Accepted by Journal of Eukaryotic Microbiology (2008.04)
JEONG ET AL.---Feeding by heterotrophic dinoflagellates on bacteria
Feeding and grazing impact by small marine heterotrophic dinoflagellates on hetertrophic
bacteria
HAE JIN JEONGa,*, KYEONG AH SEONGb, YEONG DU YOOa, TAE HOON KIMb,
NAM SEON KANGa, SHIN KIMb, JAE YEON PARKa, JAE SEONG KIMc, KWANG
HOON KIMd, JAE YOON SONGa
aSchool of Earth and Environmental Sciences, College of Natural Sciences, Seoul National
University, Seoul 151-747, Republic of Korea, and
bDepartment of Oceanography, College of Ocean Science and Technology, Kunsan National
University, Kunsan 573-701, Republic of Korea, and
cRed Tide Research Center, College of Ocean Science and Technology, Kunsan National
University, Kunsan 573-701, Republic of Korea, and
dDepartment of Biology, College of Natural Sciences, Kongju National University, Kongju 314-
701, Republic of Korea
Corresponding Author: H.J. Jeong – Telephone number: +82-2-880-6746; FAX number: +82-2-
874-9695; Email: [email protected]
ABSTRACT: We investigated the feeding of the small heterotrophic dinoflagellates Oxyrrhis
marina, Gyrodinium cf. guttula, Gyrodinium sp., Pfiesteria piscicida, and Protoperidinium bipes
on marine heterotrophic bacteria. To investigate whether they are able to feed on bacteria, we
observed the protoplasm of target heterotrophic dinoflagellate cells under an epifluorescence
microscope and transmission electron microscope. In addition, we measured ingestion rates of
the dominant heterotrophic dinoflagellate, Gyrodinium spp., on natural populations of marine
bacteria (mostly heterotrophic bacteria) in Masan Bay, Korea in 2006--2007. Furthermore, we
measured the ingestion rates of O. marina, G. cf. guttula, and P. piscicida on bacteria as a
function of bacterial concentration under laboratory conditions. All heterotrophic dinoflagellates
tested were able to feed on a single bacterium. O. marina and Gyrodinium spp. intercepted and
then ingested a single bacterial cell in feeding currents that were generated by the flagella of the
predators. During the field experiments, the ingestion rates and grazing coefficients of
Gyrodinium spp. on natural populations of bacteria were 14--61 bacteria dinoflagellate-1
h-1
and
0.003--0.972 d-1
, respectively. With increasing prey concentration, the ingestion rates of O.
marina, G. cf. guttula, and P. piscicida on bacteria increased rapidly at prey concentrations of ca.
0.7--2.2 x 106 cells ml
-1, but increased only slowly or became saturated at higher prey
concentrations. The maximum ingestion rate of O. marina on bacteria was much higher than
those of G. cf. guttula and P. piscicida. Bacteria alone supported the growth of O. marina. The
results of the present study suggest that some heterotrophic dinoflagellates may sometimes have
a considerable grazing impact on populations of marine bacteria, and that bacteria may be
important prey.
2
Key Words. Bacterivory, Feeding behavior, Food web, Graze, Ingestion, Microbial Loop, Protist
INTRODUCTION
Heterotrophic dinoflagellates (HTDs) are one of the major components in marine
planktonic food webs (Jeong 1999). They are ubiquitous protists, often abundant, and may
dominate heterotrophic protistan assemblages (Jacobson 1987; Lessard 1984). They play diverse
ecological roles in marine planktonic food webs (Hansen 1992; Lessard 1991; Sherr and Sherr
2002, 2007; Strom, Macri, and Olson 2007); HTDs are known to feed on a diverse array of prey
species, such as diatoms (Jacobson and Anderson 1986; Menden-Deuer et al. 2005),
phototrophic nanoflagellates (Jakobsen and Hansen 1997; Parrow and Burkholder 2004),
mixotrophic dinoflagellates (Adolf et al. 2007; Tillmann 2004), ciliates (Hansen 1991), and the
eggs, early naupliar stages, and adult forms of metazoans (Jeong 1994; Park et al. 2007). In turn,
HTDs are also known to be important prey for several planktonic consumers, such as ciliates
(Jeong et al. 2004a; Stoecker et al. 2002) and copepods (Koski and Riser 2006). Recently, new
prey items for HTDs, such as heterotrophic nanoflagellates, other HTDs, and fish bloods, have
been reported and thus the roles of HTDs as predators appears to be more diverse (Jeong et al.
2007a, b, c). Therefore, to understand the roles of HTDs in marine planktonic food webs,
unknown interactions between HTDs and potential prey should continue to be explored.
3
Marine heterotrophic bacteria are also one of the major components in marine ecosystems
and sometimes dominate marine plankton biomasses or particulate organic carbon (Cho and
Azam 1990; Delille 2004; Kamiyama, Itakura, and Nagasaki 2000; Selje, Simon, and Brinkhoff
2004; Sherr, Sherr, and Verity 2002). So far, red-tide algae, heterotrophic nanoflagellates
(HNFs), and small ciliates are known to be major predators of marine heterotrophic bacteria and
occasionally control populations of these bacteria (Christaki et al. 2001; Seong et al. 2006;
Vargas et al. 2007). Also, some amoebae have been known to feed on bacteria (Weekers et al.
1993). However, there are few studies on the feeding of HTDs on marine heterotrophic bacteria;
Lessard and Swift (1985) reported the feeding by natural populations of the heterotrophic
dinoflagellates Protoperidinium spp., Podolampas spp., and Diplopsalis spp. on bacteria
prelabelled with tritiated-thymidine (3H-thymidine). Also, Burkholder and Glasgow (1995)
reported that Pfiesteria piscicida fed on bacteria without detailed descriptions. However, these
studies neither reported the feeding behavior of HTDs on bacteria nor provided TEM pictures
showing ingested bacterial cells in the food vacuoles inside the protoplasm of predator cells.
Some argue that signs of bacteria inside the HTD predators might be derived from detritus to
which the bacteria are attached (i.e. by accident or byproduct). In light of this, a series of
important questions arises. (1) Are heterotrophic dinoflagellates able to feed on a single
bacterium? Many think that bacteria are too small to be eaten by large heterotrophic
dinoflagellates. (2) If heterotrophic dinoflagellates are able to feed on a single bacterium, what
are the feeding behaviors (mechanisms) of HTDs on these small bacterial cells? Also, are the
feeding behaviors of the HTDs on bacteria the same as, or very similar to, those on algal prey?
4
There are many studies on the feeding processes of heterotrophic nanoflagellates (HNFs) and
ciliates on bacteria (Fenchel 1987; Sleigh 1989; Monger and Landry 1990; Boenigk and Arndt
2000, 2002; KiØrboe et al. 2004), while there have been no reports on the feeding processes of
HTDs on bacteria. (3) What are the ingestion rates and grazing impact of HTDs on natural
populations of bacteria? Are these rates comparable to those of co-occurring red-tide algae,
HNFs, or ciliates in natural environments? What is the functional response of HTDs to bacterial
concentrations? (4) Do bacteria alone support the growth of HTDs?
To answer these questions, in the present study (1) we investigated whether five common
HTDs (i.e. the engulfment feeding dinoflagellates Oxyrrhis marina, Gyrodinium cf. guttula, and
Gyrodinium sp., the peduncle feeding Pfiesteria piscicida, and the pallium feeding
Protoperidinium bipes) are or are not able to feed on marine heterotrophic bacteria. We carefully
observed inside protoplasm of target HTD cells under an epifluorescence microscope and a TEM
after adding living bacterial cells, fluorescent labeled bacteria (FLB), bacteria-sized fluorescent
beads (0.5 μm in diameter), and/or non-fluorescent beads . Also, (2) we explored the feeding
behaviors of HTDs on bacteria using high resolution video-microscopy and by observation under
several different types of microscopes. (3) We measured ingestion rates and grazing coefficients
of the dominant heterotrophic dinoflagellates Gyrodinium spp. and co-occurring red-tide algae,
HNFs, or ciliates on natural populations of marine bacteria in Masan Bay in 2006--2007. (4) We
also examined the functional responses of heterotrophic dinoflagellates to bacteria as a function
of bacterial concentration in the laboratory and measured the growth rates of O. marina when
only bacteria were provided as prey. The results of the present study provide a basis for
5
understanding the interactions between HTDs and marine bacteria and the relative contribution
of HTDs, red-tide algae, HNFs, and ciliates as predators on bacteria.
MATERIALS AND METHODS
Preparation of experimental organisms. For the isolation and culture of Gyrodinium cf.
guttula, Gyrodinium sp., Oxyrrhis marina, Pfiesteria piscicida, and Protoperidinium bipes, using
Niskin water samplers, plankton samples were collected from the surface waters off Incheon, the
Keum River Estuary, and in Masan Bay, Korea (Table 1). The mixotrophic dinoflagellates
Amphidinium carterae and Prorocentrum minimum, or the diatom Skeletonema costatum were
provided as prey and a clonal culture of each predator species was established by two serial
single cell isolations, as described in Jeong et al. (2003, 2004b, 2006) and Kim and Jeong (2004).
These predator cultures were maintained at 20o C for > 3 month before these experiments were
conducted.
Feeding occurence. Expt 1 was designed to test whether or not each of the HTDs O.
marina, G. cf. guttula, Gyrodinium sp., P. piscicida, and P. bipes was able to feed on marine
heterotrophic bacteria under laboratory conditions. In this experiment, we observed bacterial
cells and beads inside the predators using epifluorescence microscopy and TEM, after adding
bacteria-sized fluorescent beads (0.5 μm in diameter; Polysciences, Inc., Warrington, PA, USA)
and fluorescent-labeled bacteria (FLB) for epifluorescence microscopy and bacteria-sized non-
fluorescent beads (0.5 μm) and living bacteria for transmission electron microscopy (TEM).
For the epifluorescence microscopy, the heterotrophic bacterial cells (FLB) that originated
6
from a clonal culture of a target HTD were fluorescently labeled, following Sherr, Sherr, and
Fallon (1987). To remove any aggregated FLB, the FLB were dispersed throughout the medium
using a sonicator (Bransonic cleaner 5510E-DTH, Danbury, CT, USA) for 10--30 min and then
filtered through 3-μm pore sized filter (Whatman, Polycarbonate, Maidstone, UK). In a similar
manner, the bacteria-sized fluorescent beads were also de-aggregated and then filtered. These
processes resulted in all the bacteria and > 99% of the beads being single and unattached (Fig. 1,
2). In the cases of attached beads, two or three beads were clumped. On TEM photos, the non-
fluorescent beads had a dark shell and also homogenously grey-colored contents beneath the
shell, while the bacteria had a bright grey-colored cell wall and heterogeneously grey-colored
contents with a white colored part near the center of the cell. Due to this difference, we were
easily able to distinguish beads from bacteria inside the protoplasm of HTDs under TEM (Fig. 3,
4, 16--18).
Approximately 2 x 109 FLB cells (or approximately 2 x 109 bacteria-sized fluorescent
beads) were added into each of three 270-ml PC bottles, each containing a target HTD at 3,000--
10,000 cells ml-1. One ‘bead only’ control bottle, one ‘FLB only’ control bottle, and one HTD
control bottle (without added FLB or beads) were set up for each experiment. The bottles were
placed on a plankton wheel rotating at 0.9 rpm and incubated at 20o C under a continuous
illumination of 30 µE m-2 s-1 of cool white fluorescent light. At the beginning, and after 5, 10, 20,
30 min and 1 and 2 h incubation periods, 10-ml aliquots were removed from each bottle,
transferred into 20-ml vials, and then fixed with borate-buffered formalin [final conc. = 4%
(v/v)] for FLB observation. The fixed samples were stained using 4`6`-diamidino-2-phenylindole
7
(DAPI; final conc. = 1 µM) and then filtered onto 5 µm-pore-sized PC white membrane filters.
Approximately 300--1,000 concentrated FLB cells on the membranes were observed under an
epifluorescence microscope (Ziess-Axiovert 200M, Carl Zeiss Ltd., Göttingen, Germany) at a
magnification of 1,000x and pictures were taken. For bead observation, all the contents from the
bottle were partially filtered onto 5 µm-pore-sized polycarbonate (PC) white membrane filters
until 5 ml water was retained in a filtering set. Filtered seawater was added into the retained
water and filtered again until another 5 ml water remained. The washing process was repeated 3
times. This was done to remove as many unfed beads as possible while not damaging the cells.
The last retained 5 ml water was transferred into a 20-ml vial, 1-2 drops of the water were placed
on a slide, and then examined under an epifluorescence microscope at a magnification of 640--
1,000x with oil.
For TEM, living heterotrophic bacterial cells, originating from a clonal culture of a target
HTD, and bacteria-sized non-fluorescent beads were used. Approximately 2 x 109 living bacterial
cells and/or approximately 2 x 109 beads were added into each of three 270 ml PC bottles, each
containing a target HTD of 3,000--10,000 cells ml-1. One ‘bead only’ control bottle, one ‘bacteria
only’ control bottle, and one HTD control bottle (without added bacteria or beads) were set up
for each experiment. The bottles were placed on a plankton wheel rotating at 0.9 rpm and
incubated, as described above. At the beginning, and after 5, 10, 20, 30 min and 1 and 2 h
incubation periods (for the experimental bottles), the contents of one experimental bottle from
each interval were distributed into five 50-ml centrifugal tubes and then concentrated at 20,000 g
for 10 min using a centrifuge (Vision Centrifuge VS-5500, Vision Scientific Co., Bucheon,
8
Korea). Five pellets from the 5 centrifugal tubes were then transferred into 1.5-ml tubes and
fixed for 1.5 h in 4% (w/v) glutaraldehyde in culture medium. After this, the fixative was
removed and the pellets were rinsed using a 0.2 M cacodylic acid/sodium salt solution (pH 7.4).
The pellet was then embedded in agar. After several rinses with medium, the cells were postfixed
in 1% (v/v) osmium tetroxide in deionized water. Dehydration was accomplished using a graded
ethanol series (50%, 60%, 70%, 80%, 90%, and 100% ethanol, followed by two 100% ethanol
steps). The material was embedded in Spurr’s low-viscosity resin (Spurr 1969). Sections were
obtained with a RMC MT-XL ultramicrotome (Boeckeler Instruments Inc., Tucson, AZ, USA)
and post-stained with 3% (w/v) aqueous uranyl acetate followed by lead citrate. The stained
sections were viewed with a JEOL-1010 electron microscope (JEOL Ltd., Tokyo, Japan).
For scanning electron microscopy (SEM), a 20-ml aliquot of a dense culture of O. marina
(or G. cf. guttula) was fixed with osmic acid (final concentration = 2%, w/v) in seawater for 1.5 h
and then the fixed cells were collected on a PC membrane filter (pore size = 5 µm) without
additional pressure. The fixed cells were rinsed 3 times with distilled water to remove the salt,
dehydrated through an ethanol series, and finally dried using critical point dryer (BAL-TEC,
CPD 300, Balzers, Liechtenstein, Germany). The dried filters were mounted on a stub and coated
with gold-palladium. Cells were viewed with a JSM-840A SEM (JEOL Ltd., Tokyo, Japan) and
photographed using a digital camera connected to a computer.
Feeding behaviors. Expt 2 was designed to investigate the feeding behaviors of HTDs on
bacteria. To detect the feeding process, we used living bacteria and/or bacteria-sized fluorescent
beads (0.5 μm in diameter). The feeding behaviors of O. marina, G. cf. guttula, and Gyrodinium
9
sp. were observed under light and epifluorescence microscopes with a high resolution video
system. However, the feeding behaviors of P. piscicida and P. bipes were too difficult to observe
because the feeding frequencies were low and the predators were small in size, but swam fast. To
remove aggregated beads, beads were sonicated and then filtered through a 3-μm pore sized filter,
as described above.
A dense culture of O. marina (or G. cf. guttula, Gyrodinium sp.) and living bacteria (ca. 3 x
109 cells ml-1) was added to one well of a 6-well plate chamber containing 3 ml of freshly filtered
seawater (giving a mixture of the HTD and living bacteria). Within 1 min, a 0.1 ml aliquot was
transferred onto a microscope slide and then a cover-glass was placed. We monitored the
behavior of > 100 unfed O. marina cells (or Gyrodinium spp.) with respect to the living bacteria
using a differential interference contrast (DIC) optics under an inverted epifluorescence
microscope at a magnification of 400--1,000x. The feeding process of O. marina cells (or
Gyrodinium spp.) was recorded using a video analyzing system (Sony DXC-C33, Sony Co.,
Tokyo, Japan) and also taken using a digital camera (Zeiss AxioCam MRc5, Carl Zeiss Ltd.,
GÖttingen, Germany).
By single frame playback analysis (30 frames per sec), the number of total contacts
between a predator cell and prey cells until the first ingestion of a bacterium occured (TC), the
number of contacted but non-ingested bacteria until the first ingestion of a contacted bacterium
occurred (NIB), the number of ingested bacteria (IB), the ratio of ingested bacteria to total
contact (RIB; IB/TC), elapsed time for the first bacterium to be ingested (TFBI), and handling
time (HT, elapsed time for a contacted bacterium to be ingested) were determined by monitoring
10
the behavior of 15 O. marina cells (or 15 Gyrodinium cf. guttula cells) in the presence of living
bacterial cells until it successfully ingested a prey cell or until 3 min had elapsed. Also, the
elapsed time for the second prey cells to be ingested by the same predator cell after the ingestion
of the first bacterium cell occurred was measured (n = 5 for O. marina or G. cf. guttula). For O.
marina, the number of total contacts between a predator cell and prey cells is the number of prey
cells physically coming into contact with the cingular depression of the predator (Fig. 5--7),
while for G. cf. guttula, it is the number of prey cells physically coming into contact with the
sulcus of the predator. Also, the particle speed in feeding currents flowing from near the center of
the predator’s body to the cingular depression of O. marina or to the lower part of the sulcus of G.
cf. guttula was measured.
The ingestion of bacteria-sized fluorescent beads (0.5 μm in diameter) by HTDs was also
observed. A dense culture of O. marina (or G. cf. guttula, Gyrodinium sp.) and bacteria-sized
fluorescent beads (100 μl) were added to a 42-ml polycarbonate (PC) experimental bottle (giving
a mixture of the HTD and beads). The bottle was filled to capacity with freshly filtered seawater,
capped, and then well mixed. After 10 min incubation, a 10-μl aliquot was removed from the
bottle and transferred onto a microscope slide with a cavity in the center (24 mm x 60 mm;
Superior Marienfeld, Lauda-Königshofen, Germany). We monitored the behavior of > 50 unfed
O. marina cells (or Gyrodinium spp.) with respect to the beads, using a differential interference
contrast (DIC) optics under an inverted epifluorescence microscope at a magnification of 400--
1,000x. A series of pictures showing the feeding process of O. marina cells (or Gyrodinium spp.)
was recorded using the video analyzing system and also taken using a digital camera.
11
Ingestion rates and grazing coefficients of HTDs on the natural populations of marine
bacteria in Masan Bay. Expt 3 was designed to measure the ingestion rates and grazing
coefficients of the dominant HTDs and co-occurring protists on natural populations of marine
bacteria (Table 2). Water samples were taken from the surface at one station on April 8, 2006 and
June 11, 2007 and at 2 stations on June 13, 2007 in Masan Bay, Korea by using water samplers
during red tides dominated by the euglenophyte Eutreptiella sp. (April 8, 2006) and the
raphidophyte Heterosigma akashiwo (June 11 and 13, 2007). Water temperatures and salinities in
the surface waters were measured using a YSI 30 (YSI, LA, USA), and the pH and dissolved
oxygen (DO) were measured using pH-11 (Schott Handy – Lab, Mainz, Germany) and Oxi 197i
(WTW, Weilheim, Germany), respectively. The water samples were transported into the
laboratory at our marine station within 10-30 min.
In order to determine the bacterial abundances, aliquots of the water samples were poured
into 100-ml polyethylene bottles and preserved with glutaraldehyde (final conc. = 1% v/v). Three
to twelve 1-ml fixed aliquots were stained with DAPI (final conc. = 1 µM) and then filtered onto
0.2 µm-pore-sized polycarbonate (PC) black membrane filters. The bacteria were enumerated
under an epifluorescence microscope with UV light excitation (Porter and Feig 1980).
Additionally, three 1 to 5-ml fixed aliquots were stained with DAPI and then filtered onto 0.2
µm-pore-sized PC black membrane filters. HNFs were also enumerated under an epifluorescence
microscope with UV light excitation. The HNFs could be distinguished from the PNFs
(phototrophic nanoflagellates) that exhibit orange-colored autofluorescence with blue light
excitation. Aliquots of the water samples for counting heterotrophic dinoflagellates, red-tide
12
algae, and ciliates were poured into 500-ml polyethylene bottles and preserved with acidic
Lugol’s solution. After thorough mixing, > 1000 algal cells in eight 1-ml Sedgwick-Rafter
counting chambers (SRCs) and > 150 ciliate cells in 8--12 SRCs were counted under an inverted
microscope with standard transmitted illumination.
The samples for the feeding experiments were screened gently through a 100-µm Nitex
mesh and placed in 270-ml PC bottles. Three to four days prior to these experiments, bacterial
cells collected from the same site were fluorescently labeled using the method of Sherr, Sherr,
and Fallon (1987). These experiments revealed that the fluorescently labeled bacteria (FLB) were
mostly seen as rods (cylinders) and were rarely spherical. We measured the longest (length) and
the shortest axes (width) of ca. 30--40 FLB cells for each field experiment under an
epifluorescence microscope, as in Lee and Fuhrman (1987), and then calculated the volume
according to the following equation: volume = [π (3L – W)/3 x (W/2)2] for a rod (cylinder) and
4/3 x [π R3] for a sphere, where L = length, W = width, and R = radius as in Lee (1993). The size
of the fluorescent beads (0.47 µm, size data supplied by the manufacturer, Polyscience, Inc.,
Warrington, PA, USA) was also measured to calibrate our results. The ranges of the length and
width of FLB used in this field study were 0.44--2.00 µm and 0.44--1.04 µm, respectively. The
mean (+ standard error, n) volumes of FLB were 0.35 µm3 (+ 0.04, 39) and 0.48 µm3 (+ 0.05, 28),
respectively for the field experiments at Masan Bay on April 8, 2006 and on June 11 and 13,
2007 (Table 2). FLBs were added to triplicate bottles. One control bottle (without FLB) was also
set up for each experiment. The bottles were placed on shelves and incubated at a temperature
equivalent to that of the water temperature at the sampling site under continuous illumination of
13
30 µE m-2 s-1 of cool white fluorescent light.
After 1, 5, 10, 20, and 30 min periods of incubation, 10-ml aliquots were removed from
each bottle, transferred into 20-ml vials, and then fixed with borate buffered formalin [final conc.
= 3% (v/v)]. The fixed samples were stained using DAPI (final conc. = 1 µM) and then filtered
onto 0.2 µm-pore-sized PC black membrane filters. Green inclusions (FLB) inside the
protoplasm of ca. 30 cells each of the HTDs and ciliates on the PC black membrane filters were
enumerated under an epifluorescence microscope with blue light excitation. The ingestion rate
(cells predator-1
h-1
) was then calculated by linear regression of the number of FLB per predator
cell as a function of incubation time, after Sherr, Sherr, and Fallon (1987). For comparison, the
ingestion rates of co-occurring red-tide algae, HNFs, and ciliates on bacteria were also measured.
We estimated the grazing coefficients attributable to HTDs (or red-tide algae, HNFs, and
ciliates) on bacteria by combining the data on abundances of the predators and bacteria with
ingestion rates of the predators on bacteria measured in Masan Bay as in Seong et al. (2006). The
grazing coefficients (g, d-1) were calculated as follows:
g = CR x GC x 24 (1)
where CR (ml predator-1h-1) is the clearance rate of a predator on bacteria at a given prey
concentration and GC is the predator concentration (cells ml-1). CRs were calculated as follows:
CR = IR/PC (2)
where IR (cells predator-1h-1) is the ingestion rate of the predator on bacteria and PC (cells
ml-1) is the prey concentration.
Prey concentration effect. Expt 4 was designed to investigate the ingestion rates of O.
14
marina, G. cf. guttula, and P. piscicida on heterotrophic bacteria as a function of prey
concentration. In addition, Expt 5 was designed to compare the ingestion rate of P. bipes on
bacteria at a single prey concentration where the ingestion rates of O. marina, G. cf. guttula, and
P. piscicida on bacteria were saturated in Expt 4. We did not measure the ingestion rates of P.
bipes on bacteria as a function of prey concentration because its ingestion rates were too low to
detect at lower and medium bacterial concentrations in preliminary tests. To remove any
aggregated FLB, FLB were sonicated and then filtered through a 3-μm pore sized filter, as
described above.
One or two days prior to this experiment, the bacterial cells, which originated from a
clonal culture of the target HTD, had been fluorescently labeled as described above. The
volumes of FLB and the ratios of the actual initial abundances of FLB to the abundances of non-
FLB bacteria are shown in Table 4. These experiments revealed that the FLB were mostly seen
as rods (cylinders) and were rarely spherical. We measured the volumes of FLB as described
above.
A dense culture of a target HTD species growing on algal prey and then starved for 1-2
days (until ingested prey cells were undetected inside the protoplasm of the predator cells) was
transferred into a 1-L PC bottle. Three 1-ml aliquots from the bottle were counted using a
compound microscope to determine the cell concentrations of the HTD, as described above. The
mean actual initial predator and prey concentrations are also shown in Table 4. Triplicate 80-ml
PC experimental bottles (containing mixtures of predator and prey) and triplicate predator
control bottles (containing predator only) were also established. All the bottles were then filled to
15
capacity with freshly filtered seawater, capped, placed on a shelf and incubated at 20o C under
continuous illumination of 30 µE m-2 s-1 of cool white fluorescent light. After 1, 5, 10, 20, and 30
min incubation periods, 6-ml aliquots were removed from each bottle, transferred into 20-ml
vials, and then fixed with borate buffered formalin [final conc. = 3% (v/v)]. The fixed samples
were stained using DAPI (final conc. = 1 µM) and then filtered onto 3 µm-pore-sized PC white
membrane filters. The FLB inside a HTD cell were enumerated under an epifluorescence
microscope with blue light excitation. The ingestion rates of the HTD on bacteria were
determined, as described above. Additionally, at the beginning of the experiment, a 1-ml fixed
aliquot was stained with DAPI and then filtered onto 0.2 µm-pore-sized PC black membrane
filters. Bacteria (both FLB and non-FLB) outside the HTD cells were also enumerated under an
epifluorescence microscope with UV light excitation for non-FLB, and blue light excitation for
FLB. After subsampling, the bottles were capped, placed on a shelf, and incubated again, as
described above.
Each value of the ingestion rate (cells predator-1
h-1
) was then calculated, as described
above.
All ingestion rate data were fitted to a Michaelis-Menten equation:
Imax
(x)
KIR
+ (x)IR = (3)
where Imax = the maximum ingestion rate (cells predator-1h-1); x = prey concentration (cells ml-1),
and KIR = the prey concentration sustaining ½ Imax. The prey concentration is the sum of living
16
bacteria and FLBs.
The volume of the HTDs just before Expt 4--5 were conducted were calculated by
geometric formula. The shapes of G. cf. guttula, Gyrodinium sp., and O. marina were estimated
as two cones joined at the cell equator (= maximum width of the cell). Cell volumes of these
preserved heterotrophic dinoflagellates were calculated according to the equation: volume = 1/3
x [π (cell width/2)2] x [cell length]. The shape of P. piscicida was estimated as an oval. Cell
volume of preserved P. piscicida was calculated according to the following equation: volume =
4/3 π [(cell length + cell width)/4]3. The shape of P. bipes was estimated to be two small cones
(two cones connected to each other in the bottom half of the cell: W shape) connected to a large
cone (top half of the cell: Λ shape) at the cell equator (= maximum width of the cell). Cell
volume of the preserved P. bipes was calculated according to the equation: volume = 1/3 x [π
(cell width/2)2] x [cell length/2] + 2 x {1/3 x [π (cell width/4)2] x [cell length/2]}.
Growth of HTDs fed on bacteria alone. Expt 6 was designed to investigate whether
bacteria alone support positive growth of HTDs. We measured the growth rate of O. marina on
living bacteria as a function of the prey concentration because in our preliminary test bacteria
supported the positive growth of O. marina, while they did not support that of G. cf. guttula, and
P. piscicida.
Fourteen days before these experiments were conducted, a dense culture of O. marina
growing on algal prey, was transferred into a 1-L PC bottle containing filtered autoclaved
seawater. The culture of the predator was screened gently through a 10-µm Nitex mesh and the
retained O. marina cells were quickly placed in a new 1-L PC bottle, containing filtered
17
autoclaved seawater, several times. This was done to minimize the abundance of bacteria in the
culture of the predator and also minimize possible residual growth resulting from the ingestion of
prey during batch culture. The bottles were filled to capacity with filtered seawater and placed on
rotating wheels to incubate, as above, except that illumination was provided on a 14 h:10 h light-
dark cycle.
For each HTD species, initial concentrations of the target HTD and bacteria were
established using an autopipette to deliver predetermined volumes of known cell concentrations
to the bottles. Triplicate 42-ml PC experimental bottles (mixtures of predator and prey) and
triplicate control bottles (prey only) were set up at each predator-prey combination. All the
bottles were then filled to capacity with autoclaved seawater, filtered by a 0.2-µm CP filter
(Chisso filter Co. LTD., Tokyo, Japan), and capped. To determine the actual initial and final
predator densities (cells ml-1), a 6-ml aliquot at the beginning of the experiment and a 10-ml
aliquot at the end of the experiment (at 24 h) were removed from each bottle, fixed with 5% (v/v)
Lugol’s solution, and examined with a compound microscope to determine predator abundance,
by enumerating cells in three 1-ml SRCs. The bacteria were enumerated, as described above. The
actual mean initial O. marina and prey concentrations were 770--1,230 cells ml-1
and 3.8 x 104--
2.4 x 107 cells ml
-1, respectively.
The specific growth rate µ (d-1
) of O. marina was calculated as:
GR = Ln (S1/S0) (4)
where S0 and S1 = the concentration of O. marina at 0 d and 1 d, respectively.
Data for O. marina growth rates were fitted to a Michaelis-Menten equation:
18
μmax(x - x')
KGR + (x - x')
μ = (5)
where μmax = the maximum growth rate (d-1), x = prey concentration (cells ml-1), x' = threshold
prey concentration (the prey concentration where μ = 0), and KGR = the prey concentration
sustaining ½ μmax. The data were iteratively fitted into the model using DeltaGraph® (SPSS Inc.,
Chicago, IL, USA).
RESULTS
Feeding occurence. All HTD species tested in the present study (Oxyrrhis marina,
Gyrodinium cf. guttula, Gyrodinium sp., Pfiesteria piscicida, and Protoperidinium bipes) were
able to feed on marine heterotrophic bacteria (Fig. 8--20, 23--38). Under both TEM and an
epifluorescence microscope, a single bacterium (alive before being ingested), a single bacteria-
sized bead (0.5 μm in diameter), and/or a single FLB were observed in food vacuoles inside the
protoplasm of these 4 HTDs (Fig. 8, 9, 16--18, 23, 31, 32, 37). This provides evidence that HTDs
are able to feed on a single bacterium. Also, various numbers of bacteria and beads, ranging from
one to hundreds, were observed in food vacuoles inside the protoplasm of the HTDs (Fig. 8--20,
23--38).
After 10-min incubation, only a few single bacteria or beads were observed inside the
protoplasms of O. marina cells (Fig. 8--11). After more time had passed, it was observed that
inside the protoplasm of an O. marina cell, several single bacteria or beads merged to form a
19
small package and small packages also merged to form a larger package (Fig. 12--15). After 1-h
incubation, several packages containing tens and hundreds of bacteria were observed inside the
protoplasm of most O. marina cells (Fig. 19). The size of the packages differed depending on the
number of ingested bacteria or beads inside the packages.
Like O. marina, Gyrodinium spp. had food vacuoles or packages containing various
numbers of bacteria and beads in food vacuoles inside their protoplasm (Fig. 23--30). Also,
various numbers of bacteria and beads, from one to tens, were observed in food vacuoles inside
the protoplasm of P. piscicida (Fig. 31--36), while a single or only a few bacteria and/or beads
were observed inside the protoplasm of P. bipes (Fig. 37, 38).
Feeding behaviors. Using the flagella (mainly transverse flagellum, but rarely longitudinal
flagellum), O. marina generated feeding currents. Bacterial cells from a distance 1--2 times the
body length of the predator above the predator were carried with the feeding currents toward the
cingular depression of the predator along the flow lines. Oxyrrhis marina intercepted and then
ingested a single bacterial cell in the feeding current (Fig. 39--52). Within the cingular
depression of the predator, the intercepted bacterium was carried in a spiral path (Fig. 39--48) or
linear curve (Fig. 39, 49--52). However, the bacterium inside the cingular depression was quickly
engulfed by the predator within one second (see handling time later for details). While the detail
movement of the longitudinal flagellum was easily observed (sometimes not having moved
much), that of the transverse flagellum was not observed because large parts of the flagellum
usually hid inside the cingular depression and also the transverse flagellum moved very quickly.
The transverse flagellum sometimes touched the bacterium cell, but it was unlikely to grasp the
20
prey cell. The range (mean + SE, n=15) of the number of total contacts between an O. marina
cell and prey cells until the first ingestion of a bacterium occurred (TC), the number of contacted
but non-ingested bacteria until the first ingestion of a contacted bacterium occurs (NIB), and the
ratio of ingested bacteria to total contact (RIB; IB/TC) were 4--87 (36+6), 3--86 (35+6), and
0.01--0.25 (0.05+0.02), respectively (Table 6). The range (mean+SE, n) of the elapsed time for
the first bacterium to be ingested (TFBI) was 5.9--64.0 s (32.6+4.2), while that of the handling
time (HT, time for a contacted bacterium to be ingested) was 0.2--1.0 s (0.6+0.1). The range
(mean + SE, n=5) of the elapsed time for the second prey cell to be ingested by O. marina after
the ingestion of the first bacterium cell occurred (n=5) was 7.6--26.0 s (17.1+3.0) and RIB for
the second prey cell was 0.05--0.17 (0.10+0.02) (i.e. Fig. 49--52). The range (mean + SE, n=20)
of the particle speed in feeding currents flowing from near the center of the predator’s body to
the cingular depression was 44--176 μm s-1 (100+8).
Gyrodinium cf. guttula and Gyrodinium sp. generated feeding currents by undulating the
longitudinal flagellum. Bacterial cells within a distance 1--1.5 x the body length of the predator
were carried with the feeding currents, flowing from above the epicone of the predator, via the
long and narrow sulcus, to below the hypocone of the predator or flowing along the sides of the
body of the predator, via the cingulum and the sulcus, to below the hypocone (Fig. 53).
Gyrodinium spp. ingested a single living bacterium or a single bacteria-sized bead in the feeding
currents when the prey arrived at the lower part of the sulcus, by interception (Fig. 54--64). The
range (mean + SE, n=15) of the number of total contacts between a G. cf. guttula cell and prey
cells until the first ingestion of a bacterium occurred (TC), NIB, and RIB were 29--151 (82+10),
21
28--150 (81+10), and 0.01--0.03 (0.01+0.00), respectively (Table 6). The range (mean + SE, n)
of TFBI was 24.8--134.0 s (73.3+6.9), while that of HT was 0.2--1.1 s (0.6+0.1). The elapsed
time for the second prey cell to be ingested by G. cf. guttula after the ingestion of the first
bacterium cell occurred was 43.0--104.0 s (75.2+12.3) and RIB for the second prey cell was
0.01--0.04 (0.02+0.01). The range of the particle speed in feeding currents flowing from near the
center of the predator’s body to the lower part of the sulcus was 57--270 μm s-1 (155+16).
Ingestion rates and grazing impact of protists on marine bacteria in Masan Bay. We
measured the ingestion rates of the natural populations of co-occurring HTDs, red-tide algae,
HNFs, and ciliates (< 30 μm in cell length) on natural populations of marine bacteria during red
tides dominated by the euglenophyte Eutreptiella sp. (April 8, 2006) and the raphidophyte
Heterosigma akashiwo (June 11 and 13, 2007) in Masan Bay (Table 2, 3). During all field
experiments, the water temperature ranged from 13.6--22.7 oC, while the salinity ranged from
29.2--29.5 psu (Table 2).
During the field experiments, the dominant HTDs were Gyrodinium spp. (> 95% of total
HTDs), which were observed to ingest FLB. No green inclusions were observed inside the
protoplasm of the predators in the control bottles (without FLB). During all field experiments,
the ranges of the mean abundances of Gyrodinium spp. and bacteria were 43--4,240 cells ml-1
and 4.1--7.5 x 106 cells ml-1, respectively (Table 3). The mean (+ SE, n=12) of all the ingestion
rates of Gyrodinium spp. on natural populations of bacteria was 44.9 bacteria HTD-1h-1 (4.3;
range = 14.2--60.9 bacteria HTD-1h-1; Table 3), while their mean clearance rate was 8.3 nl HTD-
1h-1 (1.3; range = 2.0--16.3 nl HTD-1h-1). The mean (+ SE, n=12) grazing coefficient of the
22
natural population of bacteria attributable to Gyrodinium spp. was 0.404 d-1 (0.113; range =
0.003--0.972 d-1; Table 3).
During all field experiments, the ranges of the mean abundances of co-occurring HNFs
and ciliates were 1,340--6,110 cells ml-1 and 22--55 cells ml-1, respectively (Table 3). The
ingestion rates of HNFs (3.7--15.6 bacteria HNF-1h-1) or mixotrophic algae on bacteria (1.3--10.6
bacteria alga-1h-1) were lower than those of Gyrodinium spp., while those of ciliates (66--638
bacteria ciliate-1h-1) were higher than those of Gyrodinium spp. (Table 3). Also, the clearance
rates of HNFs (0.4--2.8 nl HNF-1h-1) or algae on bacteria (0.3--1.6 nl alga-1h-1) were lower than
those of Gyrodinium spp., while those of ciliates (7.3--129.2 nl ciliate-1h-1) were higher than
those of Gyrodinium spp. (Table 3). The grazing coefficients of the natural population of bacteria
attributable to HNFs (0.020--0.396 d-1), ciliates (0.003--0.171 d-1), or algae on bacteria (0.001--
1.128 d-1) were lower or comparable to those of Gyrodinium spp. (Table 3).
Ingestion rates of HTDs on bacteria as a function of prey concentration. With
increasing initial prey concentration, the ingestion rates of Oxyrrhis marina, Gyrodinium cf.
guttula, and Pfiesteria piscicida on bacteria increased rapidly at prey concentrations of ca. 0.7--
2.2 x 106 cells ml
-1, and then increased only slowly or reached saturation at higher prey
concentrations (Fig. 65--67). When the data were fitted to Eq. (3), the maximum ingestion rates
of HTDs on bacteria were 71.3 cells dinoflagellate-1
h-1
for O. marina, 23.2 cells dinoflagellate-1
h-
1 for G. cf. guttula, and 13.7
cells dinoflagellate
-1h
-1 for P. piscicida (Table 5). The maximum
specific ingestion rates for O. marina, G. cf. guttula, and P. piscicida on bacteria were 2.9 x 10-2
23
h-1, 0.7 x 10-2 h-1, and 1.0 x 10-2 h-1, respectively (Table 5).
The maximum clearance rates of HTDs on bacteria were 31.3 nl dinoflagellate -1
h-1
for O.
marina, 16.1 nl dinoflagellate -1
h-1
for G. cf. guttula, and 11.4 nl dinoflagellate -1
h-1
for P.
piscicida (Table 5). The maximum volume-specific clearance rates for O. marina, G. cf. guttula,
and P. piscicida on bacteria were 6.7 x 104 h-1, 2.4 x 104 h-1, and 4.7 x 104 h-1, respectively
(Table 5).
When prey concentrations were 7.1 x 106 cells ml
-1, the ingestion and clearance rates of
Protoperidinium bipes on bacteria were 3.5 cells dinoflagellate-1
h-1
and 0.5 nl dinoflagellate -1
h-1
,
respectively (Table 5). The maximum specific ingestion rate and the maximum specific clearance
rates for P. bipes on bacteria were 0.1 x 10-2 h-1 and 5.2 x 102 h-1, respectively (Table 5).
Growth of Oxyrrhis marina fed on bacteria alone. Bacteria alone supported the growth
of Oxyrrhis marina, while they did not support that of Gyrodinium cf. guttula or Pfiesteria
piscicida. The specific growth rates of O. marina on bacteria increased with increasing initial
prey concentration up to 2.1 x 106 cells ml
-1, but were saturated at higher prey concentrations
(Fig. 68). When the data were fitted to Eq. (5), the maximum specific growth rates (μmax) and
KGR (prey concentration sustaining 0.5 µmax) were 0.592 d-1 and 2.9 x 105 cells ml
-1, respectively.
The threshold prey concentration (where net growth = 0) was 4.1 x 104 cells ml
-1.
DISCUSSION
HTD predators on heterotrophic bacteria and their feeding behaviors. All HTDs
24
tested in the present study were able to ingest marine heterotrophic bacteria. This study is the
first to provide photos of the inclusion of added heterotrophic bacteria (both non-labeled bacteria
and FLB) and bacteria-sized beads inside the protoplasm of HTDs using diverse observational
methods (light microscopy, epifluorescence microscopy, TEM) and also to report on the feeding
behaviors of HTDs on bacteria using high resolution video systems. The evidence of the present
study may help to dispel the suspicion that isotope labeled bacteria attached to the body of HTDs
might give rise to the results of Lessard and Swift (1985). Now, the species belonging to the
genera Oxyrrhis, Gyrodinium, Pfiesteria, Protoperidinium, Podolampas, and Diplopsalis have
been revealed to feed on heterotrophic bacteria. By extension, other HTD species are likely to
feed on heterotrophic bacteria. Feeding by diverse HTDs on heterotrophic bacteria may influence
our conventional view of energy flow and carbon cycling in the marine planktonic community
(see last subsection).
There has been a big debate on whether HTDs are able to feed on a single bacterium
because some have thought that HTDs might only be able to ingest bacteria attached to detritus.
Most studies on the feeding of HTDs have focused on algal prey. In these studies, HTDs have
been known to feed on algal prey by direct engulfment, pallium feeding, or by feeding tube
(Hansen and Calado 1999). To use one of these feeding behaviors, HTDs should first be
detecting, then capturing, handling, and engulfing prey cells. Therefore, some thought that
because a single bacterium is too small for HTDs to detect and capture using one of these
feeding behaviors; the lower prey size limit for prey capture of the HTDs Gyrodinium spirale
and Gyrodinium sp. has been suggested to be 3--4 μm (Hansen 1992; Jakobsen and Hansen
25
1997). Also, the lower prey size limits for pallium feeders and peduncle feeders have been
declared to be 4--8 μm and 2--4 μm, respectively (summarized by Hansen and Calado 1999).
However, the present study clearly shows, through using an epifluorescence microscope, TEM,
and video microscopy, that small HTDs are able to ingest a single living bacterium, a single
bacterium-sized bead, and a single FLB; by using feeding behaviors other than direct engulfment,
pallium feeding, or feeding tube, O. marina and Gyrodinum spp. gathered and ingested bacterial
cells (i.e. generate feeding currents carrying bacteria cells, intercepted, and then ingested the
prey cells).
Based on the classifications of the feeding behavior of the heterotrophic protists (Boenigk
and Arndt 2000; Fenchel 1987; Sleigh 1989), when bacteria are prey, O. marina and Gyrodinum
spp. are close to interception feeders (i.e. producing a feeding current and directly intercepting
food particles) because they generate a strong water current using the flagella and intercept a
single bacterium inside the feeding current. However, when algae are the prey, both O. marina
and Gyrodinum spp. are close to raptorial feeders (i.e. mobile predators that actively search for
food particles) because they capture a prey cell using a trichocyst or open the sulcus and then
engulf the prey cell (Kim and Jeong 2004); we found that O. marina captured an Isochrysis
galbana cell (ca. 4 μm in Equivalent Spherical Diameter) and larger algal prey species using
trichocysts. Therefore, O. marina and Gyrodinum spp. exhibit 2 different feeding behaviors when
feeding on pico-sized prey and when feeding on nano- or micro-sized prey. When the size ratio
of prey to predator is large, raptorial feeding is known to be effective, while when the ratio is
small, filter feeding is effective (Fenchel 1987). To increase the efficiency of gathering and
26
ingesting small bacterial cells, O. marina and Gyrodinum spp. may use a combination of filter
feeding and interception feeding: gathering prey cells in a flow generated by flagella, then
intercepting and ingesting the prey cells in feeding currents flowing in the cingular depression or
long and narrow sulcus. Therefore, O. marina and Gyrodinum spp. may have evolved to feed on
more diverse prey items by utilizing different feeding behaviors depending on the type and/or
size of prey. These HTDs may have a huge advantage in gaining energy, compared to predators
that have only one feeding behavior which limits them to being able to feed on only a certain
sized prey. Before the present study, a HTD having 2 different feeding behaviors has not been
reported (Hansen and Calado 1999), while several HNFs have been reported to exhibit different
feeding behaviors (Boenigk and Arndt 2002). Strom (1991) reported that a small Gymnodinium
sp. (600-1200 μm3) was able to feed on the cyanobacterium Synechococcus sp. (1.2 x 2.4 μm).
Gymnodinium sp. may have also different feeding behaviors when feeding on Synechococcus sp.
and when feeding on large algal prey.
In natural environments, diverse prey items for HTDs are likely to co-exist. However, in
the present study, we did not test whether HTDs prefer bacteria to larger algal prey in mixtures or
vice versa. The maximum specific ingestion rates of O. marina, G. cf. guttula, and P. piscicida on
bacteria (0.7--2.9 x 10-2 h-1) are much lower than those of O. marina, G. dominans, and P.
piscicida on optimal algal prey (2.2--3.7 x 10-1 h-1; Jeong et al. 2003, 2006; Kim and Jeong
2004). Also, the maximum volume specific clearance rates for O. marina, G. cf. guttula, and P.
piscicida on bacteria (2.4--6.7 x 104 h-1) are much lower than those for O. marina, G. dominans,
and P. piscicida on optimal algal prey (5.0 x 105 h-1 to 4.4 x 106 h-1; Jeong et al. 2003, 2006;
27
Kim and Jeong 2004). Therefore, bacteria may be selected less than the optimal algal prey when
these two prey components are both abundant. In Masan Bay, a highly eutrophicated bay, the
abundance of algal prey was sometimes very high, but sometimes very low (i.e. fluctuated by ca.
1,000-fold), while that of bacteria did not change much, maintaining high bacterial abundance
(ca. 5-fold; e.g. Jeong et al. 2005; our unpublished data). Therefore, small HTDs can obtain the
carbon for growth or maintenance of their population when the abundance of algal prey is low if
that of bacteria is still high.
Ingestion rates and grazing impact of protists on bacteria in Masan Bay. The present
study is the first study comparing the ingestion rates of the natural populations of co-occurring
HTDs, red-tide algae, HNFs, and ciliates on natural populations of marine bacteria. During the
field experiments, the ingestion and clearance rates of Gyrodinium spp. on bacteria in Masan Bay
were markedly higher than those of co-occurring HNFs or red-tide algae, while they were lower
than those of ciliates (< 30 μm). The larger cell volume of Gyrodinium spp. (ca. 950 μm3)
compared to the co-occurring HNFs (ca. 60 μm3) or red-tide algae (Heterosigma akashiwo = ca.
700 μm3; Eutreptiella sp. = ca. 280 μm3) may cause higher ingestion and clearance rates of this
HTD. Also, the heterotrophic activity of Gyrodinium spp. may lead to higher ingestion and
clearance rates compared to the mixotrophic activity of these mixotrophic algae. The smaller
volume of Gyrodinium spp. compared to the co-occurring ciliates (ca. 1,400 μm3) may cause
lower ingestion and clearance rates of this HTD. Also, the feeding behavior of Gyrodinium spp.
may be less effective than that of ciliates. Gyrodinium spp. capture and engulf a single bacterium
in water flow moving through the long sulcus one-by-one (i.e. interception feeders), while
28
ciliates generally capture many bacteria at once by filter feeding.
The grazing coefficient of the natural population of bacteria attributable to Gyrodinium spp.
in Masan Bay was 0.003--0.972 d-1 (i.e. 0.3--62% of co-occurring heterotrophic bacteria
populations were removed by Gyrodinium populations in 1 d). In this calculation, the ranges of
the mean abundances of Gyrodinium spp. and bacteria in Masan Bay were 43--4,240 cells ml-1
and 4.1--7.5 x 106 cells ml-1, respectively. The abundances of Gyrodinium spp. and bacteria in
Masan Bay in 2006--2007 were 25--4,980 cells ml-1 and 3.2--14.7 x 106 cells ml-1, respectively
(our unpublished data). The grazing coefficient is proportional to the abundance of Gyrodinium
spp. and thus, the range of the grazing coefficients calculated in the present study is typical in
Masan Bay. Therefore, the results of the present study suggest that Gyrodinium spp. can
sometimes have a considerable grazing impact on populations of marine heterotrophic bacteria in
Masan Bay.
The grazing coefficients of the natural population of bacteria attributable to Gyrodinium
spp. in Masan Bay were sometimes comparable to or higher than those of red-tide algae, HNFs,
or ciliates. For a long time, it has been thought that natural marine bacteria are removed by HNFs
and/or ciliates (Ichinotsuka, Ueno, and Nakano 2006; Sherr, Sherr, and Verity 2002; Vaque et al.
2002) and recently, red-tide algae have been added as important grazers on natural marine
bacteria (Seong et al. 2006). The present study shows that HTDs are sometimes one of the major
grazers on natural marine bacteria and thus they may compete with co-occurring red-tide algae,
HNFs, and ciliates for bacterial prey. Therefore, to investigate the total grazing impact by protists
on marine bacteria, we should measure the ingestion rates of HTDs and red-tide algae in addition
29
to HNFs and ciliates.
Prey concentration effects on ingestion rates. Before the present study, there had been
no study on the functional responses by HTDs to marine heterotrophic bacteria. The ingestion
rates of Gyrodinium cf. guttula on bacteria increased rapidly at prey concentrations of ca. 2.2 x
106 cells ml
-1, and then increased slowly at higher prey concentrations. The mean bacterial
concentrations in Masan Bay in the present study ranged from 4.1 x 106 cells ml-1 to 7.5 x 106
cells ml-1. There is a possibility that the addition of FLB into natural populations of marine
bacteria has resulted in overestimation of the ingestion rates of protists on marine bacteria. When
the ingestion rates of Gyrodinium spp. on the natural populations of marine bacteria are
calculated, using the equations in Fig. 66 and the abundances of bacteria and added FLB in Table
2, the rates obtained without FLB were theoretically lower by 1.4--11.2 % than those with FLB.
Therefore, the addition of FLBs into natural populations of marine bacteria was not likely to
result in a large overestimation of the ingestion rates of HTDs on marine bacteria.
The maximum mean ingestion rates of Gyrodinium spp. on natural populations of bacteria
(57.4 bacteria HTD-1h-1) were considerably higher than those of Gyrodinium cf. guttula on
bacteria measured in the laboratory (23.2 bacteria HTD-1h-1) (Table 3, 5). The volume of FLBs
used for the field experiments in Masan Bay (0.48 μm3) was very similar to that used for the
laboratory experiments (0.44 μm3). However, the cell volume of Gyrodinium spp. (mean +
standard error = 950 + 80 μm3, n=30) was greater than that of G. cf. guttula (660 + 80 μm3,
n=30). Therefore, the greater volume of Gyrodinium spp. may cause their higher maximum
ingestion rate compared to G. cf. guttula.
30
The Imax of O. marina on marine heterotrophic bacteria was higher than that of G. cf.
guttula, even though the cell volume of the former HTD was smaller than that of the latter one.
The ratio of ingested bacteria to total contact (RIB) for O. marina was higher than that of G. cf.
guttula, while the elapsed time for the first bacterium to be ingested (TFBI) by the former HTD
was shorter than that of the latter one. However, the particle speed in feeding currents generated
by O. marina was lower than that generated by G. cf. guttula and the handling time by the former
HTD was similar to that by the latter one. Therefore, higher RIB and shorter TFBI of O. marina
may be mainly responsible for its higher Imax on bacteria than G. cf. guttula. When feeding on
bacteria, O. marina has a more efficient feeding mechanism than G. cf. guttula.
The Imax of O. marina on marine heterotrophic bacteria, which had the highest maximum
ingestion rate among the HTD predators tested in the present study, was much higher than that of
the mixotrophic raphidophyte Chattonella ovata (25 cells alga-1
h-1
) which had the highest Imax
among the red-tide algal predators tested by Seong et al. (2006), even though the size of O.
marina (ESD = ca. 11 μm) was much smaller than that of C. ovata (40 μm). This HTD may
produce more digestive enzyme and have higher ingestion rates than the mixotrophic alga. It is
therefore worthwhile exploring the possible difference in digestive enzyme activity between
heterotrophic and mixotrophic protists. Also, when fed on bacteria, O. marina may have a more
efficient feeding behavior than C. ovata. However, the feeding behavior of C. ovata has not been
reported yet. The Imax of HTDs on marine bacteria were comparable to those of HNFs; the Imax of
O. marina on bacteria is comparable to that of the HNF Pseudobono sp. (88 cells HNF-1
h-1
;
31
Alonso et al. 2000), while Gyrodinium cf. guttula and Pfiesteria piscicida (14--23 cells HTD-1
h-
1) are comparable to those of the HNFs Bono designis and Rhynchomonas nasuta (16--19 cells
HNF-1
h-1
; Artolozaga et al. 2002). Therefore, some HTD species may sometimes compete with
several HNF species for bacterial prey, if they co-occur. Feeding by HNFs has been reported to
be affected by whether the prey are living or dead, size, their motility, and the hydrophobic and
electrostatic cell surface properties of the bacteria (Jürgens and Demott 1995; Koton-Czarnecka
and Chrost 2003; Monger, Landry, and Brown 1999; Matz and Jürgens 2001). The feeding by
HTDs on diverse bacteria may be also affected by several properties of the bacteria. Thus, it is
worthwhile exploring selective feeding by HTDs on bacteria having different properties.
Growth of Oxyrrhis marina fed on bacteria alone. The present study clearly shows that
bacteria alone supported the growth of O. marina. We maintained O. marina for more than 1
month without added algal prey. In natural environments, O. marina often lives near the bottom
where there is a large amount of detritus, which bacteria can utilize. Thus, O. marina is easily
likely to maintain or increase its own population by feeding on bacteria when bacteria
concentrations are high.
The maximum growth rate of O. marina on bacteria (0.59 d-1) is considerably lower than
that on the optimal algal prey Heterosigma akashiwo (1.43 d-1; Jeong et al. 2003). Several studies
have suggested that the optimal predator : prey size ratio, yielding the best growth would be
around 1:1--2.4:1 (Hansen 1992; Naustvoll 2000a, 2000b). Therefore, the optimal algal prey,
rather than bacteria, is likely to give best growth. However, the maximum growth rate of O.
marina on bacteria is comparable to that on the diatom Dunaliella tertiolecta, the haptophyte
32
Isochrysis galbana, or raphidophyte Fibrocapsa japonica (0.7--0.8 d-1), the suboptimal algal
prey species (Goldman, Dennett, and Gordin 1989; Jeong et al. 2003; Tillmann and Reckermann
2002) and much higher than that on the heterotrophic nanoflagellate Cafeteria sp. (0.19 d-1;
Jeong et al. 2007a). Therefore, bacteria may sometimes be an important prey for the growth of O.
marina. The relative abundance of bacteria and algae in natural environments may affect the
contribution of bacteria to the growth of O. marina. It is worthwhile investigating the selective
feeding by O. marina on bacteria and algae and calculating the relative contribution of bacteria
and algae to its growth. Utilization of both bacteria and algal prey would give a great advantage
to O. marina in competition with protists that are able to feed on only bacteria or algal prey.
Bacteria did not support positive growth in Gyrodinium cf. guttula, or Pfiesteria piscicida.
Relatively low ingestion rates and/or possibly low growth efficiency of these HTDs on bacteria
are likely not to support positive growth. Therefore, bacteria may not make a critical contribution
to the population growth of G. cf. guttula or P. piscicida in natural environments, but be
supplementary prey.
Ecological importance. The results of the present study are ecologically important for
planktonic communities for the following reasons. (1) All HTD species tested in the present
study ingested marine heterotrophic bacteria. HTDs not tested yet are also likely to feed on
heterotrophic bacteria. The pathway of material and energy transfer from heterotrophic bacteria
to HTDs may be important in marine environments, in particular in lagoons, bays, and estuaries
where bacteria and HTD concentrations are high. (2) The grazing coefficients of HTDs on
bacteria are sometimes comparable to or higher than those of co-occurring mixotrophic red-tide
33
algae, HNFs, and/or ciliates. Thus, HTDs may have a grazing impact on the populations of
bacteria comparable to that of mixotrophic red-tide algae, HNFs, or ciliates for bacterial prey.
Sometimes the abundance of HTDs is as high as 20,000 cells ml-1 (e.g. Jeong et al. 2005). Their
relative abundance among these mixotrophic and heterotrophic protists may be important for
their contribution as predators on bacteria. (3) Bacteria may be too small to be ingested by the
filter-feeding copepods, while many HTDs are ingested by the copepods (Jeong et al. 2001,
2007b; Roman, Reaugh, and Zhang 2006). Therefore, HTDs might also be a link between
bacteria and copepods and possibly some other zooplankton that are unable to directly ingest
bacteria. (4) Bacteria alone supported the growth of Oxyrrhis marina. Therefore, bacteria may
sometimes be an important prey source for the growth or maintenance of some HTDs. The
discovery of bacterivory in common HTDs in the present study may help in understanding the
cycling of materials and the energy flow in marine microbial food webs. To understand marine
planktonic food webs better, the possible roles of marine bacteria and HTDs and their
interactions with other planktonic components should be further explored.
ACKNOWLEDGEMENTS
We thank Seong Taek Kim, Jong Hyeok Kim, Soo Kyeom Kim, Jin Ah Ryu for technical
supports. This paper was funded by a grant from Korean Research Foundation (2005-070-
C00143) and a NRL grant from MOST & KOSEF (M1-0302-00-0068).
34
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Table 1. Isolation and maintenance conditions of the experimental organisms. Sampling location and time, water temperature (T, oC),
salinity (S, practical salinity units) for isolation, and prey species and concentrations (cells ml-1) for maintenance.
Organism
Location
Time
T
S
Prey species
Concentration
Reference
Pfiesteria piscicida
Off Incheon
July 2005
24.0 25.4 Amphidinium carterae 20,000 --30,000
Jeong et al. (2006)
Oxyrrhis marina
Keum Estuary
May 2001
16.0 27.7 Amphidinium carterae 8,000 Jeong et al. (2003)
Gyrodinium cf. guttula
Masan Bay
April 2003
18.5 25.0 Prorocentrum minimum 5,000 As in Kim and Jeong (2004)
Gyrodinium sp.
Keum Estuary
Nov. 2005
12.5 24.6 Prorocentrum mimimum 10,000 As in Kim and Jeong (2004)
Protoperidinium bipes
Masan Bay
April 2006
12.2 29.4 Skeletonema costatum 10,000 Jeong et al. (2004b)
32
Table 2. Water temperature (T), salinity (S), volume of fluorescent labeled bacteria (FLB), and bacterial abundance for
incubation of bacteria and Gyrodinium spp. in Masan bay, Korea in 2006--2007. Data are mean ± standard error (SE), except
those in parentheses (n). Natural: natural populations of bacteria; Abundance ratio: actual initial abundances of FLB to the
abundances of natural populations of bacteria.
Station Date
T
S Volume of FLBs Natural
FLB
Abundance ratio
(μm3) (106 cells ml–1)
(106 cells ml–1) (oC) (%) (psu)
13.6 29.5 0.35 + 0.04 (39) St 1 0.72 + 0.03 10+1 (3) April 8, 2006 7.5 + 0.78
22.6 29.3 0.48 + 0.05 (28) 3.0 + 0.46 St 1 June 11, 2007 5.4 + 0.26 57+10 (3)
22.7
0.48 + 0.05 (28) 4.1 + 0.5 3.4 + 0.7 St 1 June 13, 2007 29.2 84 +18 (3)
22.7
29.4
0.48 + 0.05 (28) 7.5 + 1.1 St 2 6.2 + 0.1
87 +12 (3)
33
Table 3. Ingestion of marine bacteria by red-tide algae, heterotrophic nanoflagellates (HNFs), and ciliates (< 30 μm) in Masan Bay, Korea during red tides in 2006-2007. Data are mean ± SE
Location Date Predator Abundance Ingestion rate Clearance rate Grazing coefficient
(ind ml–1) (cells ind–1 h–1 (nl ind–1 h–1) (d–1) )
34
St 1
April 8, 2006
Gyrodinium spp.
43 +7
31.3 + 8.7
4.1 +1.1
0.004 + 0.001
5.7 + 2.5
Eutreptiella sp. 15270+650 0.8 + 0.4 0.301 + 0.156
4170+510
8.0 + 8.7 cryptophytes 1.1 +0.2 0.072 + 0.018
1340+30
HNF 8.3 + 3.3 1.2 +0.5 0.038 + 0.018
Ciliates
22+2
90.8 + 15.6
12.6 +3.2 0.006 + 0.002
St 1
Gyrodinium spp.
7.6 +2.2
0.085 +0.034
430 + 23 June 11, 2007 40.0 + 10.2
Heterosigma akashiwo
12940 + 970 3.0 + 1.0
0.6 +0.2
0.175 +0.070
5.4 + 0.8 HNF
3340 + 550
1.0 +0.1
0.083 +0.023
618.3 + 12.2
116.0 +7.9
0.155 +0.010 Ciliates 55 + 3
Table 3 (continued)
Location Date Predator Abundance Ingestion rate Clearance rate Grazing coefficient
(cells ml–1) (cells ind–1 h–1 (nl ind–1 h–1) (d–1) )
St 1
Gyrodinium spp.
June 13, 2007 2270 + 50 57.4 + 2.4 14.3 + 1.2 0.754 + 0.060
Heterosigma akashiwo 39040 +
1990 3.0 + 1.1 0.8 + 0.3 0.655 + 0.252
HNFs
2040 + 120 5.4 + 0.8 1.4 + 0.3 0.070 + 0.021
0.053 + 0.003 ciliates
27 + 2 331.0 + 27.7 81.5 + 3.6
St 2
June 13, 2007
Gyrodinium spp.
4240 + 120
7.3 + 0.9
52.7 + 2.1 0.775+ 0.120
Heterosigma akashiwo
4.3 + 0.8
0.6 + 0.2
55490 +
3060 0.773+ 0.197
HNFs
6110 + 120
7.7 + 3.9 1.2 + 0.8 0.177 + 0.109
ciliates
31 + 2
358.9 + 59.5
52.1 + 13.7
0.039 + 0.012
35
Table 4. Volume of the predators (μm3), volume of FLB (μm3), abundance ratios (actual initial abundances of FLB to the abundances
of non-FLB bacteria, %), and mean actual initial concentrations of predator and prey in Expt 4 and 5. Initial concentrations of prey are
the sum of living bacteria and added FLB. Data are means ± SE, except those in parentheses (n).
Predator
Volume Predator
Volume FLB
Abundance
Predator
Min.
Concentration
(cells ml-1)
Max.
Prey
Min.
ratios (x105cells ml-1)
Max. Oxyrrhis marina
470 ± 50 (30)
0.43 ± 0.08 (30)
22 ± 2 (18) 734 ± 21 (3)
1270 ± 236 (3)
3.0 ± 0.1 (3)
250 ± 38 (3)
Gyrodinium cf. guttula
660 ± 80 (30)
0.44 ± 0.04 (31)
32 ± 3 (18) 150 ± 15 (3)
250 ± 25 (3)
3.0 ± 0.1 (3)
150 ± 13 (3)
Pfiesteria piscicida
240 ± 30 (30)
0.65 ± 0.11 (30)
27 ± 3 (18) 702 ± 155 (3)
1326 ± 28 (3)
2.9 ± 0.2 (3) 120 ± 7 (3)
Protoperidinium bipes
960 ± 90 (30)
0.56 ± 0.08 (31)
38 ± 6 (3)
216 ± 3 (3)
71 ± 3 (3)
36
Table 5. Ingestion rates of the HTD predators on marine bacteria in Expt 4 & 5. Parameters for Oxyrrhis marina, Gyrodinium cf.
guttula, and Pfiesteria piscicida are for functional response from Eq. (3) as presented in Fig. 65-67. *Imax and *Cmax of
Protoperidinium bipes were obtained at single prey concentrations where the ingestion rates of O. marina, G. cf. guttula, and P.
piscicida became saturated. Imax (maximum ingestion rate, cells HTD-1h-1), MSI (maximum specific ingestion rate, x 10-2 h-1),
Cmax (maximum clearance rate, nl HTD-1h-1), MSC (maximum specific clearance rate, x 104 h-1).
Figures Predator species
Imax
MSI
Cmax
MSC
65
Oxyrrhis marina
71.3
2.9
31.3
6.7
66
Gyrodinium cf. guttula
23.2
0.7
16.1
2.4
67
Pfiesteria piscicida 13.7 1.0 11.4 4.7
37
Protoperidinium bipes
3.5*
0.1
0.5*
0.05
Table 6. Quantitative data (range) on the feeding process of Oxyrrhis marina and Gyrodinium cf. guttula on living bacteria. Particle
speed in feeding currents (PS), the number of total contact between a predator cell and prey cells until the ingestion of a bacterium
occurred (TC), the number of contacted but non-ingested bacteria until the first ingestion of a contacted bacterium occurs (NIB), and
ingested bacteria (IB), the ratio of ingested bacteria to total contact (RIB; IB/TC), elapsed time for the bacterium to be ingested
(TFBI), handling time (HT, elapsed time for a contacted bacterium to be ingested). The 2nd ingestion is consecutive ingestion of
another bacterium by the same predator cell after the ingestion of the first bacterium occurred. Data in parentheses are means ± SE.
38
Predator
n
PS
(μm s-1)
TC
NIB
RIB
TFBI
(s)
HT (s)
Oxyrrhis marina (1st ingestion)
15
44--176
(100 ± 8)
4--87
(36 ± 6)
3--86
(35 ± 6)
0.01--0.25
(0.05 ± 0.02)
5.9--64.0
(32.6 ± 4.2)
0.2--1.0
(0.6 ± 0.1) Oxyrrhis marina (2nd ingestion)
5
6--21
(12 ± 2)
5--20
(11 ± 2)
0.05--0.17
(0.10 ± 0.02)
7.6--26.0 (17.1 ± 3)
0.2--0.7
(0.4 ± 0.1) Gyrodinium cf. guttula (1st ingestion)
15
57--270
(155 ± 16)
29--151
(82 ± 10)
28--150
(81 ± 10)
0.01--0.03
(0.01 ± 0.00)
24.8--134.0(73.3 ± 6.9)
0.2--1.1
(0.6 ± 0.1) Gyrodinium cf. guttula (2nd ingestion)
5
26--97
(70 ± 15)
25--96
(69 ± 15)
0.01--0.04
(0.02 ± 0.01)
43.0--104.0
(75.2 ± 12.3)
0.2--0.4
(0.3 ± 0.1)
Figure Legends
Fig. 1--4. Micrographs of bacteria-sized fluorescent beads (0.5 μm in diameter), fluorescent
labeled bacteria (FLB), and non-labeled bacteria after being sonicated and then filtered
through 3-μm pore sized filter. 1, 2. Bacteria-sized fluorescent beads and FLBs observed
under an epifluorescence microscope. 1. Fluorescent beads. 2. FLBs. 3, 4. Non-
fluorescent beads and non-labeled bacteria observed under transmission electron
microscope (TEM). 3. Non-fluorescent beads. 4. Non-labeled bacteria. Scale bars = 1
µm for Fig. 1 and 2 and 0.2 µm for Fig. 3 and 4.
Fig. 5--7. Morphology of Oxyrrhis marina. 5, 6. Scanning electron micrographs (SEM) of
an O. marina cell. 7. Drawing of an O. marina cell. tf: Transverse flagellum. lf:
Longitudinal flagellum. t: Tentacle. cd: Cingular depression. Scale bars = 2 µm for Fig.
5 and 7 and 1 µm for Fig. 6.
Fig. 8--15. Micrographs of Oxyrrhis marina with ingested fluorescent beads (0.5 μm in
diameter) and/or fluorescent labeled bacteria (FLBs) observed under an epifluorescence
39
microscope. 8. An O. marina cell with a single bead (arrow). 9. An O. marina cell with
a single FLB (inside the dashed circle). 10. An O. marina cell with 3 ingested
fluorescent beads (arrows). 11. An O. marina cell with 3 FLBs (arrows). 12. An O.
marina cell with ca. 10 ingested fluorescent beads (inside the dashed circle). 13. An O.
marina cell with ca. 10 FLBs (inside the dashed circle). 14. An O. marina cell with
several packages containing many aggregated ingested fluorescent beads (inside the
dashed circle). 15. An O. marina cell with a package containing many aggregated
ingested FLBs (inside the dashed circle). All scale bars = 10 µm.
Fig. 16--20. Transmission electron micrographs (TEM) of Oxyrrhis marina with an
ingested non-fluorescent beads (0.5 μm in diameter) and/or non-labeled bacteria. 16--18.
An O. marina cell observed under TEM after 10 min incubation (see text). 16. An O.
marina cell with a single non-fluorescent bead (inside the dashed box) and a single
bacterium (inside the dashed circle) inside food vacuoles. 17. Enlarged from Fig. 16 for
a bead inside a food vacuole. 18. Enlarged from Fig. 16 for a bacterium inside a food
vacuole. 19, 20. An O. marina cell observed under TEM after 1 h incubation (see text).
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19. An O. marina cell with 4 packages containing many aggregated ingested bacteria
(arrows). 20. Enlarged from Fig. 19 for ingested bacteria inside the largest package.
Scale bars = 1 µm for Fig. 16 and 19 and 0.2 µm for Fig. 17, 18, and 20.
Fig. 21--30. Micrographs of Gyrodinium cf. guttula without and with ingested fluorescent
beads (0.5 μm in diameter) and/or bacteria. 21. A G. cf. guttula cell without ingested
bead or bacteria observed under an epifluorescence microscope. 22. A G. cf. guttula cell
without ingested bead or bacteria observed under SEM. 23--26. G. cf. guttula cells
observed under an epifluorescence microscope. 23. A G. cf. guttula cell with an ingested
single bead (arrow). 24. A G. cf. guttula cell with 2 ingested single beads (arrow). 25. A
G. cf. guttula cell with 2 ingested FLBs (arrow). 26. A G. cf. guttula cell with several
ingested beads (arrow). 27. A G. cf. guttula cell with 4 different sized packages
containing many aggregated ingested beads (arrows). 28. A G. cf. guttula cell with ca. 2
package containing many aggregated FLBs (arrows). 29, 30. A G. cf. guttula cell
observed under TEM. 29. A G. cf. guttula cell with ca. 6 packages containing many
aggregated ingested bacteria (arrows). 30. Enlarged from Fig. 29 for ingested bacteria
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inside the largest package. Scale bars = 10 µm for Fig. 21--28, 1 µm for Fig. 29, and 0.2
µm for Fig. 30.
Fig. 31--38. Micrographs of Pfiesteria piscicida and Protoperidinium bipes with ingested
fluorescent beads (0.5 μm in diameter) and/or bacteria, observed under an
epifluorescence microscope. 31--36. P. piscicida. 31. A P. piscicida cell with an ingested
single bead (arrow). 32. A P. piscicida cell with an ingested single FLB (arrow). 33. A P.
piscicida cell with 3 ingested beads (arrow). 34. A P. piscicida cell with 2 ingested FLBs
(arrow). 35. A P. piscicida cell with a package containing many ingested beads (arrow).
36. A P. piscicida cell with a package containing several FLBs (arrow). 37, 38.
Protoperidinium bipes. 37. A P. bipes cell with an ingested single bead (arrow). 38. A P.
bipes cell with 2 ingested beads (arrows). All scale bars = 10 µm.
Fig. 39--52. Feeding processes of Oxyrrhis marina on living heterotrophic bacteria
observed under an epifluorescence microscope, recorded using high-resolution video
microscopy. 39--48. Ingestion of the first bacterium (arrows). 39. Drawing on the path of
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ingested (red and blue color) and non-ingested (black color) bacterial prey cells in
feeding currents generated by an O. marina cell. 40--48. Serial photos showing the
ingestion of the first single bacterium cell (arrow) by the O. marina cell. The bacterium
cell near and inside cingular depression was carried in a path of whirling (red color in
Fig. 39). 49--52. Consecutive ingestion of the second bacterium (arrow) by the same
predator cell after the ingestion of the first bacterium occurred. The bacterium cell near
and inside cingular depression was carried in a path of linear curve (blue color). All O.
marina cells in Fig. 40--52 were the same cell. The numbers in Fig. 40--52 are seconds
in play back frames. Scale bar = 10 µm.
Fig. 53--64. Feeding processes of Gyrodinium sp. on fluorescent labeled bacteria (FLB) and
bacteria-sized fluorescent beads (0.5 μm in diameter), observed under an
epifluorescence microscope, recorded using video microscopy. 53. Drawing on the path
of ingested (red and blue colors) and non-ingested (black color) FLBs and beads in
feeding currents generated by a Gyrodinium sp. cell. The feeding currents were
generated while the longitudinal flagellum undulated. 54--58. Ingestion of a FLB
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(arrow). 54. A FLB (white arrow) was trapped in a strong water current flowing from
above the epicone (arrow head), via the long and narrow sulcus, to below the hypocorn
of the predator. 55--57. The FLB moved by the water current flowing through the sulcus.
58. The FLB was ingested at the lower part of the sulcus at the hypocone. 59, 60. A FLB
observed inside the protoplasm of the predator. 61--64. Ingestion of a bead (arrow). 61.
Two beads (white and black arrows) were trapped in a strong water current near the
epicone (arrow head) of the predator. 62. The beads moved by the water current flowing
through the sulcus. 63. One bead (white arrow) was ingested at the lower part of the
sulcus at the hypocone, but, the other bead (black arrow) was not ingested. 64. One bead
observed inside the protoplasm of the predator. All scale bars = 10 µm.
Fig. 65. Ingestion rates (IR, cells dinoflagellate-1h-1) of Oxyrrhis marina on bacteria as a
function of the initial prey concentration (cells ml-1, x). Each value of the ingestion rate
was calculated by exploration from a linear regression curve on the number of prey
cells inside an algal predator cell over incubation time (see text for calculation).
Symbols represent treatment means + 1 S.E. The curve was fitted by a Michaelis-
44
Menten equation [Eq. (3)] using all treatments in the experiment. IR = 71.3
[x/(1,110,000+x)], r2=0.750.
Fig. 66. Ingestion rates (IR, cells dinoflagellate-1h-1) of Gyrodinium cf. guttula on bacteria
as a function of the initial prey concentration (cells ml-1, x). Each value of the ingestion
rate was calculated as in Fig. 65. Symbols represent treatment means + 1 S.E. The curve
was fitted by a Michaelis-Menten equation [Eq. (3)] using all treatments in the
experiment. IR = 23.2 [x/(1,330,000+x)], r2=0.515.
Fig. 67. Ingestion rates (IR, cells dinoflagellate -1h-1) of Pfiesteria piscicida on bacteria as a
function of the initial prey concentration (cells ml-1, x). Each value of the ingestion rate
was calculated as in Fig. 65. Symbols represent treatment means + 1 S.E. The curve was
fitted by a Michaelis-Menten equation [Eq. (3)] using all treatments in the experiment.
IR = 13.7 [x/(823,000+x)], r2=0.648.
Fig. 68. Growth rate (GR, d-1) of Oxyrrhis marina on bacteria as a function of the initial
45
prey concentration (cells ml-1, x). Symbols represent treatment means + 1 S.E. The
curve was fitted by a Michaelis-Menten equation [Eq. (5)] using all treatments in the
experiment. GR = 0.592 {[x-41,000]/[1,110,000+(x-41,000)]), r2=0.819.
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47
48
49
50
51
52
Fig. 65
53
Fig. 66
54
Fig. 67
55
Fig. 68
56