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: hjjeong@snu.ac.kr

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

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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