Biology

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Assignment: Using the paper as a guide, discuss the importance of this study relative to our discussions of endosymbiotic theory during class. We briefly discussed the Chlorarachniophyta in class; how are the chlorarachniophytes pertinent/important to the discussion of plastid acquisition? This should be a written essay, 500-1000 words in length. Thoroughly read the paper (you may need to read it more than once) and think about your response before beginning the project. Feel free to do additional research to augment your discussion. I am leaving the assignment fairly vague to give you freedom to explore the topic. I am not interested in 10 page papers—think three c’s, clear, concise, complete! Eighty percent of your grade will be based on the quality and thoughtfulness of your discussion. The other 20% will be based on the quality of your writing and on proper citations. Remember, we are using CSE, not MLA or any other format.

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http://www.elsevier.de/protis
Published online date 7 August 2006

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Correspondi

n

fax +81 29 853
e-mail iinouye
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& 2006 Elsev
doi:10.1016/j

157, 401—419, August 2006

Protist, Vol.

ORIGINAL PAPER

Hatena arenicola gen. et sp. nov., a Katablepharid
Undergoing Probable Plastid Acquisition

Noriko Okamoto2, and Isao Inouye1

Graduate School of Life and Environmental Sciences, University of Tsukuba, 1-1-1, Tennodai, Tsukuba,
Ibaraki 305-8572, Japan

Submitted February 27, 2006; Accepted May 27, 2006
Monitoring Editor: Robert A. Andersen

Hatena arenicola gen. et sp. nov., an enigmatic flagellate of the katablepharids, is described. It shows
ultrastructural affinities to the katablepharids, including large and small ejectisomes, cell covering,
and a feeding apparatus. Although molecular phylogenies of the 18S ribosomal DNA support its
classification into the katablepharids, the cell is characterized by a dorsiventrally compressed cell
shape and a crawling motion, both of which are unusual within this group. The most distinctive feature
of Hatena arenicola is that it harbors a Nephroselmis symbiont. This symbiosis is distinct from
previously reported cases of ongoing symbiosis in that the symbiont plastid is selectively enlarged,
while other structures such as the mitochondria, Golgi body, cytoskeleton, and endomembrane
system are degraded; the host and symbiont have developed a morphological association, i.e., the
eyespot of the symbiont is always at the cell apex of Hatena arenicola; and only one daughter cell
inherits the symbiont during cell division, resulting in a symbiont-bearing green cell and a symbiont-
lacking colorless cell. Interestingly, the colorless cells have a feeding apparatus that corresponds to
the location of the eyespot in symbiont-bearing cells, and they are able to feed on prey cells. This
indicates that the morphology of the host depends on the presence or absence of the symbiont. These
observations suggest that Hatena arenicola has a unique ‘‘half-plant, half-predator’’ life cycle; one cell
divides into an autotrophic cell possessing a symbiotic Nephroselmis species, and a symbiont-lacking
colorless cell, which later develops a feeding apparatus de novo. The evolutionary implications of
Hatena arenicola as an intermediate step in plastid acquisition are discussed in the context of other
examples of ongoing endosymbioses in dinoflagellates.
& 2006 Elsevier GmbH. All rights reserved.

Key words: Hatena arenicola; Katablepharidophyta/Kathablepharida; Nephroselmis symbiont; plant
evolution; plastid acquisition via secondary endosymbiosis; ultrastructure.

Abbreviations: EM ¼ electron microscopy; ER ¼
endoplasmic reticulum; ICBN ¼ International Code
of Botanical Nomenclature; ICZN ¼ International
Code of Zoological Nomenclature; LM ¼ light mi-
croscopy; SEM ¼ scanning electron microscopy;
SSU rDNA ¼ small subunit ribosomal DNA; TEM ¼
transmission electron microscopy.

g author;
4533

@sakura.cc.tsukuba.ac.jp (I. Inouye).
ess: School of Botany, University of Melbourne,
ria, Australia.

ier GmbH. All rights reserved.
.protis.2006.05.011

http://www.elsevier.de/protis

http://www.elsevier.de/protis

mailto:iinouye@sakura.cc.tsukuba.ac.jp

dx.doi.org/10.1016/j.protis.2006.05.011

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402 N. Okamoto and I. Inouye

Introduction

Eukaryotes are currently classified into five or six
supergroups (Baldauf et al. 2000; Baldauf 2003;
Bapteste et al. 2002; Nozaki et al. 2003; Simpson
and Roger 2002), and eukaryotic autotrophs (e.g.,
plants and algae) randomly scatter across those
supergroups. Eukaryotic autotrophs comprise
nine distinct divisions in cell architecture, and this
enormous diversity is explained by several en-
dosymbiotic events (Bhattacharya et al. 2004;
Falkowski et al. 2004; McFadden 2001). It is
widely accepted that a primary endosymbiosis
between a eukaryote and a cyanobacterial sym-
biont gave rise to the three extant primary
eukaryotic autotrophs, Glaucophyta, Rhodophyta,
and Viridiplantae ( ¼ land plants plus green algae)
(see Marin et al. (2005) for an alternative primary
endosymbiosis). Subsequently, secondary endo-
symbioses occurred between green or red algae
and heterotrophic eukaryotic hosts. Two algal
divisions (Euglenophyta and Chlorarachniophyta)
acquired the plastids of green algae, while four
algal divisions (Heterokontophyta, Haptophyta,
Cryptophyta, and Dinophyta) and one parasitic
phylum (Apicomplexa) acquired those of red algae
(although some Dinophyta lost their original
plastid and remained colorless or re-acquired
different plastids as discussed below). An esti-
mated two-thirds of today’s algal diversity resulted
from secondary endosymbioses (Falkowski et al.
2004; Graham and Wilcox 2000), and thus this
process is important in understanding the evolu-
tionary process of plant and algal diversification.

The transition of a symbiont to a plastid involves
a series of changes in both the host and the
symbiont (Cavalier-Smith 2003; Hashimoto 2005;
van der Giezen et al. 2003), which include the
establishment of a specific partner alga, lateral
gene transfer from the symbiont to the host’s
nucleus (Katz 2002), the development of protein-
transport machinery to carry proteins from the
host cytoplasm to the symbiont (van Dooren et al.
2001), and synchronization of cell cycles so that
the symbiont can be passed to host daughter cells
during host cell division.

Evidence about plastid integration is accumu-
lating (Andersson and Roger 2002; Archibald et al.
2003; Hackett et al. 2004b; Huang et al. 2003;
Martin and Herrmann 1998; Martin et al. 2002;
Martin 2003a, b; Nozaki et al. 2004; Stegemann
et al. 2003), however, the intermediate steps in this
process remain largely unknown. Some organisms
appear to be in an intermediate stage of plastid
acquisition, the best-known examples of which

are the Cryptophyta and Chlorarachniophyta,
whose plastids contain a vestige of the symbiont
nucleus termed a nucleomorph (e.g. Douglas et al.
2001; Gilson et al., 2006). They are thought to
represent a late stage of integration. Early
stages of plastid acquisition can be found in
the dinoflagellates (for reviews, Hackett et al.
2004a; Morden and Sherwood 2002; Schnepf and
Elbrächter 1999), where the most dramatic
changes are ongoing. The original plastids of
dinoflagellates have been of red algal origin,
though some dinoflagellates subsequently lost
their original red-algal plastids, which were re-
placed by new ones via extra secondary or tertiary
endosymbioses. These examples probably reflect
stepwise changes in symbiotic conditions during
integration (e.g. Hackett et al. 2004a), and are
useful to understand the plastid acquisition
process.

We discovered an undescribed flagellate, Hate-
na arenicola gen. et sp. nov., in October 2000, in
an intertidal sandy beach in Japan. The organism
appears to be in the process of plastid acquisition.
Most cells of H. arenicola in the natural population
have a green plastid-like structure with a red
eyespot at the cell apex, though it is inherited by
only one of the daughter cells during cytokinesis
(Okamoto and Inouye 2005a). Molecular phyloge-
netic analysis of small subunit ribosomal DNA
(SSU rDNA) and ultrastructural observations of the
plastid-like structure reveal that it is not a plastid
but an autotrophic endosymbiont belonging to the
genus Nephroselmis Stein (Prasinophyceae, Vir-
idiplantae). We previously reported the symbiotic
nature of this association (Okamoto and Inouye
2005a). This paper describes the organism as a
new genus and species of katablepharid, a group
of flagellates recently designated the phylum
Kathablepharida, division Katablepharidophyta
(Okamoto and Inouye 2005b). We compare the
symbiosis of H. arenicola with other examples of
secondary symbioses in dinoflagellates to help
elucidate the intermediate steps in the plastid
acquisition process.

Results

Description

Hatena arenicola Okamoto et Inouye gen. et sp.
nov.

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403Hatena arenicola: Halfway to a Plant?

Latin Diagnosis

Cellulae oblongae secus axem dorsiventrem valde
appresae, sine chromatophoro nec vacuola con-
tractili; 30-40mm longae; 15-20mm latae; ventraliter
subapicali cum sulco vadoso longitudinali 3-4mm
longa et ca. 2mm lata; flagellis crassis binis
inaequalibus in sulco insertis; flagello anteriore
longiore, altero posteriore breviore; ejectisomis
conspicuis distichis prope flagellas longitudinaliter
positis; plerumque cum 1-4 endosymbiontis viridis;
uni stigma endosymbionti situm ad apicem cellulae.

Holotype: Figure 1A
Type locality: Isonoura, Wakayama, Japan (Fig.

1 B-C)
Etymology:

Hatena ¼ ‘enigmatic’ in Japanese
arenicola ¼ ‘inhabiting sand’ in Latin

Light Microscopy

General Morphology: The cell is flattened along
the dorsiventral axis. In the ventral view, it is ovoid,
30-40mm long and 15-20mm wide (Fig. 1 A, D-F).

Figure 1. Hatena arenicola gen. et sp. nov. A. Ventral v
and an eyespot of the symbiont (arrowhead). B,C. Sam
showing two rows of conspicuous Type I ejectisomes
‘‘immature’’ symbiont. G-L. Cell division in Hatena arenic
symbiont. Each panel shows a different individual at a di
The scale bar is 10mm in A, D-L.

The cell has a furrow in the subapical region, 3-
4mm long and ca. 2 mm wide (Fig. 1 D). The long
anterior flagellum and shorter posterior flagellum
emerge from this furrow, and two rows of
ejectisomes are easily visible near the posterior
end of the furrow (Fig. 1 D). One large nucleus is
located in the middle posterior region of the cell,
and the rest of the cytoplasm is mostly occupied
by the plastid of the green symbiont. Cells only
rarely lacked the symbiont (Fig. 1 E), though some
symbionts were not fully developed (Fig. 1 F; see
Discussion).

Cell Division: During cell division, one daughter
cell inherits the Nephroselmis symbiont while
the other does not and becomes colorless. Figure
1 G-L shows cell division in H. arenicola (ventral
view). First, the host nucleus moves to the apex
between the flagellar insertion and the eyespot
(Fig. 1 G). The symbiont contracts to the left side of
the host cell (on the viewer’s right in figures) so that
the left half of the cell remains green, while the right
half becomes colorless (Fig. 1 G). Two new flagella
are formed, and one set moves from the right side of
the nucleus to the left (flagellar transformation;

iew of a symbiont-bearing cell showing two flagella
pling site. D. The same cell in a different focal plane,
. E. A cell lacking the symbiont. F. A cell with an
ola, where the arrowhead indicates an eyespot of the
fferent stage in cell division. N: nucleus. S: Symbiont.

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Figure 2. Hatena arenicola and its symbiont. DIC
images are shown in upper column (A, C, E, G) and
the fluorescent images of the same cells are shown
in lower column (B, D, F, H respectively). Arrow-
heads indicate the eyespot. Blue: DAPI-stained
nuclei of Hatena arenicola (large fluorescence in
the center of the cell) and of the symbiont (smaller
dots). Red: Autofluorescence of the symbiont
plastid. The scale bar is 10mm.

0

0.2

0.6

1

ab
so

rp
tio

n

450 500 550 600 650 700
wavelength

Symbiont of

Hatena arenicola

Chl. b

Chl. b

Chl. a

Chl. a

Figure 3. Microspectrophotometry of the symbiont
plastid. Spectrogram shows absorbance similar to
chlorophyll a/b-containing plastids.

404 N. Okamoto and I. Inouye

Fig. 1 H). The chromosomes separate (Fig. 1 I).
Following nuclear division (Fig. 1 J), cytokinesis
results in one green cell with the symbiont and
one colorless cell without it (Fig. 1 K-L).

Fluorescence Microscopy: Figure 2 A-H shows
DIC and fluorescence images of the same cells.
DAPI label (blue) indicates a large host nucleus in
the cell center and one to four smaller symbiont
nuclei (Fig. 2 B,D,F,H). Each symbiont nucleus is
independent of all others and is surrounded by
plastid(s), shown in red (due to autofluorescence).
Interestingly, the cell with multiple symbiont nuclei
has only a single eyespot at the apex of the host
cell (Fig. 2 E-G; arrowheads).

Microphotometry: Microphotometry of seven
cells shows an absorption pattern characteristic
of plastids with chlorophyll a/b. Average absorp-
tion is shown in Figure 3. The prominent peaks at
435 and 678 nm represent chlorophyll a absorp-
tion, while the smaller peaks at 470 and 650 nm
correspond to chlorophyll b absorption.

Uptake of Prey Cells and Symbiont Specifici-
ty: Molecular phylogenetic analysis of 16S rDNA

indicates that the symbiont is a member of
Nephroselmis (Okamoto and Inouye 2005a). In
feeding experiments using a Nephroselmis strain
(NIES1417; different from that of the symbiont in
16S rDNA sequence; data not shown), colorless
cells of H. arenicola phagocytotically engulfed the
alga (Fig. 4 A-I) and tentatively maintained it.
However, none developed fully, likely because
the strain used in the experiments was not the
exact symbiont of H. arenicola. This suggests that
symbiont specificity is at the species or strain
level. As the H. arenicola cells which engulfed
Nephroselmis NIES1417 died, it is unclear whether
the Nephroselmis cells were digested.

Crawling Motion: Hatena arenicola displays a
conspicuous crawling motion that makes it easy
to recognize this organism in a crude sample.
Figure 5 A-H illustrates cell motion and flagellar
movement. The anterior flagellum produces most
of the propulsion, while the posterior flagellum is
used to change direction. To move forward, a cell
casts the anterior flagellum, which adheres to the
substratum by its tip (Fig. 5 A-C), and pulls itself
forward (Fig. 5 D, E). It then anchors itself with the
posterior flagellum (Fig. 5 F-H), and repeats the
process. The tip of the flagellum is sharply bent
while it is attached to the substratum, as shown in
the supplemental material and in Figure 5 I. The
cell bends the posterior flagellum to change
direction (not shown).

Electron Microscopy

Electron microscopy (EM) revealed that the green
plastid-like structure in the cell is a eukaryotic

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Figure 4. Uptake of Nephroselmis (NIES1417) by Hatena arenicola. A-H were taken at 6-s intervals. I:
corresponds to Hatena arenicola and the symbiont of each frame.

Figure 5. Crawling motion of Hatena arenicola.
High-speed video images recorded at 0 ms (A),
60 ms (B), 100 ms (C), 130 ms (D), 220 ms (E),
280 ms (F), 300 ms (G), 340 ms (H), respectively. I.
SEM image showing the anterior flagellum sharply
bent at the distal end (arrowhead). The scale bar is
10 mm in I.

405Hatena arenicola: Halfway to a Plant?

endosymbiont. A single nucleus, usually dorsiven-
trally flattened, is located in the middle posterior
region of the cell, and its chromatin is always
condensed and electron dense (Fig. 6 A,B). Multi-
ple mitochondrial profiles are present throughout
the cytoplasm, though these could be different
sections of a single, large reticulate mitochon-
drion. Mitochondrial cristae are tubular (Fig. 6 C),
and there is a single large Golgi body near the
groove of flagellar insertion between the nucleus
and the flagellar apparatus (Fig. 6 D).

Ejectisomes: There are two types of ejecti-
somes beneath the membrane: large, Type I
ejectisomes sensu Vørs 1992b (Fig. 6 D-E) that
are arrayed in two rows near the flagellar insertion
(Figs 1 D, 6 F-G); and smaller Type II ejectisomes

sensu Vørs 1992b (Fig. 6 H-I) that are arrayed in
numerous rows all over the cell (Fig. 6 F-G). Each
ejectisome consists of a coiled ribbon with a
gradual depression (Fig. 6 D, H), and bound by a
single membrane. Discharged ribbons are
slightly curved (Fig. 6 J; type I ejectisome). The
ribbon does not have any kink, unlike those of
Cryptophyta.

Type II ejectisomes are situated beneath the
plasma membrane of the cell (Fig. 7 A,B) except
for a small smooth area in the apical region, below
which the eyespot is situated (asterisk in Fig. 6 G).
The ejectisomes are spaced regularly between
longitudinally oriented cytoskeletal microtubular
bundles (arrowheads in Fig. 7 A).

Hatena arenicola lacks Type III ejectisomes that
are characteristic of the genus Leucocryptos (Vørs
1992b).

Cellular and Flagellar Covering: The plasma
membrane of each cell has a characteristic
cellular covering composed of a thick inner basal
layer (asterisk in Fig. 7 B) and an outer layer of
electron-opaque material (arrowhead in Fig. 7 B).
This structure extends to the flagellar surface. The
outer layer comprises a regularly arrayed and
spiraling envelope around the whole flagellum
(Fig. 7 C).

Endoplasmic Reticulum: The endoplasmic re-
ticulum (ER) does not form a conspicuous stack
but is loosely distributed throughout the cell. The
rough ER extends beneath the surface of the cell
(double arrow in Fig. 7 B).

Flagella and Basal Bodies: The long anterior
and shorter posterior flagella emerge from a

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Figure 6. Ultrastructure of Hatena arenicola. A. A longitudinal section of a Hatena arenicola cell showing a
nucleus (N). Most of the nuclear content is condensed. The rest of the cytoplasm is occupied by the symbiont
(Sym). The symbiont has a large plastid region with multiple pyrenoids. B. A transverse section of Hatena
arenicola showing host nucleus (N) and the symbiont (Sym) with the vestigial cytoplasm (asterisk). C.
Mitochondrial profiles showing tubular cristae. D. A Golgi body near the flagellar basal bodies (arrows) and
Type I ejectisomes sensu Vørs 1992 (nearly longitudinal view). E. A transverse section of a Type I ejectisome
(sensu Vørs 1992). F. SEM image of Hatena arenicola showing Type I ejectisomes near the flagellar insertion
as well as smaller ejectisomes (Type II ejectisomes sensu Vørs 1992) regularly arrayed over the cell surface.
G. Magnified SEM image of the same cell. Note that the apical region of the cell (asterisk) lacks ejectisomes.
H. A longitudinal section of Type II ejectisome. I. A transverse section of Type II ejectisome. J. A whole mount
TEM image of a discharged type I ejectisome. The scale bar is 10 mm in A, E; 2 mm in B, G; 50 nm in C; 500 nm
in D, F; 1mm in I; 200 nm in H, I.

406 N. Okamoto and I. Inouye

ARTICLE IN PRESS

Figure 7. Surface structure and flagellar transition
region of Hatena arenicola. A. A tangential section of
the surface of Hatena arenicola. Cytoskeletal micro-
tubular bundles (arrows) are arrayed longitudinally
below the cell surface; Type II ejectisomes are
situated between them. Each pore corresponds to
the position of an ejectisome, where the surface
sheath is thin. B. A transverse section of the cell
periphery showing a bilayered surface sheath. An
arrowhead indicates the outer layer. An asterisk
indicates the thick basal layer. An arrow indicates
the plasma membrane. A double arrowhead indi-
cates a profile of rough ER. C. A tangential section of
the flagellum shows the outer layer of the surface
sheath enveloping the flagellum in a spiraling
fashion. D-J. Flagellar transition region of Hatena
arenicola. D. Diagram of flagellar transition region
reconstructed based on serial ultrathin sections.
Each letter in D indicates which part corresponds to
one of the transverse sections (E-J). The scale bar is
500 nm in A, C; 200 nm in B, E-J.

407Hatena arenicola: Halfway to a Plant?

shallow subapical furrow. The flagella are coated
by a ‘‘surface sheath’’. The diagram of the basal
body and the flagellar transition zone (Fig. 7 D) is
based on serial sections of the flagellar transition
zone (Fig. 7 E-J). At the proximal end of the basal
body, there is a cartwheel structure with a central
tube. The triplet microtubules of the basal body
terminate below the plasma membrane (Fig. 7 E),
whereas the doublet microtubules attach to the
plasma membrane by a connecting fiber (Fig. 7 F).
The flagellar transition region extends above the
plasma membrane, and an electron-dense rod

structure is present (Fig. 7 G) in the middle of this
region. Surrounding the transition region are outer
doublet microtubules lined in a loose, electron-
dense material (Fig. 7 H). The flagellar transition
region ends in a terminal plate with electron-
opaque material at the center (Fig. 7 I). The
axonemal central pair begins above the terminal
plate (Fig. 7 J).
Feeding Apparatus: As reported in Okamoto
and Inouye (2005a), colorless cells lacking the
symbiont have a complex feeding apparatus at
their apex (Fig. 8 A-C), consisting of transverse
tubular rings (arrowheads in Fig. 8 B-C) and
longitudinal microtubules arrayed in a single layer
(arrow in Fig. 8 B-C). These microtubules are
different from those that form the cytoskeleton
(double arrowheads in Fig. 8 A). Inside the micro-
tubular skeletons of the feeding apparatus are
several electron-opaque granules, some of which
are large and elongate (light gray) and others that
are smaller, granulated, and pigmented (Fig. 8 A,
C). These granules are restricted to the feeding
apparatus, never seen elsewhere in the cell.
Symbiont-bearing cells do not have a feeding
structure; the corresponding region is occupied by
the eyespot of the symbiont (Fig. 8 D-E).
Symbiont: The symbionht retains not only a
plastid but also its own cytoplasm with a nucleus
and mitochondrion(a) (Fig. 9 A-C). Most symbiont
cells contain one nucleus, which is often attached
to the membrane, and symbiont and host nuclei
often face each other (Fig. 9 A-C). The symbiont is
bounded by a single membrane (arrowhead in
Fig. 9 D), whose origin is unknown. Mitochondria
have flat, often degraded cristae, though the
extent of degradation varies among individual
cells (Fig. 9 E-F). The plastid, which is the largest
structure in the symbiont (Fig. 6 A-B), contains
multiple pyrenoids bounded by a thin starch
sheath (Fig. 9 G). The pyrenoid has shallow
invaginations of the thylakoid membrane. The
plastid has a single conspicuous eyespot, where
the morphological association between host and
symbiont is present (see below).

Free ribosomes are densely distributed through-
out the symbiont cytoplasm, though no ribosome-
bearing membrane (rough ER) is present. Occa-
sionally there are flattened, stacked membranes
next to the nucleus (Fig. 9 A). This structure would
normally be a Golgi body but it is present in a
degraded or inactive form, because no Golgi
vesicles were seen around the structure. Some
randomly shaped vacuoles of unknown origin are
also present (Fig. 9 B-C). Because each cell
division will result in half the population carrying

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Figure 8. Feeding apparatus of Hatena arenicola. A. A nearly transverse section of the feeding apparatus.
Feeding apparatus is distinct from cytoskeletal microtubules (double arrowheads). B. A magnified view
showing microtubules (arrow) regularly arrayed in a single layer along the external side of the tubular rings
(arrowheads). C. Longitudinal section of the feeding apparatus showing numerous transverse tubular rings
(arrowhead) and longitudinal microtubules arrayed in a single layer (arrows). D. An eyespot (e) of symbiont is
located at the corresponding place in the symbiont bearing cell. E. A schematic illustration of the feeding
apparatus and the corresponding place of the symbiont-bearing Hatena arenicola. The scale bar is 500 nm in
A, C-D; 250 nm in B.

408 N. Okamoto and I. Inouye

the symbiont, these morphological varieties likely
reflect degradation (see Discussion).

Cytoskeletal structures, including the flagella,
basal bodies, the flagellar apparatus, and micro-
tubular rootlets, are completely absent. These
morphological changes must affect intracellular
functions, such as protein synthesis and distribu-
tion in the symbiont (see Discussion).

The lysosome of the host cytosol is discontinuous
with the symbiont compartment. The lysosome
of some cells contains scales of Nephroselmis
and Pyramimonas (Fig. 10 A,B), indicating that
H. arenicola cells engulf other prey in addition to
its Nephroselmis partner. Because Pyramimonas
cells are digested, H. arenicola may be partly
heterotrophic (see Discussion).

Eyespot: The eyespot is composed of a single-
layered sheet of osmiophilic granules (Fig. 11 A)
that connects to the inner plastid envelope. Near
the eyespot, the plastid, symbiont, and host plasma
membranes are tightly layered (Fig. 11 A,B). In some
cells, single microtubules are aligned longitudinally
between the plasma membrane and the symbiont

membrane overlying the eyespot region (Mts in
Fig. 11 C,D). These single microtubules are distinct
from the cytoskeletal microtubular bundles (arrow-
head in Fig. 11 D), and the space between them
lacks Type II ejectisomes; this is consistent with the
smooth surface appearance of the eyespot region
(Fig. 6 F).

Molecular Phylogeny

Partial SSU rDNA sequences of H. arenicola
(AB212285) aligned with the homologues of
known eukaryotes were subjected to phylogenetic
analyses. The resulting maximum likelihood (ML)
tree (Fig. 12) showed the typical topology of the
SSU rDNA tree, and H. arenicola was included in
the katablepharid clade, which was robustly
supported with high bootstrap probability in ML
(99%), NJ (100%), and MP (99%) analyses.

Discussion

Hatena arenicola has several unique features that
suggest it is in the process of endosymbiosis with

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Figure 10. Lysosome of Hatena arenicola with
scales of prasinophytes characteristic to Pyramimo-
nas (A) and Nephroselmis (B) respectively. The scale
bar is 500 nm.

Figure 9. Ultrastructure of the symbiont. A-C. The symbiont cytoplasm, retaining a nucleus, mitochondria,
and sometimes a Golgi body-like vesicle (A) or membranes of random shape (B-C), likely in an intermediate
state of integration. D. The single symbiont-enveloping membrane (arrowhead) separates the symbiont
compartment (Sym) and host cytoplasm (H). Double membranes of the symbiont plastid are also shown
(arrows). E. Mitochondrial profiles that retain flat cristae. F. A relatively degenerated mitochondrial profile. G.
A pyrenoid, surrounded by a starch sheath. Random shallow invagination of the thylakoids. The scale bar is
1 mm in A-C; 250 nm in D-G.

409Hatena arenicola: Halfway to a Plant?

a Nephroselmis partner (Okamoto and Inouye
2005a). We will discuss the taxonomy and
classification of H. arenicola first, and then focus
on its unique endosymbiosis.

Taxonomy

Morphological and molecular analyses of H.
arenicola clearly show that it belongs to the
recently established division/phylum Katablephar-
idophyta (International Code of Botanical Nomen-
clature, ICBN)/Kathablepharida (International
Code of Zoological Nomenclature, ICZN) (Oka-
moto and Inouye 2005b). The katablepharids
comprise an ultrastructurally well-defined and
small group of heterotrophic flagellates that
includes 10 species in two genera: nine species
of Katablepharis Skuja (correct spelling in ICBN
which will be used in this paper)/Kathablepharis
Skuja (original spelling in the ICZN), and one
species of Leucocryptos (Braarud) Butcher. Kata-
blepharidaceae (ICBN) was originally described by
Skuja (1939) based on ovate or cylindrically ovate
cell shape, two flagella emerging from a subapical
depression, and conspicuous ejectisomes aligned

ARTICLE IN PRESS

Figure 11. Eyespot of the symbiont plastid. A. A longitudinal section of the eyespot (E). B. A magnified view
of another cell clearly shows the eyespot granules, the inner/outer envelop of the symbiont plastid (arrows),
the single symbiont enveloping membrane (arrowhead), and the host plasma membrane (double arrowhead)
associated with each other. C-D. Tangential sections of the eyespot region show single microtubules (Mts)
distinctive from the cytoskeletal microtubules (arrowheads) longitudinally situated between the eyespot (E)
and the plasma membrane. The scale bar is 250 nm in A; 100 nm in B; 1mm in C-D.

410 N. Okamoto and I. Inouye

in two rows near the flagellar insertion. Vørs
(1992b), and Clay and Kugrens (1999b) emended
the family by adding the following ultrastructural
features: the entire surface of the cell, including
the flagella, is coated with a bilayered surface
sheath that appears to form spiraling rows around
the cell body; tubular mitochondrial cristae; a
complex, truncated conical feeding apparatus and
cytoskeleton; a Golgi apparatus situated anteriorly
and a centrally located nucleus; and a food
vacuole in the posterior part of the cell.

Hatena arenicola shares all these characters,
except that the cell is dorsiventrally compressed
and the food vacuole is absent. We occasionally
observed a vacuole containing scales of prasino-
phytes, though its position was anterior to the
nucleus. In addition, the flagellar transition zone
containing a rod-shaped structure is fundamen-
tally the same as that of K. ovalis (Lee et al. 1992).
Based on these ultrastructural similarities and the

molecular phylogenetic data, H. arenicola un-
doubtedly belongs to the katablepharids.

Currently, all the katablepharids belong to a
single family, Katablepharidaceae, whose cells are
defined as either ‘‘oblong or cylindrically ovate’’
(Katablepharis; Clay and Kugrens, 1999b; Vørs,
1992b) or ‘‘ovate, pyriform elliptical outline’’
(Leucocryptos; Butcher 1967). The most important
distinguishing feature of Leucocryptos is the
presence of Type III ejectisomes, which have been
found only in Leucocryptos. Because H. arenicola
lacks Type III ejectisomes, it does not belong to
Leucocryptos.

Hatena arenicola is distinct from all other
katablepharids previously described in that it is
dorsiventrally compressed with a flat-oval shape,
which suggests that it does not belong to the
genus Katablepharis. Its crawling motion and the
feeding apparatus composed of single-layered
microtubules are also distinctive from the other

ARTICLE IN PRESS

0.1

Chlorarachnion CCMP242
Cercomonas longicauda

Heteromita globosa
Thaumatomonas seravini

Euglypha rotunda
Paulinella chromatophora

Ochromonas danica
Phytophthora megasperma

Skeletonema pseudocostatum
Pteridomonas danica

Prorocentrum micans
Prorocentrum mexicanum

Gymnodinium sp. MUCC284
Pfiesteria sp. B112456

Alexandrium minutum
Cryptosporidium parvum

Toxoplasma gondii
Prorodon teres

Platyophrya vorax
Emiliania huxleyi

Pavlova salina
Chlorokybus atmophyticus

Mesostigma viride
Arabidopsis thaliana

Fossombronia pusilla
Pyramimonas propulsa

Ulothrix zonata
‘Chlorella’ ellipsoidea
Tetraselmis striata

Coleochaete scutata
Closterium littorale
Cyanophora paradoxa

Cyanoptyche gloeocystis
Glaucocystis nostochinearum

Gloeochaete wittrockiana
Goniomonas truncata

Rhodomonas mariana
Hanusia phi

Geminigera cryophila
Cryptomonas ovata

Chroomonas sp. M1318
Hemiselmis brunnescens

Leukocryptos marina
Hatena arenicola

Katablepharis japonica
Heterophrys marina

Chlamydaster sterni
Raphidiophrys ambigua

Rhodella maculata
Stylonema alsidii

Bangia sp.
Porphyra umbilicalis

Acanthamoeba castellanii
Dictyostelium discoideum

Leptomyxa reticulata
Hartmannella vermiformis

Scutellospora cerradensis
Pneumocystis carinii

Saccharomyces cerevisiae
Schizosaccharomyces pombe

Basidiobolus haptosporus
Chytriomyces hyalinus

Monosiga brevicollis
Clathrina cerebrum

Cirripathes lutkeni

18S rDNA (65species 1252 sites)

100/100/99

99/83/96

64/73/62

54/86/60

58/73/-
52/88/72

53/98/67

100/89/100

100/100/100

99/100/100

99/88/56

56/67/-

100/100/100

90/97/93

86/86/58

54/61/80

100/100/100

-/73/-

100/100/100

58/63/-

99/99/97

67/72/89

72/54/74

99/100/99

95/76/81

76/-/-

100/100/100

82/85/78

75/83/74

91/79/84

100/100/100

75/88/66

99/97/83

75/64/-

65/85/89

74/-/-

76/81/84

93/-/-

-/-/54

53/-/-

50/59/-

92/77/79

ML/NJ/MP

Figure 12. Unrooted eukaryotic tree based on the SSU rDNA. The best tree of the ML method is shown.
Bootstrap propotions are shown at the internal branches, in the order of ML/NJ/MP methods. The length of
each branch is proportional to the estimated number of substitutions. Bar denotes 10% substitutions per site.
For details of the phylogenetic reconstruction methods, see text. The organisms included in the tree are listed
in Table 1. Unambiguously aligned 1252 nucleotide positions were used for the analysis.

411Hatena arenicola: Halfway to a Plant?

katablepharids (Clay and Kugrens 1999a, b;
Kugrens et al. 1994; Lee and Kugrens 1992; Lee
et al. 1991; Okamoto and Inouye 2005b; Vørs
1992a, b). Based on those features, we propose
that the organism should be assigned to a new
genus, Hatena.

The family-level taxonomy of the katablepharids
is still unclear, primarily because little molecular
sequence data exist. Until further studies eluci-
date katablepharid taxonomy, it is best to include
the genus Hatena in the family Katablepharida-
ceae.

ARTICLE IN PRESS

AH

Loss of
Feeding Appar atus

Plastid: Enlarged
Symtiont: Degraded

412 N. Okamoto and I. Inouye

Endosymbiosis

Endosymbiosis is a major driving force in plant
evolution, and thus it is important to understand
this process. Hatena arenicola may be an im-
portant model of early plastid acquisition. Sym-
biotic Nephroselmis differs from free-living
individuals in having enlarged plastids with a
greater number of pyrenoids, degraded subcellu-
lar structures, and morphologically distinct eye-
spots. Cells of known Nephroselmis species are a
maximum of 20mm in length (Nephroselmis
astigmatica; Inouye and Pienaar 1984) and pos-
sess a single plastid with a single pyrenoid. The
symbiont occupies most of the host cytoplasm,
suggesting that the symbiont plastid(s) grows
more than ten fold after being engulfed by the
host. Pyrenoids also multiply after being engulfed.
In contrast, the symbiont cytoplasm loses other
major cell components including flagellar appara-
tus and microtubular roots, endomembranes such
as the ER and transport vesicles, Golgi-like
vesicles, and amorphous membranous structures.
The dramatic growth of the plastid is in stark
contrast to the degradation of the other orga-
nelles. Because the cytoplasm is in such a
degraded state, it must be difficult to sustain the
growth and maintenance of the plastid alone. It is
likely that some metabolites from the host cell are
used to develop and maintain the plastid.

B

DE

F

G
C

Predator phase Plant phase

Formation of
Feeding apparatus

Uptake of
the partner

Figure 13. Half-plant, half-predator-hypothesis. The
solid line indicates a witnessed process, and the
broken line indicates a hypothetical process. A-D: A
green cell with the symbiont, lacking the feeding
apparatus (A) divides (B) into one green (C) and one
colorless cell (D). E-G: The colorless cell should form
a feeding apparatus de novo and engulfs a Ne-
phroselmis cell. G-H: The symbiont plastid selec-
tively grows in the host cytoplasm. Because cell
division of a colorless cell or a cell with an
‘‘immature’’ symbiont (H) has never been observed,
uptake and the subsequent changes in both
host and symbiont apparently occur within one
generation.

Eyespot Morphology

The morphology of the eyespot is most suggestive
of a host-symbiont coordination. The tight layering
of the four distinct membranes of different origin
implies functional cooperation. The eyespot is an
important component of a photo-sensing complex
found in various algal groups (Melkonian and
Robenek 1984; Gualtieri 2001). It effectively
regulates the light received by a photoreceptor
located on a nearby membrane, thereby allowing
the alga to detect light direction. Preliminary
observations have shown that lateral, and not
vertical, incidence, is important for the behavior of
H. arenicola. Hence, we speculate that the
photoreceptor and the regulatory mechanism
would be laterally aligned in the cell, so that the
eyespot can effectively shade the photorecepter
from the lateral incidence. Assuming that the
eyespot is functional in H. arenicola, the photo-
receptor should be situated in either of the outer or
inner plastid membrane, the single endosymbiont
envelope, or the plasma membrane. Their mor-
phological association can be explained as a

consequence of functional collaboration. Because
H. arenicola crawls two-dimensionally, a photo-
tactic response to laterally projected light is
plausible. To test whether a functional association
exists, further investigation of the threshold and
the efficiency of phototaxis in both colored and
colorless H. arenicola cells are required.

Morphological Changes of the Host and
‘‘Half-plant, half-predator’’ Model

Hatena arenicola cells without a symbiont have a
complex feeding apparatus in place of an eye-
spot, indicating that symbiont acquisition is
accompanied by drastic morphological changes
in both the symbiont and the host.

ARTICLE IN PRESS

413Hatena arenicola: Halfway to a Plant?

Such structural changes would necessitate life
cycle changes. Based on observations presented
in this paper, we propose the ‘‘half-plant, half-
predator’’ hypothesis, where H. arenicola switches
its lifestyle between that of a plant and a predator
(Fig. 13; see also Okamoto and Inouye 2005a for
detailed explanation). This proposed life cycle is
also supported by the observation that the extent
of degradation of symbiont mitochondria and
membranous structures varies among individual
cells. The presence of prasinophyte scales in a H.
arenicola lysosome suggests that H. arenicola
lives heterotrophically to some extent, perhaps
until it engulfs its symbiotic partner. This repre-
sents an intermediate state of trophic alteration.
Although there are several assumptions in the
model, it promises to lead to new insights and
helps elucidate the plastid integration process.

Evolutionary Implications

Extra secondary and tertiary endosymbioses in
dinoflagellates may represent evolutionary events
that occurred during plastid acquisition (for re-
views, see Hackett et al. 2004a; Morden and
Sherwood 2002; Schnepf and Elbrächter 1999).
The earliest stage is represented by the crypto-
phyte symbiont, in which only cytoskeletal com-
ponents are lost, as in Amphidinium latum Lebour
(Horiguchi and Pienaar 1992), A. poecilochroum
Larsen (Larsen 1988), and Gymnodinium acidotum
Nygaard (Fields and Rhodes 1991; Wilcox and
Wedemayer 1984). The cell cycles of the host and
symbiont are not synchronized, and the host cell
must repeatedly capture symbionts. The next
stage is represented by the symbiont of diatom
origin, in which various subcellular structures are
lost, except the nucleus, mitochondrion(a) and
plastid, as in Durinskia baltica (Levander) Carty et
Cox (formerly Peridinium balticum; Chesnick and
Cox 1987, 1989; Chesnick et al. 1997; Eschbach
et al. 1990; Tippit and Pickett-Heaps 1976; Tomas
and Cox 1973) and Kryptoperidinium foliaceum
(Stein) Lindemann (Dodge 1971; Eschbach et al.
1990). At this stage, the symbiont divides syn-
chronously with the host cell (Chesnick and Cox
1987, 1989; Tippit and Pickett-Heaps 1976), so
that the association between the host and
symbiont becomes permanent, and repeated
uptake of the symbiont is no longer necessary.
Finally, the symbiont cytoplasm is reduced, as
seen in Lepidodinium viride (Watanabe et al. 1987;
Watanabe et al. 1990), Gymnodinium chlorophor-
um (Elbrächter and Schnepf 1996), both with
symbionts of prasinophyte origin, and Karenia

brevis (Davis) Hansen et Moestrup, Karenia
mikimotoi (Miyake et Kominami ex Oda) Hansen
et Moestrup, Karlodinium veneficum (Leadbeater
et Dodge) Larsen with symbionts of haptophyte
origin (Daugbjerg et al., 2000; Inagaki et al., 2000;
Tangen and Björnland, 1981). The symbiont of
Karenia and Karlodinium species has no remnant
of cytoplasm, and can therefore be recognized as
an integrated plastid. Based on cytoplasm reduc-
tion and the lack of cell cycle synchronization, the
symbiosis of H. arenicola can be placed between
the cryptophyte and diatom types of symbiosis
mentioned above. Nevertheless, their morpholo-
gical association suggests an intimate host—sym-
biont relationship. Previous studies on symbiosis
in dinoflagellates have focused on symbiont
degradation only. In this study, we demonstrated
that major morphological changes also occur in
the host, suggesting that plastid acquisition is not
merely ‘‘enslavement’’ where the symbiont is
degraded, but also a process during which the
host itself changes to establish a new association
with the symbiont.

The rigid pattern of asymmetrical inheritance of
the symbiont is also suggestive of a partly
regulated association. The symbiont always
comes to the left side of the host cell (ventral
view) before the migration of the host nucleus,
implying an interaction between the symbiont
compartment and the host cytoskeleton. If the
symbiont moved to the division plane, and not to
one side of the cell, the symbiont would co-
segregate, just as in the division of D. baltica,
where the center-positioned symbiont co-segre-
gates upon host cytokinesis (Tippit and Pickett-
Heaps 1976).

Another question is whether the association of
H. arenicola and the symbiont has developed
genetic modification. The process of endosym-
biosis is hypothesized to include genetic changes
such as lateral gene transfer (LGT) from symbiont
to host, coupled with evolution of a protein
transport machinery from host to symbiont (e.g.
Gilson and McFadden 2002). It is unclear when
those changes start and how they integrate.
Considering host—symbiont intimacy, H. arenico-
la would have already experienced or may be
experiencing some of these changes. Therefore,
the study of LGT or of a protein transport
machinery in H. arenicola would be an interesting
topic for future studies.

Recently, Marin et al. (2005) reported another
example of primary endosymbiosis in Paulinella
chromatophora Lauterborn, a freshwater thecate
amoeba that bears a cyanobacterium-like

ARTICLE IN PRESS

414 N. Okamoto and I. Inouye

structure. They reported that the symbiosis of P.
chromatophora is a more recent event than the
origin of all other plastids, based on the molecular
phylogeny of ribosomal DNA operon sequences.
This is consistent with a morphological feature of
the symbiont, namely, a peptidoglycan layer that
must have originated from the cell wall of an
ancestral cyanobacterium.

The symbiotic relationships of H. arenicola, P.
chromatophora, and the dinoflagellates probably
represent different intermediate steps in plastid
acquisition via primary or secondary endosymbio-
sis. Continued study and comparison of these
groups should provide further insight into plastid
evolution.

Concluding Remarks

Hatena arenicola gen. et sp. nov. is likely in the
process of plastid acquisition via secondary
endosymbiosis. Although it is in an early inter-
mediate stage of acquisition, the two organisms
have already established an intimate association
in ultrastructure and likely in metabolic function.
Based on behavioral and ultrastructural observa-
tions, we propose a ‘‘half-plant, half-predator’’ life
cycle. Because H. arenicola shows an early
intermediate state of plastid acquisition, it should
provide further insight into plant evolution. This
study provides a foundation for future studies on
the topic.

Methods

Sampling and temporary maintenance in the
laboratory: Because it is not possible to culture
Hatena arenicola in the laboratory, we used crude
samples from the natural habitat. Cells were
collected at Isonoura Beach, Wakayama Prefec-
ture, Japan (Fig. 1 B,C), April—December from
2000 to 2004. Samples were maintained in the
laboratory at room temperature in f/2 medium,
under ca. 10mmol photons m�2 s�1, and the
light—dark cycle was L:D ¼ 5:19 h.

Morphological observations: Light micro-
scopy (LM) and fluorescence microscopy (FM)
was conducted using a Leica DMR light micro-
scope (Leica Wetzlar GmbH, Wetzlar) and the LM
image was taken with a Keyence VB6010 digital
chilled CCD camera (Keyence, Osaka). For FM,
40,6-Diamidino-2-phenylindole (DAPI) was used to
stain the nucleus. The DAPI fluorescence along
with the autofluorescence of the plastid were

observed using a D filter cube (Leica Wetzlar
GmbH, Wetzlar).

Microspectrophotometry was performed by
majoring three different regions of the symbiont
in each of seven cells. Each absorption spectrum
was recorded in the range of 300—800 nm with a
light microscope (ECLIPSE, Nikon, Tokyo)
equipped with a high-resolution multichannel
photodetector (MCPD 7000, Otsuka Eelectronics,
Osaka) at Okazaki National Institute for Basic
Biology, Japan. The average of the three measure-
ments was considered the representative absor-
bance of each cell. Because the absorptions
obtained were almost uniform across the seven
cells, their average is shown.

A unialgal culture of Nephroselmis sp.
(NIES1417) was established from the same sam-
ple site by micropipette isolation and maintained
in f/2 medium at 20 1C under ca. 10mE light
intensity, and a light-dark cycle of L:D ¼ 14:10 h.
The uptake of Nephroselmis sp. (NIES1417) was
photographed under a CKX31 inversed light
microscope (Olympus, Tokyo) equipped with a
COOLPIX 990 digital camera (Nikon, Tokyo) at 6-s
intervals.

High-speed video images were recorded at 200
frames per second using an OPTIPHOT micro-
scope (Nikon, Tokyo), equipped with an MHS-200
high-speed video capturing system (Nac Inc.,
Tokyo). The images were digitized on a Macintosh
computer using an NIH imaging program (public
domain, developed at the US National Institute of
Health; available at http://rsb.info.nih.gov/nih-im-
age/) for analysis of the cellular and flagellar
motion.

Preparation for transmission electron micro-
scopy (TEM) and scanning electron microscopy
(SEM) was performed as described elsewhere
(Moriya et al. 2000; Okamoto and Inouye 2005b).
The observations were made with a JEOL 100CXII
electron microscope (JEOL, Tokyo) and a JSM-
6330 scanning electron microscope (JEOL, To-
kyo).

Molecular phylogeny: To avoid contamination
of the prey genome, single cells of Hatena
arenicola were isolated with micropipette into a
0.2-ml PCR tube containing 10 ml of sterilized
double distilled water, and immediately frozen at
�80 1C for more than 15 min to completely disrupt
the cells. The first and the second nested PCR
were performed with existing degenerate primer
sets (Moriya et al. 2000) using rTaq (TOYOBO,
Osaka). Thermal cycling for the first PCR con-
sisted of 33 cycles. Annealing temperatures
ranged from 50 to 47 1C (six cycles decreasing

http://rsb.info.nih.gov/nih-image/

http://rsb.info.nih.gov/nih-image/

ARTICLE IN PRESS

Table 1. Sequences used for the phylogenetic
analyses

Organism Accession
number

SSU rDNA
Katablepharidophyta/Kathablepharida

Hatena arenicola AB212285
Katablepharis japonica AB231617
Leucocryptos marina AB193602

Metazoa
Cirripathes lutkeni AF052902
Clathrina cerebrum U42452
Monosiga brevicollis AF084618

Fungi
Basidiobolus haptosporus AF113413
Chytriomyces hyalinus M59758
Pneumocystis carinii L27658
Saccharomyces cervisiae V01335
Schizosaccharomyces pombe Z19578
Scutellospora cerradensis AB041344

Amoebae
Acanthamoeba castellanii M13435
Hartmannella vermiformis M95168
Leptomyxa reticulata AF293898
Dictyostelium discoideum K02641

Cercozoa
Cercomonas longicauda AF101052
Chlorarachnion CCMP242 U03479
Euglypha rotunda X77692
Heteromita globosa U42447
Paulinella chromatophora X81811
Thaumatomonas seravini AF411259

Viridiplantae
Arabidopsis thaliana X16077
‘Chlorella’ ellipsoidea D13324
Chlorokybus atmophyticus M95612
Closterium littorale AF115438
Coleochaete scutata X68825
Fossombronia pusilla X78341
Mesostigma viride AJ250108
Pyramimonas propulsa AB017123
Tetraselmis striata X70802
Ulothrix zonata Z47999

Heterokontophyta
Ochromonas danica M32704
Pteridomonas danica L37204
Skeletonema pseudocostatum X85394
Phytophthora megasperma X54265

Alveolata
Cryptosporidium parvum X64340
Toxoplasma gondii X75429
Platyophrya vorax AF060454
Prorodon teres X71140
Alexandrium minutum U27499
Gymnodinium sp. MUCC284 AF022196
Pfiesteria sp. B112456 AF218805

Table 1. (continued )

Organism Accession
number

Prorocentrum mexicanum Y16232
Prorocentrum micans M14649

Cryptophyta
Chroomonas sp. M1318 AJ007279
Cryptomonas ovata AJ421147
Geminigera cryophila U53124
Hanusia phi U53126
Hemiselmis brunnescens AJ007282
Rhodomonas mariana X81373
Goniomonas truncata U03072

Glaucophyta
Cyanophora paradoxa X68483
Cyanoptyche gloeocystis AJ007275
Glaucocystis nostochinearum X70803
Gloeochaete wittrockiana X81901

Haptophyta
Emiliania huxleyi L04957
Pavlova salina L34669

Rhodophyta
Bangia sp. AF043362
Porphyra umbilicalis AB013179
Rhodella maculata U21217
Stylonema alsidii L26204

Centroheliozoa
Chlamydaster sterni AF534709
Heterophrys marina AF534710
Raphidiophrys ambigua AF534708

415Hatena arenicola: Halfway to a Plant?

the temperature by 0.5 1C for each cycle, and 27
cycles at a constant temperature). An extension
was performed at 72 1C for 1 min, and denaturing
was done at 94 1C for 30 s. The final extension
period was at 72 �C for 7 min. Thermal cycling for
the second PCR consisted of 33 cycles with an
annealing step at 53 1C for 30 s, an extension step
at 72 1C for 1 min, and denaturing at 94 1C for 30 s,
with a final extension period at 72 1C for 7 min. The
sequences were determined by direct sequen-
cing. Cycle sequencing reaction was performed
using a DYEnamic ET terminator cycle sequencing
kit (Amersham biosciences, Buckinghamshire), as
per the manufacturer’s instructions. Sequencing
was conducted with an ABI PRISM 377 DNA
Sequencer (Applied Biosystems, California), and
sequences were confirmed free of contaminants
and not of haptophyte prey origin by comparing
at least two cells or performing a BLAST search
at the National Center for Biotechnology Informa-
tion (NCBI) server (http://www.ncbi.nlm.nih.gov/

http://www.ncbi.nlm.nih.gov/BLAST/

ARTICLE IN PRESS

416 N. Okamoto and I. Inouye

BLAST/). The sequence (AB212285) was depos-
ited to GenBank.

The SSU sequence was manually aligned to the
existing alignments of global eukaryotes (Okamo-
to and Inouye 2005b), which include 65 taxa for
SSU rDNA (Table 1). To avoid the long-branch
attraction (LBA) artifact in SSU rDNA analysis,
organisms with an extraordinary evolutionary rate,
such as the Euglenozoa, the diplomonads, and
the palabasalids, were excluded after confirming
that they are not a sister group of the katablephar-
ids. A total of 1252 unambiguously aligned
nucleotide positions were selected for phyloge-
netic analyses.

The maximum likelihood (ML), neighbor joining
(NJ), and maximum parsimony (MP) methods for
phylogenetic analysis were applied to the data set
using PAUP* ver.4.0 b 10 (Swofford 2003). In the
ML analysis, the evolutionary model was selected
using the Akaike information criterion (AIC) test in
Modeltest ver.3 (Posada and Crandall 1998),
which selected the general time reversible (GTR)
model with rate variation among sites and
invariant sites (GTR+I+gamma). The estimated
gamma-shape parameter (alpha) of the discrete
gamma-distribution was 0.5645, and the propor-
tion of invariant sites was 0.3257. A further tree
search with GTR+I+gamma model with eight site
rate categories that approximates site rate was
used to produce the optimal tree. Bootstrap
proportion (BP) values for internal branches of
the optimal tree of the ML analysis were obtained
using PAUP* through 300 bootstrap resamplings
for the ML method and 1000 resamplings for the
NJ and MP methods.

Acknowledgments

We thank Dr. Masakatsu Watanabe and Dr.
Shigeru Matsunaga (Graduate University for Ad-
vanced Studies, Japan) for their dedication to the
microspectrophotometric work and assistance in
sampling, and Dr. Hiroshi Kawai (Research Center
for Inland Seas, University of Kobe) for use of his
laboratory equipment. We appreciate Dr. Tetsuo
Hashimoto and Dr. Miako Sakaguchi for their
assistance and advices in molecular phylogeny.
We are grateful to Dr. Jeremy Pickett-Heaps for
taking movie images of H. arenicola. This research
was supported in part by Japan Society for the
Promotion of Sciences (JSPS) Grants
RFTF00L0162 (to I.I.) and 1612007 (to N.O.).
N.O. was also supported by a JSPS Research

Fellowship for Young Scientists as a JSPS
Research Fellow.

Appendix A. Supplementary materials

Supplementary data associated with this article
can be found in the online version at doi:10.1016/
j.protis.2006.05.011

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• Hatena arenicola gen. et sp. nov., a Katablepharid Undergoing Probable Plastid Acquisition

Introduction
Results
Description
Latin Diagnosis
Light Microscopy
General Morphology
Cell Division
Fluorescence Microscopy
Microphotometry
Uptake of Prey Cells and Symbiont Specificity
Crawling Motion
Electron Microscopy
Ejectisomes
Cellular and Flagellar Covering
Endoplasmic Reticulum
Flagella and Basal Bodies
Feeding Apparatus
Symbiont
Eyespot
Molecular Phylogeny
Discussion
Taxonomy
Endosymbiosis
Eyespot Morphology
Morphological Changes of the Host and ’’Half-plant, half-predator’’ Model
Evolutionary Implications
Concluding Remarks
Methods
Acknowledgments
Supplementary materials
References