A, B), eggplant (C), and tobacco (D) 28 to 35 dpi. Nu = nucleus. Direct comparison of light microscopy (E) and SEM (F) images of a feeding tube isolated from a giant-cell in a gall from an eggplant root. Inset: Section of the feeding tube showing the collapsed lumen.

The cytoplasm collected from giant-cells fed on by late parasitic second-stage juveniles was not as dense and granular as the cytoplasm of giant-cells fed on by egg-laying adult females. In total, we examined giant-cell cytoplasm from 45 galls representing life stages spanning later parasitic second-stage juveniles through non-feeding molting adult females and never observed FTs (SI Appendix, Fig. S4 A and C), even after treating with 0.1% sodium deoxycholate. However, FTs were present in cytoplasm collected from giant-cells fed on by young adult females prior to laying eggs and their length varied depending on whether the female was only slightly swollen (FTs average 23 µm in length) or had an oval body (FTs average 63 µm in length). (SI Appendix, Fig. S4 B and D).

Electron Microscopy.

To gain additional insight into FT ultrastructure, we reanalyzed previous electron micrographs of cross-sections of FTs (12), and further describe the electron-dense, crystalline pattern of the wall, which appears to be formed from a patchwork of randomly arranged bands of striations consisting of alternating electron-dense stripes and electron-translucent channels. We measured the stripes to be 6.9 nm (6.4 to 8.0 nm) wide and the channels to be 7.9 nm (6.6 to 8.6 nm) wide (12; SI Appendix, Fig. S5 A and B). In some parts of the wall, the striations were oriented directly across the wall connecting the FT lumen with the giant-cell cytoplasm and the membrane system enveloping the FTs. The majority of the striations, however, did not completely traverse the wall. Thus, the wall appeared to have a uniform consistency formed by the ends of the striations and in longitudinal sections of FT walls where only a few of the striations were oriented to be visible, (SI Appendix, Fig. S5 B and C). This complex, matrix-like design of the FT wall suggests that it might consist of one or more proteins.

The FT wall is likely formed from secretory proteins produced in the dorsal esophageal gland cell of actively feeding adult females (9, 16). Within the dorsal gland cell, secretory proteins are packaged into secretory granules that are transported anteriorly via microtubules in the long cytoplasmic extension and accumulate in the ampulla, a collecting reservoir for secretory granules (16). The ampulla is connected to the esophageal lumen near the nematode stylet knobs via a short terminal duct that extends about 1 µm into the ampulla and forms an elaborate multibranched valve (SI Appendix, Fig. S5D and 16). The cuticularized distal ends of the branches of the valve are covered with an expandable membranous end-sac that during the secretory process becomes filled with secretory components released by exocytosis from secretory granules to be secreted from the nematode stylet (SI Appendix, Fig. S5 D and E and 14). Remarkably, a reexamination of previously unpublished electron micrographs revealed fine structure of some of the secretory components that had an electron-dense, striated crystalline pattern identical to that present in the FT wall (12), not only in the dilated membranous end-sac of an open dorsal gland valve as previously described (16), but also in the cuticularized esophageal lumen connecting the valve to the stylet (SI Appendix, Fig. S5 E, Inset). This observation suggests that preassembled FT wall-like material is pliable and can pass through the dorsal gland valve to be released from the stylet to form FTs in the cytoplasm of the giant-cell.

Establishment of the Feeding Tube Enrichment Method.

To obtain insight into the composition of FTs, a method was developed and optimized for their enrichment and solubilization for MS-based proteomics analysis. A schematic of sample collection and processing is illustrated in Fig. 2A. This involved a series of experiments to first determine the appropriate conditions for fractionation, enrichment, and solubilization of FTs, as well as to establish the amount of material necessary to achieve detection by LC-MS. The large size of most FTs (Table 1) enabled semipurification and concentration from microquantities of cytoplasm microaspirated from giant-cells by centrifugation and washing. Upon high-speed centrifugation of collected cytoplasm, nuclei and other organelles lysed, which allowed for the removal of the majority of their contents and other soluble cytoplasmic components by discarding the supernatant and following this with a wash step of the pelleted FTs and other insoluble material. A number of detergents were tested and it was ultimately determined that a low concentration of SDS (0.1% SDS) resulted in the complete solubilization of dried FTs within 15 s (SI Appendix, Fig. S6). An immediate radial and longitudinal expansion of the FT was observed likely due to protein denaturation leading to the breakage of intramolecular linkages providing the underlying structure to the FTs. Solubilization was followed by high-speed centrifugation to pellet any insoluble material and the supernatant containing FT-enriched proteins was polymerized in a tube gel to minimize sample loss and retain compatibility with MS analysis. The ability to enrich and solubilize FTs from giant-cell cytoplasm was essential for furthering our analyses to identify RKN proteins that may be core components of feeding tubes.

Fig. 2.

MS–based proteomic analysis of M. incognita FTs. Schematic representation of the workflow (A). A 4-wk-old tomato gall induced by M. incognita from which the cortex has been stripped away and the adult female removed intact to expose the giant-cells. Stripped root sections with the vascular cylinder enclosing the giant-cell complex were manually cut under a stereomicroscope and the cytoplasmic contents were removed by microaspiration using a glass Pasteur pipette pulled to a fine beveled tip. FTs were pelleted by high-speed centrifugation, washed, solubilized, and digested prior to MS and data analysis. Overview of proteome discoverer/SEQUEST-HT database search results (B). PSM = peptide spectral match.

MS-based Proteomics Analysis of Isolated Feeding Tubes.

To identify M. incognita gene(s) that encode the protein(s) that make up the FT, we conducted proteomics analysis on four FT-enriched samples spanning two host plants, including tobacco (Nicotiana tabacum) and tomato (Solanum lycopersicum). The same set of abundant proteins was consistently detected across biological replicates of the two host species (Dataset S1). Proteomic analysis of the samples resulted in a combined average of 8,503 high confidence proteins identified from a combined average of 41,874 peptide spectral matches (PSM) from a total of 526,701 MS/MS scans (Fig. 2B). The N. tabacum samples (MGM5, MGM6) averaged 11,239 proteins and 45,494 PSMs from a total of 280,053 MS/MS scans after excluding contaminants. The results stratified by organism, averaged 10,743 N. tabacum proteins and 489 M. incognita proteins. Those proteins were derived from an average of 45,050 N. tabacum PSMs and 1,061 M. incognita PSMs. The S. lycopersicum samples (MGM1, MGM7) averaged 5,767 proteins and 38,254 PSMs from a total of 246,648 MS/MS scans after excluding contaminants. The results stratified by organism, averaged 5,404 S. lycopersicum proteins and 357 M. incognita proteins. Those proteins were derived from an average of 37,464 S. lycopersicum PSMs and 1,072 M. incognita PSMs. Collectively, our method enabled the LC-MS identification of a list of RKN proteins present in FT-enriched samples isolated from giant-cell cytoplasm being actively fed upon by adult females.

Filtering Pipeline.

Intraspecies comparison identified high confidence PSMs to 406 and 184 M. incognita (Minc) proteins from tobacco and tomato, respectively (Fig. 3A). An interspecies comparison identified 160 Mincs that were shared between host plants. Of these, 29 Minc proteins were deemed to be M. incognita-specific and met a set threshold of >1 peptide detected (Fig. 3B). These 29 Minc proteins were further grouped into 13 protein families based on >50% amino acid (aa) sequence identity (Dataset S2). A cross-comparison of the 29 corresponding Minc gene sequences with our previously identified list of 83 Minc genes determined to be upregulated in the dorsal gland of M. incognita adult females (Fig. 3C; 18) found 11 candidate Mincs in common (Fig. 3D). Of the remaining 18 Minc genes, eight were determined to be paralogues of these 11 Mincs, which appear to have been misannotated in the genome and therefore filtered out of our earlier study (18) due to the lack of a signal peptide (Dataset S2). Together, these 19 Minc genes code for proteins spanning 7 protein families (Fig. 3E; paralogues not shown). These 7 protein families contained members with a predicted N-terminal secretion signal peptide, lacked transmembrane domains, and did not share homology to any other proteins in current protein databases. All but one of these genes, Minc3s00310g09909, was previously confirmed by in situ hybridization to be specifically expressed in the dorsal gland of actively feeding adult female RKNs (18). We reasoned that the expression of genes coding for proteins involved in FT formation would not only be exclusively expressed in the dorsal gland of adult females, but would also exhibit a high level of expression in this life stage relative to preparasitic second-stage juveniles (ppJ2), which are not feeding and therefore unlikely to be expressing high levels of genes coding for proteins contributing to FT formation. Thus, we extracted these data from life-stage specific RNAseq datasets for a comparison of the expression level of the identified candidates between ppJ2, female heads, and dorsal glands (DG) enriched from adult female heads (18). For all 11 candidates, few, if any reads were detected in ppJ2; however, read counts were high in adult female heads and exhibited even higher counts in the dorsal gland-enriched samples. Minc genes encoding proteins in families 3 and 7 exhibited the highest level of expression in the adult female dorsal gland-enriched samples (Fig. 3E). Proteins in these two families were also two of the most abundant proteins in FT-enriched samples based on the number of peptides and PSMs (Dataset S1).

Fig. 3.

Identification of FT-associated proteins through MS-based proteomics analysis. Proteins identified in FT-enriched samples from tobacco and tomato (A). Two biological replicates were conducted for each host plant species. Intraspecies comparisons identified 406 and 184 high confidence Minc proteins from tobacco and tomato, respectively. An interspecies comparison identified 160 high confidence proteins shared between host plants. These candidates were further filtered to 29 Minc proteins belonging to 13 protein families representing M. incognita-specific peptides for which >1 peptide was detected (B). A comparison between the 29 identified Minc genes and the 83 Minc genes determined to be upregulated in the adult RKN female dorsal gland transcriptome (C, 18) identified eleven Minc genes in common encoding proteins belonging to 7 protein families. Of the remaining 18 Minc genes, eight were determined to be paralogues of these 11 Mincs, which appear to have been misannotated in the genome and therefore filtered out of our earlier study (18) due to the lack of a signal peptide (D). Mean expression of the eleven Minc genes in J2, female heads, and enriched DG extracted from ref. 18 (E). Lists of Minc genes and proteins described in this figure can be found in Dataset S2.

Considering that FTs are required for RKN parasitism, we reasoned that candidate FT proteins should be highly conserved across RKN species. Therefore, we mined the existing genomes of eight Meloidogyne species for orthologues of the seven M. incognita protein families. Of these Minc protein families, only two (3 and 7) were conserved across all species. Family 3 included two members (Minc01309, Minc03994) (SI Appendix, Fig. S7 A–D and Dataset S2) whereas Family 7 included five members (Minc03784, Minc20083, Minc10218, Minc03997, Minc01312) (SI Appendix, Fig. S8 A–F and Dataset S2). Thus, by coupling FT enrichment methodology with a microproteomics profiling platform, we identified a short-list of adult RKN stylet-secreted proteins that represented high-confidence candidates with potential roles in FT structure and/or function (Dataset S2).

Localization of Selected Proteins to the Dorsal Gland of Adult Females and Feeding Tubes.

Protein family 7 grouped into two subfamilies based on sequence similarity and was one of the most highly expressed gene families in the dorsal gland of adult females (Fig. 3E; 18). The predicted proteins do not share homology with any proteins of known function from any other organism making them unique to RKNs. Subfamily 1 included Minc03784, Minc20083, and Minc10218, which shared 97% aa identity (SI Appendix, Fig. S8D). The Minc03784, Minc20083, and Minc10218 proteins are 676-amino-acids in length including a predicted 20-aa N-terminal SP and have a predicted molecular weight of ~75 kDa. Subfamily 2 included Minc03997 and Minc01312, which shared 97% aa identity (SI Appendix, Fig. S8E). The Minc03997 and Minc01312 proteins are 646- and 648-amino-acids in length, respectively, including a predicted 21-aa N-terminal SP and nuclear localization signal, and have a predicted molecular weight of ~72 kDa. A protein sequence alignment between Minc03784 and Minc03997, chosen as representative members of each of these two subfamilies, indicated that these two proteins only share 66% amino acid identity (SI Appendix, Fig. S8F). Minc03997 was expressed to a lesser extent than Minc03784 in the dorsal gland of the adult female (Fig. 3E; 18). A developmental expression time course of Minc03784 and Minc03997 revealed distinct expression patterns during the M. incognita life cycle (SI Appendix, Fig. S9). Though both genes were found to be significantly upregulated in adult females relative to egg and preparasitic second-stage juvenile stages (18), we show here that Minc03997 expression peaks in the earlier parasitic stages (7 to 14 days postinoculation, dpi) and declines in adult females (4 wpi) (SI Appendix, Fig. S9 A, C, E, and G), whereas Minc03784 expression is barely detectable in earlier parasitic stages and is strongly upregulated in adult females (SI Appendix, Fig. S9 B, D, F, and H).

For the aforementioned reasons, we sought to confirm whether the Family 7 proteins were localized in feeding tubes. For this, a recombinant N-terminal 6xHIS-tagged Minc03784 protein (minus the signal peptide sequence) was produced in Escherichia coli, affinity-purified over a nickel column and used to produce rabbit polyclonal antibodies (SI Appendix, Fig. S10 A–D). The polyclonal antisera produced was confirmed to recognize 1) the purified Minc03784 recombinant protein (SI Appendix, Fig. S10 C and D) and 2) a predominant protein of the same size in FT-enriched protein samples by SDS-PAGE and western blotting (SI Appendix, Fig. S10 E and F). No signal was detected on blots probed with the purified Rabbit IgG control antisera (SI Appendix, Fig. S10F). The polyclonal antiserum was then used to localize production of the Family 7 proteins within adult female specimens. The Minc03784 antibody labeled proteins in the secretory granules of the dorsal gland lobe, extension, and ampulla of adult RKN female specimens of M. incognita (Fig. 4 A–C). No fluorescence was detected in specimens probed with the purified Rabbit IgG control antisera (SI Appendix, Fig. S11 A–C). To determine if the proteins were in fact located in FTs, we carried out immunohistochemistry on isolated giant-cell cytoplasm containing FTs spread and dried on glass slides. No fixation of the FTs was necessary. The Minc03784 antibody specifically labeled the FTs in the cytoplasmic spreads (Fig. 4 D–F and SI Appendix, Fig. S12). No fluorescence was detected in feeding tubes on glass slides probed with purified Rabbit IgG control antisera (SI Appendix, Fig. S11 D–F). In addition, the Minc03784 antibody cross-reacted with protein produced in the dorsal esophageal gland of adult RKN female specimens of Meloidogyne arenaria (SI Appendix, Fig. S13 A and B) further confirming conservation of this effector protein family across species. Taken together, these results provided direct evidence of this family of stylet-secreted effector proteins as a core component of FTs.

Fig. 4.

Immunofluorescence of a M. incognita adult female head and a feeding tube isolated from giant-cell cytoplasm in galls from tobacco roots using anti-Minc03784 polyclonal antisera. An immunofluorescence image using anti-Minc03784 antibody on an excised head of an adult female nematode isolated 4 wpi. DG, dorsal gland; DGA, dorsal gland ampulla. [Scale bar, 50 µm (A–C).] Immunofluorescence images using anti-Minc03784 on a feeding tube isolated from giant-cell cytoplasm fed upon by an adult female at 4 wpi. Feeding tube length = 124 µm (D–F). Bright-field (differential interference contrast, DIC), fluorescence, and image overlays are shown.

In Silico Structural Prediction.

Protein family 7 also harbored several putative glycosylation sites (NXS/T) with 10 conserved cysteines spaced throughout their sequences that may influence protein folding, stability, solubility, and interactions (SI Appendix, Fig. S8F). The Minc03784 amino acid sequence was used to generate a predicted 3D structural model via AlphaFold 3 (19) (SI Appendix, Fig. S14), which indicates several folded domains with intervening disordered regions. Three potential disulfide bonds were identified in the structure. Models of the folded domains were compared with known structures in the Protein Data Bank to identify potential functional homologs, but no proteins with significant structural similarity were found.

Discussion

The structure and composition of the unique RKN FT is only now beginning to be resolved for a better understanding of RKN parasitism of plants. Collection of the dense golden-tinted cytoplasm from giant-cells fed on by adult RKN females from individual galls 28 to 35 dpi was very effective for isolating hundreds of FTs from multiple host species facilitating development of methodology for their enrichment and solubilization for MS-based proteomic analysis. Full-length FTs that had tapered sealed distal ends varied in size and were frequently considerably longer than previously observed in giant-cell sections (12). The greatest number of long FTs (≥100 µm) were isolated from giant-cells in galls on tobacco roots. Some isolated FTs were coiled and revealed the incredible flexibility of the FT wall. The coiling of FTs might occur when it is no longer being used by the adult female, since a coiled FT may affect the function of the FT as an extension of the stylet and thereby limit its access to the giant-cell cytoplasm. The structure of the crystalline FT wall with alternating electron-dense stripes (~7 nm wide) and electron-translucent channels (~8 nm wide) is striking and unique to RKN FTs. Although FTs are produced by other sedentary endoparasitic nematodes (e.g., Rotylenchulus, Heterodera, Globodera), their FTs have thin homogenous walls that are uneven along the lumen and are not enclosed by a compact membrane system (20, 21), unlike RKN FTs that are immediately enveloped by the membrane system (12). The unique association of the membrane system with newly formed and functional FTs indicates that the membrane system is critical for RKN FT function. Certainly, determining how and what triggers the rearrangement of the giant-cell endomembrane system to immediately envelop newly formed RKN FTs will significantly advance our understanding of FT function.

The dense syrup-like cytoplasm collected from the giant-cell cluster in single galls, in addition to yielding FTs, contained large numbers of round nuclei that varied in size and typically had multiple nucleoli. FTs were frequently observed embedded in clusters of nuclei of various sizes in the granular cytoplasm, but did not appear to be directly connected to nuclei. This observation is consistent with a recent array tomography study evaluating nuclear clustering architecture in giant-cells (22), which identified thick tube-like structures resembling FTs crossing and exiting a nuclear cluster without any connections. The nuclear clustering may have biological significance in giant-cells, as they appear to be anchored by the cytoskeleton and interconnected and likely communicate via nucleotubes, which are not visible using conventional light microscopy.

We also collected cytoplasm from giant-cells in young galls to precisely determine how early in the parasitic cycle FTs are formed. The cytoplasm collected from these giant-cells was less dense with fewer round nuclei, which were smaller than the nuclei present in the cytoplasm collected from older giant-cells. Interestingly, FTs were not observed in the cytoplasm collected from the giant-cells fed on by RKN parasitic second-stage juveniles, or from giant-cells from which we hand dissected non-feeding J3, J4, and molting adult females, indicating parasitic second-stage juveniles were able to obtain nutrients essential for their development from the cytoplasm without forming FTs. Consistent with these observations, in a microscopy study of giant-cells in tomato infected with M. incognita and Meloidogyne hapla, Paulson and Webster (23) described crystalline protein inclusion bodies that we now know were FTs as only being observed in the giant-cell cytoplasm after 10 dpi. In contrast, parasitic second-stage juveniles of the sedentary endoparasitic cyst nematodes, Heterodera species, form FTs by 1 to 2 dpi (21, 24). This indicates that FTs are necessary for adult RKN females to withdraw sufficient soluble assimilates from the cytosol of the giant-cells during the reproductive phase of their life cycle when the demand for nutrients would be greatest while they are producing hundreds of eggs. Furthermore, the production of a long FT enables the adult female to have direct access to nutrients throughout the cytoplasm when the giant-cells have greatly expanded in size.

The previously observed differences between RKN FTs and FTs of other sedentary endoparasitic nematodes, including the timing of their formation (late vs. early), wall structure (crystalline vs. irregular), and membrane association (enveloped vs. not enveloped), coupled with the fact that nematodes belonging to these different groups produce distinct tubes when infecting the same host plant, suggested FT wall components are of nematode origin (6, 25). Here, by coupling FT enrichment with MS-based proteomic analysis, we identified candidate FT proteins of nematode origin and used immunohistochemistry to demonstrate a family of stylet-secreted effector proteins as a core component of RKN FTs. This effector protein family is conserved across RKN species, but searches to the existing proteomes of other plant-parasitic nematodes known to produce FTs did not identify a similar family of proteins, again supporting the unique wall structure of RKN FTs. We demonstrated that members of this family of secretory proteins were produced in the dorsal gland of adult females and packaged via Golgi bodies into secretory granules that accumulated in the ampulla. These secretory proteins when transferred from the granules via exocytosis to the membranous end-sac of the dorsal gland valve may in fact polymerize into the small bands with a FT wall-like crystalline pattern that we observed in FT electron micrographs and pass through the ducts of the valve to enter the esophageal lumen for secretion from the stylet to form the FT. Future use of immunogold electron microscopy will be essential to specifically localize the effector proteins within the FT wall-like material observed within the dorsal gland end-sac. Nevertheless, how this material assembles into a perfectly round FT following secretion from the stylet remains a mystery, although it may require a component of the giant-cell cytoplasm as previously speculated (6, 26), possibly aided through interactions between FT proteins and components of the giant-cell endomembrane and cytoskeleton.

Future structural studies will shed more light on the mechanism underlying FT assembly, which could involve interactions among members of the same effector family and/or one or more other effector protein families. Indeed, we identified a second abundant dorsal gland produced effector protein family in our FT-enriched samples with conservation across Meloidogyne species that we are investigating further for a role in FT structure and function. Additionally, the candidate effector proteins identified in the FT-enriched sample not found to be highly conserved across RKN species, but shown to be expressed in the dorsal gland of adult females, may represent M. incognita stylet-secreted effectors with important roles in other aspects of giant-cell function pulled down during semipurification and enrichment of FTs from the giant-cell cytoplasm. Also Minc03997, which despite sharing 66% amino acid identity with Minc03784, contains a predicted nuclear localization signal, lacks a cis-acting DOrsal Gland promoter motif (“DOG box”) (27, 28), and as shown herein, exhibits a distinct expression profile during parasitism, presenting the intriguing possibility of functional diversification within this RKN effector protein family that warrants further exploration.

In summary, our short list of FT-associated proteins provides an exceptional exploratory tool that paves the way to a much clearer understanding of FT biology. Further, these findings open the door to an approach to disrupt RKN parasitism by interfering with the establishment of a structure essential for feeding as a means to engineer broad-spectrum RKN crop plant resistance that could help to reduce the billions of dollars in annual yield losses by these important agricultural pathogens.

Materials and Methods

Feeding Tube Isolation.

Meloidogyne incognita was cultured on greenhouse-grown Rutgers tomato (Lycopersicon esculentum Mill.), Black Beauty eggplant (Solanum melongena L.) and Xanthi SX tobacco (N. tabacum L.) and M. arenaria was cultured on greenhouse grown Black Beauty eggplants, which were inoculated with eggs (29). Since plants were inoculated with eggs and infection was not synchronized, giant-cell age was based on nematode stage of development and dpi (3). Root sections with individual galls with single egg masses were excised from infected plants 28 to 35 dpi and transferred to a Petri dish with water. Under a stereomicroscope, the egg sac, cortical tissue, and intact adult female RKN were carefully stripped from the galled root sections, leaving only the vascular cylinder enclosing the giant-cell complex. Root sections in which the cytoplasm of the giant-cells had a golden tint were selected for further processing (SI Appendix, Fig. S1A). The giant-cell complex from tobacco was elongated and larger than the giant-cell complex from tomato which was compact and round in shape and from eggplant which was small and elongated. The stripped root sections were transferred with a very small amount of water to the cavity on a microscope depression concave slide. The giant-cells in the vascular cylinder were cut directly in half with a double-edged razor blade. The cut ends of the vascular cylinder sections with the exposed giant-cells were squeezed with forceps to expel the dense giant-cell cytoplasm with the FTs in a drop of water in the cavity on a new slide. The samples were viewed with Nomarski differential-interference contrast objectives (10×, 20×) on an Olympus BH-2 Microscope (Olympus, USA) equipped with a Canon EOS M50 camera. Isolated FTs were photographed and their length (µm) was determined digitally with IMT iSolution Lite imaging software. For scanning electron microscopy (SEM), the cytoplasm from the giant-cells in the cut vascular cylinder root sections was expelled onto 12 mm diameter glass coverslips coated with 0.1% poly-L-Lysine. After the giant-cell cytoplasm air dried, the coverslips were transferred to 12 mm aluminum SEM stubs. The samples were then sputter coated (Structure Probe, Inc. West Chester PA) with approximately 15 nm of gold. Imaging was performed on a Teneo FE-SEM (Thermo Fisher Sci, Hillsboro, OR) at 5 kV with the secondary electron detector.

Cytoplasm was also collected from giant-cells in tomato and tobacco galls containing late parasitic second-stage juveniles through young adult females to determine how early in the RKN life cycle FT were formed. The cortex on the galls was stripped as described above and the cytoplasm was collected from giant-cell complexes. Specimens were staged according to ref. 3.

Electron Microscopy.

Electron micrographs were generated previously and reanalyzed in this study to reveal unique details of the FT wall and dorsal gland secretions. In brief, M. incognita-infected tissue was obtained from monoxenic cultures of Impatiens balsamifera L. cv. Tom Thumb (30) 28 to 35 dpi. Galled root tissue was fixed in 4% paraformaldehyde in 0.05 M phosphate buffer (pH 7.2) for 4 h at room temperature followed by 3% glutaraldehyde in the same buffer at 4 °C overnight and processed as previously described (12). Adult female RKN obtained from tomato 26 to 32 dpi were fixed 2 h at room temperature in 3% glutaraldehyde in 0.05 M phosphate buffer, pH 6.8, and processed as previously described (16).

Enrichment and Solubilization of Feeding Tubes.

Only stripped root sections from which intact female RKN were carefully removed from the galls were selected for proteomics analysis to avoid nematode contamination which would result from ruptured female nematodes. The cytoplasm from giant-cells in the cut vascular cylinder of 10 to 20 root sections was microaspirated using a glass Pasteur pipette pulled to a fine beveled tip and collected in an Eppendorf Lo-Bind 0.5 mL microcentrifuge tube. Samples were stored no more than 2 h at RT and then frozen at −80 °C until processed. The frozen giant-cell cytoplasm sample was thawed and 50 mM ammonium bicarbonate buffer (ABC; Sigma S-2454) was added to adjust the sample volume to 20 µL. The cytoplasm was centrifuged at 21,000 × g (15,000 rpm) for 5 min at RT to lyse organelles and pellet the FTs and any remaining insoluble material. The tube was rotated 180 degrees and centrifuged again at 21,000 × g (15,000 rpm) for 5 min at RT. The supernatant was carefully removed avoiding a very faint pellet in the bottom of the tube. The pellet was washed with 20 µL 50 mM ABC in a sonication bath for 30 s to disrupt the pellet and wash collected FTs. The sample was centrifuged at 21,000 × g (15,000 rpm) for 5 min at RT to pellet the FTs. The supernatant was again discarded avoiding the pellet. The tube was placed in a 37 °C incubator with the lid open to dry the pellet for ~30 min for efficient solubilization. Following drying, 10 µL of 0.1% SDS (Fisher BP1311)/50 mM ABC was added to the pellet. The tube was incubated for 5 min at RT with vortexing to solubilize FTs. The sample was centrifuged at 21,000 × g (15,000 rpm) for 5 min at RT to pellet any insoluble material. Approximately 10 µL of solubilized FT protein was transferred to a new Eppendorf Lo-Bind 0.5 mL microcentrifuge tube; total protein was estimated to be ~500 to 1,000 ng using the Micro BCA protein assay kit (Pierce).

The sample was adjusted to 20 µL with 50 mM ABC and reduced by addition of 3.33 µL of reducing agent stock (10 mM DTT Sigma D9779/5 mM TCEP Sigma C-4706 stock) followed by mixing and incubation for 20 min at 70 °C (or 5 min at 100 °C). For alkylation, 3.33 µL of alkylating agent stock (100 mM IAA, Sigma I-1149) was added. The sample was mixed well and incubated in the dark for 20 min at RT, followed by addition of 12 µL acrylamide/bisacrylamide (40%, v/v, 29:1, Sigma A-7802). The sample was mixed well and the gel was polymerized by addition of 1.67 µL 10% APS (Sigma A-3678) and 0.67 µL TEMED (Sigma T-7024) and incubated for 30 min at RT (12% gel).

UHPLC grade water suitable for MS (Sigma 900682) and acetonitrile (ACN; Sigma 900667) were used in all following steps. The tube gel was washed with 300 µL 20 mM ABC for 15 min. Excess liquid was removed. The tube gel was washed with 300 µL 20 mM ABC/ACN (1:1 v/v) for 15 min. If the tube gel was not turning white, the tip of a 200 uL pipette tip was heat sealed and used as a pestle to break the tube gel apart. Excess liquid was removed. To dehydrate the gel pieces, 100 µL 100% ACN was added to the pieces and incubated for 5 min and gel pieces turned white. Excess liquid was removed. The gel pieces were rehydrated in 300 µL 20 mM ABC for 10 min. Excess liquid was removed. The gel pieces were dehydrated again with 100 µL 100% ACN for 5 min. If gel pieces were not white, excess liquid was removed and 100 µL 100% ACN added and allowed to incubate for 5 min. Excess liquid was removed. The gel pieces were rehydrated in 300 µL 20 mM ABC for 10 min. Excess liquid was removed. The gel pieces were dehydrated with 200 µL ACN for at least 5 min. Excess liquid was removed. Gel pieces were dehydrated with 200 µL ACN for at least 5 min and turned white before removing the ACN. Excess acetonitrile was removed and gel pieces were dried in a speed vac for 25 min at RT. The lids were quickly closed and the tubes wrapped in parafilm and stored in a plastic bag with desiccant until shipped to Proteome Sciences (San Diego) for LC-MS analysis. The number of galls used for cytoplasm collection for the submitted samples were tobacco MGM5 (n = 12), MGM6 (n = 20), and tomato MGM1 (n = 20), MGM7 (n = 10).

LC-MS Analysis.

Digestion was performed using 2.5 ng/µL of trypsin by rehydrating the dried tube gel pieces in 70 µL of digestion buffer and incubating at 37 °C for 4 h. Peptides were extracted in two washes of equal volume of 25%:0.1% acetonitrile:formic acid and a final wash of 100% acetonitrile. Supernatants were pooled and lyophilized. Samples were resuspended and desalted as per manufacturer instructions for ZipTips, following which the samples were lyophilized and resuspended in 4 µL 5%:0.1% acetonitrile:formic acid for introduction to LC-MS. Samples were injected in technical duplicates on a 25-centimeter C18 pulled silica column heated to 50 °C with mobile phases consisting of (A) HPLC grade water with 0.1% formic acid and (B) 80% HPLC grade acetonitrile with 0.1% formic acid flowing at 350 nanoliters per minute with peptides eluted by increasing the concentration of mobile phase B linearly over 85 min. The mass spectrometer was operated in a data-dependent fashion, with initial high-resolution MS1 occurring in the orbitrap (120 K) and MS2 peptide fragmentation spectra collected in the ion trap at rapid acquisition speed.

All raw files were processed in Proteome Discoverer 3.0. Spectra were searched against the following databases: Tomato: S. lycopersicum OX4081 (downloaded from Uniprot 03/14/2024), Tobacco: N. tabacum 4097 (downloaded from Uniprot 04/25/2024), Nematode: M. incognita_PRJEB8714 (downloaded from www.parasite.wormbase.org on 3/14/2024), and a database of common MS contaminants, using SEQUEST-HT. Up to two missed tryptic cleavage sites were allowed during the searches. Precursor mass tolerance was set to 10 ppm and 0.6 Da was set for fragment tolerance. Carbamidomethylation of cysteine was set for static modifications, while methionine oxidation was set as dynamic modifications both at the peptide level, with acetylation of N termini set as dynamic modification for proteins. All peptide spectral matches were filtered with a strict 1% FDR for identification.

Protein Expression, Purification, and Antibody Production.

A codon-optimized Minc03784 gene sequence minus the predicted N-terminal secretion signal sequence was synthesized commercially (GenScript) in a pET30a expression vector with an N-terminal His₆-affinity tag. Proteins were recombinantly expressed in E. coli BL21 Star^TM (DE3). Cell cultures were grown in LB broth supplemented with 50 µg mL⁻¹ kanamycin and protein expression was induced with 0.5 mM IPTG. The protein was insoluble (inclusion bodies) whether cells were grown at 15 °C for 16 h or 37 °C for 4 h. The protein was solubilized and refolded in a buffer containing 50 mM Tris-HCl, 150 mM NaCl, 0.5 M L-Arginine, 1 mM TCEP, and 10% glycerol at pH 8.0 followed by affinity-purification to a purity of 80% in PBS, pH 7.2 using the NI-NTA method. Custom rabbit polyclonal antibodies (GenScript) were generated in New Zealand rabbits.

Sodium-Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunoblotting.

SDS-PAGE was performed using TGX Stain-free gels 4 to 15% (BioRad) and 1X running buffer (25 mM Tris, 193 mM glycine, 0.1% SDS, pH 8.3). Recombinant Minc03784 protein samples were mixed with 4× Laemmli sample buffer (1 M Tris–HCl, 1% Bromophenol Blue, 100% glycerol, pH 6.8). Electrophoresis was performed at 100 V constant for 2 h at room temperature. Protein was visualized directly on a BioRad Chemidoc after electrophoresis and transferred to nylon membrane using a Trans-Blot Turbo Transfer System (BioRad). Membranes were blocked overnight in 5% nonfat milk, incubated for 1 h in a 1:1,000 dilution of primary antibody [THE™ His Tag Antibody, mAb, Mouse (GenScript, A00186) or anti-Minc03784, pAb, Rabbit], washed five times with TBS-T (25 mM Tris, 0.15 M NaCl, 0.05% Tween-20), incubated 1 h in a 1:10,000 dilution of secondary goat anti-rabbit (or anti-mouse) IgG-HRP (Invitrogen 31462/31448), washed five times with TBS-T, and then incubated with Clarity Western ECL Substrate (BioRad) and imaged with a BioRad Chemidoc.

Immunofluorescence.

Giant-cell cytoplasm was microaspirated as described above and spread onto positively charged slides and allowed to dry at room temperature. Slides were washed twice for 15 min with 1× PBS (137 mM NaCl, 1.4 mM KH₂PO₄, 2.7 mM KCl, 4.3 mM Na₂HPO₄.7H₂0, pH 7.4) followed by incubation in 10% goat serum in 1× PBS amended with protease inhibitors (0.001 mM leupeptin, 1 mM Na2EDTA, 0.02 mM iodoacetamide, 0.02 mM phenylmethylsulfonyl fluoride, 0.001 mM pepstatin A) for 1 h at room temperature on a platform shaker at low speed. Slides were then incubated with a 1:50 dilution of either Minc03784 rabbit polyclonal antisera or purified Rabbit IgG control (GenScript A01008) in 10% goat serum amended with protease inhibitors for 1 h at room temperature. Following three washes for 5 min each with 1× PBS, slides were incubated in the dark at room temperature with a 1:500 dilution of goat anti-rabbit Alexa-488 secondary antibody (Invitrogen). Slides were washed three times for 5 min each with 1× PBS. Slowfade Diamond Antifade Mountant (Invitrogen) was used for mounting and specimens viewed with an Olympus BX53 epifluorescence microscope equipped with a Jenoptik Gryphax ARKTUR 8MP (4KHD) USB3 color camera with Gryphax software.

Four-week-old adult females were individually extracted from the roots and placed in a cavity slide containing 200 µL of molecular grade ethanol. Following dehydration, a needle was used to poke a hole in the perineal region and the nematode head was manually cut from the body with a scalpel. Pools of 20 heads were collected in siliconized 1.5 mL microcentrifuge tubes containing 2% paraformaldehyde in M9 buffer [Na₂HPO₄-7H₂0, KH₂PO₄, NaCl, MgSO₄-7H₂0] and incubated for 18 h at room temperature. After fixation, the heads were washed 2× with M9 buffer and stored at −80 °C until use. Upon processing, the heads were incubated in 0.5 mg/mL Proteinase K (Invitrogen) for 30 min on a nutator at room temperature. Samples were washed with 1 mL M9 buffer, then frozen on dry ice for 15 min. Heads were resuspended in 1 mL cold (−20 °C) methanol and placed on dry ice for 30 s. Heads were pelleted and suspended in cold (−20 °C) acetone and placed on dry ice for 1 min. Acetone was removed and heads were rehydrated by addition of dH₂0. Heads were washed with PBS-T (1× PBS, 0.5% Triton X-100) and incubated in 10% goat serum amended with protease inhibitors for 8 h on a nutator at room temperature. This was followed by overnight incubation in 1:300 dilution of primary antibody in 10% goat serum amended with protease inhibitors. Heads were washed three times for 5 min in 1 mL of PBS-T followed by incubation in 1:500 dilution of goat anti-rabbit IgG, IgM (H + L) secondary antibody, Alexa Fluor™ 488 in Tris-Saline-BSA (0.15 M NaCl, 0.01 M Tris, pH 7.2, 0.2% Triton X-100, 3% BSA) for 3 h at room temperature in the dark. Heads were washed twice with PBS-T and once with dH₂0 before being mounted on slides in a drop of anti-fade mountant and observed as above.

Isolation of RKN Life Stages.

M. incognita eggs were collected using the sodium hypochlorite method (29). Eggs were collected on a 25 µm (no. 500) sieve, sterilized in 0.02% sodium azide, and hatched on the antibiotic solution at 27 °C for 3 d to collect preparasitic second-stage juveniles. To isolate parasitic juveniles of M. incognita, 6-wk-old Xanthi SX tobacco (N. tabacum L.) plants were inoculated with 200,000 eggs. Infected roots were harvested at 7- and 14-dpi for the collection of mixed parasitic J2-J3/J4, and mixed parasitic J2-young molting adult females, respectively. The roots were homogenized for 30 s at medium speed in a kitchen blender and the slurry was washed through a nested stack of sieves with pore sizes of 250, 180, 150, and 25 µm. Nematodes were purified from the material collected from the 25 µm sieve by sucrose centrifugation. To further purify nematodes from small root debris, samples were passed through a 106 µm sieve onto a 25 µm (7 dpi) or through a 150 µm sieve onto a 106 µm sieve (14 dpi), pelleted, flash-frozen in liquid nitrogen, and stored at −80 °C until RNA extraction was performed. Adult females were hand dissected from galls at 4 wpi. Three sets of specimens of each life stage (eggs, ppJ2, 7dpi pJ2-J3/J3, 14dpi pJ2-YAF, and 4wpi AF) were independently harvested at different times for a total of three biological replicates.

Gene Expression Analysis.

Total RNA was extracted from frozen nematode pellets by grinding in a 200 µL glass mortar and pestle using a NucleoSpin miRNA Kit (Macherey-Nagel) following the manufacturer’s instructions. Total RNA was treated with Turbo DNase (Invitrogen) to remove contaminating genomic DNA following the manufacturer’s instructions. Total RNA was used to synthesize cDNA using a PrimeScript first-strand cDNA synthesis kit (Takara Bio, San Jose, CA) following the manufacturer’s instructions. Real-time qRT-PCR was conducted on a CFX96 Real-Time System (Bio-Rad, Hercules, CA) using PowerUp SYBR Green Master Mix (Applied Biosystems, Waltham, MA) with the following conditions: 50 °C for 2 min, followed by 95 °C for 2 min, then 40 cycles of 95 °C for 15 s and 60 °C for 1 min, and then a melt-curve analysis from 65 to 95 °C with a ramp rate of 0.5 °C/s. Conditions used to amplify full-length cDNA sequences were as follows: 94 °C for 2 min, then 35 cycles of 94 °C for 30 s, 58 °C for 30 s, and 72 °C for 2.5 min. Gene-specific primer sequences used in PCR analysis are listed in Dataset S3. Reactions were conducted in triplicate with a no-template control for each gene tested. The expression level of each gene tested was normalized to the M. incognita disulfide-isomerase (DISU) gene (31). Relative fold change was calculated using the 2-^ΔΔCt method (32). The experiment was conducted three times using cDNA synthesized from three independent sets of specimens.

Supplementary Material

Appendix 01 (PDF)

Dataset S01 (XLSX)

Dataset S02 (XLSX)

Dataset S03 (XLSX)

Data, Materials, and Software Availability

The MS proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (33) partner repository with the dataset identifier PXD065741. All other data are included in the manuscript and/or supporting information.