Since Poghossian (1966) established the genus Meloidoderita Poghossian, 1966 to accommodate the new species Meloidoderita kirjanovae Poghossian, 1966, two other Meloidoderita species have been described. Meloidoderita kirjanovae was isolated from roots of mint (Mentha longifolia (L.) Huds.) from the Mergi region in Armenia. Poghossian (1966) placed Meloidoderita within Heteroderidae Filipjev & Schuurmans Stekhoven, 1941 (Skarbilovich 1947) on the basis of cyst induction with a pattern of spine-like structures. Wouts and Sher (1971) considered Meloidoderita as genus inquirenda in the subfamily Heteroderinae Filipjev & Schuurmans Stekhoven, 1941. One year later Wouts (1972) reported that in the previous study, due to a lack of type material and an insufficient description, they “could not establish the exact status of the genus Meloidoderita”. Afterwards, after examining five females identified as Meloidoderita kirjanovae and on the basis of the presence of a large egg-sac (gelatinous matrix), short stylet, the absence of a cyst, and pronounced galls in the observed roots, Wouts (1972) considered Meloidoderita as a valid genus belonging in Meloidogynidae Skarbilovich, 1959 (Wouts 1973).

Kirjanova and Poghossian (1973) re-described Meloidoderita kirjanovae and established a newly erected family, Meloidoderitidae, within Criconematidea Taylor, 1936 (1914) (Thorne 1949). Moreover, Poghossian (1975) reported that the material examined by Wouts probably had been contaminated by Meloidogyne hapla Chitwood, 1949.

Meloidoderita kirjanovae has been recorded parasitizing on Mentha spp. (mint and water mint) and Utrica dioica L. (common nettle) (Poghossian 1966, Narbaev 1969, Cohn and Mordechi 1982, Vovlas et al. 2006).

Siddiqi (1985, 2000) classified Meloidoderita in the subfamily Meloidoderitinae Kirjanovae & Poghossian, 1973, family Sphaeronematidae (Raski & Sher, 1952) Geraert, 1966, superfamily Tylenchuloidea (Skarbilovich, 1974) Raski & Siddiqui, 1975 and suborder Criconematina Siddiqi, 1980.

The second species of Meloidoderita, Meloidoderita safrica, was described by Van den Berg and Spaull (1982) from soil and root samples of sugarcane (Saccharum hybrid) in South Africa.

Golden and Handoo (1984) described Meloidoderita polygoni from USA. Previously, Golden (1976) and Andrews et al. (1981) reported the occurrence of a population of Meloidoderita sp. from roots of smartweed (Polygonum hydropiperoides Michx.), which was not able to infect mint and nettle.

During a nematode survey conducted in Mont-Saint-Michel Bay in France, a Meloidoderita population was isolated from soil and roots of the halophyte Atriplex (= Halimione) portulacoides (L.) Aellen. This nematode was infecting roots of sea purslane (Atriplex portulacoides) growing in a muddy soil salt marsh region. Preliminary morphological and molecular analyses (G. Karssen, unpublished) indicated that the population differed from all three known described species of Meloidoderita and represented a new species. This was the first Meloidoderita species collected from a salt marsh environment.

The main objectives of the present study were to:i) describe a new species of Meloidoderita isolated from soil and roots samples of Atriplex portulacoides from a salt marsh region in France and provide a detailed morphological description based on LM and SEM; ii) characterize Meloidoderita species by means of small subunit rDNA sequencing; iii) determine the phylogenetic position of Meloidoderita within the suborder Criconematina.

Specimens for light microscopy (LM) were fixed in heated (70^°C) TAF (2 ml triethanolamine, 7ml formaldehyde and 91 ml distilled water (Courtney et al. 1955)), and processed to anhydrous glycerin following the method of Seinhorst (1966). Fixed specimens including second-stage juveniles, males, females, cystoids, egg-masses and eggs were mounted in a small drop of desiccated glycerin with the paraffin wax method on Cobb slides (Southey 1986).

Measurements and drawings were performed on a light microscope Olympus BH-2 equipped with Nomarski Differential Interference Contrast (DIC).

Specimens were drawn with a drawing tube, scanned and modified using Photoshop software version CS 5.1.

Light micrographs of specimens were taken with a Leica DC 300 F camera attached to a Zeiss Axio Imager M1 microscope. The original descriptions of closely related species (Table 1) were used for morphological and morphometrical comparison.

For SEM observation nematodes were fixed in 3% glutaraldehyde buffered with 0.05M phosphate buffer (pH 6.8) for 1.5 h and post-fixed with 2% osmium tetroxide for 2h at 22^°C. The specimens were dehydrated in a seven-graded ethanol series of 15-25-35-50-70-95 and 100% (Wergin 1981), critical point dried with carbon dioxide, and sputter coated with a layer of 4–5 nm Pt in a dedicated preparation chamber (CT 1500 HT, Oxford Instruments). The nematodes were examined and photographed with a field emission electron microscope Jeol 6300 F, at 5 kV (Karssen 1996, 1998).

Single nematodes (five individuals in total) were transferred to a 0.2 ml Eppendorf vial containing 25 µl of sterile water. An equal volume of lysis buffer containing 0.2 M NaCl, 0.2 M Tris-HCl (pH 8.0), 1% (vol/vol) β-mercaptoethanol, and 800 µg/ml of proteinase K was added. Lysis took place in a Thermomixer (Eppendorf, Hamburg, Germany) at 65°C and 750 rpm for 2 h followed by a 5 min incubation at 100°C (to inactive proteinase). Lysate was immediately used or stored at –20°C. SSU rDNA was amplified as two partially overlapping fragments using three universal and one nematode-specific primer (1912R). The latter was included to avoid amplification of non-target eukaryotic SSU rDNA. For the first fragments, either the primer 988F (5'-ctc aaa gat taa gcc atg c-3') or the primer 1096F (5'-ggt aat tct gga gct aat ac-3') was used in combination with the primer 1912R (5'-ttt acg gtc aga act agg g-3'). The second fragment was amplified with primers 1813F (5'-ctg cgt gag agg tga aat-3') and 2646R (5' -gct acc ttg tta cga ctt tt-3'). PCR was performed in a final volume of 25 µl containing 3 µl of 100 times-diluted crude DNA extract, 0.1 µM of each PCR primer and a ready-To-Go PCR bead (GE Healthcare, Little Chalfont, UK). The following PCR program was used: 94°C for 5 min; 5× (94°C, 30 s; 45°C, 30 s; 72°C, 70 s) followed by 35× (94°C, 30 s; 54°C, 30 s; 72°C, 70 s), and 72°C for 5 min. Gel-purified amplification products (Marligen, Ijamsville, MD) were cloned into a TOPO-TA vector (Invitrogen, Carlsbad, CA) and sent off for sequencing using standard procedures (Holterman et al. 2009). The newly generated SSU rDNA sequences were deposited at GenBank under accession numbers FJ969126 and FJ969127.

SSU rDNA-obtained sequences were aligned using the ClustalW algorithm as implemented in the program BioEdit 7.0.1 (Hall 1999). Manual improving and editing the alignment was then performed using arthropod secondary structure information (http://www.psb.ugent.be/rRNA/secmodel/index.html) according to Wuyts et al. (2000). Outgroup taxa and those nematodes compared with the sequence of the new Meloidoderita were chosen in accordance with Holterman et al. (2009). The final alignment included 39 SSU rDNA sequence and contained 1883 aligned position including gaps.

The phylogenetic tree was constructed using Bayesian inference (MrBayes 3.1.2 (Ronquist and Huelsenbech 2003)) and a fast maximum likelihood method (RAxML-VI- HPC v.4.0.0 (Stamatakis 2006)). Modeltest 3.06 (Posada and Crandall 1998) identified the general time reversible (GTR) model with invariable sites and a gamma-shaped distribution of substitution rates as the best substitution model. Bayesian analysis was performed with a random starting tree and four Markov chains. The programme was run for 5 × 10⁶ generations with a sampling frequency of 1, 000 generations. Two independent runs were performed for each analysis. After discarding the ‘burn-in’ samples of 500, 000 generations, sampled trees were combined to generate a 50% majority rule consensus tree, which represents posterior probabilities.

The second phylogenetic tree was constructed with a fast maximum likelihood method. The SSU rDNA alignment was analysed at a distant server (CIPRES, http://www.phylo.org) running the program, RAxML-VI-HPC v.4.0.0 using the same GTR model. One hundred bootstrap replicates were performed.

Meloidoderita salina sp. n. was described from a salt marsh area at Mont-Saint-Michel Bay in France, parasitizing the halophyte plant Atriplex portulacoides. On average, this area has a salinity of about 34–35g/L which usually increases after submersion by the tides. The presence of a well sclerotized S-E duct is a noticeable character, especially in adult males and matured females of Meloidoderita salina sp. n. which could be correlated with their saline environment and their halophytic host plant. The presence of a strongly sclerotized S-E duct has been also reported in the genus Halenchus N.A. Cobb in M.N. Cobb, 1933 as the only known marine Tylenchomorpha. The genus Halenchus with three species is exclusively marine parasitic nematode which produces galls on sea algae (Siddiqi 2000). The “widened and sclerotized excretory duct, exclusively marine, and parasitic on sea algae” are the key characters that have been applied by Siddiqi (2000) in support of the subfamily Halenchinae with its single genus Halenchus in Anguinidae Nicoll, 1935 (1926). Considering the sclerotization of S-E duct in both Meloidoderita salina sp. n. and Halenchus, more physiological studies will probably clarify the role of this structure in these genera.

Spiegel and Cohn (1985) and Vovlas et al. (2006) reported secretion of gelatinous matrix from the vulva slit in Meloidoderita kirjanovae. Vovlas et al. (2006) considered it as a discriminating character for differentiation between “Meloidoderita kirjanovae and that of other tylenchulids such as Tylenchulus and Trophonema which secret the gelatinous matrix from the secretory-excretory pore”. They discussed that “this physiological characteristic may confirm the result of phylogenetic analysis” as inferred by Subbotin et al. (2005, 2006) and Sturhan and Geraert (2005), who studied the phylogeny of Tylenchuloidea. Nevertheless, no evidence (e.g. the present of the vulval glands) was observed to support their opinion regarding formation of the gelatinous matrix. In Meloidoderita salina sp. n. the S-E pore is a well-developed structure connected to a markedly sclerotized duct running posteriorly. It is possible that this prominent structure could be also involved in the production of the gelatinous matrix.

Poghossian (1966) classified Meloidoderita under the family Heteroderidae. However, some years later Kirjanova and Poghossian (1973) established the new family Meloidoderitidae to accommodate Meloidoderita, and placed it within the superfamily Criconematoidea. Siddiqi (1985, 2000) proposed the new subfamily Meloidoderitinae to accommodate its single genus, namely Meloidoderita and the type species Meloidoderita kirjanovae, under the family Sphaeronematidae and the suborder Criconematina on the basis of “the lack of the neck; uterine walls form a protective cystoid body for eggs” (Siddiqi 2000).

Siddiqi (2000) described the genus Meloidoderita as mature females with a swollen body, without neck or tail, and males without bursa. Andrassy (2007) also described the Meloidoderita adult female as “without neck”. Regardless, Kirjanova and Poghossian (1973), Van den Berg and Spaull (1982), and Golden and Handoo (1984) who reported the presence of an irregularly shaped neck region modified by root tissue and influenced by the cellular root structures. We also observed in Meloidoderita salina sp. n. females a well-defined and twisted neck region (Figs 5, 8).

Siddiqi (2000) described the family Sphaeronematidae as “ectoparasite” in which the juveniles “attack and feed on roots ectoparasitically”. However, it was Siddiqi who wrote in 1985: “Meloidoderita kirjanovae is reported to be endoparasitic in Mentha longifolia roots, becoming secondarily exposed as the growing female ruptures the root epidermis”. Andrassy (2007) also defined the genus Meloidoderita as “ectoparasitic” nematodes. In addition to Cohn and Mordechai (1982) and Andrews et al. (1981) who reported Meloidoderita kirjanovae and Meloidoderita sp. respectively as semi-endoparasitic, Vovlas et al. (2006) recently reported, “Severe infections of Meloidoderita kirjanovae were detected on young roots of Mentha aquatica. Adult females of Meloidoderita kirjanovae protruded from the surface of all infected root segments occurring individually or in clusters, but did not cause distortion of the entire root diameter. Eggs were laid in a gelatinous matrix regularly protruding from the root surface but cystoid body was often located within the root cortex”. Andrews et al. (1981) reported that juveniles migrated intracellularly through the cortex. Further studies are needed to examine the biology, life-cycle and histopathology of Meloidoderita sp. and to clarify their parasitic behavior.

Cohen and Mordechai (1982), while studying the biology of Meloidoderita kirjanovae, observed several males attached to or enveloped by old second-stage juveniles cuticle. They reported that it “could obviously be identified as offspring of the particular female beneath the egg-mass, rather than having migrated from outside. Furthermore, often more than one molting cuticle was present at the same time, indicating that development of juveniles into adult males was a relatively short process and apparently did not necessitate feeding on the host tissues”. These enveloped males in second-stage juveniles cuticle have been reported by Van den Berg and Spaull (1982). In the present study these enveloped males were also described and we did not observed any J3 or J4 male stages.

In the classification scheme proposed by Siddiqi (2000) the suborder Criconematina was described as “phasmids absent”. Andrassy (2007) has also emphasized that “the absence of phasmids” is one of “the main distinguishing characteristics of this suborder”.

Recently Sturhan and Geraert (2005) assessed the presence of phasmids in Tylenchulidae. They observed phasmid-like structures in Sphaeronema, Meloidoderita, Tylenchulus, Trophotylenchulus. However, they did not found phasmids in examined species of Criconematidae, Hemicycliophora sp., Paratylenchus, Cacopaurus and Tylenchocriconema. Our observation (LM and SEM) confirmed the presence of phasmids in both juveniles and males of Meloidoderita salina sp. n.

Phylogenetic studies done by Subbotin (2005, 2006) Vovlas et al. (2006) Palomares-Ruis et al. (2010) and our phylogenetic analysis showed that Meloidoderita together with Sphaeronema form a clade and are placed as stem taxa at the base of the Criconematina phylogenetic trees. These morphological observations and molecular studies show that the lack of phasmids in other taxa of Criconematina could be considered as an apomorphic character (Sturhan and Geraert 2005). Hence, within Criconematina those taxa without phasmids could be probably defined by the autapomorphism of the absence of phasmids.

Based on the distribution of the type host Atriplex portulacoides in tidal salt marshes in France, it may be expected that Meloidoderita salina sp. n. is more widely distributed in West-European salt marshes. Sturhan and Geraert (2005) reported an unknown Meloidoderita sp. and also an undescribed Sphaeronema species isolated from Atriplex portulacoides, both from northern Germany. We suggest further sampling along the North Sea coast (France, Belgium, Germany and UK) to characterize the distribution of this species.

Human consumption is currently one of the most important aspects for cultivation of Atriplex spp. It has a salty taste when it is eaten raw or cooked, and is presently served in luxury restaurants. Atriplex portulacoides has an important role in primary production, and in the food web in salt marsh ecosystems (Bouchard et al. 1998, Neves et al. 2007, 2008). Atriplex spp.is also used for other agricultural and environmental aspects such as dune stabilization, land reclamation, or as livestock fodder and ornamental plant (Aronson 1986, Khan et al. 2000, Daoud et al. 2001). The effect of Meloidoderita salina sp. n. on the host plant Atriplex portulacoides is unknown and needs to be studied.

It is interesting to report that during this study we found a unique sub-cuticular hexagonal beaded pattern in the cystoids of Meloidoderita salina sp. n. This specific pattern can be seen on the surface of the cystoid and displays symmetrical hexagons (Figs 5H, I, 8D–F). This pattern reported in this study is probably the first to be observed among all the identified species of nematodes so far.