Hydrodynamic stimulation of dinoflagellate bioluminescence: a computational and experimental study
1 Scripps Institution of Oceanography, University of California San Diego,
9500 Gilman Drive, La Jolla, CA 92093-0202, USA
2 Mechanical and Aerospace Engineering Department, University of California,
Irvine, CA 92697, USA
3 SSC San Diego, 53560 Hull Street, D363, San Diego, CA 92152,
USA
* Author for correspondence (e-mail: mlatz{at}ucsd.edu)
Accepted 11 March 2004
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: acceleration, bioluminescence, Ceratocorys, dinoflagellate, flow, Lingulodinium, numerical simulation, shear
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Flow-stimulated dinoflagellate luminescence is well known anecdotally from
numerous observations of oceanic bioluminescence associated with breaking
surface waves, swimming organisms and moving ships (e.g.
Hobson, 1966;
Staples, 1966
;
Tett and Kelley, 1973
;
Rohr et al., 1998
).
Ecologically, dinoflagellate bioluminescence is thought to serve an
anti-predatory role (Esaias and Curl,
1972
; White, 1979
;
Buskey et al., 1983
,
1985
;
Mensinger and Case, 1992
;
Fleisher and Case, 1995
).
Laboratory studies of dinoflagellate bioluminescence indicate that fluid shear
stress is an important stimulatory component
(Latz et al., 1994
;
Latz and Rohr, 1999
), although
fluid acceleration has also been suggested as a stimulus for bioluminescence
(Anderson et al., 1988
). The
objective of this study was to examine the response of luminescent
dinoflagellates in an independent flow field that allowed assessment of the
relative stimulatory contributions of acceleration and shear stress. In this
context, acceleration was associated with velocity gradients along a
streamline and not unsteady or angular velocity.
The present study used a smooth converging nozzle, providing a laminar flow
field, to assess the importance of shear stress and acceleration in
stimulating dinoflagellate bioluminescence. Advantages of this flow field are:
(1) regions of relatively high shear stress and acceleration are spatially
separated, (2) properties of the flow field change continually along the
downstream direction and (3) the governing hydrodynamic equations are exactly
known. Moreover, compared with previous flow fields used to study
bioluminescence stimulation (Latz et al.,
1994; Latz and Rohr,
1999
), the nozzle flow is unique in that there is no depletion of
bioluminescence capacity prior to measurement.
Laboratory tests were performed over a range of flow rates for two
dinoflagellate species, Lingulodinium polyedrum (formerly
Gonyaulax polyedra) and Ceratocorys horrida. L. polyedrum,
approximately 35 µm in diameter and common in coastal waters
(Lewis and Hallett, 1997), is
the most well-characterized dinoflagellate in terms of its luminescent
response to flow (e.g. Anderson et al.,
1988
; Latz et al.,
1994
; Latz and Rohr,
1999
). The oceanic species C. horrida, endemic to warm
oligotrophic regions (Graham,
1942
), is approximately twice as large as L. polyedrum
and possesses prominent antapical spines
(Zirbel et al., 2000
). Both
species have similar flash durations of approximately 150 ms
(Latz and Lee, 1995
),
resulting in a pathline illuminating the cell trajectory in the flow
(Latz et al., 1995
). Previous
studies with fully developed pipe flow indicate that the threshold luminescent
response of C. horrida occurs in flows with shear stress levels
approximately one order of magnitude less than for L. polyedrum
(Nauen, 1998
).
The following hypotheses were tested:
The position of cell stimulation and the corresponding flow properties at those locations were determined using a combination of video observations of individual flashes within the nozzle and numerical simulations of the flow field for identical flow conditions as the experiments.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() | (1) |
Flow from the head tank through the nozzle was driven by gravity and
controlled by a manual valve at the pipe exit. The Reynolds number
(Re) of each flow rate
(Re=UavgD/µ, where
is
fluid density and µ is fluid dynamic viscosity) was calculated based on
average flow velocity (Uavg) at the nozzle exit diameter
D (where D=2Ye), determined from
measurements of volumetric flow rate made at the beginning, end and at 1-min
intervals throughout each experiment. Values of Re were rounded off
to the nearest hundred. Observations of injected dye confirmed that the flow
remained laminar for all flow rates tested.
Experimental approach
Cultures of Lingulodinium polyedrum Stein Dodge and
Ceratocorys horrida Stein were grown in half-strength f/2 medium
(Guillard and Ryther, 1962)
minus silicate and maintained at 20±0.5°C in an environmental
chamber on a 12 h:12 h light:dark cycle as previously described
(Latz and Rohr, 1999
).
Two types of experiments were performed. Cell suspension experiments, in which a homogenous distribution of organisms was present in the flow field, involved no a priori assumptions about the position of cell stimulation. Cell injection experiments introduced cells at specific radial positions at the nozzle inlet to verify the position of stimulated cells as determined by the cell suspension experiments.
Prior to the end of the light phase, when cells are mechanically
inexcitable (Biggley et al.,
1969; Latz and Lee,
1995
), subsamples of the cultures were diluted with 0.45 µm
filtered seawater if necessary and, depending on the experiment, added to the
head tank for cell suspension experiments or loaded into syringes for cell
injection experiments. At the beginning of the dark phase, the room was
darkened and cells were thereafter subjected to short periods of dim red light
only. Testing commenced 2.54 h into the dark phase, when stimulated
bioluminescence is maximal (Biggley et al.,
1969
; Latz and Lee,
1995
). Room temperature was 1920°C and varied by
<0.5°C during each experiment.
Bioluminescence within the nozzle was imaged with an intensified SIT video camera (Cohu Inc. model 55) or intensified CCD video camera (Dage GenIISYS), each fitted with a Fujinon 25 mm lens used at f/0.85 or f/1.4 and fitted with a +4 diopter lens. Video frame rate was 30 Hz. The 12 cm-wide camera field of view encompassed the entire nozzle, with the focal plane centered on the nozzle centerline.
For cell suspension experiments, cultures of either L. polyedrum or C. horrida were diluted into filtered seawater to give a final volume of 16 liters and a calculated cell concentration of 15 or 30 cells ml-1, respectively. Random swimming by both species helped maintain a nearly homogenous distribution of organisms; thus, it was assumed that mean cell concentration did not vary within a test. A single daily experiment consisted of one filling of the head tank and tests with several flow rates.
Cell injection experiments examined the response of cells introduced at known radial positions at the nozzle inlet. Cells were injected at the nozzle inlet along a radius in a plane perpendicular to the axis of the camera. Cells were loaded into 60 ml plastic syringes fitted to TeflonTM tubing coupled to a plastic pipette tip with a 0.04 cm orifice. The tip orifice was positioned flush with the inlet of the nozzle, with the radial position of the tip controlled by a micromanipulator. Cells were injected at the nozzle inlet at a rate of 0.008 ml s-1. Dye studies were first performed using the identical injection apparatus to visualize the trajectory of injected material at different radial positions. The dye stream was observed to remain in the injection plane throughout its passage through the nozzle, confirming that the radial position of injected material could be measured directly from the two-dimensional video record.
Two types of cell injection experiments were performed. First, using L.
polyedrum, a series of eight radial positions between wall and centerline
was tested at Re2500 (Table
1). Second, cells of L. polyedrum and C. horrida
were injected at centerline to verify that, because of their response latency,
they were not responding out of view of the initial camera position. This
concern is greatest for cells moving along centerline, where flow velocity is
highest and cells stimulated within the nozzle could respond as much as
several centimeters downstream of the nozzle. To verify that cells were not
being stimulated downstream out of the camera view, the camera position was
moved so that it imaged 20 cm downstream of the nozzle to account for a
response latency as high as 0.1 s, five times the estimated response latency
of 20 ms (Widder and Case,
1981
). Cells were injected at centerline for the highest flow rate
tested in the cell suspension experiments. Periodically, the injector position
was moved to the wall to verify cell stimulability.
|
Video analysis
The objective of the video analysis of cell suspension and injection
experiments was to provide the position of each flash response so that
subsequent numerical simulations could calculate the position of cell
stimulation and the hydrodynamic parameters at that location. Although L.
polyedrum and C. horrida can produce more than one flash
(Latz and Lee, 1995), during
the brief residence time in the nozzle no more than 1 flash cell-1
was observed. Flashes typically lasted for 34 successive video frames
and, because the cells were moving, appeared as streaks within each video
frame (Fig. 1). Approximately
40 individual flashes were analyzed for each flow rate using single-frame
playback of the video record on a video monitor. The precision for measuring
the position of a flash on the screen of the video monitor was 0.05 cm. Unless
otherwise stated, values represent the arithmetic mean with one standard error
of the mean. In some cases, median values were used for comparison between
flow rates. Statistical tests were performed using Statview software (SAS
Institute, Inc., Cary, NC, USA).
|
The downstream position, X1, of flash initiation was always obtained directly from the individual video frames. For cell injection experiments, the radial position of flash initiation, Y1, was also measured directly from the video record because flashes occurred in the injection plane normal to the camera. For cell suspension experiments, flash radial position could not be measured directly because flashes could occur azimuthally at any angle within the nozzle; therefore, a different analysis method was required. For this case, the flash radial position, Y1, was estimated from average flash velocity, determined by the change in position of the leading edge of a flash streak in consecutive video frames. Given X1 and flash velocity, Y1 was calculated from the numerical flow simulations. For both types of experiments, once the downstream and radial positions of flash initiation (X1, Y1) were determined, numerical simulations adjusted for flash latency to provide the position of cell stimulation (X0 and Y0) and the values of acceleration and shear stress at that position (see next section).
As a check for accuracy, both methods of determining radial position, Y1, were compared for a subset of flashes recorded during the cell injection experiments. As previously described, for injected cells the radial position of flash initiation can be determined directly from each video frame by direct measurement. These radial positions were compared with those estimated from the average flash speed analysis using numerical simulations.
Numerical simulations
Numerical simulations of the nozzle flow field allowed for high-resolution
mapping of the position of organism stimulation and the flow parameters at
that position. This approach was especially important because flow properties
changed continuously in the downstream direction. Numerical simulations served
three purposes: (1) to obtain the radial position of flash initiation,
Y1, for cell suspension experiments; (2) to account for
the response latency of organisms within the developing flow field, such that
the position of cell stimulation was upstream of that for flash initiation;
and (3) to calculate values of acceleration and shear stress at the position
of stimulated cells.
For these simulations, it was assumed that the organisms behaved as fluid
particles and that their presence had no effect on the flow. This assumption
is quite plausible because the volume fraction of the organisms, approximately
10-7, was sufficiently low
(Elghobashi, 1994), their
density is only slightly greater than that of the liquid water
(Kamykowski et al., 1992
) so
they are almost neutrally buoyant, and their local swimming speed
(Kamykowski et al., 1992
) is
much less than the carrier flow velocity. Previous pipe flow experiments at
these organism concentrations demonstrated no effect on the Newtonian nature
of the flow (Latz and Rohr,
1999
). Moreover, video recordings of flash and dye trajectories
from cell and dye injection studies showed no apparent differences, suggesting
that, for the purpose of calculating the position of stimulation, individual
cells followed fluid streamlines.
The numerical method computed the properties (velocity and pressure) of the flow inside the nozzle. This flow was laminar, incompressible and axisymmetric. The governing equations were the NavierStokes and continuity equations, which are expressions of conservation of momentum and mass, respectively. These partial differential equations were discretized on a boundary-fitted orthogonal curvilinear grid generated by solving a system of Laplace equations with suitable boundary conditions to satisfy orthogonality. Prescribed boundary conditions were: no-slip along the nozzle wall, axisymmetry along centerline, uniform velocity profile at the nozzle inlet plane, and zero axial velocity at the exit plane.
For the two-dimensional nozzle geometry, there were two coordinate
variables, X and Y. The geometry was mapped from the
physical domain X,Y onto a uniformly spaced orthogonal ,
domain (Mobley and Stewart,
1980
). The resulting grid allowed the governing equations to be
discretized using the finite volume method in which the solution domain was
divided into contiguous quadrilateral curvilinear control volumes. A modified
SIMPLE algorithm (Patankar,
1980
) was used to solve the discretized equations and obtain the
two components of velocity and pressure at the grid nodes.
The following steps were taken to obtain the flow conditions responsible
for stimulation of the organism (Table
2). The flow field was computed for similar Reynolds numbers as
the experiments. The downstream (X1) and radial
(Y1) position of flash initiation was obtained as
described in the previous section. The final position (X0
and Y0) of organism stimulation was calculated from
X1 and Y1 by following the streamline
upstream an additional 20 ms to account for the response latency
(Widder and Case, 1981). The
hydrodynamic properties at the stimulation position (X0
and Y0) were calculated using cubic-spline interpolation
of the properties at the fixed grid nodes.
|
The effect of steady and changing pressure on bioluminescence stimulation
was not considered in this study because dinoflagellates are known to be
relatively insensitive to pressure (Gooch
and Vidaver, 1980; Swift et
al., 1981
; Krasnow et al.,
1981
; Donaldson et al.,
1983
). If shear stress thresholds of 0.1 N m-2 for
luminescent dinoflagellates were the same for pressure, above-threshold levels
would exist essentially throughout the entire ocean
(Rohr et al., 2002
).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Cell suspension experiments
Despite being distributed throughout the flow volume, flash responses of
L. polyedrum and C. horrida were only observed at the nozzle
throat (Fig. 1). For both
species, the downstream position, X1, of flash initiation
(Fig. 3) shifted significantly
upstream with increasing flow rate (one-way ANOVA: L. polyedrum,
F=313.1, d.f.=2, 119, P<0.0001; C. horrida, F=136.5,
d.f.=3, 160, P<0.0001). The differences between species were
consistent with the higher flow sensitivity of C. horrida
(Nauen, 1998). First, the
minimum flow rate to which C. horrida responded (Re=400) was
lower than for L. polyedrum (Re=800). Second, at similar
flow rates, flashes of C. horrida were initiated higher in the nozzle
than for L. polyedrum. For example, at the highest flow rate tested
(Re=5100) the mean downstream position of 3.9±0.08 cm for
C. horrida was significantly different from that of 4.8±0.03
cm for L. polyedrum (t-test, t=9.94, d.f.=81,
P<0.0001).
|
Greater than 99% of all cells were stimulated within the boundary layer;
four responses by L. polyedrum attributed to cells outside the
boundary layer may have been spontaneous flashes
(Sweeney and Hastings, 1958;
Latz and Lee, 1995
) and not
flow stimulated. For all flow rates, L. polyedrum cells were
stimulated at a mean position of <0.035 cm from the wall, within the
boundary layer (data not shown). Values of acceleration at the position of
stimulated cells were always <4 m s-2 and frequently near zero,
especially at the lowest flow rate (Fig.
4). The minimum value of shear stress at these cell positions
within the boundary layer was 0.7, 0.6 and 0.6 N m-2 for
Re=800, 2500 and 5100, respectively.
|
All C. horrida cells were stimulated within the boundary layer,
with a mean of <0.028 cm from the wall (data not shown). At
Re=400, the minimum stimulatory flow rate tested, values of fluid
acceleration at the position of stimulated cells were <0.59 m
s-2 (<0.1 g)
(Fig. 5). At these positions,
shear stress was 0.080.44 N m-2. Even at the highest flow
rate tested (Re=5100), all stimulated cells were located near the
wall where acceleration was <2 m s-2 (0.2
g). The minimum value of shear stress at the cell positions
was 0.08, 0.17, 0.05 and 0.09 N m-2 for Re=400, 900, 2300
and 5100, respectively.
|
To assess the relative stimulatory effects of acceleration and shear stress, values at the position of stimulated cells (X0, Y0) were normalized to maximum levels along the nozzle radius, Y, for that downstream X0 position. For both species, mean values of shear stress were 79100% of maximum when mean values of acceleration were only 321% of maximum (Fig. 6).
|
Calculation of the position of a stimulated cell is sensitive to the value
of response latency used in the numerical simulations. For example, a longer
response latency will translate the position of stimulation further upstream
from the position of flash initiation. Because the response latency of L.
polyedrum and C. horrida to flow stimulation is unknown, the
value of 20 ms, obtained for mechanical stimulation of the dinoflagellate
Pyrocyctis fusiformis (Widder and
Case, 1981), was used. To examine the sensitivity of the
computational results to the chosen value of response latency, differences in
the calculated position of stimulated cells of L. polyedrum as a
function of response latency were assessed for three flow rates. Latency
values of 5, 10, 15 and 25 ms were tested in addition to the 20 ms standard.
At the lowest flow rate tested (Re=800), there were no resolvable
differences in downstream cell position and the ranges of acceleration and
shear stress at the position of stimulated cells for the different latency
values. At the intermediate flow rate of Re=2500, increasing latency
values `pushed' the position of stimulated cells further upstream, resulting
in decreases in values of shear stress at the cell position but resulting in
minimal differences in acceleration. At the highest flow rate
(Re=5100), the value of response latency affected both the cell
downstream position and shear stress at the position of stimulated cells.
However, regardless of the chosen value of response latency, cells were always
stimulated in the boundary layer where shear stress levels were high compared
with elsewhere in the flow field.
Cell injection experiments
To further confirm that organisms were stimulated within the wall shear
layer and not in the region of high acceleration, cells of L.
polyedrum were injected at the nozzle inlet in the plane normal to the
axis of the camera at different radial positions from centerline to the inlet
wall. No flashes were observed at the injector, suggesting that there was
minimal pre-stimulation of cells due to the injection procedure. Only cells
injected at >0.7Yin were stimulated within the nozzle
(Fig. 7A).Stimulated cells had
a mean downstream position, X0, of 5.5 cm and were located
approximately 0.01 cm from the wall, within the boundary layer (data not
shown). The maximum response rate of 71% of injected cells, for an injection
position of 0.8Yin, indicated that most cells were
responding. Shear stress levels at the position of stimulated cells were near
maximum for that downstream (X) position, while acceleration was
<10% of maximum levels outside the boundary layer
(Fig. 7B).
|
Cell injection experiments also allowed comparison of the numerical simulation estimates of Yo with those from direct video analysis. Analysis of 40 flashes, from an experiment in which cells were injected at 0.8Yin at the nozzle inlet at Re=2500, showed little difference between the two methods. Video analysis gave a radial position Y1=0.23±0.03 cm at a downstream position X1=5.31±0.19 cm; the numerical simulations calculated a radial position Y0=0.22±0.02 cm at X0=5.46±0.18 cm. The difference between methods was less than the uncertainty associated with determining the flash initiation position on the video monitor. Although each method has its own assumptions and limitations, both predict flash positions within the boundary layer and yield similar response trends.
Because centerline velocities were as high as 2 m s-1, a response latency of 20 ms could result in flashes occurring as much as 4 cm downstream from where they were stimulated, possibly outside the initial camera field of view. To account for this possible bias, cells injected along centerline were monitored up to 20 cm downstream of the nozzle exit. Dye experiments showed that the flow remained laminar throughout this region. At Re=5100, the highest flow rate tested in this study, no flashes from >3000 cells of L. polyedrum injected at centerline were observed either within the nozzle or downstream of the nozzle within the exit pipe. For almost 3000 cells of C. horrida injected at centerline for Re=5100, 3% of the cells were observed responding within the exit pipe. During these experiments, cell viability was confirmed by periodically moving the injection point to the inlet wall, where numerous flashes were observed.
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Changes in flow rate affected the levels of acceleration and shear stress
within the flow field but did not alter the general flow pattern. For both
species, there was a significant change in the downstream position of
stimulated cells, as the stimulation position moved upstream with increasing
flow rate. This response was consistent with the higher levels of shear stress
within the upstream boundary layer as flow rate increased. A comparison of the
responses of the two species indicated that differences in the spatial pattern
of stimulation reflected the species flow sensitivity. C. horrida
stimulated cells were positioned higher in the flow field, for equivalent flow
rates, and responded at lower flow rates than for L. polyedrum,
consistent with the former's greater flow sensitivity
(Nauen, 1998).
For three completely independent flow fields simple Couette flow,
fully developed pipe flow and nozzle flow the luminescent response is
consistent with a mechanism of stimulation based on fluid shear. These first
two flow fields are dominated by shear. For Couette flow in the gap between
concentric cylinders, with the outer cylinder rotating, there is a nearly
linear velocity gradient (and thus nearly constant shear) between the outer
and inner cylinders; the mean shear stress in the gap changes as a function of
angular rotation (van Duuren,
1968). Although there is centripetal acceleration, there is no
acceleration along velocity gradients. In fully developed laminar pipe flow,
the parabolic velocity distribution across the pipe radius results in a
gradient of shear stress, with maximum shear at the wall and zero shear at
centerline (Schlichting,
1979
). In this flow field there is no mean acceleration and the
mean shear profile across the pipe is balanced by the pressure gradient. In
nozzle flow, used in the present study, most of the volume is dominated by
acceleration, with only a thin shear layer near the wall at the nozzle throat.
Nozzle flow is also different from the other flow fields in that it presents a
developing flow field where flow parameters change dramatically throughout the
volume.
For all three flow fields, response thresholds are determined based on the
minimum flow condition in which bioluminescence is stimulated. In the present
study with nozzle flow, a response threshold was obtained for each flow rate
based on the minimum shear stress value at the position of all stimulated
cells. For L. polyedrum, the response threshold for shear stress was
0.6 N m-2. This response threshold is similar to, although somewhat
higher than, the shear stress threshold of 0.3 N m-2 obtained for
fully developed pipe flow at similar concentrations
(Latz and Rohr, 1999). For
C. horrida, the shear stress response threshold obtained for nozzle
flow was 0.05 N m-2 while that for fully developed pipe flow is
0.02 N m-2 (Nauen,
1998
). Considering that a difference in flash location of as small
as 0.01 cm can result in significantly different flow properties, as well as
the uncertainty in organism response latency, the experimental results using
nozzle flow were remarkably consistent with those from other flow fields.
Overall, these results demonstrate that organisms are responding to specific,
quantitative, hydrodynamic aspects of the flow, regardless of the flow field
used.
Flow sensing
Dinoflagellate bioluminescence is considered to have an anti-predator
function by decreasing grazing pressure
(Esaias and Curl, 1972;
White, 1979
) through altering
predator swimming behavior (Buskey et al.,
1983
,
1985
). However, the response
threshold for dinoflagellate bioluminescence, occurring in flows with shear
stress levels in the order of 0.1 N m-2
(Latz et al., 1994
;
Nauen, 1998
;
Latz and Rohr, 1999
; present
study), equivalent to fluid strain rates of approximately 100 s-1,
is several orders of magnitude higher than response thresholds for
flow-stimulated predator avoidance behaviors by other planktonic organisms
(reviewed by Kiørboe et al.,
1999
; Jakobsen,
2001
). Are levels of fluid strain sufficient to stimulate
bioluminescence present in the feeding current of a predator? Using siphon
flow as a mimic of a grazer feeding current, a feeding current flow rate of
0.279 ml s-1 (table
2 of Jakobsen,
2001
) and equation 2 of Kiørboe et al.
(1999
), bioluminescence
stimulation is estimated to occur at a distance of only 0.06 cm from the
siphon inlet. If the siphon flow field represents the feeding current of a
filtering dinoflagellate predator, then dinoflagellate bioluminescence would
only be stimulated very close to the predator. Video observations of the
interactions between dinoflagellates and their copepod grazers verify that
bioluminescence is associated with feeding activities and not flow disturbance
(Buskey et al., 1985
).
Therefore, it is unlikely that dinoflagellate bioluminescence is stimulated by
the flow within a predator feeding current, as are the escape behaviors of
copepods, rotifers and ciliates. Rather, stimulation may occur either by
direct handling of the cell by a grazer or by the shear stress produced by a
moving organism. Bioluminescence generated by swimming animals in a
`minefield' of luminescent dinoflagellates can allow visual predators to
better find prey (Mensinger and Case,
1992
; Fleisher and Case,
1995
). Oceanic conditions providing supra-threshold shear levels
include not only the boundaries of swimming organisms
(Rohr et al., 1998
) but also
flow at the ocean boundaries and in highly turbulent events such as breaking
waves (Rohr et al., 2002
).
Under what conditions would accelerating flows be stimulatory? In a study
of L. polyedrum (Anderson et al.,
1988), bioluminescence was not stimulated by steady centripetal
acceleration as high as 100 g or changing centripetal
acceleration of 1 g s-1. Only variable centripetal
acceleration of the order of 10 g s-1, associated
with abrupt starts and stops of a rotating chamber, were considered
stimulatory, but it is unclear if the stimulation resulted from the
acceleration change or from vibration or start/stop transients. In the present
study, accelerations as high as 20 g, with estimated rates of
change of >10 g s-1, were not stimulatory to
L. polyedrum. It is unlikely that accelerations above this magnitude
are prevalent in the ocean, so acceleration is not an ecologically relevant
stimulus of dinoflagellate bioluminescence.
Nevertheless, highly accelerating flows may be important tools in
understanding mechanotransduction. Flow stimuli are believed to elicit
bioluminescence as a result of cell deformation, leading to a signaling
process involving a calcium-mediated second messenger pathway
(von Dassow and Latz, 2002).
The initial step may involve changes in membrane fluidity
(Mallipattu et al., 2002
) or
activation of plasma membrane proteins as in other flow-sensitive organisms
(Gudi et al., 1996
;
Labrador et al., 2003
). This
process leads to generation of an action potential at the vacuole membrane
(Eckert, 1966
) that results in
proton flux into the cytoplasm, activating the luminescent chemistry
(Fritz et al., 1990
). Flow
conditions with equivalent levels of fluid strain are expected to cause
identical amounts of cell deformation, whether the strain is due to elongation
within an accelerating fluid (due to the velocity gradient along a streamline)
or shear (due to the velocity gradient across streamlines). Bioluminescence
should therefore be stimulated in highly accelerating flows in which the
elongation stress is greater than the known threshold for shear stress. The
equivalence of elongation and shear in eliciting a biological response was
experimentally validated for escape swimming of the copepod Acartia
tonsa (Kiørboe et al.,
1999
). The issue is difficult to investigate for bioluminescence
stimulation because of the very high flow rates necessary to obtain
sufficiently high levels of fluid strain from acceleration.
Dinoflagellate flow visualization
Dinoflagellate bioluminescence is a powerful tool for flow visualization
under conditions not amenable to conventional methods
(Latz et al., 1995;
Rohr et al., 1998
). Organisms
are the size of typical flow markers
(Irani and Callis, 1973
), can
remain in suspension due to cell swimming or near-neutral buoyancy, respond
nearly instantaneously within regions of above threshold shear stress, and can
be used at cell concentrations that have no effect on the Newtonian nature of
the fluid. A further advantage is that dinoflagellate bioluminescence is
easily visualized and quantified. Unlike visualization of conventional flow
markers, bioluminescence can be detected by low-light camera systems at
distances in the order of meters, without the need to optically discriminate
individual organisms. The level of average bioluminescence intensity, a
function of species abundance, stimulatory volume and level, and the number of
organisms responding (Rohr et al.,
1998
; Latz and Rohr,
1999
), is readily measured by extremely sensitive photomultiplier
detectors. Demonstrated uses of dinoflagellate flow visualization include
visualization of boundaries and flow separation
(Latz et al., 1995
;
Rohr et al., 1998
) and
pinpointing regions of high shear in bioreactors
(Chen et al., 2003
). Possible
applications of luminescent flow visualization include mapping regions of high
shear in prosthetic heart valve flows
(Yoganathan et al., 1998
) and
identifying highly dissipative regions of flow in the ocean
(Rohr et al., 2002
).
The present study successfully incorporated computational and experimental
fluid approaches to examine a transient biological process. Even though the
organisms were advected through this flow field of the order of seconds, the
near-instantaneous luminescent response and use of numerical simulations
allowed high-resolution mapping of the stimulatory regions of the flow field.
Computational approaches have tremendous potential in determining the forces
on, and deformation of, small organisms in both laminar and turbulent flows
(e.g. Jiang et al., 2002).
When coupled with numerical simulations, the experimental use of
dinoflagellate bioluminescence is an effective technique to resolve flow
properties at the small spatial and temporal scales relevant to the organism
and holds promise for providing new understanding of complex flow phenomena
(Stokes et al., 2004
).
![]() |
Acknowledgments |
---|
![]() |
Footnotes |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Anderson, D. M., Nosenchuck, D. M., Reynolds, G. T. and Walton, A. J. (1988). Mechanical stimulation of bioluminescence in the dinoflagellate Gonyaulax polyedra Stein. J. Exp. Mar. Biol. Ecol. 122,277 -288.[CrossRef]
Biggley, W. H., Swift, E., Buchanan, R. J. and Seliger, H.
H. (1969). Stimulable and spontaneous bioluminescence in the
marine dinoflagellates, Pyrodinium bahamense, Gonyaulax polyedra, and
Pyrocystis lunula. J. Gen. Physiol.
54, 96-122.
Buskey, E., Mills, L. and Swift, E. (1983). The effects of dinoflagellate bioluminescence on the swimming behavior of a marine copepod. Limnol. Oceanogr. 28,575 -579.
Buskey, E. J., Reynolds, G. T., Swift, E. and Walton, A. J. (1985). Interactions between copepods and bioluminescent dinoflagellates: direct observations using image intensification. Biol. Bull. 169,530 .
Chen, A. K., Latz, M. I. and Frangos, J. A. (2003). The use of bioluminescence to characterize cell stimulation in bioreactors. Biotechnol. Bioeng. 83, 93-103.[CrossRef][Medline]
Donaldson, T. Q., Tucker, S. P. and Lynch, R. V. (1983). Stimulation of bioluminescence in dinoflagellates by controlled pressure changes. Naval Res. Lab. Report 8772.
Eckert, R. (1966). Excitation and luminescence in Noctiluca miliaris. In Bioluminescence in Progress (ed. F. Johnson and Y. Haneda), pp.269 -300. Princeton, NJ: Princeton University Press.
Elghobashi, S. E. (1994). On predicting particle-laden turbulent flows. Appl. Sci. Res. 52,309 -329.
Esaias, W. E. and Curl, H. C., Jr (1972). Effect of dinoflagellate bioluminescence on copepod ingestion rates. Limnol. Oceanogr. 17,901 -906.
Fields, D. M. and Yen, J. (1996). The escape behavior of Pleuromamma xiphias in response to a quantifable fluid mechanical disturbance. Mar. Freshw. Behav. Physiol. 27,323 -340.
Fields, D. M. and Yen, J. (1997). Implications of the feeding current structure of Euchaeta rimana, a carnivorous pelagic copepod, on the spatial orientation of their prey. J. Plankton Res. 19,79 -95.[Abstract]
Fleisher, K. J. and Case, J. F. (1995).
Cephalopod predation facilitated by dinoflagellate luminescence.
Biol. Bull. 189,263
-271.
Fritz, L., Morse, D. and Hastings, J. W. (1990). The circadian bioluminescence rhythm of Gonyaulax is related to daily variations in the number of light-emitting organelles. J. Cell Sci. 95,321 -328.[Abstract]
Gooch, V. D. and Vidaver, W. (1980). Kinetic analysis of the influence of hydrostatic pressure on bioluminescence of Gonyaulax polyedra. Photochem. Photobiol. 31,397 -402.
Graham, H. W. (1942). Studies in the morphology, taxonomy, and ecology of the Peridiniales. Scient. Results Cruise VII Carnegie, Biol. Ser. 3, 1-129.
Gudi, S. R. P., Clark, C. B. and Frangos, J. A.
(1996). Fluid flow rapidly activates G proteins in human
endothelial cells. Involvement of G proteins in mechanochemical signal
transduction. Circ. Res.
79,834
-839.
Guillard, R. R. L. and Ryther, J. H. (1962). Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt, and Detonula confervacea (Cleve) Gran. Can. J. Microbiol. 8,229 -239.
Hobson, E. S. (1966). Visual orientation and feeding in seals and sea lions. Nature 214,326 -327.
Irani, R. R. and Callis, C. F. (1973). Particle Size: Measurement, Interpretation, and Application. New York: Wiley.
Jakobsen, H. H. (2001). Escape response of planktonic protists to fluid mechanical signals. Mar. Ecol. Progr. Ser. 214,67 -78.
Jiang, H., Meneveau, C. and Osborn, T. R.
(2002). The flow field around a freely swimming copepod in steady
motion. Part II: Numerical simulation. J. Plankton
Res. 24,191
-213.
Juhl, A. R. and Latz, M. I. (2002). Mechanisms of fluid shear-induced inhibition of population growth of a red-tide dinoflagellate. J. Phycol. 38,683 -694.[CrossRef]
Juhl, A. R., Velazquez, V. and Latz, M. I. (2000). Effect of growth conditions on flow-induced inhibition of population growth of a red-tide dinoflagellate. Limnol. Oceanogr. 45,905 -915.
Juhl, A. R., Trainer, V. L. and Latz, M. I. (2001). Effect of fluid shear and irradiance on population growth and cellular toxin content of the dinoflagellate Alexandrium fundyense.Limnol. Oceanogr. 46,758 -764.
Kamykowski, D., Reed, R. E. and Kirkpatrick, G. J. (1992). Comparison of sinking velocity, swimming velocity, rotation and path characteristics among six marine dinoflagellate species. Mar. Biol. 113,319 -328.
Kiørboe, T., Saiz, E. and Visser, A. (1999). Hydrodynamic signal perception in the copepod Acartia tonsa. Mar. Ecol. Progr. Ser. 179,97 -111.
Krasnow, R., Dunlap, J., Taylor, W., Hastings, J. W., Vetterling, W. and Haas, E. (1981). Measurements of Gonyaulax bioluminescence including that of single cells. In Bioluminescence Current Perspectives (ed. K. H. Nealson), pp. 52-63. Minneapolis: Burgess Publishing.
Labrador, V., Chen, K.-D., Li, Y.-S., Muller, S., Stoltz, J.-F. and Chien, S. (2003). Interactions of mechanotransduction pathways. Biorheology 40, 47-52.[Medline]
Latz, M. I. and Lee, A. O. (1995). Spontaneous and stimulated bioluminescence in the dinoflagellate, Ceratocorys horrida (Peridiniales). J. Phycol. 31,120 -132.
Latz, M. I. and Rohr, J. (1999). Luminescent response of the red tide dinoflagellate Lingulodinium polyedrum to laminar and turbulent flow. Limnol. Oceanogr. 44,1423 -1435.
Latz, M. I., Case, J. F. and Gran, R. L. (1994). Excitation of bioluminescence by laminar fluid shear associated with simple Couette flow. Limnol. Oceanogr. 39,1424 -1439.
Latz, M. I., Rohr, J. and Hoyt, J. (1995). A novel flow visualization technique using bioluminescent marine plankton Part I: Laboratory studies. IEEE J. Oceanic Eng. 20,144 -147.
Lewis, J. and Hallett, R. (1997). Lingulodinium polyedrum (Gonyaulax polyedra) a blooming dinoflagellate. Oceanogr. Mar. Biol. Annu. Rev. 35, 97-161.
Mallipattu, S. K., Haidekker, M. A., von Dassow, P., Latz, M. I. and Frangos, J. A. (2002). Evidence for shear-induced increase in membrane fluidity in the dinoflagellate Lingulodinium polyedrum. J. Comp. Physiol. A 188,409 -416.
Mensinger, A. F. and Case, J. F. (1992). Dinoflagellate luminescence increases susceptibility of zooplankton to teleost predation. Mar. Biol. 112,207 -210.
Mobley, C. D. and Stewart, R. J. (1980). Numerical generation of boundary-fitted orthogonal curvilinear coordinate systems. J. Comput. Phys. 34,124 -135.
Nauen, J. C. (1998). Biomechanics of two aquatic defense systems. Ph.D. Dissertation. University of California, San Diego.
Patankar, S. V. (1980). Numerical Heat Transfer and Fluid Flow. Washington: Hemisphere Pub. Corp.; New York: McGraw-Hill.
Peters, F. and Marrasé, C. (2000). Effects of turbulence on plankton: an overview of experimental evidence and some theoretical considerations. Mar. Ecol. Prog. Ser. 205,291 -306.
Rohr, J., Latz, M. I., Fallon, S., Nauen, J. C. and Hendricks,
E. (1998). Experimental approaches towards interpreting
dolphin-stimulated bioluminescence. J. Exp. Biol.
201,1447
-1460.
Rohr, J., Hyman, M., Fallon, S. and Latz, M. I. (2002). Bioluminescence flow visualization in the ocean: an initial strategy based on laboratory experiments. Deep-Sea Res. I 49,2009 -2033.
Schlichting, H. (1979). Boundary-Layer Theory. New York: McGraw-Hill.
Staples, R. F. (1966). The distribution and characteristics of surface bioluminescence in the oceans. Naval Oceanographic Office Technical Report No. 184.
Stokes, M. D., Deane, G. B., Rohr, J. and Latz, M. I. (2004). Bioluminescence imaging of wave-induced turbulence. J. Geophys. Res. 109,C01004 .
Sweeney, B. M. and Hastings, J. W. (1958). Rhythmic cell division in populations of Gonyaulax polyedra. J. Protozool. 5,217 -224.
Swift, E. Meunier, V. A., Biggley, W. H., Hoarau, J. and Barras, H. (1981). Factors affecting bioluminescent capacity in oceanic dinoflagellates. In Bioluminescence Current Perspectives (ed. K. H. Nealson), pp.95 -106. Minneapolis, MN: Burgess Publishing Co.
Tett, P. B. and Kelley, M. G. (1973). Marine bioluminescence. Oceanogr. Mar. Biol. A. Rev. 11, 89-173.
Thomas, W. H. and Gibson, C. H. (1990). Quantified small-scale turbulence inhibits a red tide dinoflagellate, Gonyaulax polyedra Stein. Deep-Sea Res. 37,1583 -1593.
Thomas, W. H. and Gibson, C. H. (1992). Effects of quantified small-scale turbulence on the dinoflagellate Gymnodinium sanguineum (splendens): contrasts with Gonyaulax (Lingulodinium) polyedra, and the fishery implication. Deep-Sea Res. 39,1429 -1437.
Titelman, J. (2001). Swimming and escape behavior of copepod nauplii: implications for predator-prey interactions among copepods. Mar. Ecol. Prog. Ser. 213,203 -213.
van Duuren, F. A. (1968). Defined velocity gradient model flocculator. J. Sanitary Eng. Div. 94,671 -682.
von Dassow, P. and Latz, M. I. (2002). The role
of Ca2+ in stimulated bioluminescence of the dinoflagellate
Lingulodinium polyedrum. J. Exp. Biol.
205,2971
-2986.
White, H. H. (1979). Effects of dinoflagellate bioluminescence on the ingestion rates of herbivorous zooplankton. J. Exp. Mar. Biol. Ecol. 36,217 -224.[CrossRef]
Widder, E. A. and Case, J. F. (1981). Bioluminescence excitation in a dinoflagellate. In Bioluminescence Current Perspectives (ed. K. H. Nealson), pp.125 -132. Minneapolis, MN: Burgess Publishing Co.
Yen, J. and Fields, D. M. (1992). Escape responses of Acartia hudsonica (Copepoda) nauplii from the flow field of Temora longicornis (Copepoda). Ergebnisse Limnol. 36,123 -134.
Yoganathan, A. P., Ellis, J. T., Healy, T. M. and Chatzimavroudis, G. P. (1998). Fluid dynamic studies for the year 2000. J. Heart Valve Disease 7, 130-139.
Zirbel, M. J., Veron, F. and Latz, M. I. (2000). The reversible effect of flow on the morphology of Ceratocorys horrida (Peridiniales, Dinophyta). J. Phycol. 36,46 -58.