Characterization of Vittatidera zeaphila (Nematoda: Heteroderidae) from Indiana with molecular phylogenetic analysis of the genus

Abstract

In the summer of 2016, a field of corn (Zea mays) in Spencer County, Indiana was observed with heavily stunted plants, and from the
affected roots a large number of cysts were recovered. Soil samples
were submitted to one of us (JF), who extracted the nematode cysts
and sent them to the USDA-ARS, Mycology and Nematology Genetic Diversity and Biology Laboratory (MNGDBL), Beltsville, MD for
morphological and molecular identification. Cysts and the recovered
second-stage juveniles (J2) that were examined morphologically conformed to the measurements of Vittatidera zeaphila, the goose cyst
nematode originally described from Tennessee, USA in 2010. The
molecular analysis of J2 showed the sample from Spencer County
matched exactly with V. zeaphila according to ribosomal DNA markers ITS, 28S, and 18S, and with mitochondrial cytochrome oxidase
I (COI). The nuclear marker heat shock protein 90 (Hsp90) was also
analyzed for the first time from the Indiana population of V. zeaphila.
Similarities to existing cyst nematode sequences are reported herein.
Geographically, although the county is across the Ohio River from
Kentucky, the previously reported Hickman County, Kentucky location and Indiana detection are approximately 200 miles apart. To the
best of our knowledge, this is the first report of V. zeaphila in Indiana.

Full text

1
JOURNAL OF NEMATOLOGY
e2020-24 | Vol. 52
Article | DOI: 10.21307/jofnem-2020-024
© 2020 Authors. This is an Open Access article licensed under the Creative
Commons CC BY 4.0 license, https://creativecommons.org/licenses/by/4.0/
Characterization of
Vittatidera zeaphila
(Nematoda:
Heteroderidae) from Indiana with molecular phylogenetic
analysis of the genus
Andrea M. Skantar1,*,
Zafar A. Handoo1, Mihail R. Kantor1,
Lynn K. Carta1, Jamal Faghihi2 and
Virginia Ferris2
1Mycology and Nematology
Genetic Diversity and Biology
Laboratory, USDA, ARS,
BARC-West, Bldg. 010A, Rm. 111,
Beltsville, MD, 20705.
2Department of Entomology,
Purdue University, 901 West State
St. West Lafayette, IN, 47907-2089.
*E-mail: andrea.skantar@usda.gov
This paper was edited by
Erik J. Ragsdale.
Received for publication July 29,
2019.
Abstract
In the summer of 2016, a field of corn (Zea mays) in Spencer Coun-
ty, Indiana was observed with heavily stunted plants, and from the
affected roots a large number of cysts were recovered. Soil samples
were submitted to one of us (JF), who extracted the nematode cysts
and sent them to the USDA-ARS, Mycology and Nematology Ge-
netic Diversity and Biology Laboratory (MNGDBL), Beltsville, MD for
morphological and molecular identification. Cysts and the recovered
second-stage juveniles (J2) that were examined morphologically con-
formed to the measurements of Vittatidera zeaphila, the goose cyst
nematode originally described from Tennessee, USA in 2010. The
molecular analysis of J2 showed the sample from Spencer County
matched exactly with V. zeaphila according to ribosomal DNA mark-
ers ITS, 28S, and 18S, and with mitochondrial cytochrome oxidase
I (COI). The nuclear marker heat shock protein 90 (Hsp90) was also
analyzed for the first time from the Indiana population of V. zeaphila.
Similarities to existing cyst nematode sequences are reported herein.
Geographically, although the county is across the Ohio River from
Kentucky, the previously reported Hickman County, Kentucky loca-
tion and Indiana detection are approximately 200 miles apart. To the
best of our knowledge, this is the first report of V. zeaphila in Indiana.
Keywords
Cyst nematode, 18S rDNA, 28S rDNA, Taxonomy, Vittatidera zeaphila.
Globally, cyst nematodes species cause serious dam-
age to a wide variety of economically important crops.
The need for information on cyst-forming nematode
species has been instrumental in stimulating growth
of nematology worldwide. The cyst nematode group
currently contains eight genera, with a total of 121 valid
species (Handoo and Subbotin, 2018). The general
morphology and molecular taxonomy and phylogeny
of cyst nematodes have been given in detail in two
recent review articles (Baldwin and Handoo, 2018;
Subbotin and Skantar, 2018). Vittatidera zeaphila
(Bernard et al., 2010), the goose grass cyst nematode,
was first described from Obion County, Tennessee,
USA, parasitizing corn and goosegrass. Later, host
range studies and potential sources of resistance to
V. zeaphila were published by Donald et al. (2012).
In the summer of 2016, a field of corn (Zea mays) in
Spencer County, Indiana was observed with heavily
stunted plants, and from the affected roots a large
number of cysts were recovered. Soil samples
were submitted to one of us (JF), who extracted the
nematode cysts and sent them to the USDA-ARS,
Mycology and Nematology Genetic Diversity and
Biology Laboratory (MNGDBL), Beltsville, MD for
morphological and molecular identification. Cysts and
second-stage juveniles (J2) conformed morphologically
and morphometrically to V. zeaphila. We report here
the first occurrence of this species in Indiana, thus
representing the third state after Tennessee, and
Kentucky in the United States.

2
Vittatidera zeaphila
from Indiana:
Skantar et al.
Materials and methods
Various stages
Cysts, white females, and J2 were obtained from soil
and roots associated with corn plants from Spencer
County, Indiana. Juveniles were separated from soil
by sieving and Baermann funnel extraction or were
collected from cysts removed from fresh roots and
kept in water in watch glasses. Juveniles were fixed
in 3% formaldehyde and processed to glycerin with
a formalin glycerin method (Hooper, 1970; Golden,
1990). Females and some cysts were removed from
roots after fixation for 12 hr in 3% formaldehyde
solution.
Photomicrographs of cyst vulval cones, and J2
were made with an automatic 35-mm camera at-
tached to a compound microscope having an inter-
ference contrast system. Roots and whole cysts were
photographed under a dissecting microscope Nikon
SMZ18, and light microscopic images of fixed nem-
atodes were taken on a Nikon Eclipse Ni compound
microscope using a Nikon DS-Ri2 camera. Meas-
urements were made with an ocular micrometer on
a Leica WILD MPS48 Leitz DMRB compound micro-
scope. All measurements are in micrometers unless
otherwise stated.
Nematode DNA preparation
The molecular identification was performed using
DNA extracted from single nematodes as template
in PCR reactions. Single juveniles were mechanically
disrupted with sharp forceps tips in 20 µl nematode
extraction buffer (500 mM KCl, 100 mM Tris-Cl
(pH8.3), 15 mM MgCl2, 10 mM dithiothreitol (DTT),
4.5% Tween 20 and 0.1% gelatin) (Thomas et al.,
1997) and stored at −80°C until needed. To prepare
DNA extracts, frozen nematodes were thawed,
1 µl proteinase K (from 2 mg/ml stock solution) was
added, and the tubes were incubated at 60°C for
60 min, followed by 95°C for 15 min to deactivate the
proteinase K. Two or five microliters of extract were
used for each PCR reaction.
PCR amplication and cloning
ITS: Amplification of the internal transcribed spacer
region ITS1&2 rDNA contained 0.2 µM each primer,
TW81 (Joyce et al., 1994) and AB28 (Howlett et al.,
1992), 1.5 mM MgCl2, 0.2 mM dNTPs, 1U Platinum
Taq DNA polymerase (Invitrogen, Carlsbad, CA), 3 µl
nematode DNA extract, and supplied enzyme reaction
buffer in a total volume of 25 µl. Cycling included one
step of 95°C for 2 min, followed by 35 cycles of 95°C
for 30 sec, 55°C for 30 sec, and 72°C for 90 sec,
finished with one cycle at 72°C for 5 min (Skantar et
al., 2007).
28S: Amplification of the 28S large ribosomal sub-
unit (LSU) D2-D3 expansion segment included the
primers D2A [5′-ACAAGTACCGTGAGGGAA AGTT-3′]
and D3B [5′-TCGGAAGGAACCAGCTACTA-3′] and
were amplified as previously described (De Ley et al.,
2005; Ye et al., 2007).
18S: The 18S (small subunit: SSU) sequence was
amplified with primers in one fragment with forward
primer 18S-CL-F3: [5′-CT TGTCTCAAAGATTAAGC-
CATGCAT-3′] (Carta and Li, 2018) and reverse prim-
er 1912R according to Holterman et al. (2006). Re-
actions contained 2 µl nematode DNA extract, 0.5 μ l
10 μ M primers, 0.5 μ l 10 mM dNTP, 1U Platinum Taq
DNA polymerase (Invitrogen, Carlsbad, CA), 0.75 μ l
50 mM MgCl2, and 2.5 µl PCR buffer in a total vol-
ume of 25 µl. PCR cycling conditions were 95°C for
3 min, 35X (94°C for 30 sec, 50°C for 40 sec, 72°C for
70 sec), 72°C for 5 min, 4°C until finish.
COI: Mitochondrial cytochrome oxidase I (COI) was
amplified with primers Het-coxiF [5′-TAGT TGATCG-
TAAT TT TAATGG-3′] and Het-coxiR [5′-CCTAAAA-
CATA ATG AA AATGW G C - 3 ′]. Amplifications were per-
formed in 25 µl reactions with 1x PicoMaxx (Agilent)
buffer, 0.2 mM dNTPs, 0.3 µM each primer, 0.125 µl
Dream Taq DNA Polymerase (Thermo Fisher), 0.5 µl
PicoMaxx Taq, and 3 µl DNA extract. Cycling condi-
tions were as described previously (Subbotin, 2015).
Hsp90: Heat shock protein 90 (Hsp90) frag-
ments were amplified with degenerate primers U288
[5′-G AYAC VG GV AT YG GN ATG A CYA A -3 ′] and L1110
[5′-TCRCARTTVTCCATGATRAAVAC-3′] (Skantar and
Carta, 2004). Cycling was performed with 1× Pico-
Maxx reaction buffer, 0.2 mM dNTPs, 1.5 mM MgCl2,
0.3 µM each primer, 1.25 U PicoMaxx Taq, 1 U Plat-
inum Taq, and 3 µl nematode DNA extract. PCR cy-
cling conditions were 94°C 2 min, followed by 45 cy-
cles of [94°C 20 sec, 65°C 5 sec, 60°C 5 sec, 55°C
5 sec, 45°C 5 sec, 68°C 3 min], ending with 1 cycle of
68°C for 15 min.
PCR products were analyzed by electrophoresis
on 2% agarose with 1X SB (sodium borate-EDTA)
buffer. Gels were stained with ethidium bromide and
visualized using the U:Genius gel documentation
system (Syngene, Frederick, MD). Hsp90 fragments
were cloned using the Strataclone PCR Cloning
Kit (Agilent, Santa Clara, CA) according to
manufacturer’s instructions. Plasmid clone DNA was
prepared with the QiaPrep Spin Miniprep Kit (Qiagen,
Valencia, CA) and digested with Eco RI to verify the
presence of insert. Sequencing was performed by

3
JOURNAL OF NEMATOLOGY
Genewiz, Inc. Direct sequencing of PCR amplicons
was used to obtain the 28S sequences (assigned
GenBank accession numbers MK121965-MK121968),
the ITS 1&2 rDNA sequence (MK121952), the 18S
rDNA sequence (MK182465), and COI sequences
(MK253554-MK253558). Accession numbers were
assigned for new sequences from cloned Hsp90
of V. zeaphila (MK580824-MK580829), Heterodera
avenae (MH484608, MH484609, and MH484611), and
the outgroup Helicotylenchus digonicus (MK580830).
Phylogenetic inference and
tree visualization
Raw sequence reads were processed in Sequencher
5.4.6 (Genecodes, Inc., Ann Arbor, MI). Multiple DNA
sequence alignments were created using Geneious
Prime 2019.0.3 (www.geneious.com) with built-in
parameters or the MAFFT plug-in, with auto-selection
of best algorithm depending on data. Alignments of
Hsp90 intron regions were adjusted further using
Geneious Alignment with free end gaps and identity
(1.0/0.0) cost matrix and inspected visually to ensure
preservation of exon boundaries and reading frames.
Phylogenetic analysis using Bayesian inference
(Huelsenbeck and Ronquist, 2001) was performed
via the CIPRES Gateway (Miller et al., 2010), except
where noted otherwise. The model of nucleotide
evolution was determined by jModelTest 2.1.3
(Darriba et al., 2012), with the best-fit model GTR+I+G
selected for Hsp90. The parameters for base
frequency, proportion of invariable sites, and gamma
distribution shape, and substitution rates according
to Akaike Information Criteria (AIC) generated in
a custom command block implemented in MB.
Bayesian analysis was run with four chains for 2 × 106
generations, with Markov chains sampled at intervals
of 500 generations. Two runs were performed for
each analysis. After burn-in samples were discarded
and convergence evaluated, the remaining samples
were retained for further analysis. Topologies were
used to generate 50% majority rule consensus trees
with posterior probabilities given on appropriate
clades. Trees were visualized in Geneious. Alternative
alignments of Hsp90 genomic sequences or coding
regions were made using Clustal Omega (Sievers
et al., 2011) and MUSCLE (Edgar, 2004) plug-ins for
Geneious, and corresponding trees were made in
MB or IQ-TREE (Nguyen et al., 2015) to investigate the
effect of removing introns or partitioning the data on
tree topology and support values.
COI alignments were likewise made within
Geneious, using Atalodera carolynae as the outgroup.
Phylogenetic reconstruction based on COI alignments
were performed in IQ-TREE, within which ModelFinder
(Kalyaanamoorthy et al., 2017) determined the best-fit
model to be K3Pu + F + I + G4. The consensus tree was
constructed from 1,000 bootstrap trees using UFBoot2
(Hoang et al., 2018) with bootstrap values indicated on
appropriate branches. Alternative MB trees were made
for comparison, including runs where rate parameters
were independently specified for codon position.
Results and discussion
Description
Morphology and measurements
Photomicrographs of V. zeaphila from Spencer
County, Indiana are shown in Figure 1. Measurements
of second-stage juveniles (n = 10) included length of
body (range = 332-385 μ m, me an = 351 μ m). Lip region
rounded, slightly set off with three to four annules,
stylet short, well developed (15.0-16.0 μ m, 15.8 μ m)
with rounded knobs, lateral field with four distinct lines,
tail elongate conoid with narrowly rounded terminus
(37.0-48.0 μ m, 42.1 μ m), and hyaline tail terminus
(11-15 μ m, 13.1 μ m). Shapes of the tail, tail terminus,
and stylet knobs were consistent with V. zeaphila.
The cysts were lemon-shaped, dark to reddish light
brown in color; vulval cone was not protuberant, with
large egg masses (Fig. 1K); mature cysts had vulval
cone circumfenestrate, underbridge, bullae absent.
Morphometrics of cysts (n = 5) included body length (L)
including neck (520-866 μ m, 69 6 μ m); body width (W)
(320-495 μ m, 399.8 μ m; L/W (1.4-2.1, 1.7 μ m); neck
length (60-100 μ m, 74.0 μ m) and width (45-55 μ m,
50.0 μ m); fenestra length (52-65 μ m, 58.4 μ m) and
width (32-40 μ m, 37.4 μ m). The cysts had light to wavy
line type of cyst wall cuticular pattern (Fig. 1A,B); anus
opening was prominent; punctations often present in
terminal area of cyst; morphometrics of cysts were
also consistent with V. zeaphila. Lateral field distinct
in both females and cyst stages, arched, represented
by short transvers lines between neck and cone. In
female perineal region phasmid present.
Living nematode juveniles (J2) collected from
the cysts were examined morphologically and
molecularly for species identification. Observations
of morphological characters critical for identification
(Fig. 1C-J) indicated that the specimens agreed
with the previous Vittatidera zeaphila description by
Bernard et al. (2010), except for minor morphometric
difference in the head end to excretory pore distance
which is closer to 76.7 (70-80 μ m) vs 148 (142-161 μ m)
in the original description, and which according to
scale bar measurements should be 74 (71-80.5).

4
Vittatidera zeaphila
from Indiana:
Skantar et al.
Molecular characterization
Three sequences of ITS 1&2 rDNA obtained from
separate J2 were assembled into a 936 bp alignment
of identical sequences that overlapped with a
540 bp region from JF741961 previously described
from the TN population of V. zeaphila (Bernard et
al., 2010). Except for a few ambiguous positions in
the latter sequence (possible sequencing artifacts),
the ITS sequences from both populations were
identical. 28S rDNA sequences obtained from four
J2 were assembled into a 762 bp alignment along
with JF741960 previously described from the TN
population of V. zeaphila, differing at 0 to 2 bp from
each other and from the TN sequence. The 18S
rDNA sequence was obtained from amplification of
the gene from a single J2. A 985 bp alignment with
the TN V. zeaphila sequence (JF741962) showed that
the 18S sequences were identical. For Hsp90, six
sequences were obtained from cloning the amplicons
from a single J2. Sequence lengths were 1412 bp with
four introns. Excluding the degenerate primer regions
at each end, the sequences varied from 4 to 11 bp
among the clones, mostly within introns or at third
codon positions. No Hsp90 sequence was available
from the TN population for comparison.
Phylogenetic relationships among heteroderid
nematodes were inferred from analysis of 35 partial
Hsp90 sequences from 14 species, in a genomic
alignment of 1887 bp (Fig. 2). Vittatidera zeaphila
sequences had two introns (273 bp and 200 bp) that
were significantly longer than the corresponding
introns from other cyst nematodes in the alignment.
Vittatidera zeaphila formed a sister group to the
Figure 1: Photomicrographs of Vittatidera zeaphila population on Corn (Zea mays) from Spencer
County, Indiana. (A, B): light micrograph of vulval cones of V. zeaphila showing fenestra area with
arrow showing the anal area, (C, D): males anterior ends showing head and esophageal regions,
(E, F): second-stage juveniles anterior ends showing head and esophageal regions, (G, H):
second-stage juveniles posterior ends showing tail and tail terminus, (I): male posterior end
showing the spicule, (J): lateral field with four lines at mid-body, (K): cysts with attached egg
masses on corn roots with arrows showing second-stage juveniles feeding on corn roots.

5
JOURNAL OF NEMATOLOGY
Figure 2: Phylogenetic relationships of Vittatidera zeaphila and other selected cyst nematodes, as
inferred from a 1887 bp alignment of partial Hsp90 genomic DNA sequences, with
Helicotylenchus digonicus as the outgroup. A 50% majority rule consensus tree obtained from
Bayesian analysis was generated using the GTR + I + G model of nucleotide substitution. Branch
support values above 50% are shown on appropriate branches. New sequences are highlighted
in bold.
Globodera clade, although support was moderate
(0.82). Consistent with previously published 18S,
28S, and ITS trees (Bernard et al., 2010), the
Hsp90 tree placed V. zeaphila genetically distant
from the other cyst nematode of corn, Heterodera
zeae (Fig. 2). Cactodera cacti grouped apart from
the other Punctoderinae, appearing in a clade that
included the Goettingiana group, but with weak
support (0.63). To assess the effect of additional
taxa on topologies, another tree was constructed
from a shorter, 594 bp genomic alignment of 50
sequences; this tree resulted in weaker support for

6
Vittatidera zeaphila
from Indiana:
Skantar et al.
the position of V. zeaphila relative to the Globodera
clade and it did not further resolve the other clades
(not shown). This is most likely due to the shorter
alignment containing fewer informative characters.
Moreover, a third alignment that included only
coding regions with introns removed contained
essentially the same topology and branch support
as seen in Figure 2, even when codon positions
were modeled independently (not shown). In
previous studies with trees inferred from ITS rDNA,
V. zeaphila was not resolved in relation to the rest of
the Punctoderinae (Subbotin and Skantar, 2018), so
Figure 3: Phylogenetic relationships of Vittatidera zeaphila and other select cyst nematodes, as
inferred from a 373 bp alignment representing 33 taxa, with Atalodera carolynae as the outgroup.
The tree was constructed in IQ-TREE using model K3Pu + F + I + G4; the consensus tree was
constructed from 1,000 bootstrap trees using UFBoot2. Bootstrap values are included on
appropriate branches. New sequences are highlighted in bold.

7
JOURNAL OF NEMATOLOGY
the long Hsp90 fragment analyzed here performed
as good or better, considering the relative lack of
available sequences for analysis compared to ITS.
COI sequences obtained from five J2 were
assembled into a 456 bp alignment. All sequences
were identical to each other and were 100% match
to V. zeaphila from Tennessee (MK093060). BlastN
showed that the next closest sequence matches
had maximum similarity of only 82% to H. glycines
and other Heterodera species. In total, 46 selected
cyst nematode COI fragments were assembled in
a trimmed 373 bp alignment representing 33 taxa,
with the resulting IQ-TREE shown in Figure 3. In this
analysis, V. zeaphila appeared as a sister taxon to the
Cyperi group of Heteroderinae, with 97% support.
Taxa within the Avenae and Schachtii groups were
also strongly supported and agreed with trees based
upon ITS rDNA (Subbotin and Skantar, 2018). A
number of additional analyses using MB parameters
under the GTR+I+G model adjusted to include
more or fewer taxa left H. filipjevi and H. latipons in
unresolved positions relative to the rest of the Avenae
group, which disagreed with previous phylogenies
(Subbotin and Skantar, 2018). Relationships among
Globodera spp. were not conclusively resolved by
COI, possibly due to gene duplications and variability
within the multipartite mitochondrial genomes in
those species (Armstrong et al., 2000; Phillips
et al., 2016). The relative position of V. zeaphila
relative to Cactodera spp. and Globodera spp. was
best supported in the tree shown in Figure 3. In a
prior study focused primarily on the relationship of
H. medicaginis relative to populations of H. glycines,
a COI tree placed V. zeaphila in a weakly supported
clade with Meloidodera spp., but its relationship
relative to Punctodera spp. and Cactodera spp.
was not well resolved (Powers et al., 2019). An 18S
tree from another study placed V. zeaphila in a
clade with a single C. betulae sequence, but could
not resolve its relationship to the clade containing
Globodera spp. and Heterodera spp. (DeLuca et
al., 2013). The 28S tree from Bernard et al. (2010)
contained a wider range of heteroderid genera for
comparison, with V. zeaphila as a sister taxon to
Heterodera, Cactodera, Punctodera, Globodera,
and Dolichodera. The relationship of V. zeaphila to
the other circumfenestrate nematodes remains fluid
and will be strengthened as additional sequences
become available.
Although evidence of damage to host plants from
V. zeaphila was previously established (Donald et al.,
2012), there remains an urgent need to investigate the
economic impact of this nematode on corn in order
to develop sustainable management strategies.
Acknowledgments
M. K. was supported in part by an appointment to
the Research Participation Program at the Mycology
and Nematology Genetic Diversity and Biology Lab-
oratory, USDA, ARS, Northeast Area, Beltsville, MD,
administered by the Oak Ridge Institute for Science
and Education through an interagency agreement be-
tween the US Department of Energy and USDA-ARS.
The authors thank Daniel Emmert for sample collec-
tion, and Maria Hult and Shiguang Li of MNGDBL
for technical assistance. Mention of trade names or
commercial products in this publication is solely for
the purpose of providing specific information and
does not imply recommendation or endorsement by
the US Department of Agriculture. USDA is an equal
opportunity provider and employer.
References
Armstrong, M. R., Blok, V. C. and Phillips, M. S. 2000.
A multipartite mitochondrial genome in the potato cyst
nematode Globodera pallida. Genetics 154: 181–92.
Baldwin, J. G. and Handoo, Z. A. 2018. General
morphology of cyst nematodes. In Perry, R. N., Moens,
M. and Jones, J. T. (Eds), Cyst nematodes CABI, Wall-
ingford, Oxfordshire, 337–64.
Bernard, E. C., Handoo, Z. A., Powers, T. O., Don-
ald, P. A. and Heinz, R. D. 2010. Vittatidera zeaphila
(Nematoda: Heteroderidae), a new genus and species
of cyst nematode parasitic on corn (Zea mays). Journal
of Nematology 42:139– 50.
Carta, L. K. and Li, S. 2018. Improved 18S small
subunit rDNA primers for problematic nematode ampli-
fication. Journal of Nematology 50:533–42.
Darriba, D., Taboada, G., Doallo, R. and Posada, D.
(2012), jModelTest 2: more models, new heuristics and
parallel computing. Nature Methods, 9:772.
De Ley, P., Tandingan De Ley, I., Morris, K.,
Abebe, E., Mundo-Ocampo, M., Yoder, M., Heras,
J., Waumann, D., Rocha-Olivares, A., Burr, A. H. J.,
Baldwin, J. G. and Thomas, W. K. 2005. An integrated
approach to fast and informative morphological
vouchering of nematodes for applications in molecular
barcoding. Philosophical Transactions of the Royal
Society B 360:1945–58.
DeLuca, F., Vovlas, N., Lucarelli, G., Troccoli, A.,
Radicci, V., Fanelli, E., Cantalapiedra-Navarrete, C.,
Palomares-Ruis, J. E. and Castillo, P. 2013. Heterodera
elachista the Japanese cyst nematode parasitizing corn
in Northern Italy: integrative diagnosis and bionomics.
European Journal of Plant Pathology 136:857–72.
Donald, P. A., Heinz, R., Bernard, E., Hershman,
D., Hensley, D., Flint-Garcia, S. and Joost, R. 2012.

8
Vittatidera zeaphila
from Indiana:
Skantar et al.
Distribution, host status and potential sources of re-
sistance to Vittatidera zaeaphila. Nematropica 42:91–5.
Edgar, R. C. 2004. MUSCLE: multiple sequence
alignment with high accuracy and high throughput.
Nucleic Acids Research 32:1792–7.
Golden, A. M. 1990. Preparation and mounting
nematodes for microscopic observations. In Zuckerman, B.
M., Mai, W. F. and Krusberg, L. R. (Eds), Plant nematology
laboratory manual University of Massachusetts Agricultural
Experiment Station, Amherst, MA, pp. 197–205.
Handoo, Z. A. and Subbotin, S. A. 2018. Taxono-
my, identification and principal species. In Perry, R. N.,
Moens, M. and Jones, J. T. (Eds), Cyst nematodes
CABI, Wallingford, Oxfordshire, pp. 365–98.
Hoang, D. T., Chernomor, O., von Haeseler, A.,
Minh, B. Q. and Vinh, L. S. 2018. UFBoot2: improving
the ultrafast bootstrap approximation. Molecular Biolo-
gy and Evolution 3 5:518 –2 2.
Holterman, M., van der Wurff, A., van den Elsen, S.,
van Megen, H., Bongers, T., Holovachov, O., Bakker,
J. and Helder, J. 2006. Phylum-wide analysis of SSU
rDNA reveals deep phylogenetic relationships among
nematodes and accelerated evolution toward crown
clades. Molecular Biology and Evolution 23:17 9 2– 8 00.
Hooper, D. J. 1970. Handling, fixing, staining, and
mounting nematodes. In Southey, J. F. (Ed.). Laboratory
methods for work with plant and soil nematodes 5th ed.,
Her Majesty’s Stationery Office, London, pp. 39–54.
Howlett, B. J., Brownlee, A. G., Guest, D. I., Adcock,
G. J. and McFadden, G. I. 1992. The 5S ribosomal RNA
gene is linked to large and small subunit ribosomal
RNA genes in the oomycetes, Phytophthora vignae,
P. cinnamomi, P. megaspera f. sp. glycinae and
Saprolegnia ferax. Current Genetics 22:455–61.
Huelsenbeck, J. P. and Ronquist, F. 2001.
MRBAYES: Bayesian inference of phylogenetic trees.
Bioinformatics 17:754– 5 .
Joyce, S. A., Reid, A., Driver, F. and Curran, J.
1994. Application of polymerase chain reaction
(PCR) methods to identification of entomopathogenic
nematodes. In Burnell, A. M., Ehlers, R. -U. and
Masson, J. P. (Eds), COST 812 Biotechnology:
Genetics of Entomopathogenic Nematode-Bacterium
Complexes. Proc. Symp. Workshop, St. Patrick’s
College, Maynooth, Co. Kildare, Ireland. Luxembourg:
European Commission, DG XII, pp. 178–87.
Kalyaanamoorthy, S., Minh, B. Q., Wong, T. K. F.,
von Haeseler, A. and Jermiin, L. S. 2017. ModelFinder:
fast model selection for accurate phylogenetic estimates.
Nature Methods 14:5 87– 9.
Miller, M. A., Pfeiffer, W. and Schwartz, T. 2010.
Creating the CIPRES Science Gateway for inference
of large phylogenetic trees. Proceedings of the Gate-
way Computing Environments Workshop (GCE), New
Orleans, LA, 14 November, pp. 1–8.
Nguyen, L.-T., Schmidt, H. A., von Haeseler, A.
and Minh, B. Q. 2015. IQ-TREE: a fast and effective
stochastic algorithm for estimating maximum likelihood
phylogenies. Molecular Biology and Evolution 32:26 8 –74 .
Phillips, W. S., Brown, A. M. V., Howe, D. K., Peetz,
A. B., Blok, V.C., Denver, D. R. and Zasada, I. A. 2016.
The mitochondrial genome of Globodera ellingtonae is
composed of two circles with segregated gene content
and differential copy numbers. BMC Genomics 17:
706, doi: 10.1186/s12864-016-3047-x.
Powers, T., Skantar, A., Harris, T., Higgins, R., Mullin,
P., Hafez, S., Handoo, Z., Todd, T. and Powers, K.
2019. DNA barcoding evidence for the North American
presence of alfalfa cyst nematode, Heterodera medi-
caginis. Journal of Nematology 51:e2019–16.
Sievers, F., Wilm, A., Dineen, D. G., Gibson, T. J.,
Karplus, K., Li, W., Lopez, R., McWilliam, H., Remmert,
M., Söding, J., Thompson, J. D. and Higgins, D. G.
2011. Fast, scalable generation of high-quality protein
multiple sequence alignments using Clustal Omega.
Molecular Systems Biology 7:5 3 9.
Skantar, A. M. and Carta, L. K. 2004. Molecular
characterization and phylogenetic evaluation of the
Hsp90 gene from selected nematodes. Journal of
Nematology 36:466–80.
Skantar, A. M., Handoo, Z. A., Carta, L. K. and
Chitwood, D. J. 2007. Morphological and molecular
identification of Globodera pallida associated with po-
tato in Idaho. Journal of Nematology 39:133 – 4 4.
Subbotin, S. A. 2015. Heterodera sturhani sp. n.
from China, a new species of the Heterodera avenae
species complex (Tylenchida: Heteroderidae). Russian
Journal of Nematology 23:14 5 –52.
Subbotin, S. A. and Skantar, A. M. 2018. Molecular
taxonomy and phylogeny. In Perry, R. N., Moens, M.
and Jones, J. T. (Eds), Cyst nematodes CABI, Walling-
ford, Oxfordshire, pp. 399–418.
Thomas, W. K., Vida, J. T., Frisse, L. M., Mundo, M.
and Baldwin, J. G. 1997. DNA sequences from formalin-
fixed nematodes: integrating molecular and morpho-
logical approaches to taxonomy. Journal of Nematology
29:250 –4.
Ye, W., Giblin-Davis, R. M., Davies, K. A., Purcell,
M. F., Scheffer, S. J., Taylor, G. S., Center, T. D., Mor-
ris, K. and Thomas, W. K. 2007. Molecular phyloge-
netics and the evolution of host plant associations
in the nematode genus Fergusobia (Tylenchida: Fer-
gusobiinae). Molecular Phylogenetics and Evolution
45:123 – 41.