Harmful Algae 28 (2013) 126–139
Contents lists available at SciVerse ScienceDirect
Harmful Algae
journal homepage: www.elsevier.com/locate/hal
Acquired phototrophy in Mesodinium and Dinophysis – A review of
cellular organization, prey selectivity, nutrient uptake and
bioenergetics
Per Juel Hansen a,b,*, Lasse Tor Nielsen a,b, Matthew Johnson c, Terje Berge a,b,
Kevin J. Flynn d
a
Marine Biological Section, Biological Institute, University of Copenhagen, Strandpromenaden 5, DK-3000 Helsingør, Denmark
Centre for Ocean Life, National Institute for Aquatic Resources, Technical University of Denmark Kavalergården 6, DK-2920 Charlottenlund, Denmark
c
Woods Hole Oceanographic Institution, Department of Biology Watson Building, MS#52, Woods Hole, MA 02543, USA
d
Centre for Sustainable Aquatic Research, Swansea University, Singleton Park, Swansea SA2 8PP Wales, UK
b
A R T I C L E I N F O
A B S T R A C T
Article history:
Received 8 March 2013
Received in revised form 13 June 2013
Accepted 13 June 2013
Acquired phototrophy, i.e. the use of chloroplasts from ingested prey, can be found among some species
of dinoflagellates and ciliates. The best studied examples of this phenomenon in these groups are within
the ciliate genus Mesodinium and the dinoflagellate genus Dinophysis, both ecologically important genera
with a worldwide distribution. Mesodinium species differ considerably in their carbon metabolism. Some
species rely almost exclusively on food uptake, while other species rely mostly on photosynthesis. In
Mesodinium with acquired phototrophy, a number of prey organelles in addition to chloroplasts may be
retained, and the host ciliate has considerable control over the acquired chloroplasts; Mesodinium
rubrum is capable of dividing its acquired chloroplasts and can also photoacclimate. In Dinophysis spp.,
the contents of ciliate prey are sucked out, but only the chloroplasts are retained from the ingested prey.
Some chloroplast house-keeping genes have been found in the nucleus of Dinophysis and some
preliminary evidence suggests that Dinophysis may be capable for photoacclimation. Both genera have
been claimed to take up inorganic nutrients, including NO3 , indicating that processes beyond
photosynthesis have been acquired. M. rubrum seems to depend upon prey species within the Teleaulax/
Plagioselmis/Geminigera clade of marine cryptophytes. Up until now, Dinophysis species have only been
maintained cultured on M. rubrum as food, but other ciliates may also be ingested. Dinophysis spp. and M.
rubrum are obligate mixotrophs, depending upon both prey and light for sustained growth. However,
while M. rubrum only needs to ingest 1–2% of its carbon demand per day to attain maximum growth,
Dinophysis spp. need to obtain about half of their carbon demand from ingestion for maximum growth.
Both Mesodinium and Dinophysis spp. can survive for months in the light without food. The potential role
for modeling in exploring the complex balance of phototrophy and phago-heterotrophy, and its
ecological implications for the mixotroph and their prey, is discussed.
ß 2013 Elsevier B.V. All rights reserved.
Keywords:
Acquired phototrophy
Ciliates
Dinoflagellates
Mixotrophy
Symbiosis
1. Introduction
1.1. Acquired phototrophy in Mesodinium and Dinophysis
Acquired phototrophy, i.e. the use of chloroplasts from ingested
phototrophic prey or hosting of endosymbiotic algae, can be
found among some species of marine ciliates and dinoflagellates
* Corresponding author at: University of Copenhagen, Marine Biological Section,
Biological Institute, Strandpromenaden 5, 3000 Helsingør, Denmark.
Tel.: +45 35 32 19 85.
E-mail address: [email protected] (P.J. Hansen).
1568-9883/$ – see front matter ß 2013 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.hal.2013.06.004
(Esteban et al., 2010; Stoecker et al., 2009; Johnson, 2011a,b;
Hansen, 2011). The best studied examples of this phenomenon in
the two groups are found within the ciliate genus Mesodinium and
the dinoflagellate genus Dinophysis, both ecologically important
genera with a worldwide distribution. Mesodinium rubrum
(=Myrionecta rubra) is renowned for its ability to form ‘‘red tides’’
in coastal areas all around the world (Lindholm, 1985), while
Dinophysis spp, are known for their production of diarrheic
shellfish toxins (DSP: Okadaic acid, and DTX), and pectenotoxins
(see review by Reguera et al., 2012). Both genera contain species
that are dependent upon the utilization of chloroplasts of
cryptophyte origin, which they integrate with their own metabolism in different ways.
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Our knowledge of acquired phototrophy among Mesodinium
and Dinophysis has increased substantially over the past 12 years. A
major breakthrough occurred in 2000, when the first culture of
Mesodinium rubrum was established from Antarctic waters, by
Gustafson et al. (2000) feeding the ciliate the cryptophyte
Geminigera cryophila. Now cultures of M. rubrum are established
and maintained from a variety of different places around the world
(see Garcı́a-Cuetos et al., 2012). The first successful culture of a
Dinophysis species (Dinophysis acuminata) was established from
Korean waters in 2006, following the discovery of its trophic link
with M. rubrum (Park et al., 2006). Now, at least six Dinophysis
species are in culture worldwide (see Garcı́a-Cuetos et al., 2012;
127
Nielsen et al., 2013). Both genera (sensu lato) contain species with
varying degree of dependency on acquired phototrophy. The
taxonomy of both genera has changed recently and thus we will
briefly describe the most important changes and their implications
for our understanding of the evolution of acquired phototrophy
among them.
1.2. The genus Mesodinium
Mesodinium forms a well-defined group within the order
Cyclotrichiida (Class Litostomatea; Lynn, 2010), with several
unique morphological characteristics not found in other ciliates
Fig. 1. Mesodinium. Phylogeny based on nuclear small subunit (SSU) rDNA sequences inferred from Bayesian analysis. Five freshwater Mesodinium environmental sequences
constituted the outgroup. Branch support was obtained from Bayesian posterior probabilities and bootstrap (100 replicates) in maximum likelihood analyses. At internodes,
posterior probabilities (<1) are written first followed by bootstrap values (in percentage) from ML. The black dots represent the highest possible posterior probability (1.0)
and bootstrap value (100%). Species in bold face were sequenced in this study. [Correction added after online publication July 3, 2012: The GenBank accession number of M.
rubrum 41273 was changed to 412736.].
After Garcı́a-Cuetos et al. (2012) with permission.
128
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
(Fig. 1; discussed in Garcı́a-Cuetos et al., 2012). Further, all
Mesodinium spp. studied thus far have been shown to possess small
subunit (SSU) rRNA gene sequences that are divergent from other
ciliates, due to extensive deletions and elevated substitution rates
(Johnson et al., 2004). The species of Mesodinium that retain
ingested cryptophytes have all, until recently, been referred to as
either Mesodinium rubrum or Myrionecta rubra. However, the genus
Myrionecta is currently considered invalid based on the arrangement of the basal bodies forming the cirri (bundled cilia) (Garcı́aCuetos et al., 2012). Currently, the genus Mesodinium includes at
least three species, which are able to retain ingested cryptophytes:
Mesodinium rubrum, Mesodinium major and Mesodinium chamaeleon (Garcı́a-Cuetos et al., 2012). In addition, another six species
that do not retain ingested prey cells have also been described:
Mesodinium acarus, Mesodinium cinctum, Mesodinium fimbriatum,
Mesodinium pulex, Mesodinium pupula and Mesodinium velox (see
Moestrup et al., 2012).
The genus Mesodinium is found worldwide in all types of
aquatic environments from freshwater to marine and from tropical
to Arctic/Antarctic waters (e.g. Lindholm, 1985). Mesodinium
acarus, Mesodinium fimbriatum and Mesodinium pulex have all
been observed in freshwater, but at least Mesodinium acarus and
Mesodinium pulex have also been observed in brackish and marine
waters (e.g. Foissner et al., 1999). The remaining species are
commonly found in brackish and marine waters (Lindholm, 1985;
Foissner et al., 1999). The work by Garcı́a-Cuetos et al. (2012) using
both molecular and microscopic techniques revealed at least four
marine clades within the genus Mesodinium: Clade 1: Mesodinium
pulex, clade 2: Mesodinium pupula, clade 3: Mesodinium chamaeleon, clade 4: Mesodinium rubrum and Mesodinium major. An
additional clade may be made up of some of the Mesodinium pulexlike ciliates from freshwater (Bass et al., 2009; Garcı́a-Cuetos et al.,
2012). Added to this is the molecular study of Herfort et al. (2011),
which indicates there might be additional phycoerythrin-containing symbiotic species yet to be described.
1.3. The genus Dinophysis
More than 100 species have been assigned to the genus
Dinophysis (e.g. Gómez, 2005). There have been some dispute
concerning the relationship between the genera Dinophysis and
Phalachroma Stein – the latter considered a synonym of Dinophysis
(Abé, 1967; Balech, 1976). The genus Phalacroma was used to
identify species with an elevated epitheca visible in lateral view
and narrow horizontal projected cingular lists (Hallegraeff and
Lucas, 1988; Steidinger and Tangen, 1996). Recent molecular
analysis of LSU rDNA of a large group of Dinophysiales supports the
view that Phalacroma is a separate genus (Handy et al., 2009;
Jensen and Daugbjerg, 2009; Fig. 2). Phalacroma includes mainly
heterotrophic species, but at least one species, Phalacroma mitra,
contains chloroplasts of haptophyte origin (Koike et al., 2005).
Recently, chloroplast genes from a variety of algal origins were
found in P. mitra (Nishitani et al., 2012); however to what extent
ingested chloroplasts are functional is unknown at present.
Accepting the separation of Phalacroma and Dinophysis, it seems
likely that all species of Dinophysis rely on acquired phototrophy.
2. Cellular organization of acquired symbiotic organelles
2.1. Mesodinium rubrum
The cellular organization of Mesodinium rubrum (Fig. 3), with
respect to the incorporation of ingested prey cells (from now on
termed ‘‘reduced symbionts’’) has been studied in detail on several
occasions using TEM, light and epifluoresence microscopy on
material collected from natural samples (Taylor et al., 1971;
Hibberd, 1977; Oakley and Taylor, 1978) as well as from laboratory
cultures (Johnson et al., 2006, 2007; Hansen and Fenchel, 2006;
Garcı́a-Cuetos et al., 2010, 2012; Moestrup et al., 2012; Nam et al.,
2012). With respect to the fine structure of M. rubrum and its
symbionts, published results are generally in accordance with each
other. The reduced symbionts seem to be delimited from the ciliate
cytoplasm by a single membrane. The symbionts have been
estimated to constitute 36% of the volume of the total
Mesodinium cell volume (Hansen and Fenchel, 2006). The
cytoplasm of the ingested prey can be recognized by the
mitochondria; these are small, cylindrical or flattened and with
flat cristae. In contrast, the ciliate mitochondria are larger, ovoid
and with tubular cristae. Chloroplast numbers range from 6 to 36
per cell. The chloroplasts are flattened and somewhat curved and
located in the periphery of the ciliate and are concentrated in the
anterior and posterior ends of the cell; they are absent from the
middle part of the Mesodinium cell (Taylor et al., 1971; Hibberd,
1977; Hansen and Fenchel, 2006). The thylakoids are arranged in
triplets. Pyrenoids are stalked and the chloroplasts include a
nucleomorph. Vacuoles containing accumulation (‘‘storage’’)
products are often present.
There is usually an extra nucleus in Mesodinium rubrum, the socalled ‘‘symbiont nucleus’’ or ‘‘prey nucleus’’, which measures ca.
4.5 5.7 mm. However, in dividing M. rubrum cells (Kattegat
strain) there may be two symbiont nuclei present (Hansen and
Fenchel, 2006), suggesting that the ciliate can induce division of
this acquired nuclear material. The Antarctic strain does not seem
to be able to divide the cryptophyte nucleus, and tend to lose it
through cell division upon starvation (Johnson and Stoecker,
2005; Johnson et al., 2007). Thus, in this M. rubrum strain the
acquisition of the cryptophyte nucleus has been referred to as
‘‘karyoklepty’’ (Johnson et al., 2007). Ultrastructural features of
the free-living prey organisms tend to differ in important aspects
from that of the symbionts inside M. rubrum, indicating that some
morphological changes of the chloroplasts occur after ingestion
and integration. The single chloroplast in the cryptophyte prey
is typically smaller and the pyrenoids are not stalked in
comparison with the appearance inside M. rubrum. Also, the
individual nuclei in the cryptophyte prey are substantially smaller
than the symbiont nucleus (Hansen and Fenchel, 2006; Johnson
et al., 2007).
2.2. Other Mesodinium species
Only two other species of Mesodinium species have been
studied with regard to the cellular organization of the symbionts:
Mesodinium major and Mesodinium chamaeleon (Moestrup et al.,
2012; Garcı́a-Cuetos et al., 2012). The cellular organization of
Mesodinium major with respect to its reduced symbionts is quite
similar to Mesodinium rubrum. The symbionts of Mesodinium major
are delimited by a single membrane and consist of cryptophyte
chloroplasts, a ‘‘symbiont nucleus’’, mitochondria, and some
cytoplasm. However, Mesodinium major is a larger species and it
contains many more chloroplasts and mitochondria than does
Mesodinium rubrum. It also differs from Mesodinium rubrum in that
it can occasionally transfer some of the chloroplasts into so-called
‘‘flaps’’, which are cytoplasmic extensions at the anterior end of
the cell.
In Mesodinum chamaeleon the ‘‘symbiont nucleus’’, so typical for
both Mesodinum rubrum and Mesodinum major, is completely
absent. Newly ingested cryptophyte cells could be found
completely intact inside food vacuoles of the ciliate, similar to
what can be found in many entirely heterotrophic ciliates. Ingested
cryptophyte cells will, after some time, be reduced (partly
digested) to mainly single chloroplast with the nucleomorph
remaining intact and a cryptophyte nucleus. Each food vacuole
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
129
Fig. 2. Phylogeny of 31 members of the Dinophysiales based on nuclear-encoded LSU rDNA sequence including domains D1–D6 (1441 bp) and inferred from Bayesian
analysis. Two Prorocentroids (Prorocentrum micans and Prorocentrum minimum) constituted the outgroup. Branch support was obtained from Bayesian posterior probabilities
and bootstrap (100 replicates) in maximum-likelihood (ML) analyses. At internodes, posterior probabilities (£1) are written first, followed by bootstrap values (in percentage)
from ML. d, indicates the highest possible posterior probability (1.0) and bootstrap value (100%). Asterisks indicate type species. Species in boldface were determined twice.
The dinophysioids have been divided into three clades (A–C). Clade C has been further subdivided into seven subclades.
After Jensen and Daugbjerg (2009) with permission.
contains one cryptophyte cell and the food vacuoles are not
connected to each other but rather distributed at the periphery of
the ciliate cell (Moestrup et al., 2012).
2.3. Dinophysis spp.
The cellular organization of Dinophysis species (sensu stricto)
has been studied using transmission electron microscopy (TEM) in
three species: Dinophysis acuminata, Dinophysis caudata, and
Dinophysis fortii, using both wild and cultured material (Hallegraeff
and Lucas, 1988; Nagai et al., 2008; Garcı́a-Cuetos et al., 2010;
Kim et al., 2012). All studied species have chloroplasts of
cryptophyte origin and genetic studies indicate that the chloroplasts are closely related or identical to chloroplasts of species
within the Teleaulax/Plagioselmis/Geminigera clade (the TPG clade)
(e.g. Takishita et al., 2002; Janson, 2004; Minnhagen and Janson,
2006; Nagai et al., 2008; Park et al., 2008; Nishitani et al., 2010;
Garcı́a-Cuetos et al., 2010). All other cell organelles of cryptophyte
origin are lost in Dinophysis spp. (Hallegraeff and Lucas, 1988;
Nagai et al., 2008; Garcı́a-Cuetos et al., 2010; Kim et al., 2012). The
chloroplasts are located in clusters varying in numbers from two
clusters in a small species like Dinophysis acuminata to up to five
clusters in Dinophysis acuta. The chloroplasts inside these clusters
generally have terminal pyrenoids and the thylakoids are usually
stacked in doublets (Fig. 4). This is very different from what is
found in species of the TGP clade and from the ‘‘symbionts’’ inside
Mesodinium rubrum. In these organisms the chloroplasts have
central pyrenoids and the thylakoids are arranged in triplets. The
starkly contrasting chloroplast structure within these groups led to
some controversy concerning whether the chloroplasts in Dinophysis are permanent or whether they come from ingestion of M.
rubrum prey (Myung et al., 2006; Park et al., 2006, 2010; Nagai et
al., 2008; Minnhagen et al., 2008, 2011; Garcı́a-Cuetos et al., 2010;
Nishitani et al., 2010).
Species within the genus Phalacroma are generally considered
heterotrophic, feeding on ciliates (Hansen, 1991; Elbrächter, 1991).
So far only one species, Phalacroma mitra, has been shown to
incorporate chloroplasts and those are of haptophyte, not of
cryptophyte origin (Koike et al., 2005). Like in Dinophysis, only the
chloroplasts are sequestered, other cell organelles from ingested
prey seem to be missing. However, the chloroplasts are placed
along the periphery of the P. mitra cells and not in specific centers
or clusters as in Dinophysis. Also, the pyrenoids of the sequestered
haptophyte chloroplasts are centrally placed, not terminally as
130
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Fig. 3. Mesodinium rubrum, grown in culture. (A and B) Light micrograph of a starved cell of M. rubrum prior to extraction. (C) Light micrograph of a fed cell of M. rubrum. (D)
Longitudinal section through the cell illustrating the cilia, the profiles of the chloroplast (Chl), a pyrenoid (Pyr) and starch grains (St). (E and F) Details of chloroplast
thylakoids, which are always arranged in 3-thylakoid lamellae (tThy). (G) Chloroplast showing the lateral position of the pyrenoid. The pyrenoid is surrounded by a few starch
grains. Mitochondria of two types are also visible in the figure, from the ciliate (hMit) and from the cryptophyte endosymbiont (eMit), respectively, but the cristae of the latter
cannot be distinguished at this low magnification.
After Garcı́a-Cuetos et al. (2010) with permission.
are the pyrenoids of cryptophyte origin in Dinophysis spp. (Koike
et al., 2005).
Recently, Nishitani et al. (2012) found rbcL sequences in fieldcollected Phalacroma mitra cells that belong to haptophyceae
(73.7%), particularly from within the genus Chrysochromulina.
However, Nishitani et al. (2012) also found rbcL sequences from
other algal groups: prasinophytes (5.2%), dictyochophytes (4.5%),
pelagophytes (4.5%), bolidophytes (1.0%) and diatoms (0.3%). To
what extent chloroplasts from these other algal groups are
incorporated and functionally used by P. mitra is yet unknown;
they might just be recently ingested cells. Cultures of P. mitra are
not yet available, so the confirmation or rejection of other types of
chloroplasts than those from haptophytes remains speculative.
One possibility is that P. mitra acquires its chloroplasts from
mixotrophic oligotrich ciliates, which have been shown to harbor
chloroplasts from a variety of algal sources (Stoecker et al., 1987).
This may be likely due to the propensity of the Phalacroma genus to
feed on several different types of ciliates.
3. Prey capture and prey selection
3.1. Prey capture and prey selection in Mesodinium
Prey cells are captured by means of the tentacles, which are
positioned at the anterior end in all Mesodinium species. Prey cells
are caught at the tip of the tentacles, which have mucocysts
associated with them (Lindholm et al., 1988; Moestrup et al., 2012;
Garcı́a-Cuetos et al., 2012). The tentacles with the adhered prey are
then retracted into the oral region bringing the prey into the oral
cavity. The tentacles in different species are built upon the same
principle, but differ in the number of microtubules: 13 in
Mesodinium chamaeleon, 14 in Mesodinium rubrum and Mesodinium
major and 23 in heterotrophic Mesodinium pulex and Mesodinium
pupula (Moestrup et al., 2012; Garcı́a-Cuetos et al., 2012).
Mesodinium spp sense swimming prey at a certain distance from
the cell; the cirri seem to play a role in the detection of
hydromechanical signals made by the swimming prey (Jakobsen
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
131
Fig. 4. Dinophysis acuminata, grown in culture. (A) Light micrograph of a starved cell of D. acuminata prior to extraction. (B) Lightmicrograph of a well fed cell of D. acuminata.
(C) Epifluorescence micrograph of a starved cell of D. acuminata illustrating the compound pyrenoids (cPyr) of both stellate chloroplasts. (D) Paired thylakoids within the
chloroplasts. (E) Longitudinal sections through two cells, showing general appearance of the cell, nucleus (N) and, in the cell on the left, the posterior chloroplast complex with
its compound pyrenoid (cPyr). (F) The posterior chloroplast complex, located immediately behind the nucleus (N), with many close-packed pyrenoids (cPyr), from which the
individual chloroplasts extend. Three food vacuoles are also present (Va).
After Garcı́a-Cuetos et al. (2010) with permission.
et al., 2006). Detailed laboratory studies on exploitation of different
types of prey are only available for Mesodinium rubrum and
Mesodinium pulex (Jakobsen et al., 2006; Park et al., 2007; Myung
et al., 2011; Tarangkoon and Hansen, 2011; Hansen et al., 2012),
while little is known about the prey exploited by Mesodinium
chameleon (Garcı́a-Cuetos et al., 2012).
Mesodinium pulex is an omnivorous species, which ingests a
large variety of different prey cells, including cryptophytes,
dinoflagellates and ciliates. It may even ingest prey of its own
size (i.e., the ciliate Metanophrys sp., Dolan and Coats, 1991) and it
has recently been shown to be cannibalistic in prey deplete
cultures (Moestrup et al., 2012). Critical for its ability to locate (and
thence capture) prey is that the prey cells are motile. However,
other factors matter as well. M. pulex will respond to the presence
of a fast swimming organism, like Gymnodinium simplex, with an
escape response even though the prey size is in the correct range
for capture and ingestion (Jakobsen et al., 2006). Also, certain
cryptophytes, like Rhodomonas salina, are not efficiently captured
by M. pulex, simply because the mucocysts do not stick to the cell
surface of these cryptophytes (Jakobsen et al., 2006). Finally, many
small dinoflagellates and cryptophytes may respond to attack from
M. pulex by making escape responses, by which they can ‘‘jump’’
away from the predator.
Several attempts have been made to cultivate Mesodinium
rubrum on other clades of cryptophytes and also on species
belonging to other algal groups. Park et al. (2007) studied the
growth response of starved M. rubrum (Korean strain), when
offered four cryptophytes belonging to the TPG clade and two
cryptophytes belonging to the ‘Rhodomonas clade’ (CR-MAL03 and
CR-MAL06) in the light. Except for CR-MAL06, positive growth
rates were observed during the incubation time studied (6 d), and
all rates were significantly different from the unfed control. The
results indicate that MR-MAL01 can grow on species not closely
related to Teleaulax, although the observed growth rate on the CRMAL03 strain was low (ı` = 0.16 d 1) at least for 6 days, which was
the duration of the experiment. This observation was confirmed by
Myung et al. (2011), who found low growth rates of the same strain
of M. rubrum when starved cells were fed CR-MAL03 over a longer
period (10 d). Hansen et al. (2012) exposed an isolate of M. rubrum
from the Kattegat (Denmark) to one dinoflagellate and eight
species of cryptophytes belonging to five different clades of
cryptophytes and could only find sustained growth for at least 12 d
when supplied with Teleaulax acuta and Teleaulax amphioxeia as
prey. Growth was observed in some of the cases, where M. rubrum
was fed other prey types, but the rates were not significantly
different from the unfed controls.
132
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Ingestion rates of Mesodinium rubrum on cryptophytes outside
TPG clade have not been provided in any of the three studies done
so far. However, data presented on development of prey
populations in mixed cultures with M. rubrum compared to in
mono culture by Myung et al. (2011) and Hansen et al. (2012) could
indicate that M. rubrum ingests preys of various types. Both these
studies clearly demonstrated that M. rubrum was not able to
control the prey populations, irrespective of initial concentrations
of predator and prey. In comparison, M. rubrum can control prey
populations of cryptophytes when fed species within the TPG clade
under certain initial predator:prey concentration ratios (e.g.
Gustafson et al., 2000; Johnson and Stoecker, 2005; Hansen and
Fenchel, 2006; Hansen et al., 2012). There is a clear need to study
direct uptake in M. rubrum when subjected to prey outside the TPG
clade in the future to elucidate whether M. rubrum are discriminate
feeders (i.e. selection occurs before actual ingestion) and it only
ingests and incorporates preys within the TPG clade. Nevertheless,
it is possible to say that all strains of M. rubrum studied so far can
only exploit prey species from the TPG clade for sustained and/or
rapid growth.
Since Mesodinium chameleon is not yet in culture, little is known
about its prey preferences. However, it has been shown to feed on a
variety of different cryptophytes belonging to different clades, and
it will change its color (red or blue-green) depending upon
whether it feeds on red cryptophytes (Teleaulax spp.) or blue-green
cryptophytes (Chroomonas spp.) (Moestrup et al., 2012). Thus, it
seems to be able to use a wider variety of different clades of
cryptophytes than Mesodinium rubrum.
3.2. Prey capture and prey selection in Dinophysis and Phalacroma
Dinophysis and Phalacroma species appear to all feed exclusively
on ciliates (Hansen, 1991; Elbrächter, 1991; Park et al., 2006;
Nishitani et al., 2008a,b; Nagai et al., 2008; Riisgaard and Hansen,
2009; Rodriguez et al., 2012). Six species of Dinophysis (Dinophysis
acuminata, Dinophysis acuta, Dinophysis caudata, Dinophysis fortii,
Dinophysis infundibulus, and Dinophysis tripos) have been cultured
so far and all these cultures have been maintained on Mesodinium
rubrum as prey (Park et al., 2006; Nishitani et al., 2008a,b; Nagai
et al., 2008; Kim et al., 2011; Rodriguez et al., 2012). The prey
capture and handling of M. rubrum cells by Dinophysis acuminata
and Dinophysis acuta have been observed by one of us (PJH) and
will be described below.
The swimming behavior of Mesodinium rubrum is very
characteristic and has been studied in great detail using high
speed video by Fenchel and Hansen (2006) and Riisgård and Larsen
(2009). M. rubrum usually sit completely still for some seconds
followed by a single jump, where after the ciliate remains
essentially motionless. Each jump is initiated by the movement
of the cirri to the front of the cell, where they remain until the jump
is completed. After the movement of the cirri, the large numbers of
cilia in the equatorial belt will beat; the travelled distance depends
upon the duration of their movement. M. rubrum displays this
behavior at all times even if not disturbed; the typical periodicity of
jumps is 1–15 s, the speed is up to a maximum of 12 mm s 1 and
the average distance of travel is 160 mm (Fenchel and Hansen,
2006; Riisgård and Larsen, 2009). However, the frequency and
duration of jumping may be greater if the ciliate detects strong
hydromechanical signals, such as disturbances from predators like
copepods (Jonsson and Tiselius, 1990; Fenchel and Hansen, 2006;
Riisgård and Larsen, 2009). Analysis of the hydrodynamic
disturbance created by M. rubrum jumps suggests that its
propulsion, which is dependent upon an equatorial ciliary belt,
generates a weak and spatially limited hydrodynamic disturbance
(Jiang, 2011). Thus M. rubrum may be difficult for certain predators,
such as copepods, to detect and capture.
In contrast to Mesodinium spp., Dinophysis spp. are fairly slow
swimmers. However, they are able to detect Mesodinium rubrum
cells at a distance from which the ciliate cannot detect the
Dinophysis cell. They do this either by chemical sensing or due to a
more sensitive sensing of the hydromechanical signal elicited by
M. rubrum. Upon detection of a potential prey cell, the Dinophysis
cell starts to circle around the ciliate and at some point of time
shoots off a capture filament that will adhere to the ciliate. For a
short period (seconds) the ciliate attempts to swim away, before it
gets immobilized. The immobilized ciliate is then drawn close to
the dinoflagellate cell and a peduncle enters the ciliate before the
contents are sucked out (tube feeding).
To what extent Dinophysis species are able to catch other ciliates
with a similar swimming behavior as Mesodinium rubrum is
unknown. Potential prey organisms to look for are the ciliates
Tontonnia, Laboea, Strobilidium, because they elicit a swimming
behavior similar to M. rubrum, i.e. periods of time where the cell is
immobile followed by fast jumps (e.g. Stoecker et al., 2009).
Tintinnids, like Favella ehrenbergii and Schmidingerella arcuata have
been tested as prey for Dinophysis acuminata and Dinophysis acuta,
but none of the Dinophysis species were observed to feed or grow
upon any of the tintinnids (PJH, unpublished data).
The number of potential prey species so far reported for
Phalacroma species is quite limited. The only species for which a
little is known is for the heterotrophic Phalacroma rotundata (as
Dinophysis rotundata). This species has been observed to feed upon
ciliates including the prostomatid ciliate Tiarina fusus (Hansen,
1991) and some tintinnids (Elbrächter, 1991), potentially indicating a different prey precapture behavior than Dinophysis species.
Tiarina and tintinnids all swim continuously and fast in a stretched
out helix interrupted by tumbles, which are short periods where
the ciliates stop and swim slightly backwards followed by forward
swimming. They never exhibit periods of non-motility as seen in
Mesodinium species. Precapture behavior has never been observed
for Phalacroma species, but it might well be different from
Dinophysis spp. taking into account the swimming behavior of the
different prey types of the two genera. Phalacroma spp. might
actually trap their ciliate prey when they try to ingest a Phalacroma
cell; the prey becomes the predator. Indeed this may explain why
T. fusus can ingest, for instance, Dinophysis acuminata cells, while it
in turn is ingested by Phalacroma rotundata (Hansen, 1991). It is
still uncertain from where the plastidic Phalacroma mitra derives
it’s mainly haptophyte chloroplasts (Koike et al., 2005).
4. Incorporation and replacement of cell organelles, lateral
transfer of chloroplast house-keeping genes, nutrient uptake
and photoacclimation
Substantial molecular evidence support that chloroplast gene
sequences from certain free-living TPG cryptophytes are identical
to those of both Mesodinium rubrum and Dinophysis spp.; thence it
has been suggested that the chloroplasts are sequestered
continuously from ingested prey cells (Hackett et al., 2003; Janson,
2004; Minnhagen and Janson, 2006; Nagai et al., 2008; Minnhagen
et al., 2011). While chloroplast gene sequences were quite similar
or identical in all three organisms, direct proof of continuous
organelle sequestration have been lacking until very recently.
Detailed transmission electron microscopy of all three organisms
revealed some important structural differences of the chloroplasts
in Dinophysis compared to that of the prey M. rubrum, and this
raised questions as to whether they came from ingested M. rubrum
cells (Garcı́a-Cuetos et al., 2010). Below we consider the evidence
that support the transfer of chloroplasts and other cell organelles
from cryptophytes to M. rubrum and further into Dinophysis spp.,
and the dependency of a constant supply of the cell organelles in
the two genera.
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Johnson et al. (2006) found identical nucleomorph 18s gene
sequences between cultured Mesodinium rubrum (CCMP 2563) and
its prey Geminigera cryophila (CCMP 2564), and used quantitative
(q) PCR for the gene to demonstrate division of the chloroplast in
the ciliate. However, the first direct indications that M. rubrum
receives its symbionts via ingestion came from a study by Park
et al. (2010) using a chloroplast gene (psbA) as a means to track the
uptake of different types of cryptophyte prey. They found that
chloroplasts inside M. rubrum cells switched from Teleaulax
amphioxeia chloroplasts to Teleaulax acuta chloroplasts when the
prey was switched from Teleaulax amphioxeia to Teleaulax acuta.
However, no numbers of copies of chloroplast genes or an exact
time frame was provided. In a later study, Hansen et al. (2012)
confirmed these preliminary results of Park et al. (2010) and
demonstrated that a M. rubrum culture that have been kept on a
diet of Teleaulax amphioxeia had chloroplasts that were identical to
those of Teleaulax amphioxeia. When such cultures were then
switched to a diet consisting entirely of Teleaulax acuta cells, the
chloroplasts of M. rubrum cells were gradually replaced over the
course of 35 days to become almost entirely of the Teleaulax acuta
type (Fig. 5). There is thus clear evidence now that M. rubrum is
dependent upon the continuous uptake and replacement of
cryptophyte symbionts and that it can utilize chloroplasts from
the TPG clade.
Fig. 5. Mesodinium rubrum. Cellular concentrations of chlorophyll a cell 1 in (A) high
light (HL) acclimated M. rubrum and (B) low light (LL) acclimated M. rubrum. Data
points represent individual sample replicates. Prey was depleted after 28 days.
After Johnson and Stoecker (2005) with permission.
133
The focus in the literature has so far mainly been on the
sequestration of chloroplasts in Mesodinium rubrum. However,
Johnson et al. (2007) provided some evidence that M. rubrum also
are able to sequester cryptophyte nuclei. They demonstrated that
cryptophytes nuclei were retained for up to 30 d in an Antarctic
strain of M. rubrum, with a half-life of 10 days. The nuclei were
transcriptionally active and apparently able to service plastids
derived from multiple cryptophyte cells of the same species.
Johnson et al. (2007) found that the presence of the cryptophyte
nucleus in M. rubrum corresponded to higher growth rates,
enhanced photochemical efficiency, and its ability to divide
plastids. However, exactly how the cryptophyte nucleus regulates
plastid activity is still unknown. Also, what is the fate of the
cryptophyte nuclei within M. rubrum, when prey is switched from
one species to another? Does the ciliate simultaneously use two
distinct symbiont nuclei? Future experiments should address the
exact role of the symbiont nucleus and how the ciliates control the
functioning and division of chloroplasts from different prey
species.
In the case of Dinophysis, only the chloroplasts from cryptophytes remain; all other cryptophyte cell organelles are lost. The
first solid evidence that the chloroplasts of Dinophysis were in fact
sequestered from the prey (Mesodinium rubrum) came from a
recent study by Kim et al. (2012). They demonstrated that the
chloroplasts of newly ingested M. rubrum cells were separated out
from the rest of the ingested material and transferred to the
preexisting clusters of chloroplasts. During the first 24 h the
chloroplasts undergo quite extraordinary structural transformation, including the movement of the pyrenoids from a lateral to a
terminal position of the chloroplast and a decrease in the number
of thylakoids from three to two (Fig. 6). The digestion of
cryptophyte nuclei and nucleomorphs by the dinoflagellate shortly
after prey ingestion raises the question of the functional controls of
the chloroplasts? Evidence suggests that these regulatory functions are taken over by the host (Wisecaver and Hackett, 2010).
They identified five proteins, complete with plastid-targeting
peptides, encoded in the nuclear genome of Dinophysis acuminata
that function in photosystem stabilization and metabolite
transport. Of the five nuclear-encoded, plastid proteins identified
in D. acuminata, only the photosystem II subunit M (psbM) appears
to be of cryptophyte origin. The psbM protein is thought to be
involved in photosystem dimer formation (Ferreira et al., 2004).
Two of the plastid-related proteins, the triose-phosphate transporter (TPT) and ferredoxin (petF) grouped with peridinin
dinoflagellates (i.e., the ancestral dinoflagellate pigment) in
phylogenetic analyses. The plastid TPT is involved in transport
of fixed carbon out of the plastid (Martinez-Duncker et al., 2003)
and presumably provides a mechanism by which D. acuminata
benefits from the photosynthesis. Ferredoxin (petF) is responsible
for distributing the electrons generated by photosystem I to
various reactions in the plastid stroma (Arnon, 1988). The
remaining two proteins, a light harvesting protein (LHP) and
psbU, appear to have been derived over evolutionary time from
either haptophytes or fucoxanthin containing dinoflagellates (i.e.,
dinoflagellates that have replaced the peridinin plastid with one
derived from haptophytes and containing the photopigment
fucoxanthin). LHPs shuttle the light energy captured by chlorophyll and accessory pigments to the photosystems (Durnford et al.,
2007) and may be involved in stabilizing the photosystem in
response to heat or photodamage (Eppard et al., 2000; Richard
et al., 2000; Elrad and Grossman, 2004). PsbU is specifically
involved in protecting the photosystem from heat and photodamage and may have an increased functional interaction with
photosystem II (Enami et al., 2008). Thus D. acuminata seems to
have only a few chloroplast house-keeping genes and appears to be
unable to replicate (divide) the plastids. This, along with the
134
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Fig. 8. Mesodinium rubrum. Symbiont exchange experiment. The percentage of
symbiont nucleomorph large subunit (LSU) copies in an experiment in which prey
was switched from Teleaulax amphioxeia to Teleaulax acuta at the start of the
experiment. Values are treatment means SE, n = 4, except for at Day 35, where
n = 3.
After Hansen et al. (2012) with permission.
Fig. 6. Mesodinium rubrum. Estimated chlorophyll a (chl a) production and chl a
gained by ingestion of cryptophyte prey each 2 weeks period. Histograms represent
sample means.
After Johnson and Stoecker (2005) with permission.
absence of a cryptophyte nucleus, suggests that D. acuminata is
dependent upon a continuous supply of chloroplasts. This
contrasts with the situation in M. rubrum, which has the potential
to synthesize and replicate new chloroplasts at least 3–5 times
when starved of prey (Hansen and Fenchel, 2006).
This raises the next question, as to the extent to which
photoacclimation (i.e. the change in pigmentation in response to
changes in irradiance) can occur in Mesodinium rubrum and
Dinophysis spp. Photoacclimation has been shown to take place in
M. rubrum (Fig. 7; Johnson et al., 2006; Moeller et al., 2011). It has
been shown that cellular chlorophyll a and phycoerythin
concentrations range from 28–73 to 15–82 mg cell 1, respectively
over a range of 1.7–100 mmol photons m 2 s 1, with the highest
levels obtained at low light. Results from similar studies for
Dinophysis have not been published, but epifluorescence micrographs of Dinophysis acuminata and Dinophysis acuta clearly
indicate that chloroplasts are larger when cells are grown in
low irradiance, suggesting that some level of photoacclimation
may be taking place in these organisms (Fig. 8; Nielsen et al., 2012).
Further studies are needed to fully understand the level of
chloroplast control in Dinophysis spp.
Little is known about the utilization of dissolved inorganic and
organic nutrients by Mesodinium and Dinophysis. A few studies
have indicated that Mesodinium and Dinophysis may be able to take
up both inorganic and organic nitrogen. In all cases, 15N tracer
techniques were used to study uptake of nitrogen (Packard et al.,
1978; Wilkerson and Grunseich, 1990; Seeyave et al., 2009).
Fig. 7. Dinophysis acuta. Epifluorescence micrographs of cells incubated in A) low light (15 mmol photons m
Nielsen, orginal.
2
s
1
) and B) high light (130 mmol photons m
2
s
1
). Lasse Tor
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Nothing is known about the ability of Dinophysis and Mesodinium
to assimilate inorganic or organic phosphate.
Mesodinium rubrum is the only Mesodinium species for which
inorganic and organic nitrogen uptake has been studied. It has
been shown to take up nitrogen in the form of nitrate, ammonium
and DON. Light seems to be required for nitrate assimilation and to
a lesser extent for ammonium assimilation, while dark assimilation of ammonium occurred in all experiments (Packard et al.,
1978; Wilkerson and Grunseich, 1990). No effect of light on DON
uptake was observed.
In Dinophysis, uptake of inorganic and organic nitrogen has only
been studied for Dinophysis acuminata (Seeyave et al., 2009).
Results have indicated that it can utilize nitrate, ammonium and
urea. Apparently it was best at utilizing ammonium and urea, and
that it has a high affinity for these N-substrates (Seeyave et al.,
2009).
The reported data should however be interpreted with caution
as the studies were all carried out using natural assemblages where
these species dominated. It can thus not be excluded that the
tracers may have been taken up (1) by accompanying nano- and
picoplankton organisms, which are quite abundant and may have
been overseen in samples (2) or via prey particles and thus not
directly. Now that cultures are available of both genera, future
studies should address more carefully the direct uptake of
inorganic and organic nitrogen and phosphorous in cultures
without prey.
5. Effects of light on food uptake, chloroplast division and
growth rates
5.1. Effects of light on Mesodinium spp
Mesodinium rubrum cannot grow in the dark even if supplied
with plenty of food, and it cannot grow indefinitely in the light
without its preferred cryptophyte prey. It is an obligate mixotroph
that depends heavily upon phototrophy. Maximum growth rates
135
range from 0.5 to 0.7 d 1 for temperate isolates at 15–20 8C
(Hansen and Fenchel, 2006; Park et al., 2007; Smith and Hansen,
2007) and about 0.15–0.2 d 1 for the Antarctic strain (Gustafson
et al., 2000; Johnson and Stoecker, 2005; Johnson et al., 2006,
2007). Growth rates are highly dependent upon irradiance, with
maximum growth rates obtained at irradiances above 20 and
50 mmol photons m 2 s 1 for polar and temperate strains, respectively, but positive and sustained growth rates can be obtained at
very low irradiance levels if prey is available. Moeller et al. (2011)
demonstrated sustained phototrophic growth in the Antarctic M.
rubrum strain at irradiances as low as 1.7 mmol photons m 2 s 1.
Prey uptake rates are generally quite low (Hansen and Fenchel,
2006; Smith and Hansen, 2007). Temperate strains of M. rubrum
ingest up to ca. 4 cryptophytes d 1, but the ingestion of just one
cryptophyte cell d 1 will sustain maximum growth rates (m = 0.5
d 1), which equals an assimilation of phagocytized prey-C of less
that 1–2% of the daily carbon need (Fig. 9). Thus, M. rubrum is
predominantly a phototroph.
The only non-symbiotic Mesodinium species in culture is so far
Mesodinium pulex (Jakobsen et al., 2006; Tarangkoon and Hansen,
2011). Unlike the symbiotic species, Mesodinium pulex can grow in
the dark as a pure heterotroph, but at decreased growth rates
compared to in the light. Maximum growth rates are typically
higher than in the symbiotic Mesodinium rubrum, and maximum
growth rates of 0.7–1.0 d 1 have been reported (Tarangkoon and
Hansen, 2011). Differences in growth rates in the light and in
darkness observed in M. pulex cannot be explained by photosynthesis. Well fed M. pulex cells photosynthesize, but measured rates
can only account for at best 4% of the total carbon needs for
growth and thus likely do not represent a form of mixotrophy
(Tarangkoon and Hansen, 2011). Instead, measurements show that
food uptake increases in the light compared to in the dark. Reasons
for increased prey uptake and growth of non-symbiotic Mesodinium cells in the light are currently unknown but warrant
exploration. A study by Strom (2001) has shown that light appears
to aid digestion, grazing rates, and growth in herbivorous protists,
Fig. 9. Morphological transformation of the retained Mesodinium rubrum chloroplasts and coalescence to stellate compound chloroplasts. (A) Light micrograph of Dinophysis
caudata 30 min after feeding. The ingested reddish-brown chloroplasts have moved to the periphery of the cell. (B) TEM image of the chloroplast indicated by dotted circle in
A. The chloroplast has a terminal positioned pyrenoid (Pyr), but the thylakoids are oriented along the long axis of the organelle. (C) Light micrograph of D. caudata 21 h after
feeding. The ingested reddish-brown chloroplasts form four stellate compound chloroplasts. (D) Ultrastructure of the stellate compound chloroplast indicated by black dotted
box in C. Note the seven loosely packed pyrenoids (Pyr) and the thylakoids (arrow) running parallel to the margin of one pyrenoid. (E) Ultrastructure of two chloroplasts
indicated by white circle in C, showing thylakoids arranged as a mix of stacks of two (double arrows) and three (triple arrows).
After Kim et al. (2012) with permission.
136
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
and suggests that photooxidation may help in breaking down
organic matter in food vacuoles.
5.2. Effects of light on Dinophysis spp
All species of Dinophysis studied so far require light for growth
and they cannot grow in darkness even when given an ample food
supply (Fig. 10; Kim et al., 2008, 2012; Nishitani et al., 2008a,b;
Nagai et al., 2008; Riisgaard and Hansen, 2009; Nielsen et al., 2012,
2013). Maximum growth rates range from 0.4 to 0.9 d 1 at
temperatures of 15–20 8C for temperate isolates. Growth rates are
highly dependent upon irradiance, with maximum growth rates
obtained at irradiances above 50 mmol photons m 2 s 1, but
sustained growth rates can be attained at very low irradiance
levels (Nielsen et al., 2012, 2013) (Fig. 11).
Prey uptake rates in Dinophysis spp. can be quite high (Kim et al.,
2008; Riisgaard and Hansen, 2009). Temperate strains of
Dinophysis acuminata may ingest >10 Mesodinium rubrum cells
per day at high prey concentrations, which corresponds to 70–90%
of the gross daily C uptake rate (Kim et al., 2008; Riisgaard and
Hansen, 2009). However, at low prey concentrations (<200 M.
rubrum cells ml 1), only 0–55% of the gross C uptake may be
derived from phagotrophy (Riisgaard and Hansen, 2009). These
data could indicate that in natural environments, D. acuminata may
often be food limited, and, in this situation, photosynthesis may
not only be a supplement to the basic nutrition, but rather the
primary C source (Velo-Suárez et al., 2013). As there is no evidence
at present that suggest that Dinophysis spp. can divide its acquired
chloroplasts (Minnhagen et al., 2008, 2011; Wisecaver and
Hackett, 2010), it is to be expected that ingestion rates in
Dinophysis species are much higher than those found in M. rubrum.
5.3. Survival in Mesodinium and Dinophysis
The symbiotic Mesodinium rubrum and Dinophysis species are all
excellent survivors in the absence of prey. Several reports suggest
that they can survive for up to 3 months in the light (Johnson and
Stoecker, 2005; Hansen and Fenchel, 2006; Smith and Hansen,
2007; Nielsen et al., 2012, 2013). The reason for this ability lies in
the fact that well fed M. rubrum and Dinophysis spp. cells only
divide 3–4 times if subjected to sudden starvation. This hinders
dilution of chloroplasts by continuous cell division and allows the
cells to maintain a reduced rate of photosynthesis, which is high
enough to meet basic respiration requirements of survival.
Research on an Antarctic strain of Mesodinium rubrum indicates
that cells kept cool and in darkness have a cell-specific half-life
time of at least 77 d, thus underscoring the resilience of this species
and its ability to overwinter during the Antarctic winter (Moeller
et al., 2011). In low light, respiration rates in this strain decrease,
and the quantum yield of growth is greatest, indicating that the
ciliate is highly efficient in utilizing low levels of light (Moeller
et al., 2011). At higher growth irradiances, the Antarctic strain of M.
rubrum will respire up to 50% of its photosynthate (Moeller et al.,
2011).
6. Modeling symbiotic Mesodinium and Dinophysis
Fig. 10. Mesodinium rubrum. (A) Growth of M. rubrum under two irradiances as a
function of prey (Teleaulax sp.) concentration. (B) Ingestion rate of Teleaulax sp. cells
as a function of prey concentration (no difference between curves, p > 0.05). (C)
Contribution of ingested prey carbon to growth (for cell volume to carbon content
conversions), assuming 33% growth efficiency as a function of prey concentration.
All data points are means of triplicates (SE). Broken lines denote 99% confidence
intervals. LL: low light; HL: high light. Where no error bars are shown, the error was
smaller than the symbol.
After Smith and Hansen (2007) with permission.
Models are intended to present simplifications of the real world,
enhancing our understanding of it by removing extraneous detail.
Modeling mixotrophs is a particular challenge because of the
complexity of the organisms and because inclusion of the
‘‘extraneous’’ detail may actually prove to be essential. Most effort
has been expended on models that describe mixotrophs that are
constitutive phototrophs (species with permanent chloroplasts).
Only the model of Flynn and Mitra (2009) provides a platform for
exploring the contrasts between constitutive mixotrophs, and
those (such as Mesodinium and Dinophysis) that are specialist nonconstitutive mixotrophs (thus acquiring chloroplasts from
ingested prey). While the Flynn and Mitra (2009) model is not
ideal for this particular application, its operation reveals a need to
explicitly simulate acquired phototrophy (Mitra and Flynn, 2010).
Indeed, considering the dynamics of the interaction between an
organism that depends on a specific ‘‘prey’’ species, not only as a
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
137
Fig. 11. Dinophysis acuminata. Short-term experiments showing mixotrophic growth rates and (b) ingestion rate of the dinoflagellate as a function of light intensity. Data
points are shown as mean SE for three replicates. Curves were fitted by a modified Michaelis–Menten equation (Eq. 2) for growth rates and a Michaelis–Menten equation for
ingestion rates. Mixotrophic growth rate (mmix, d 1) = 0.78(x 0.38)/[21.50 + (x 0.38)], r2 = 0.92. Ingestion rate (pg C Dinophysis 1 d 1) = 2265.46x/(27.54 + x), r2 = 0.98.
After Kim et al. (2008) with permission.
source of nutrients but for basal physiological capacity, requires
not only a model of the mixotroph, but also of its prey. In this
capacity we lack information to configure a model describing the
growth of the specialist cryptophyte prey for the Mesodinium, and
of course for Dinophysis we lack the Mesodinium model.
The basis of models brings our ignorance of the processes into
sharp relief. The currency for plankton models is either elements or
some combination of elements and cell number (thus involving cell
size). The immediacy of the challenge for mixotrophs can be judged
by reference to the figures accompanying this manuscript; as is
common for work in this arena, units are often referenced to cells,
with no capacity for deriving the mass-specific rate data required
for models. A need to follow cell size can be readily made for
considering the growth of these organisms, but this introduces
practical problems associated with determining the contribution
of prey biomass (within food vacuoles, or for these organisms,
distributed within the mixotroph proper) to the whole mixotroph
biomass as an experimental biologist would measure it.
As the predator releases nutrients that are re-assimilated into
the prey, thus affecting prey quality, there is considerable scope for
a series of trophic interactions with positive and/or negative
interactions (Mitra and Flynn, 2006). To understand this in its
entirety would require a full parameterization with respect to cells,
C, N, P, Chl and perhaps even Fe, for the cryptophyte first.
Cryptophytes of known nutrient history (nutrient and illumination
history) would then need to be supplied to the Mesodinium and the
resultant interaction followed with changes in predator and prey
numbers and size, external nutrients, and as far as possible internal
elemental and pigment content, all followed over time. Similar
interactions would be required starting with different prey status
(cryptophyte plus/minus other items), fed to Mesodinium of
different initial nutrient history. For Dinophysis, all this would
be required first, before considering the effect of providing
Mesodinium and other prey to Dinophysis itself.
Because of the dynamics surrounding the fate of the acquired
plastids, models intended for a full mechanistic description of
these mixotrophs need not only to be multi-element (i.e. C, N, P,
(Fe)), but they require a modification of the description of
photosynthesis and photosystem regulation from that which is
commonly employed (Geider et al., 1998; Flynn, 2001). Thus, the
state variable describing Chl:C, with its attendant constants, are
now acquired from consumption of the prey, and over time decay
with the dynamics seen in cells of the mixotroph. One approach
here may be to consider that photoacclimation is allowed up to a
maximum value (set by Chl:Cmax), but over time, and perhaps as
functions of nutrient history and light exposure, the value of
Chl:Cmax decays. It is restored by feeding on the specialist prey
(cryptophyte or Mesodinium for Mesodinium or Dinophysis,
respectively). How the quality of the Mesodinium prey affects
the acquired phototrophic potential of Dinophysis is a question that
comes from such an interpretation.
The challenges of the experimental regime described above
invites an additional approach, that of using phenomenological
information to guide model construction and testing. This
approach ensures that model structure and operation are
consistent with understanding. In this context, the current
understanding, as reviewed in this article, provides a good
platform for future modeling work. Without doubt, the construction of models for these organisms will then drive a cycle of
experimental and conceptual/theoretical thinking that will further
enhance science in this arena. In natural conditions match and
mismatch of Mesodinium with prey and predator, together with
physical processes (e.g. Sjöqvist and Lindholm, 2011), all conspire
to further complicate the issue. However, the first step remains to
construct a model capable of describing interactions in a welldefined set of conditions in the laboratory.
Acknowledgments
This work was financially supported by a Leverhulme Trust
International Workshop and by a grant project no. 2101-07-0084
from The Danish Council for Strategic Research to PJH and LTN.
[SS]
References
Abé, T.H., 1967. The armoured dinoflagellata: II. Prorocentridae and dinophysidae
(B)—dinophysis and its allied genera. Publication Seto Marine Biological Laboratory 15, 37–78.
Arnon, D.I., 1988. The discovery of ferredoxin: the photosynthetic path. Trends in
Biochemical Sciences 13, 30–33.
Balech, E., 1976. Some Norwegian Dinophysis species (Dinoflagellata). Sarsia 61, 75–
94.
Bass, D., Brown, N., Mackenzie-Dodds, J., Dyal, P., Nierzwicki-Bauer, S.A., Vepritskiy,
A.A., Richards, T.A., 2009. Molecular perspective on ecological differentiation
and biogeography of cyclotrichiid ciliates. Journal of Eukaryotic Microbiology
56, 559–567.
Dolan, J.R., Coats, D.W., 1991. A study of feeding in predacious ciliates using prey
ciliates labelled with fluorescent microspheres. Journal of Plankton Research 13
(3) 609–627.
Durnford, D., Koziol, A., Borza, T., Ishida, K., Keeling, P., Lee, R., 2007. Tracing the
evolution of the light-harvesting antennae in chlorophyll a/b containing organisms. Photosynthesis Research 91, 271–279.
Elbrächter, M., 1991. Food uptake mechanisms in phagotrophic dinoflagellates and
classification. In: Patterson, D.J., Larsen, J. (Eds.), The Biology of Free-Living
Heterotrophic Flagellates. Systematics Association Special, Vol. 45. Clarendon
Press, Oxford, pp. 303–312.
Elrad, D., Grossman, A.R., 2004. A genome’s-eye view of the light-harvesting
polypeptides of Chlamydomonas reinhardtii. Current Genetics 45, 61–75.
Enami, I., Okumura, A., Nagao, R., Suzuki, T., Iwai, M., Shen, J.R., 2008. Structures and
functions of the extrinsic proteins of photosystem II from different species.
Photosynthesis Research 98, 349–363.
138
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Eppard, M., Krumbein, W.E., von Haeseler, A., Rhiel, E., 2000. Characterization of
fcp4 and fcp12, two additional genes encoding light harvesting proteins of
Cyclotella cryptica (Bacillariophyceae) and phylogenetic analysis of this complex
gene family. Plant Biology 2 (3) 283–289.
Esteban, G.F., Fenchel, T., Finlay, B.J., 2010. Mixotrophy in Ciliates. Protist 161 (5)
621–641.
Fenchel, T., Hansen, P.J., 2006. Swimming behaviour in Mesodinium rubrum. Marine
Biology Research 2, 33–40.
Ferreira, K.N., Iverson, T.M., Maghlaoui, K., Barber, J., Iwata, S., 2004. Architecture of
thephotosynthetic oxygen-evolving center. Science 303 (5665) 1831–1838.
Flynn, K.J., 2001. A mechanistic model for describing dynamic multi-nutrient, light,
temperature interactions in phytoplankton. Journal of Plankton Research 23 (9)
977–997.
Flynn, K.J., Mitra, A., 2009. Building the perfect beast: modelling mixotrophic
plankton. Journal of Plankton Research 31 (9) 965–992.
Foissner, W., Berger, H., Schaumburg, J., 1999. Identification and ecology of limnetic
plankton ciliates.In: Informationsberichte des Bayerischen Landesamtes für
Wasserwirtschaft, Heft 3/99. , pp. 793.
Garcı́a-Cuetos, L., Moestrup, O., Hansen, P.J., Daugbjerg, N., 2010. The toxic dinoflagellate Dinophysis acuminata harbors permanent chloroplasts of cryptomonad
origin, not kleptochloroplasts. Harmful Algae 9, 25–38.
Garcı́a-Cuetos, L., Moestrup, Ø., Hansen, P.J., 2012. Studies on the genus Mesodinium
II. Ultrastructural and molecular investigations of five marine species help
clarifying the taxonomy. Journal of Eukaryotic Microbiology 59 (4) 374–400.
Geider, R.J., MacIntyre, H.L., Kana, T.M., 1998. A dynamic regulatory model of
phytoplanktonic acclimation to light nutrients and temperature. Limnology
and Oceanography 43 (4) 679–694.
Gómez, F., 2005. A list of free-living dinoflagellate species in the world’s oceans.
Acta Botanica Croatia 64, 129–212.
Gustafson, D.E., Stoecker, D.K., Johnson, M.D., Van Heukelem, W.F., Sneider, K., 2000.
Cryptophyte algae are robbed of their organelles by the marine ciliate Mesodinium rubrum. Nature 405 (6790) 1049–1052.
Hackett, J.D., Maranda, L., Yoon, H.S., Bhattacharya, D., 2003. Phylogenetic evidence
for the cryptophyte origin of the plastid of Dinophysis (Dinophysiales, Dinophyceae). Journal of Phycology 39 (2) 440–448.
Hallegraeff, G.M., Lucas, I.A.N., 1988. The marine dinoflagellate genus Dinophysis
(Dinophyceae): photosynthetic, neritic and non-photosynthetic, oceanic species. Phycologia 27 (1) 25–42.
Handy, S.M., Bachvaroff, T.R., Timme, R.E., Coats, D.W., Kim, S., Delwiche, C., 2009.
Phylogeny of four dinophysiacean genera (Dinophyceae, Dinophysiales) based
on rDNA sequences from single cells and environmental samples. Journal of
Phycology 45, 1163–1174.
Hansen, P.J., 1991. Dinophysis: a planktonic dinoflagellate genus which can act both
as a prey and as a predator of a ciliate. Marine Ecology Progress Series 69, 201–
204.
Hansen, P.J., Fenchel, T., 2006. The bloom-forming ciliate Mesodinium rubrum
harbours a single permanent endosymbiont. Marine Biology Research 2,
169–177.
Hansen, P.J., 2011. The role of photosynthesis and food uptake for the growth of
marine mixotrophic dinoflagellates. Journal of Eukaryotic Microbiology 58 (3)
203–214.
Hansen, P.J., Moldrup, M., Tarangkoon, W., Garcı́a-Cuetos, L., Moestrup, Ø., 2012.
Direct evidence for symbiont sequestration in the marine red tide ciliate
Mesodinium rubrum. Aquatic Microbial Ecology 66, 63–75.
Herfort, L., Peterson, T.D., McCue, L.A., Crump, B.C., Prahl, F.G., Baptista, A.M.,
Campbell, V., Warnick, R., Selby, M., Roegner, G.C., Zuber, P., 2011. Myrionecta
rubra population genetic diversity and its cryptophyte chloroplast specificity in
recurrent red tides in the Columbia River estuary. Aquatic Microbial Ecology 62,
85–97.
Hibberd, D.J., 1977. Ultrastructure of the cryptomonad endosymbiont of the redwater ciliate Mesodinium rubrum. Journal of Marine Biological Association UK
57, 45–61.
Jakobsen, H.H., Everett, L.M., Strom, S.L., 2006. Hydromechanical signaling between
the ciliate Mesodinium pulex and motile protist prey. Aquatic Microbial Ecology
44, 197–206.
Janson, S., 2004. Molecular evidence that plastids in the toxinproducing dinoflagellate genus Dinophysis originate from the free-living cryptophyte Teleaulax
amphioxeia. Environmental Microbiology 6 (10) 1102–1106.
Jensen, M.A., Daugbjerg, N., 2009. Molecular phylogeny of selected species of the
order Dinophysiales (Dinophyceae) – testing the hypothesis of a Dinophysioid
radiation. Journal of Phycology 45 (5) 1136–1152.
Jiang, H., 2011. Why does the jumping ciliate Mesodinium rubrum possess an
equatorially located propulsive ciliary belt? Journal of Plankton Research
33, 998–1011.
Johnson, M.D., Tengs, T., Oldach, D., Stoecker, D.K., 2004. Highly divergent SSU rRNA
genes found in the marine ciliates Myrionecta rubra and Mesodinium pulex.
Protist 155, 347–359.
Johnson, M.D., Stoecker, D.K., 2005. Role of feeding in growth and photophysiology
of Myrionecta rubra. Aquatic Microbial Ecology 39, 303–312.
Johnson, M.D., Tengs, T., Oldach, D., Stoecker, D.K., 2006. Sequestration, performance, and functional control of cryptophyte plastids in the ciliate Myrionecta
rubra (Ciliophora). Journal of Phycology 42, 1235–1246.
Johnson, M.D., Oldach, D., Delwiche, C.F., Stoecker, D.K., 2007. Retention of transcriptionally active cryptophyte nuclei by the ciliate Myrionecta rubra. Nature
445, 426–428.
Johnson, M.D., 2011a. The acquisition of phototrophy: adaptive strategies of hosting
endosymbionts and organelles. Photosynthesis Research 107, 117–132.
Johnson, M.D., 2011b. Acquired phototrophy in ciliates: a review of cellular interactions and structural adaptations. Journal of Eukaryotic Microbiology 58 (3)
185–195.
Jonsson, P., Tiselius, P., 1990. Feeding behaviour, prey detection and capture
efficiency of the copepod Acartia tonsa feeding on planktonic ciliates. Marine
Ecology Progress Series 60, 35–44.
Kim, M., Kim, S., Yih, W., Park, M.G., 2011. The marine dinoflagellate genus
Dinophysis can retain plastids of multiple algal origins at the same time.
Harmful Algae 13, 105–111.
Kim, M., Nam, S.W., Shin, W., Coats, D.W., Park, M.G., 2012. Dinophysis caudata
(Dinophyceae) sequesters and retains plastids from the mixotrophic ciliate prey
Mesodinium rubrum. Journal of Phycology 48 (3) 569–579.
Kim, S., Kang, Y.G., Kim, H.S., Yih, W., Coats, D.W., Park, M.G., 2008. Growth and
grazing responses of the mixotrophic dinoflagellate Dinophysis acuminata as
functions of light intensity and prey concentration. Aquatic Microbial Ecology
51, 301–310.
Koike, K., Sekiguchi, H., Kobiyama, A., Kawachi, M., Ogata, T., 2005. A novel type of
kleptoplastidy in Dinophysis (Dinophyceae): presence of haptophyte-type plastid in Dinophysis mitra. Protist 156, 225–237.
Lindholm, T., 1985. Mesodinium rubrum—a unique photosynthetic ciliate.
Advances in Aquatic Microbiology 3, 1–48.
Lindholm, T., Lindros, P., Mörk, A.-C., 1988. Ultrastructure of the photosynthetic
ciliate Mesodinium rubrum. Biosystems 21 (2) 141–149.
Lynn, D.H., 2010. Ciliated Protozoa: Characterization, Classification, and Guide to
the Literature. Springer, Dordrecht, pp. 605.
Martinez-Duncker, I., Mollicone, R., Codogno, P., Oriol, R., 2003. The nucleotide
sugar transporter family: a phylogenetic approach. Biochimie 85 (3–4) 245–
260.
Minnhagen, S., Janson, S., 2006. Genetic analyses of Dinophysis spp. support kleptoplastidy. FEMS Microbial Ecology 57 (1) 47–54.
Minnhagen, S., Carvalho, W.F., Salomon, P.S., Janson, S., 2008. Chloroplast DNA
content in Dinophysis (Dinophyceae) from different cell cycle stages is consistent with kleptoplasty. Environmental Microbiology 10 (9) 2411–2417.
Minnhagen, S., Kim, M., Salomon, P.S., Yih, W., Graneli, E., Park, M.G., 2011. Active
uptake of kleptoplastids by Dinophysis caudata from its ciliate prey Myrionecta
rubra. Aquatic Microbial Ecology 62, 99–108.
Mitra, A., Flynn, K.J., 2006. Promotion of harmful algal blooms by zooplankton
predatory activity. Biology Letters 2, 194–197.
Mitra, A., Flynn, K.J., 2010. Modelling mixotrophy in harmful algal blooms: more or
less the sum of the parts? Journal of Marine Systems 83, 158–169.
Moeller, H.V., Johnson, M.D., Falkowski, P.G., 2011. Photoacclimation in the phototrophic marine ciliate Mesodinium rubrum (Ciliophora). Journal of Phycology 47
(2) 324–332.
Moestrup, Ø., Garcı́a-Cuetos, L., Hansen, P.J., Fenchel, T., 2012. Studies on the genus
Mesodinium I. Ultrastructure and description of M. chamaeleon sp. nov., a
benthic marine species with green or red chloroplasts. Journal of Eukaryotic
Microbiology 59 (1) 20–39.
Myung, G., Yih, W., Kim, H.S., Park, J.S., Cho, B.C., 2006. Ingestion of bacterial cells by
the marine photosynthetic ciliate Myrionecta rubra. Aquatic Microbial Ecology
44, 175–180.
Myung, G., Kim, H.S., Park, J.S., Park, M.G., Yih, W., 2011. Population growth and
plastid type of Myrionecta rubra depend on the kinds of available cryptomonad
prey. Harmful Algae 10 (5) 536–541.
Nagai, S., Nitshitani, G., Tomaru, Y., Sakiyama, S., Kamiyama, T., 2008. Predation by
the toxic dinoflagellate Dinophysis fortii on the ciliate Myrionecta rubra and
observation of sequestration of ciliate chloroplasts. Journal of Phycology 44 (4)
909–922.
Nam, S.W., Shin, W., Coats, D.W., Park, J.W., Yih, W., 2012. Ultrastructure of the Oral
Apparatus of Mesodinium rubrum from Korea. Journal of Eukaryotic Microbiology 59 (6) 625–636.
Nielsen, L.T., Krock, B., Hansen, P.J., 2012. Influence of irradiance and food availability on DSP toxin production, growth and photosynthesis in Dinophysis
acuminata. Marine Ecology Progress Series 471, 34–50.
Nielsen, L.T., Krock, B., Hansen, P.J., 2013. Production and excretion of okadaic acid,
pectenotoxin-2 and a novel dinophysistoxin from the DSP-causing marine
dinoflagellate Dinophysis acuta—effects of light, food availability and growth
phase. Harmful Algae 23, 34–45.
Nishitani, G., Nagai, S., Sakiyama, S., Kamiyama, T., 2008a. Successful cultivation of
the toxic dinoflagellate Dinophysis caudata (Dinophyceae). Plankton Benthos
Research 3, 78–85.
Nishitani, G., Nagai, S., Takano, Y., Sakiyama, S., Baba, K., Kamiyama, T., 2008b.
Growth characteristics and phylogenetic analysis of the marine dinoflagellate
Dinophysis infundibulus (Dinophyceae). Aquatic Microbial Ecology 52, 209–
221.
Nishitani, G., Nagai, S., Baba, K., Kiyokawa, S., Kosaka, Y., Miyamura, K., Nishikawa,
T., Sakurada, K., Shinada, A., Kamiyama, T., 2010. High-Level congruence of
Myrionecta rubra prey and Dinophysis species plastid identities as revealed by
genetic analyses of isolates from Japanese coastal waters. Applied and Environmental Microbiology 76 (9) 2791–2798.
Nishitani, G., Nagai, S., Hayakawa, S., Kosaka, Y., Sakurada, K., Kamiyama, T.,
Gojobori, T., 2012. Multiple plastids collected by the dinoflagellate Dinophysis
mitra through kleptoplastidy. Applied and Environmental Microbiology 78 (3)
813–821.
P.J. Hansen et al. / Harmful Algae 28 (2013) 126–139
Oakley, B.R., Taylor, F.J.R., 1978. Evidence for a new type of endosymbiotic organization in a population of the ciliate Mesodinium rubrum from British Columbia.
BioSystems 10, 361–369.
Packard, T., Blasco, D., Barber, R., 1978. Mesodinium rubrum in the Baja California
upwelling system. In: Boje, R., Tomczak, M. (Eds.), Upwelling Ecosystems.
Springer-Verlag, Berlin, pp. 73–89.
Park, M.G., Kim, S., Kim, H.S., Myung, G., Kang, Y.G., 2006. First successful culture of
the marine dinoflagellate Dinophysis acuminata. Aquatic Microbial Ecology 45,
101–106.
Park, M.G., Park, J.S., Kim, M., Yih, W., 2008. Plastid dynamics during survival of
Dinophysis caudata without its ciliate prey. Journal of Phycology 44 (5) 1154–
1163.
Park, M.G., Kim, M., Kim, S., Yih, W., 2010. Does Dinophysis caudata (Dinophyceae)
have permanent plastids? Journal of Phycology 46 (2) 236–242.
Park, J.S., Myung, G., Kim, H.S., Cho, B.C., Yih, W., 2007. Growth responses of the
marine photosynthetic ciliate Myrionecta rubra to different cryptomonad
strains. Aquatic Microbial Ecology 48, 83–90.
Reguera, B., Velo-Suarez, L., Raine, R., Park, M.G., 2012. Harmful Dinophysis species: a
review. Harmful Algae 14, 87–106.
Richard, C., Ouellet, H., Guertin, M., 2000. Characterization of the LI818 polypeptide
from the green unicellular alga Chlamydomonas reinhardtii. Plant Molecular
Biology 42, 303–316.
Riisgaard, K., Hansen, P.J., 2009. Role of food uptake for photosynthesis, growth and
survival of the mixotrophic dinoflagellate Dinophysis acuminata. Marine Ecology
Progress Series 381, 51–62.
Riisgård, H.U., Larsen, P.S., 2009. Ciliary-propelling mechanism, effect of temperature and viscosity on swimming speed, and adaptive significance of ‘jumping’ in the ciliate Mesodinium rubrum. Marine Biology Research 5, 585–
595.
Rodriguez, F., Escalera, L., Reguera, B., Rial, P., da Silva, T.D., 2012. Morphological
variability, toxicology and genetics of the dinoflagellate Dinophysis tripos
(Dinophysiaceae, Dinophysiales). Harmful Algae 13, 26–33.
Seeyave, S., Probyn, T.A., Pitcher, G.C., Lucas, M.I., Purdie, D.A., 2009. Nitrogen
nutrition in assemblages dominated by Pseudo-nitzschia spp., Alexandrium
139
catenella and Dinophysis acuminata off the west coast of South Africa. Marine
Ecology Progress Series 379, 91–107.
Sjöqvist, C.O., Lindholm, T., 2011. Natural co-occurrence of Dinophysis acuminata
(Dinoflagellata) and Mesodinium rubrum (Ciliophora) in thin Layers in a coastal
inlet. Journal of Eukaryotic Microbiology 58, 365–372.
Smith, M., Hansen, P.J., 2007. Interaction between Mesodinium rubrum and its prey:
importance of prey concentration, irradiance and pH. Marine Ecology Progress
Series 338, 61–70.
Steidinger, K.A., Tangen, K., 1996. Dinoflagellates. In: Tomas, C.R. (Ed.), Identifying
Marine Diatoms and Dinoflagellates. Academic Press, San Diego, pp. 387–598.
Stoecker, D.K., Johnson, M.D., de Vargas, C., Not, F., 2009. Acquired phototrophy in
aquatic protists. Aquatic Microbial Ecology 57, 279–310.
Stoecker, D.K., Michaels, A.E., Davis, L.H., 1987. Large proportion of marine planktonic ciliates found to contain functional chloroplasts. Nature 326, 790–792.
Strom, S.L., 2001. Light-aided digestion, grazing and growth in herbivorous protists.
Aquatic Microbial Ecology 23, 253–261.
Takishita, K., Koike, K., Maruyama, T., Ogata, T., 2002. Molecular evidence for plastid
robbery (kleptoplastidy) in Dinophysis, a dinoflagellate causing diarrhetic shellfish poisoning. Protist 153, 293–302.
Tarangkoon, W., Hansen, P.J., 2011. Prey selection, ingestion and growth responses
of the common marine ciliate Mesodinium pulex in the light and in the dark.
Aquatic Microbial Ecology 62, 25–38.
Taylor, F.J.R., Blackbourn, D.J., Blackbourn, J., 1971. Red water ciliate Mesodinium
rubrum and its incomplete symbionts: review including new ultrastructural
observations. Journal of Fisheries Research Board Canada 28, 391–407.
Velo-Suárez, L., González-Gil, S., Pazos, Y., Reguera, B., 2013. The growth season of
Dinophysis acuminata in an upwelling system embayment: a conceptual model
based on in situ measurements. Deep-Sea Research II (in press).
Wilkerson, F.P., Grunseich, G., 1990. Formation of blooms by the symbiotic ciliate
Mesodinium rubrum: the significance of nitrogen uptake. Journal of Plankton
Research 12, 973–989.
Wisecaver, J.H., Hackett, J.D., 2010. Transcriptome analysis reveals nuclear-encoded
proteins for the maintenance of temporary plastids in the dinoflagellate Dinophysis acuminata. BMC Genomics 11, 366–375.