Download molecular phylogeny of the haplosporidia based on

J. Parasitol., 90(5), 2004, pp. 1111–1122
q American Society of Parasitologists 2004
MOLECULAR PHYLOGENY OF THE HAPLOSPORIDIA BASED ON TWO INDEPENDENT
GENE SEQUENCES
Kimberly S. Reece, Mark E. Siddall*, Nancy A. Stokes, and Eugene M. Burreson
Department of Environmental and Aquatic Animal Health, Virginia Institute of Marine Science, The College of William and Mary, Gloucester
Point, Virginia 23062. e-mail: [email protected]
ABSTRACT: The phylogenetic position of the Haplosporidia has confounded taxonomists for more than a century because of the
unique morphology of these parasites. We collected DNA sequence data for small subunit (SSU) ribosomal RNA and actin genes
from haplosporidians and other protists for conducting molecular phylogenetic analyses to help elucidate relationships of taxa
within the group, as well as placement of this group among Eukaryota. Analyses were conducted using DNA sequence data from
more than 100 eukaryotic taxa with various combinations of data sets including nucleotide sequence data for each gene separately
and combined, as well as SSU ribosomal DNA data combined with translated actin amino acids. In almost all analyses, the
Haplosporidia was sister to the Cercozoa with moderate bootstrap and jackknife support. Analysis with actin amino acid sequences
alone grouped haplosporidians with the foraminiferans and cercozoans. The haplosporidians Minchinia and Urosporidium were
found to be monophyletic, whereas Haplosporidium was paraphyletic. ‘‘Microcell’’ parasites, Bonamia spp. and Mikrocytos
roughleyi, were sister to Minchinia, the most derived genus, with Haplosporidium falling between the ‘‘microcells’’ and the more
basal Urosporidium. Two recently discovered parasites, one from abalone in New Zealand and another from spot prawns in
British Columbia, fell at the base of the Haplosporidia with very strong support, indicating a taxonomic affinity to this group.
Haplosporidia is composed of histozoic and coelozoic parasites in a variety of freshwater and marine invertebrates; some
species are significant pathogens of commercially important
molluscs (Burreson et al., 2000). Since the discovery of the first
species in the late 1800s, the Haplosporidia have been a troublesome group for taxonomists and phylogeneticists. Historically, the taxon has been treated as a last resort for a diversity
of spore-forming parasites that have multinucleated naked cells
(plasmodia) in their life cycles and that were not readily classifiable elsewhere (Sprague, 1979).
There have been numerous taxonomic schemes proposed for
placement of the group within the protists and for appropriate
taxa within the Haplosporidia. Caullery and Mesnil (1899) established Haplosporidium for 2 parasites of marine annelids and
placed the genus in the new order Haplosporidia in the class
Sporozoa of the phylum Protozoa. Caullery (1953) recognized
6 genera in the order, and Kudo (1971) recognized 7 genera. A
major change in the classification of the Haplosporidia was the
separation of the Haplosporidia and the Paramyxea from other
‘‘sporozoa’’ by establishment of the new phylum Ascetospora
(Sprague, 1979). This scheme proposed 2 classes: Stellatosporea, for the families Marteiliidae and Haplosporidiidae, and Paramyxea, for the family Paramyxidae. The family Haplosporidiidae contained only 3 genera, i.e., Haplosporidium, Minchinia, and Urosporidium. Desportes and Nashed (1983) recognized that the family Marteiliidae belonged in the class
Paramyxea, not Stellatosporea, because of its development cycle, and proposed 2 classes in the phylum Ascetospora, i.e.,
Haplosporea and Paramyxea. However, they suggested that the
2 classes probably should be raised to phylum rank because of
very different developmental sequences. Recently, the phylum
Ascetospora has been abandoned, and Haplosporidia and Paramyxea have each been elevated to phylum rank (Desportes
and Perkins, 1990; Perkins, 1990, 1991; Cavalier-Smith, 1993).
Separate phylum rank for Haplosporidia and Paramyxea has
been accepted by most researchers; however, Corliss (1994) re-
Received 5 March 2003; revised 12 December 2003; accepted 17
February 2004.
* Department of Invertebrate Zoology, American Museum of Natural
History, New York, New York 10024.
tained the phylum Ascetospora for both Haplosporidia and Paramyxea.
The earliest molecular phylogenetic analyses for the Haplosporidia (Siddall et al., 1995; Flores et al., 1996) placed the
phylum as a monophyletic group within the Alveolata and as a
taxon of equal rank with the other alveolate phyla. A more
recent analysis, with much more sequence data available for a
variety of taxa, has placed the Haplosporidia as sister taxon to
the Dictyosteliida (Berthe et al., 2000). The analysis by Berthe
et al. (2000) also provided molecular phylogenetic support for
separate phylum rank for Haplosporidia and Paramyxea. The
most recent phylogenetic analysis involving the Haplosporidia
(Cavalier-Smith and Chao, 2003) hypothesizes that the group
includes the Paramyxea and falls within the Cercozoa.
The phylum Haplosporidia was recently described by Perkins
(2000) as a group of parasitic protists that form ovoid, walled
spores with an orifice covered externally by a hinged lid or
internally by a flap of wall material. There are 31 recognized
species in 3 genera, i.e., Minchinia, Haplosporidium, and Urosporidium. However, recent molecular phylogenetic analyses
supported the inclusion of the enigmatic genus Bonamia and
Mikrocytos roughleyi, which are not known to form spores,
within the Haplosporidia (Carnegie et al., 2000; CochennecLaureau et al., 2003).
For this study, we used nucleotide sequence data of 2 genes,
actin and the small subunit (SSU) ribosomal RNA (rRNA)
gene, to investigate the position of the Haplosporidia among
eukaryotes, to determine the taxonomic composition of the
group, and to assess the monophyly of recognized genera.
MATERIALS AND METHODS
Sample collection
Samples of haplosporidian-infected host tissues were collected in
U.K., France, Spain, and Australia, and in Mississippi, Virginia, and
Michigan in the United States (Table I). Cercozoan culture samples
ATCC50317 and ATCC50318 were obtained from ATCC for DNA isolation and amplification of actin gene fragments.
DNA isolation
Spores of haplosporidian species were isolated after degradation of
infected host tissue, and genomic DNA was extracted by methods previously described (Flores et al., 1996). DNA was isolated from the
1111
1112
THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 5, OCTOBER 2004
TABLE I. Collection sites and dates of samples collected for this study.
Parasite species
Haplosporidium nelsoni
Host
Sample collection site
Minchinia teredinis
Undescribed parasite
Crassostrea virginica (Eastern
oyster)
Panopeus herbstii (mud crab)
Crassostrea virginica (Eastern
oyster)
Helcion pellucidus (limpet)
Physa parkeri (snail), Stagnicola
emarginalus (snail)
Microphallid trematode in Callinectes sapidus (blue crab)
Stictodora lari (trematode) in Battilaria australis (whelk)
Lepidochitona cinereus (chiton)
Ruditapes decussatus (carpet shell
clam)
Teredo navalis (shipworm)
Pandalus platyceros (spot prawn)
Undescribed haplosporidian
Cyrenoida floridana (marsh clam)
Haplosporidium louisiana
Haplosporidium costale
Haplosporidium lusitanicum
Haplosporidium pickfordi
Urosporidium crescens
Urosporidium sp.
Minchinia chitonis
Minchinia tapetis
cercozoan cultures with the QIAamp DNA mini-kit (Qiagen, Carlsbad,
California) according to the manufacturer’s protocol for tissue samples.
Polymerase chain reaction amplification
The SSU rRNA genes for most of the haplosporidian species were
amplified in the polymerase chain reaction (PCR) using ‘‘universal’’
eukaryotic primers (Medlin et al., 1988) and conditions previously described (Flores et al., 1996). Internal SSU rRNA gene primers Urocomp280SSU (Reece and Stokes, 2003) and Uro280SSU (TAAYTGTTCGGATCGCATGGC) were paired with the eukaryotic primers
16S-A and 16S-B (Medlin et al., 1988), respectively, to amplify the
Australian Urosporidium sp. rather than host DNA. Amplification conditions were same as for eukaryotic primers above. Generation of the
SSU rRNA gene sequence for Haplosporidium pickfordi required several amplifications using specific primers: HAP-F1 (Renault et al., 2000)
and 16S-B; 16S-A and MpickSSU391 (GCTTATTCAATCGGTAGGAGC); 16S-A and MpickSSU280 (CAATCGTCTATCCCCACTTG);
Mpick125F (AACCGTGGTAACTCCAGGG) and Mpick1450R (TTATTGCCCCACGCTTCC); Mpick250F (CATTCAAGTTTCTGCCCTA
TCAG) and Mpick1220R (TGTCTGGTAAGTTTTCCCGTGTTG);
HAP-F1 and HAP-R3 (Renault et al., 2000). Reaction mixtures for PCR
were same as above except with Mpick125F1 Mpick1450R and
Mpick250F1 Mpick1220R, where buffer and polymerase were from
the Expand High Fidelity PCR system (Roche Diagnostics, Indianapolis, Indiana). Cycling parameters consisted of an initial denaturation of
4 min at 94 C followed by 35 cycles of 30 sec at 94 C for denaturation,
30 sec at 48 C for HAP-F1116S-B and HAP-F11HAP-R3, 55 C for
16S-A1MpickSSU391 and 16S-A1MpickSSU280, 59 C for
Mpick125F1Mpick1450R and Mpick250F1Mpick1220R for annealing, 1.5 min at 72 C for extension and a final extension of 5 min at 72 C.
A central coding region of actin genes from the haplosporidians, their
hosts, and cercozoans was amplified using various combinations of 2
forward (480, 481) and 2 reverse (482, 483) universal actin gene primers (Carlini et al., 2000) using amplification parameters and conditions
previously described (Reece et al., 1997). Amplification products were
cloned into either the plasmid vector pNoTA/T7 using the Prime PCR
Cloner Cloning System (5 Prime-3 Prime, Inc., Boulder, Colorado) or
into pCR2.1 using the TA Cloning Kit (Invitrogen, Carlsbad, California)
according to the manufacturers’ instructions.
DNA sequencing
DNA clone inserts of SSU and actin gene fragments were sequenced
manually as previously described (Reece et al., 1997) or by automated
sequencing using either unidirectional or simultaneous bidirectional cy-
Year collected
Virginia
1992–1997
Gloucester Point, Virginia
Wachapreague, Virginia
1991
1996
Cap de la Hague, France
Douglas Lake, Michigan
1998
1999, 2000
Wachapreague, Virginia
1994
Sydney, New South Wales,
Australia
Plymouth, U.K.
Galicia, Spain
2000
Wachapreague, Virginia
Malaspina Strait, British Columbia, Canada
Ocean Springs, Mississippi
1991, 1993, 1997
1999
1996
1996
1999
cle sequencing. Reactions for automated analysis were done as previously described (Reece and Stokes, 2003). Sequences were imported
into MacVector 7.0 Sequence Analysis Software (Oxford Molecular
Ltd., Oxford, U.K.) for trimming vector sequences, alignment, and for
the actin gene sequences, coding region analyses. Actin gene introns
(see Table II) were located, and the splice junction points were identified
by using the MacVector 7.0 package to translate the DNA sequences in
all 3 reading frames followed by coding region analyses and alignment
to haplosporidian and other protistan actin gene fragments that did not
contain introns.
Sequence and phylogenetic analyses
Occasionally, host or other contaminant SSU rRNA and actin genes
were amplified rather than the targeted haplosporidian sequences. Identification of the DNA source for all amplified gene fragments involved
BLAST (Altschul et al., 1990) searches of the National Center for Biotechnology Information (NCBI) GenBank database, as well as frequent
intermediate phylogenetic analyses, which included previously obtained
host, haplosporidian, and other protozoan sequences. In some cases,
amplification of haplosporidian sequences from several infected host
individuals was done to demonstrate that identical or highly similar
(.98% sequence identity) sequences could be obtained from independent samples.
Phylogenetic analyses were conducted in 2 series. The first concerned
the assessment of the relationship of the Haplosporidia to other eukaryotic groups and comprised 121 taxa for the SSU rDNA sequences and
50 taxa for the actin sequences ranging across Eukaryota. Considerations at that level were limited principally to support for the groupings
of and among various clades of protists, as opposed to support within
each grouping. The second series concerned specific analysis of the
phylogenetic relationships of genera and species within Haplosporidia
using the closest relatives of the group, as determined from the first
series of analyses. Sequences were aligned for each of these series independently.
The SSU rRNA and actin gene sequences used in the phylogenetic
analyses, along with their GenBank accession numbers, are listed in
Table III. Sequences were aligned using the CLUSTALW algorithm
(Thompson et al., 1994) in the MacVector 7.0 package using a variety
of open and extend gap penalties in the ranges of 4–20 and 2–10,
respectively. The SSU rRNA gene alignments were compared with secondary structure–based alignments done through the Ribosomal Database Project II website (http://rdp.cme.msu.edu/html/, Maidak et al.,
2001). Final alignments used in the analyses were accomplished with
gap penalties of 8 for insertions and 3 for extensions both in pairwise
REECE ET AL.—MOLECULAR PHYLOGENY OF THE HAPLOSPORIDIA
1113
TABLE II. Actin gene introns, locations, and lengths. Sequences in bold were used in the phylogenetic analyses presented here.
Parasite species
No. of unique
actin sequences
No. of DNA
identified
clones sequenced
Haplosporidium costale
Haplosporidium louisiana
1
4
4
7
Haplosporidium nelsoni
Minchinia chitonis
1
3
4
8
Minchinia tapetis
3
6
Minchinia teredinis
3
5
Urosporidium crescens
Undescribed parasite CfP
Undescribed parasite SPP
1
1
1
4
1
5
Gene
designation
No. of introns
within amplified
fragment
HcAc1
HlAc1
HlAc5
HlAc8
HlAc11
HnAc3
McAc5
McAc11
McAc42
MtaAc1
MtaAc7
MtaAc9
MteAc2
MteAc5
MteAc9
UcAc1
CfhapAc64
SppAc6
2
0
0
0
3
0
1
3
1
0
2
0
2
2
0
0
1
4
Intron positions* (intron
lengths in bp)
131, 172 (18, 17)
131, 172, 224 (177, 63, 83)
304 (124)
131, 172, 224 (25, 25, 28)
304 (114)
131, 172 (33, 77)
172, 224 (22, 60)
131, 172 (23, 25)
172 (25)
131, 172, 224, 304 (50, 60, 222, 185)
* Intron positions are designated relative to the homologous amino acid position in the human aortic actin gene (GenBank NMp005159).
and in multiple alignment phases. In addition, aligned sequences were
analyzed in various combinations to determine the sensitivity of various
groups to the relative inclusion and exclusion of potentially phylogenetically informative sites. In the first series of analyses, which concerned the relative position of Haplosporidia within Eukaryota, it was
noted that there were large inserted regions in the SSU rDNA sequences
of some taxa, including some of the haplosporidians, e.g., the prawn
parasite and Urosporidium species. In light of this, a total of 798 sites
in the alignment were targeted for removal to determine the effect that
removal or inclusion of these sequences had on results. Similarly, in
terms of the actin nucleotide data, the relative contribution of third
codon positions to the predicted phylogenies was assessed, both through
their exclusion and by using translated amino acid sequences.
All combinations of SSU rDNA nucleotides, with and without the
798 sites targeted for exclusion, alone, and in combination with actin
nucleotides, with and without third codon positions, or using translated
amino acid sequences alone or in combination with the SSU rDNA data,
were examined in terms of their relative support for higher-level relationships among the included eukaryotic taxa. Insofar as the specifics
of species-level relationships were of only passing interest in the analyses of all Eukaryotic taxa, groupings and relative levels of support
were determined through unweighted parsimony jackknife methods
(Farris et al., 1996) for nucleotide combinations using XAC (Farris,
1998). Unweighted parsimony jackknife and bootstrap in PAUP 4.0b10
(Swofford, 2002) were used for those analyses concerning amino acid
sequences with the ‘‘emulate Jac’’ option. In each case, 1,000 replicates
were used, each with 5 random additions and with a subtree-pruningregrafting branch swapping algorithm. After determinations of jackknife
support in these higher-level Eukaryota analyses, we also searched for
optimal trees on the combined SSU rDNA and actin amino acid sequences because jackknife values from this data combination were intermediate among the 6 combinations tried. This search used 30 random
taxon addition search sequences followed by branch breaking (TBR
branch swapping).
With respect to the secondary analyses in which we were concerned
with the specifics of the relationships within Haplosporidia, sequences
were realigned for the 26 included taxa for the combined analysis according to the same parameters as above. In these analyses, all sites for
both nucleotide sets were included and combinations of data considered
SSU rDNA alone, as well as those data in combination with actin nucleotides or with translated actin amino acid sequences. Thorough heuristic searches for most parsimonious trees (as opposed to jackknife
support trees) were conducted with PAUP 4.0b10 (Swofford, 2002) for
all the secondary data sets. In each case, searches involved 30 random
taxon addition search sequences followed by branch breaking (TBR
branch swapping).
RESULTS
From some samples, novel host as well as parasite SSU
rRNA or actin gene sequences (or both) were identified. Host
and parasite SSU rRNA and actin gene sequences that were
confidently identified as either of host or parasite origin by
BLAST searches and phylogenetic analyses in this study were
deposited in GenBank. Additional sequences whose origin
could not be determined were obtained from many samples and
were presumed to be from contaminating organisms.
Jackknife values obtained from the various data combinations
used across Eukaryota for determination of the closest extant
relatives of Haplosporidia are listed in Table IV. Bootstrap values were comparable with the jackknife values in all analyses,
never varying by more than 3%, and usually bootstrap support
values were slightly higher. With all but 1 of the data set combinations, the Haplosporidia were supported as having arisen
from a common ancestor with the Cercozoa. With the complete
combined nucleotide data set, which included all the SSU rRNA
and actin gene nucleotides, the Haplosporidia were not supported as having arisen from a common ancestor with the Cercozoa. This complete nucleotide data set, however, failed to find
greater than 50% support for the Haplosporidia grouping with
any of the other Eukaryotic clades used in this analysis. Parsimony analysis with the actin amino acid data set alone
grouped the Haplosporidia with the Foraminifera and most
closely to the type 1 foraminiferan actins (Keeling, 2001). The
foraminiferan and haplosporidian clade was sister to the cercozoans. The support for these groupings, however, was below
50%. All the remaining data sets found that Haplosporidia and
Cercozoa are clades with support levels ranging from 61% with
the SSU rDNA alone up to 81% with the SSU rDNA sequences
1114
THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 5, OCTOBER 2004
TABLE III. Taxa included in phylogenetic analysis and the GenBank accession numbers of their SSU rRNA and actin gene sequences. Sequences
determined for this study are shown in bold.
Taxon
SSU rRNA gene
Actin gene
Bonamia ostreae
Bonamia sp.
Haplosporidium costale
Haplosporidium louisiana
Haplosporidium lusitanicum
Haplosporidium nelsoni
Haplosporidium pickfordi
Mikrocytos roughleyi
Minchinia chitonis
Minchinia tapetis
Minchinia teredinis
Urosporidium crescens
Urosporidium sp.
Undescribed Cyrenoida floridana parasite (CfP)
Undescribed Haliotis iris parasite (NZAP)
Undescribed Pandalus platyceros parasite (SPP)
Allogromia sp.
Ammonia sp.
Ammonia beccarii
Reticulomyxa filosa
Cercomonas longicauda
Cercomonas sp.—ATCC50317
Cercomonas sp.—ATCC50318
Chlorarachnion reptans
Chlorarachnion sp.
Euglypha rotunda
Heteromita globosa
Massisteria marina
Unidentified cercozoan species
Phagomyxa bellerocheae
Phagomyxa odontellae
Plasmodiophora brassicae
Achlya bisexualis
Bolidomonas pacifica
Cafeteria roenbergensis
Cafeteria sp.
Chrysosaccus sp.
Chrysosphaera parvula
Chrysoxys sp.
Ciliophrys infusionum
Costaria costata
Fucus vesiculosus
Hyphochytrium catenoides
Lagynion scherffelii
Ochromonas danica
Phytophthora megasperma
Phytophthora infestans
Rhizidiomyces apophysatus
Skeletonema costatum
C9G
Labyrinthula sp.
QPX
Ulkenia profunda
Ajellomyces capsulatus
Apusomonas proboscidea
Candida glabrata
Kluyveromyces lactis
Neurospora crassa
Pneumocystis carinii
Saccharomyces cerevisiae
Acanthamoeba castellanii
AF192759, AF262995
AF337563
AF387122
U47851
AY449713
U19538
AY452724
AF508801
AY449711
AY449710
U20319
U47852
AY449714
AY449712
AF492442
AY449715–716
X86093†
NA
U07937†
AJ132367†
AF101052
U42449
U42450
U03477
NA
X77692
U42447
AF174374
UEU130858
AF310903†
AF310904†
U18981†
M32705
AF167154
AF174364
AF174366
AF123300
AF123299
AF123302
L37205
X53229
U97110
AF163294
AF123288
M32704
X54265
NA
AF163295
X85395
AF474172
AB022105
AF261664
L34054
Z75307
L37037
X51831
X51830
X04971
S83267
J01353
AF251938
NA*
NA
AY450407
AY450408–411
NA
AY450412
NA
NA
AY450413–415
AY450416–418
AY450419–421
AY450422
NA
AY450423
NA
AY450424
AJ132370–371
AJ132372–373
NA
AJ132374–375
NA
AF363534
AF363536
NA
AF363528
NA
NA
NA
NA
NA
NA
NA
X59936
NA
NA
NA
NA
NA
NA
NA
X59937
X98885
NA
NA
NA
NA
M59715
NA
NA
NA
NA
NA
NA
U17498
NA
AF069746
M25826
U78026
L21183
V01288
V00002, J01016
REECE ET AL.—MOLECULAR PHYLOGENY OF THE HAPLOSPORIDIA
TABLE III. Continued.
Taxon
SSU rRNA gene
Actin gene
Chilomonas paramecium
Chlamydomonas reinhardtii
Emiliania huxleyi
Nitella flexilis
Chondrus crispus
Stylonema alsidii
Marteilia refringens
Alexandrium fundyense
Amphidinium carterae
Gonyaulax spinifera
Hematodinium sp.
Lepidodinium viride
Pfiesteria piscicida
Prorocentrum minimum
Symbiodinium corculorum
Symbiotic dinoflagellate BBSR 323
Perkinsus chesapeaki
Perkinsus marinus
Perkinsus olseni
Babesia bovis
Caryospora bigenetica
Colpodella sp.
Cryptosporidium parvum
Eimeria tanella
Parvilucifera infectans
Sarcocystis hominis
Theileria parva
Toxoplasma gondii
Anophyroides haemophila
Didinium nasutum
Entodinium caudatum
Euplotes crassus
Frontonia vernalis
Oxytricha nova
Oxytricha trifallax
Paramecium tetraurelia
Prorodon viridis
Strombidium purpureum
Tetrahymena pyriformis
Tetrahymena thermophila
Acanthometra sp.
Arthracanthid 206
Haliommatidium sp.
Encephalitozoon cuniculi
Encephalitozoon hellum
Entamoeba histolytica
Euglena gracilis
Trichomonas foetus
Trichomonas vaginalis
Trypanosoma brucei
Haliotis iris
Crassostrea virginica
Cyrenoida floridana
Helcion pellucidus
Lepidochitona cinereus
Pandalus platyceros
Physa parkeri
Ruditapes decussatus
Stictodora lari
Teredo navalis
L28811
M32703
M87327
U05261
Z14140
AF168633
AJ250699†
U09048
AF009217
AF052190
AF286023
AF022199
AF077055
Y16238
L13717
U52356
AF042707
AF042708
L07375
L19078
AF060975
AY142075, AY449717
AF115377
U40264
AF133909
AF006470
L02366
U03070
U51554
U57771
U57765
AY007438
U97110
X03948
AF164121
X03772
U97111
U97112
M98021
X56165
AF063240
AF063239
AF018159
L17072
M87327
X65163
M12677
M81842
U17510
AJ009141
AF492441
L78851†
NA
NA
NA
NA
NA
AF29512†
NA
NA
NA
D50839
S64188
NA
U03676
NA
NA
NA
U84289
NA
NA
NA
NA
U84290
NA
NA
NA
U84287
NA
NA
NA
NA
M86241
NA
NA
NA
NA
U10429
NA
NA
AF078106
J04533
NA
M22480
NA
AF043608
NA
NA
X05195
M13939
NA
NA
NA
NA
AF031701
M19871
AF057161
NA
AF237734
M20310
NA
X75894†
AY452511–512†
AY452513†
AY452514–515†
AY452516†
AY485722†
AY452517–518†
AY452519–521†
AY452522†
* NA, sequence not available.
† Sequences used for analyses, but not included in trees presented.
1115
1116
THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 5, OCTOBER 2004
TABLE IV. Summary of jackknife support values for phylogenetic analyses with various data sets.*
Ciliophora
Dinozoa
Apicomplexa
Alveolata
Stramenopiles
Labyrinthulids
Fungi
Chlorophytes
(FUN 1 CHL)
Haplosporidia
Cercozoa
(HAP 1 CERC)
SSU
SSUX
SSU1ACT
SSUX1ACTX
SSU1ACTX
SSUX1ACT
SSU1ACTaa
SSUX1ACTaa
100
100
—
52
67
100
97
90
78
91
100
61
99
95
—
73
69
91
88
86
77
67
100
64
95
100
—
54
80
96
93
97
—
83
100
—†
98
94
—
78
77
86
72
—
53
90
100
79
100
100
—
64
78
94
92
74
68
94
100
63
88
96
—
—
75
97
68
91
—
92
99
75
100
100
—
70
72
94
98
86
79
95
100
74
99
97
—
77
75
94
84
71
66
87
100
81
* SSU, all SSU rDNA nucleotides aligned by CLUSTALW; SSUX, SSU rDNA nucleotides aligned by CLUSTALW, with 798 poorly aligned nucleotide positions
removed; ACT, all actin gene nucleotides aligned by CLUSTALW; ACTX, actin gene nucleotides aligned by CLUSTALW, with the third positions of codons
removed.
† No alternative grouping of the Haplosporidia was supported by this analysis.
combined with actin amino acid sequences but excluding the
798 sites corresponding to SSU rDNA insertions in the haplosporidian sequences. Intermediate between these values were the
analyses using all SSU rDNA data and either the actin amino
acid sequences with 74% support for the cercozoan–haplosporidian clade or the actin nucleotide sequences with third positions excluded with 63% support for the cercozoan–haplosporidian clade. Overall, both these analyses rendered high levels
of support for other known eukaryotic groups (Table IV). The
SSU rDNA data combined with the actin amino acid sequences,
when subjected to a full parsimony search, yielded 4 trees with
equal length of 17,977, a consistency index (CI) of 0.306, and
a retention index (RI) of 0.557, the strict consensus of which
is illustrated in Figure 1. Foraminifera SSU rRNA gene sequences could not be confidently aligned with the other sequences and, therefore, were not used in these analyses. The
consensus tree of 122 equally parsimonious trees with a length
of 1,153 (CI 5 0.440; RI 5 0.555) resulting from analysis with
the actin amino acid data set is shown in Figure 2.
For parsimony analyses examining the in-group relationships
of the Haplosporidia, 8 cercozoan taxa were used as the outgroup (Fig. 3). Use of SSU rDNA data alone yielded a single
tree with length 3,838 (CI 5 0.600; RI 5 0.658). Use of SSU
rDNA in combination with translated actin amino acids also
rendered a single tree, identical in topology to the SSU rDNA
tree, with length 4,172 (CI 5 0.607; RI 5 0.649). Use of SSU
rDNA in combination with actin nucleotides yielded 2 trees
each with length 5,213 (CI 5 0.572; RI 5 0.589). The consensus of those 2 trees disagreed with the other analyses in moving
a clade composed of H. pickfordi and Haplosporidium lusitanicum to a sister group position with Haplosporidium louisiana
as opposed to with Haplosporidium costale. The undescribed
haplosporidian parasite from Cyrenoida floridana consistently
grouped within the described Minchinia species. All combined
data set analyses agreed on a variety of other findings including
that (1) Minchinia is monophyletic (if the parasite from C. floridana is considered a Minchinia species); (2) Urosporidium is
monophyletic; (3) the nonspore-forming ‘‘microcell’’ parasites
within the Haplosporidia, e.g., Bonamia spp. and M. roughleyi,
are monophyletic and sister to Minchinia; (4) Haplosporidium
as currently constituted is paraphyletic; and (5) the earliest lineages in Haplosporidia are the yet to be described parasite species recently obtained from prawns and from abalone.
In several cases, more than 1 actin gene type was amplified
from DNA of a haplosporidian species. Multiple paralogs were
found in H. louisiana, Minchinia chitonis, Minchinia teredinis,
and Minchinia tapetis. Only a single type of actin gene was
identified from Haplosporidium nelsoni, H. costale, and Urosporidium crescens, and no actin gene that could be confidently
identified as a parasite sequence was successfully amplified
from H. lusitanicum, H. pickfordi, or from the Australian Urosporidium sp. found in Stictodora lari (trematode) of Battilaria
australis (whelk), although 10–27 DNA clones were sequenced
from each of these samples. As sequences were obtained, phylogenetic analyses were conducted to identify orthologous
genes for inclusion in the overall phylogenetic analyses. Many
haplosporidian actin gene sequence fragments were found to
contain introns. Introns were found in at least 1 actin gene paralog from each of the haplosporidian taxa from which actin
genes were able to be amplified, except from H. nelsoni, and
U. crescens. Orthologs and paralogs of parasite origin were
identified by phylogenetic analyses with known actin gene sequences from hosts and other protozoans. The genes with introns as well as the single intronless actin genes isolated from
H. nelsoni and U. crescens were determined to be orthologous
(Table II) and were used in the comprehensive phylogenetic
analyses. Intron positions were conserved among the haplosporidian actin genes, with 4 different intron positions identified
at positions corresponding to the amino acids 131, 172, 224,
and 304 of the human aortic actin gene (GenBank NMp005159).
The total number of introns within the amplified gene fragment
ranged from 1 to 4 (Table II). Individual actin genes from the
described haplosporidian species contained 1–3 introns, and the
introns were found at all 4 of the positions in the actin gene of
the haplosporidianlike spot prawn parasite. Intron lengths
ranged from very short introns of 17–33 nucleotides, up to 222
nucleotides. The introns were flanked by typical splice junction
sequences with ‘‘GT’’ at the 59 end and ‘‘AG’’ at the 39 end of
all introns.
REECE ET AL.—MOLECULAR PHYLOGENY OF THE HAPLOSPORIDIA
1117
FIGURE 1. Strict jackknife consensus of 4 equal length trees resulting from parsimony analysis with the SSU rDNA and actin amino acid data
set. Analysis was done on the complete taxonomic data set with 798 poorly aligned nucleotide positions in the SSU rDNA removed. Jackknife
support values are given at the nodes. Dashed lines indicate clades that did not have jackknife support values above 50.
1118
THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 5, OCTOBER 2004
FIGURE 2. Strict jackknife consensus of 122 equal length trees resulting from parsimony analysis with the actin amino acid data set. Analysis
was done on actin sequences from 47 taxa. Jackknife support values are given at the nodes. Dashed lines indicate clades that did not have
jackknife support values above 50.
REECE ET AL.—MOLECULAR PHYLOGENY OF THE HAPLOSPORIDIA
1119
FIGURE 3. Strict jackknife consensus trees of parsimony analyses conducted with the reduced taxonomic data set that included the cercozoans
and haplosporidians for examination of relationships within the phylum Haplosporidia. Jackknife support values are given at the nodes. A. Tree
resulting from analysis of the SSU rDNA sequence data. Tree statistics: length (L) 5 3,838, consistency index (CI) 5 0.600, retention index (RI)
5 0.658. B. Strict consensus of 2 trees of equal length resulting from analysis of the SSU rDNA and actin gene sequence data. Tree statistics: L
5 5,213, CI 5 0.572, RI 5 0.589. C. Tree resulting from analysis of the SSU rDNA and actin amino acid sequence data. Tree statistics: L 5
4,172, CI 5 0.607, RI 5 0.649.
1120
THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 5, OCTOBER 2004
DISCUSSION
This combined actin–SSU rRNA data set, including the data
for several haplosporidian taxa collected during this study and
additional data already in GenBank, places Haplosporidia in a
position of recent common ancestry with cercozoans and not
within the Alveolata as previously hypothesized on the basis of
the SSU rRNA gene data alone (Siddall et al., 1995; Flores et
al., 1996). This study includes not only an additional gene and
many additional haplosporidian taxa but also several more protistan SSU rRNA and actin gene sequences that were not available for those studies. The haplosporidians grouped with the
foraminiferans and cercozoans in analyses based on actin sequences alone, although jackknife support for this relationship
was below 50% (Fig. 2). We did not, however, include the foraminiferan SSU rDNA sequences in any of the analyses because we did not have confidence in alignments that included
these divergent sequences. Keeling (2001) found that the Foraminifera and Cercozoa were closely related in actin and tubulin gene phylogenies. Any more definitive conclusions regarding the relationships between Haplosporidia and Foraminifera, however, will require additional data.
Although previous studies have suggested that Haplosporidia
and Cercozoa are closely related (Cavalier-Smith, 2002; Cavalier-Smith and Chao, 2003), this is the first study to demonstrate
moderate support (JK 5 74) for this affinity. Cavalier-Smith
(2002) stated that the Haplosporidia are members of the Cercozoa but presented absolutely no data on which to evaluate
this claim. Cavalier-Smith and Chao (2003) had only weak support (BS 5 20 with Marteilia refringens included in the analysis
and BS 5 60 with it removed) for inclusion of the Haplosporidia within the Cercozoa. Although that study also suggested
that M. refringens was related to the haplosporidians, the results
of this study were consistent with the results of Berthe et al.
(2000), in that we found no support for this relationship in any
of our analyses. The fact that there is strong support in this
study for many other accepted taxonomic groupings, including
the stramenopiles, alveolates, labyrinthulids, fungi, chlorophytes and cercozoans (see Table IV), lends support to the validity
of these results. Moreover, the use of combined data from these
2 loci provided more support for grouping Haplosporidia with
Cercozoa, than either locus did alone and with support values
similar to those seen for Alveolata and Stramenopiles arising
at comparable depths in the resulting tree. In addition, if the
Cercozoa are accepted as a phylum, then the results of this
study hypothesize that the Haplosporidia are also a distinct phylum, rather than included within the Cercozoa as hypothesized
by Cavalier-Smith and Chao (2003).
This phylogenetic study based on molecular data from 2 independent DNA sequences, namely SSU rRNA and actin gene
sequences, generally supports the earlier taxonomic assignments within the group Haplosporidia that were largely based
on spore morphologies (Sprague, 1979; Ormières, 1980; McGovern and Burreson, 1990; Perkins, 1990, 1991, 2000). Monophyly of Urosporidium and Minchinia was supported in this
study. Haplosporidium is paraphyletic and falls intermediate to
the more basal Urosporidium and the most derived genus Minchinia, as previously hypothesized by Ormières (1980).
Many of the more recent observations regarding taxa found
to have haplosporidian affinities and reassignments within the
group were supported by this study. The Bonamia spp. and M.
roughleyi fell within the Haplosporidia, as had been observed
in previous studies (Carnegie et al., 2000; Cochennec-Laureau
et al., 2003). These ‘‘microcells’’ grouped together in this analysis as a sister group to Minchinia. The results of this study
strongly suggest that the undescribed C. floridana parasite is a
Minchinia species because it fell between M. chitonis and M.
teredinis within a monophyletic Minchinia clade in all the analyses. In another study, the freshwater parasite Minchinia pickfordae was assigned recently to Haplosporidium on the basis
of spore ornamentation (Burreson, 2001) and renamed H. pickfordi. Its removal from Minchinia was supported here. A haplosporidianlike abalone parasite (Diggles et al., 2002; Hine et
al., 2002; Reece and Stokes, 2003) fell at the base of the Haplosporidia, sister to another haplosporidianlike parasite recently
identified in the spot prawn, Pandalus platyceros (Bower and
Meyer, 2002). Overall, there was strong support (jackknife support value 5 95) for monophyly of the group Haplosporidia,
provided that Bonamia spp. and M. roughleyi are included.
There was also strong support (jackknife support value 5 95)
for the placement of the abalone and spot prawn parasites at
the base of the haplosporidian clade.
It is interesting to note that introns were found within actin
genes from many of the haplosporidian species in this study.
The presence and the position of introns often lent further support to results of the phylogenetic analyses. Four intron positions were identified within the haplosporidian actin genes, with
the number of introns within a particular gene fragment ranging
from 1 to 3 within genes from the described species. At least
1 actin gene paralog with introns was isolated from each of the
Minchinia species. The actin gene from the undescribed C. floridana parasite contained an intron, consistent with its placement
within this genus. Introns were also found in the spot prawn
parasite actin gene at each of the 4 conserved positions found
in haplosporidian actin genes, lending additional support to the
phylogenetic affinity of this spot prawn parasite with Haplosporidia. Several of these introns were surprisingly short (17–33
bp). Even these short introns, however, had characteristic splice
junction sequences and began at their 59 ends with the dinucleotide GT and ended at their 39 ends with the dinucleotide AG.
In addition, when the intron sequences were removed, the remaining exon sequences could be translated in frame to yield
typical actin amino acid sequences, further supporting the validity of even the shortest introns. Introns were also found in
actin genes from a cercozoan species (Keeling, 2001), the group
found to be sister to Haplosporidia in the combined data set
analysis, although the intron positions differed from those seen
in the haplosporidian actin genes. To our knowledge, actin
genes have not yet been isolated from Bonamia spp., M. roughleyi, or the abalone parasite, and although we attempted, we did
not successfully isolate actin genes from the Australian Urosporidium sp., H. pickfordi, or H. lusitanicum. It is quite likely
that several actin gene paralogs from these parasites have yet
to be isolated. Phylogenetic analyses with these sequences as
well as characterization of the intron structure may lend additional insight into relationships within the group.
Overall, the results of this molecular study raise interesting
questions regarding spore formation for taxa within the group
Haplosporidia. Spores have not been observed for either the
spot prawn or abalone parasites that lie at the base of the clade
REECE ET AL.—MOLECULAR PHYLOGENY OF THE HAPLOSPORIDIA
(Bower and Meyer, 2002; Diggles et al., 2002; Hine et al.,
2002), suggesting that the ancestral state included a lack of the
ability to form spores and that spore formation arose along the
lineage to the Urosporidium, Haplosporidium, and Minchinia
genera. Spores have not been observed, however, for the Bonamia spp. and M. roughleyi, which in the phylogenetic analyses
fell between the spore-forming genera Minchinia and Haplosporidium. This hypothesizes a loss of the ability to form spores
in the microcell group. It is possible, however, that the life
stages of these undescribed basal parasites identified to date and
the microcells are intermediate stages of previously undiscovered haplosporidians and that the spore stages have not yet been
observed. Alternatively, the ancestor to the Haplosporidia had
the ability to form spores, but this was lost in the lineages to
the spot prawn and abalone parasites, as well as in the lineage
to the microcells, whereas the other taxa have retained this ability.
Relationships within the group Haplosporidia hypothesized
by the molecular phylogenetic analyses provide a framework
for assessing proposed relationships within the group based on
morphology. Perkins (2000) assigned species to Minchinia if
they had spore extensions that were visible with the light microscope and to Haplosporidium if the spore ornamentation was
not visible with the light microscope. Most other researchers
have followed Ormières’ (1980) criteria and assigned species
to Minchinia if the spore ornamentation was composed of epispore cytoplasm and to Haplosporidium if the spore ornamentation was derived from the spore wall (McGovern and Burreson, 1990; Hine and Thorne, 1998; Azevedo et al., 1999; Burreson, 2001). The molecular phylogenetic analyses presented
here support the importance of ornamentation origin. All the
species included in the analysis that have ornamentation composed of epispore cytoplasm (Minchinia spp.) formed a monophyletic group. The species that have ornamentation derived
from the spore wall (Haplosporidium spp.) formed a paraphyletic clade, suggesting that more genera are necessary to encompass the morphological diversity in species with spore
wall–derived ornamentation. Unfortunately, at present, there are
insufficient data on spore wall ornamentation for many of the
haplosporidian species to propose new generic assignments. As
knowledge increases, it will be interesting to assess the concordance between molecular and morphological data sets.
ACKNOWLEDGMENTS
The authors thank Brenda S. Kraus, Kathleen Apakupakul, and Karen
Hudson for technical support and Antonio Villalba, Barbara Nichols,
Susan Bower, Gary Meyer, Ben Diggles, and John Walker for providing
samples. This work was supported by grant BIO-DEB-9629487 from
the U.S. National Science Foundation. VIMS contribution 2617.
LITERATURE CITED
ALTSCHUL, S. F., W. GISH, W. MILLER, E. W. MYERS, AND D. J. LIPMAN.
1990. Basic local alignment search tool. Journal of Molecular Biology 215: 403–410.
AZEVEDO, C., J. MONTES, AND L. CORRAL. 1999. A revised description
of Haplosporidium amoricanum, parasite of Ostrea edulis L. from
Galicia, northwestern Spain, with special reference to the sporewall filaments. Parasitology Research 85: 977–983.
BERTHE, F. C. J., F. LE ROUX, E. PEYRETAILLADE, P. PEYRET, D. RODRIGUEZ, M. GOUY, AND C. P. VIVARÉS. 2000. Phylogenetic analysis of
the small subunit ribosomal RNA of Marteilia refringens validates
1121
the existence of phylum Paramyxea (Desportes and Perkins, 1990).
Journal of Eukaryotic Microbiology 47: 288–293.
BOWER, S. M., AND G. R. MEYER. 2002. Morphology and ultrastructure
of a protistan pathogen in the haemolymph of shrimp (Pandalus
spp.) in the northeastern Pacific Ocean. Canadian Journal of Zoology 80: 1055–1068.
BURRESON, E. M. 2001. Spore ornamentation of Haplosporidium pickfordi Barrow, 1961 (Haplosporidia), a parasite of freshwater snails
in Michigan, USA. Journal of Eukaryotic Microbiology 48: 622–
626.
———, N. A. STOKES, AND C. S. FRIEDMAN. 2000. Increased virulence
in an introduced pathogen: Haplosporidium nelsoni in the eastern
oyster Crassostrea virginica. Journal of Aquatic Animal Health 12:
1–8.
CARLINI, D. B., K. S. REECE, AND J. E. GRAVES. 2000. Actin gene family
evolution and the phylogeny of coleoid cephalopods (Mollusca:
Cephalopoda). Molecular Biology and Evolution 17: 1353–1370.
CARNEGIE, R. B., B. J. BARBER, S. C. CULLOTY, A. J. FIGUERAS, AND D.
L. DISTEL. 2000. Development of a PCR assay for detection of the
oyster pathogen Bonamia ostreae and support for its inclusion in
the Haplosporidia. Diseases of Aquatic Organisms 42: 199–206.
CAULLERY, M. 1953. Appendice aux Sporozoaires. Classe des haplosporidies. In Protozoaires: Rhizopodes, Actinopodes, Sporozoaires,
Cnidospories, P. P. Grassé (ed.). Traité de Zoologie, vol. I, no. 2.
Masson et Cie, Paris, France, p. 922–934.
———, AND F. MESNIL. 1899. Sur le genera Aplosporidium (nov.) et
l’ordre nouveau des Aplosporidies. Compte rendu des seances de
la Societe de Biologie et de ses Filiales 51: 789–791.
CAVALIER-SMITH, T. 1993. Kingdom Protozoa and its 18 phyla. Microbiological Reviews 57: 953–994.
———. 2002. The phagotrophic origin of eukaryotes and phylogenetic
classification of Protozoa. International Journal of Systematic and
Evolutionary Biology 52: 297–354.
———, AND E. E. CHAO. 2003. Phylogeny of Choanozoa, Apusozoa
and other Protozoa and early eukaryote megaevolution. Journal of
Molecular Evolution 56: 540–563.
COCHENNEC-LAUREAU, N., K. S. REECE, F. C. J. BERTHE, AND P. M. HINE.
2003. Revisiting Mykrocytos roughleyi taxonomic affiliation leads
to the genus Bonamia (Haplosporidia). Diseases of Aquatic Organisms 54: 209–217.
CORLISS, J. O. 1994. An interim utilitarian (‘‘user-friendly’’) hierarchical
classification and characterization of the protists. Acta Protozoologica 33: 1–51.
DESPORTES, I., AND N. N. NASHED. 1983. Ultrastructure of sporulation
in Minchinia dentali (Arvy), an haplosporean parasite of Dentalium
entale (Scaphopoda, Mollusca): Taxonomic implications. Protistologica 19: 435–460.
———, AND F. O. PERKINS. 1990. Phylum Paramyxea. In Handbook of
Protoctista, L. Margulis, J. O. Corliss, M. Melkonian, and D. J.
Chapman (eds.). Jones and Bartlett Publishing, Boston, Massachusetts, p. 30–35.
DIGGLES, B. K., J. NICHOL, P. M. HINE, S. WAKEFIELD, N. COCHENNECLAUREAU, R. D. ROBERTS, AND C. S. FRIEDMAN. 2002. Pathology of
cultured paua (Haliotis iris) infected by a novel haplosporidian parasite, with some observations on the course of disease. Diseases of
Aquatic Organisms 50: 219–231.
FARRIS, J. S., V. A. ALBERT, M. KÄLLERSJÖ, D. LIPSCOMB, AND A. G.
KLUGE. 1996. Parsimony jackknifing outperforms neighbor-joining.
Cladistics 12: 99–124.
———. 1998. Xac. Program and documentation. Swedish Museum of
Natural History, Stockholm, Sweden.
FLORES, B. S., M. E. SIDDALL, AND E. M. BURRESON. 1996. Phylogeny
of the Haplosporidia (Eukaryota: Alveolata) based on small subunit
ribosomal RNA gene sequence. Journal of Parasitology 82: 616–
623.
HINE, M., AND T. THORNE. 1998. Haplosporidium sp. (Haplosporidia) in
hatchery-reared pearl oysters, Pinctada maxima (Jameson, 1901),
in north Western Australia. Journal of Invertebrate Pathology 71:
48–52.
HINE, P. M., S. WAKEFIELD, B. K. DIGGLES, V. L. WEBB, AND E. W.
MAAS. 2002. Ultrastructure of a haplosporidian containing Rickettsiae, associated with mortalities among cultured paua Haliotis iris.
Diseases of Aquatic Organisms 49: 207–19.
1122
THE JOURNAL OF PARASITOLOGY, VOL. 90, NO. 5, OCTOBER 2004
KEELING, P. J. 2001. Foraminifera and Cercozoa are related in actin
phylogeny: Two orphans find a home? Molecular Biology and Evolution 18: 1551–1557.
KUDO, R. R. 1971. Protozoology, 5th ed. Charles C. Thomas Publishing,
Springfield, Massachusetts, 1,174 p.
MAIDAK, B. L., J. R. COLE, T. G. LILBURN, C. T. PARKER JR., P. R.
SAXMAN, R. J. FARRIS, G. M. GARRITY, G. J. OLSEN, T. M. SCHMIDT,
AND J. M. TIEDJE. 2001. The RDP-II (Ribosomal Database Project).
Nucleic Acids Research 29: 173–174.
MCGOVERN, E. R., AND E. M. BURRESON. 1990. Ultrastructure of Minchinia sp. from shipworms (Teredo spp.) in the Western North Atlantic, with a discussion of taxonomy of the Haplosporidiidae. Journal of Protozoology 37: 212–218.
MEDLIN, L., H. J. ELWOOD, S. STICKEL, AND M. L. SOGIN. 1988. The
characterization of enzymatically amplified eukaryotic 16S-like
rRNA-coding proteins. Gene 71: 491–499.
ORMIÈRES, R. 1980. Haplosporidium parisi n. sp., Haplosporidie parasite de Serpula vermicularis L. étude ultrastructurale de la spore.
Protistologica 16: 467–474.
PERKINS, F. O. 1990. Phylum Haplosporidia. In: Handbook of Protoctista, L. Margulis, J. O. Corliss, M. Melkonian, and D. J. Chapman
(eds.). Jones and Bartlett Publishing, Boston, Massachusetts, p. 19–
29.
———. 1991. ‘‘Sporozoa’’: Apicomplexa, Microsporidia, Haplosporidia, Paramyxea, Myxosporidia, and Actinosporidia. In Microscopic
anatomy of invertebrates, vol. 1, F. W. Harrison and J. O. Corliss
(eds.). Wiley-Liss, New York, New York, p. 261–331.
———. 2000. Phylum Haplosporidia Caullery and Mesnil, 1899. In
Illustrated guide to Protozoa, 2nd ed., J. J. Lee, G. F. Leedale, and
P. Bradbury (eds.). Society of Protozoologists, Lawrence, Kansas,
p. 1328–1341.
REECE, K. S., M. E. SIDDALL, E. M. BURRESON, AND J. E. GRAVES. 1997.
Phylogenetic analysis of Perkinsus based upon actin gene sequences. Journal of Parasitology 83: 417–423.
———, AND N. A. STOKES. 2003. Molecular analysis of a haplosporidian parasite from cultured New Zealand abalone Haliotis iris. Diseases of Aquatic Organisms 53: 61–66.
RENAULT, T., N. A. STOKES, B. CHOLLET, N. COCHENNEC, F. BERTHE, AND
E. M. BURRESON. 2000. Haplosporidiosis in the Pacific oyster, Crassostrea gigas, from the French Atlantic coast. Diseases of Aquatic
Organisms 42: 207–214.
SIDDALL, M. E., N. A. STOKES, AND E. M. BURRESON. 1995. Molecular
phylogenetic evidence that the phylum Haplosporidia has an alveolate ancestry. Molecular Biology and Evolution 12: 573–581.
SPRAGUE, V. 1979. Classification of the Haplosporidia. Marine Fisheries
Review 41: 40–44.
SWOFFORD, D. L. 2002. PAUP*: Phylogenetic analysis using parsimony
(*and other methods), version 10. Sinauer Associates, Sunderland,
Massachusetts.
THOMPSON, J. D., D. G. HIGGINS, AND T. J. GIBSON. 1994. Improving the
sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix
choice. Nucleic Acids Research 22: 4673–4680.