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Physiological and ecological consequences of environmental temperature on Antarctic protists

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Physiological and ecological consequences of environmental temperature on Antarctic protists

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Content PHYSIOLOGICAL AND ECOLOGICAL CONSEQUENCES OF ENVIRONMENTAL
TEMPERATURE ON ANTARCTIC PROTISTS

by

Julie Marie Rose

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MARINE ENVIRONMENTAL BIOLOGY)

May 2007

Copyright 2007 Julie Marie Rose
Dedication

For my father
ii
Acknowledgements
I would like to thank my family and friends for all of their help and support
throughout the past six years. My family has provided an amazing support network for
me, even from thousands of miles away. In particular, I want to thank my father, Michael
Rose, for all of his time, patience and unwavering support of my life decisions. My dad
has always been my biggest cheerleader and it has been a wonderful comfort to know that
he has always stood behind me. I doubt very much that without his help and love I would
have ever finished my degree. I want to thank my advisor, David Caron. Without his
help, guidance and support this dissertation would not have been possible. Thanks to my
committee for all of their advice and support. I would also like to thank my co-workers
past and present at the University of Southern California. Becky Schaffner and Pete
Countway have been my surrogate big sister and big brother ever since I was a silly kid
fresh out of undergrad and without their help and guidance my life in California would
have been much more difficult and much less fun. Beth Stauffer, Steffi Moorthi, Ilana
Gilg, Adriane Jones, Astrid Schnetzer, Tim McLean, Matt Travao, Pratik Savai and
Patrick Vigil have all been wonderful co-workers that I have also been lucky enough to
count as friends. While I didn’t get to see Rebecca Gast, Bob Sanders, Dawn Moran and
Mark Dennett as frequently, their contribution to my life and work as collaborators and
friends has been just as important. I especially want to thank Becky Gast for being not
just a role model to me as a successful, bright, sane female scientist but also a wonderful
friend and confidante. Thanks to the two sharp undergraduates, Neil Vora and Vanessa
Sippel, who worked with me as reu students and helped with experiments for two
iii
summers. Finally thanks to the captain and crew of the RVIB Nathaniel B. Palmer and
the other scientists in the MIXURS, GRINCHES, REMIXURS and CORSACS cruises.
iv

Table of Contents

Dedication ii

Acknowledgments iii

List of Tables vii

List of Figures viii

Abstract xiii

Dissertation Introduction 1

Chapter 1: Counting heterotrophic nanoplanktonic protists in cultures
and aquatic communities by flow cytometry 5
Introduction 5
Methods 8
Results 17
Discussion 32
Conclusions 46

Chapter 2: Does low temperature constrain the growth rates of
heterotrophic protists? Evidence and implications for algal blooms
in cold waters 47
Introduction 47
Methods 52
Results and Discussion 53

Chapter 3: Effects of temperature and prey type on growth rate and
gross growth efficiency of an Antarctic bacterivorous protist 71
Introduction 71
Methods 75
Results 80
Discussion 86
Conclusions 92

Chapter 4: Effect of temperature and prey type on nutrient
regeneration by an Antarctic bacterivorous protist 93
Introduction 93
Methods 96
v
Results 99
Discussion 109
Conclusions 114

Chapter 5: Low temperature constraints on grazing by
Antarctic microzooplankton in culture and in natural
assemblages 115
Introduction 115
Methods 118
Results 126
Discussion 137
Conclusions 146

Dissertation Conclusions 147

Bibliography 155

Appendices 167
vi
List of Tables

Table 1-1: Uptake of LysoTracker Green
®
by two phototrophic
eukaryotes and one phototrophic prokaryote. Samples were
preserved with 0.5% paraformaldehyde in the “Dead” treatment. 27

Table 4-1: Changes in concentration of particulate carbon and
particulate and dissolved nitrogen and phosphorus before and after
bacterial grazing by Paraphysomonas imperforata. 104

Table 4-2: Comparison of percent of initial particulate nutrients
regenerated as ammonium or soluble phosphorus at the time of
peak protistan abundance for strains of bacterivorous
Paraphysomonas imperforata examined at different temperatures. 108

Table 5-1: Chlorophyll concentration, phytoplankton abundance,
abundance of total microzooplankton and subgroups of
microzooplankton in late austral summer at four stations in the
Ross Sea, Antarctica. 127

Table 5-2: Ingestion rates and specific ingestion rates of Antarctic
ciliates in culture and in natural plankton assemblages. 133

vii
List of Figures

Figure 1-1. Effect of incubation time (A,C) and stain concentration (B,D)
on cytometric counts of the ciliate Uronema sp. (A,B) and the heterotrophic
flagellate Bodo caudatus (C,D). Large dashed lines (A-D) indicate
microscopical count at time zero and circles indicate cytometric counts.
Small dashed lines (A,C) represent an exponential curve fit to the protistan
counts based on cytometry. 18

Figure 1-2. Effect of LysoTracker Green on the green fluorescence of three
heterotrophic protists and one heterotrophic bacterium in unstained cultures
(A,D,G,J), in cultures preserved with 1% glutaraldehyde and then stained
(B,E,H,K) or in cultures stained live (C,F,I,L). Species examined were
Paraphysomonas bandaiensis (A-C), Pteridomonas sp. (D-F), Uronema sp.
(G-I) and Halomonas halodurans (J-L). 20

Figure 1-3. An example showing detection and discrimination of a
heterotrophic protistan species in a bacterized culture. (A-D) Cytograms
demonstrate Pteridomonas sp. population growth over time. All samples
were stained live with LysoTracker Green before cytometric analysis.
Population growth is shown by steadily increasing events within the
protistan polygon at 24 hours (A), 48 hours (B), 72 hours (C) and 96 hours
(D). 22

Figure 1-4. Comparison of microscopical counts and cytometric counts of
five actively growing heterotrophic protistan populations over a range of
abundances. Solid line indicates regression of data. Dashed line indicates
1:1 ratio. ( ) Uronema sp., ( S ),Paraphysomonas butcheri, ( ) Bodo
caudatus, ( z ) Pteridomonas sp., and ( T ) Paraphysomonas bandaiensis. 23

Figure 1-5. (A-C) Comparison of growth curves of three heterotrophic
protists enumerated by cytometric counts (closed symbols) and
microscopical counts (open symbols). Growth rates were calculated from
the slope of the line of population growth during the exponential growth
phase. Solid lines correspond to growth rates calculated from cytometric
counts. Dashed lines correspond to growth rates calculated from
microscopical counts. (D) Comparison of microscopical counts and
cytometric counts over all phases of population growth from (A-C). ,
Paraphysomonas vestita, Ì , Cafeteria roenbergensis and { ,
Paraphysomonas imperforata. 25

viii
Figure 1-6. Effect of ingestion of algal prey on the fluorescent signal of a
heterotrophic protist. A small unidentified chrysophyte was used as prey
for a heterotrophic protist (Uronema sp.) Cytograms show red fluorescence
(FL3) vs. forward scatter for the chrysophyte (A), late exponential cultures
of Uronema sp. (B) and cultures of Uronema sp. thirty minutes after the
addition of the chrysophyte (C). Regions were drawn around the algal
population in (A) to include at least 97% of the events. 28

Figure 1-7. Cytometric analysis of a natural water sample taken from
Boothbay Harbor, ME, for the enumeration of heterotrophic nanoplanktonic
protists. (A) Phototrophs were detected based on high relative chlorophyll
fluorescence in cytograms of red fluorescence (FL3) vs. forward scatter
(FSC) (red events gated in A). (B) Detrital particles were identified based
on relatively high side scatter and relatively low LysoTracker Green
fluorescence in cytograms of green fluorescence (FL1) vs. side scatter
(SSC) (yellow events gated in B). (C) 2.5 μm beads were used for size
estimation. They were identified based on their high green fluorescence
and relatively low red fluorescence in cytograms of red fluorescence (FL3)
vs. green fluorescence (FL1). (D) Populations identified in (A-C) were
removed from the plot of LysoTracker Green fluorescence vs. forward
scatter, and remaining events larger than 2.5 μm were counted as
heterotrophic nanoplanktonic protists (blue events gated in D). 30

Figure 1-8. Comparison of microscopical and cytometric counts for 26
natural water samples. Solid line indicates regression of data. Dashed line
indicates 1:1 ratio. Open circles indicate samples taken from highly
eutrophic locations (Newport Beach Harbor and Los Angeles Harbor). 31

Figure 2-1. Summary of the relationship between published growth rates
of phototrophic protists and temperature. Solid line represents the Eppley
(1972) curve of maximal predicted growth rates of phototrophic protists.
Dashed line represents the curve from Brush et al. (2002). 54

Figure 2-2. Summary of the relationship between growth rates and
temperature for bacterivores (A), and herbivores (B). Solid line represents
the Eppley curve for phototrophic protists from Figure 1. 57

Figure 2-3. Effect of temperature on the growth rate of bacterivorous (A)
and herbivorous (B) Paraphysomonas spp. Solid line represents the
Eppley curve. 60

Figure 2-4. Growth rates of herbivorous and bacterivorous protists as a
function of cell volume. 62

ix
Figure 2-5. Growth rates of phototrophic protists (A), bacterivorous protists
(B), herbivorous protists (C) and copepods (D) as a function of temperature.
Solid line in (A) represents the Eppley curve, and dashed line represents
the Brush curve. Solid line in (B-D) represents predicted maximal growth
rates at any temperature. Solid larger symbols indicate the maximal values
within each data set used to calculate the regression equations, open
smaller symbols indicate values not used in calculation of the regression
equation. 64

Figure 2-6. Comparison of maximal growth rate predictor lines for
phototrophic protists, herbivorous protists and copepods generated in
Figure 2-5. The thick solid line represents the Eppley curve, the thin solid
line represents the Brush curve, the dashed line represents maximal
growth rates of herbivorous protists and the dotted line represents maximal
growth rates of copepods. 66

Figure 3-1. Transmission electron micrograph of Paraphysomonas
imperforata (negative stain using 1% uranyl acetate). A) Whole cell view
showing flagellation and surface scales, 5,000x. Marker bar = 2 μm B)
High magnification showing scale morphology, 10,000x . Marker bar =
0.5 μm. 76

Figure 3-2. Growth dynamics of an Antarctic strain of P. imperforata
(filled symbols) and removal of bacterial prey (open symbols) at three
temperatures and on three strains of bacteria (A-E): Antarctic bacterial
strain A as food at 10˚C (A), 5˚C (B) and 0˚C (C); Antarctic bacterial
strain B at 0˚C (D); H. halodurans at 0˚C (E). Composite of information
from (A)-(C) comparing the growth of P. imperforata at 10
o
C ( ), 5
o
C
( S ), and 0
o
C ( z ) (F). 81

Figure 3-3. Gross growth efficiencies of the Antarctic strain of
P. imperforata based on measurements of cell volume (solid) and carbon
(hatched). Protists were grown at three temperatures and on three strains
of bacteria, as indicated. Error bars represent one standard deviation. 82

Figure 3-4. Relationship between temperature and growth rate (A) or gross
growth efficiency (B) for an Antarctic strain of P. imperforata. Error bars
represent one standard deviation. 84

Figure 3-5. Relationship between temperature and growth rate (A) or gross
growth efficiency (B) for strains of Paraphysomonas spp. observed in this
study ( z ), and in published reports for temperate and polar strains ( Ì ).
Solid lines in (B) indicate mean values and dashed lines indicate median
values for each 5˚C temperature interval. 85
x

Figure 4-1. Changes in protistan abundance (A, B) and particulate carbon
concentrations (C, D) in cultures of Paraphysomonas imperforata grown
at three temperatures (A, C) and on three different strains of bacteria at a
single temperature (B, D). Symbols indicate particulate carbon
concentration (C, D) and lines indicate protistan abundance (A, B).
(A, C) 0˚C, Antarctic bacterial strain A ( , solid line); 5˚C, Antarctic
bacterial strain A ( z , dashed line); 10˚C, Antarctic bacterial strain A ( S ,
dotted line). (B, D) 0˚C, Antarctic bacterial strain A ( , solid line); 0˚C,
Antarctic bacterial strain B ( , dashed line); 0˚C, Halomonas halodurans
( T , dotted line). 100

Figure 4-2. Changes in particulate nitrogen (solid symbols) and ammonium
(open symbols) during bacterial grazing by Parapyhysomonas imperforata
on Antarctic bacterial strain A at 0˚C (a), 5˚C (b) and 10˚C (c), and at 0˚C
on Antarctic bacterial strain B (d) and Halomonas halodurans (e).
Changes in ammonium concentration in all treatments (f). Note different
ranges for x and y axes in (a-f). 101

Figure 4-3. Changes in particulate phosphorus (solid symbols) and
dissolved phosphorus (open symbols), including dissolved organic
phosphorus ( Ì ), soluble reactive phosphorus ( { ) and total dissolved
phosphorus ( ) (a-e). Grazing by Paraphysomonas imperforata on
Antarctic bacterial strain A at 0˚C (a), 5˚C (b) and 10˚C (c), and at 0˚C on
Antarctic bacterial strain B (d) and Halomonas halodurans (e). Changes in
phosphorus concentration in all treatments (f). Note different ranges for x
and y axes in (a-f). 102

Figure 4-4. Regeneration of ammonium normalized to the natural
logarithm abundance of Paraphysomonas imperforata in the different
treatments. 0˚C, Antarctic bacterial strain A ( ); 5˚C, Antarctic bacterial
strain A ( { ); 10˚C, Antarctic bacterial strain A ( Ì ); 0˚C, Antarctic
bacterial strain B ( ◊); 0˚C, Halomonas halodurans ( ∇). 105

Figure 4-5. Changes in particulate and dissolved fractions of initial total
nitrogen (a) and phosphorus (b). 0˚C, Antarctic bacterial strain A (solid
bars); 5˚C, Antarctic bacterial strain A (horizontal stripes); 10˚C, Antarctic
bacterial strain A (diagonal stripes); 0˚C, Antarctic bacterial strain B
(dotted bars); 0˚C, Halomonas halodurans (white bars). Bars correspond
to particulate or dissolved nutrient concentrations as a percentage of the
initial total nutrient concentration (particulate plus dissolved). 107

xi
Figure 5-1. Ingestion rates with different abundances of fluorescently
labeled algae. Average number of ingested Thalassiosira cells
microzooplankton
-1
at four stations in late austral summer in the Ross Sea,
Antarctica. Station 1 (A), Station 2 (B), Station 3 (C) and Station 4 (D). 129

Figure 5-2. Ingestion rates with different abundances of fluorescently
labeled algae. Average number of ingested Thalassiosira cells per
Eutintinnus (A, B) or per Codonellopsis (C, D) at Station 2 (A, C) and
Station 4 (B, D). 131

Figure 5-3. Ingestion rates of the cultured Antarctic Strombidium sp.
feeding on the Antarctic alga Thalassiosira sp. 1 after starvation ( ), after
2 days of feeding on a dense culture of Thalassiosira sp. 1 ( z ) and after 4
days of feeding ( S). 134

Figure 5-4. Comparison of specific ingestion rates measured in this study
to values for temperate ciliates reported in Hansen et al (1997). Cultures
of Antarctic Strombidium sp. ( z ), field populations of Eutintinnus sp ( Ì ),
field populations of Codonellopsis sp. ( ◊) and literature values for cultures
of temperate ciliates ( ). 136

Figure 5-5. Sample growth rate of the Antarctic Strombidium sp. feeding
on Pyramimonas sp. (A). A comparison of growth rates of the Antarctic
Strombidium sp. feeding on six species of Antarctic algae (B):
Pyramimonas sp. (a), Thalassiosira sp. 1 (b), Thalassiosira sp. 2 (c),
Mallomonas sp. (d), Chlamydomonas sp. (e), Polarella glacialis (f). 138

xii
Abstract
The Ross Sea, Antarctica experiences one of the world’s largest annual
phytoplankton blooms at exceptionally low environmental temperature (-1.5-0.5˚C).
Chlorophyll concentrations during the bloom can exceed 15 μg l
-1
. Heterotrophic protists
seasonally dominate biomass within Antarctic marine ecosystems, and function as
important players in nutrient remineralization and carbon flow. However, these potential
algal grazers do not prevent the formation of phytoplankton blooms in this ecosystem.
Low temperature represents a constant potential limit on maximal growth of all Antarctic
species, yet the effect of temperature on growth of heterotrophic protists is not well
characterized. Existing data for growth rates and growth efficiencies of polar protists are
conflicting. I investigated top-down control of algal and bacterial standing stocks at
ambient Ross Sea temperatures through a literature review of temperature effects on
protistan growth rate, laboratory experiments with cultured Antarctic protists and field
estimates of grazing rates by assemblages of Antarctic herbivorous protists. Growth rates
of phototrophs and heterotrophs at ambient Antarctic temperatures were universally low,
but growth rates of heterotrophs appeared to be more strongly affected at low temperature
than phototrophs. Maximum growth rates of herbivores were equivalent to algae at 20
o
C,
but a quarter of maximum algal growth rates at 5
o
C. Growth and nutrient
remineralization rates of Antarctic bacterivorous protists were low at 0
o
C and increased
sharply with increasing temperature. Growth efficiency and nutrient remineralization
efficiency by Antarctic bacterivorous protists were comparable to temperate conspecifics
at all temperatures examined. Specific ingestion rates of Antarctic herbivorous protists in
xiii
culture and in field assemblages were extremely low when compared with temperate
congeners. The estimated grazing impact of assemblages of Antarctic microzooplankton
on phytoplankton standing stocks was low at four stations within the Ross Sea, even
though microzooplankton abundance was high. These results suggest that strong
constraints by temperature on top-down control (protozooplankton growth and grazing)
in the Ross Sea may contribute to bloom formation.
xiv
Dissertation Introduction
Heterotrophic protists are important players in aquatic ecosystems. These
organisms comprise a significant portion of the total living biomass (Stoecker et al. 1994,
Caron et al. 1995). They serve as an important source of mortality for bacteria, algae and
other heterotrophic protists (Sanders et al. 1992, Sherr and Sherr 1994). Heterotrophic
protists also function as important recyclers of carbon, phosphorus and nitrogen through
their grazing activities, and as such play a major role in nutrient flow in aquatic
environments (Strom 2000, Sherr and Sherr 2002).
Heterotrophic protists are also important in Antarctic marine ecosystems,
although less is known about their functional roles in this environment. The abundance
and distribution of heterotrophic protists in Antarctic coastal waters has been thoroughly
documented (Garrison and Gowing 1993 and references cited therein). Heterotrophic
nanoflagellates have been shown to dominate the total microbial biomass (excluding
bacteria) in the coastal waters of the Ross Sea in austral autumn and may thus exhibit
significant top-down control of bacterial assemblages during this season (Dennett et al.
2001). Heterotrophic microzooplankton have also been reported at high abundance in
Antarctic coastal ecosystems (Garrison et al. 1986, Garrison 1991, Gowing and Garrison
1992, Stoecker et al. 1993, Caron et al. 2000). Ciliates and dinoflagellates are especially
abundant in mid- to late summer (Caron et al. 2000). Abundances have been reported on
the order of 10
4
cells l
-1
for ciliates and dinoflagellates in the Ross Sea (Caron et al.
2000), 10
3
heterotrophic dinoflagellates l
-1
in the Weddell Sea (Garrison and Buck 1989),
1
10
4
ciliates l
-1
in the Weddell Sea (Buck and Garrison 1983) and 10
5
ciliates l
-1
in Prydz
Bay (Davidson and Marchant 1992).
The high abundance of heterotrophic microzooplankton in Antarctic marine
ecosystems has led some to speculate that Antarctic microzooplankton exert strong
grazing pressure on primary production by Antarctic phytoplankton (Klaas 1997).
However, the coastal waters surrounding the continent of Antarctica are the location of
annual algal blooms that are both massive and spatially extensive (Smith and Nelson
1985, Smith et al. 2000, Smith et al. 2003). The presence of these annual blooms
suggests that grazing pressure by heterotrophs in early to mid-spring during bloom
formation is not strong enough to control the growth of phototrophs in this ecosystem.
Estimates of physiological rates (including growth rates, ingestion rates and grazing
rates) of Antarctic heterotrophic microzooplankton are few, but published values are all
low. Burkill et al. (Burkill et al. 1995) measured low rates of herbivory in the
Bellingshausen Sea, Antarctica. Caron et al. (2000) measured rates of herbivory during
four cruises spanning three seasons in the polynya of the Ross Sea, Antarctica. Only 13
of 51 experiments yielded detectable rates of herbivory using the dilution method, and the
highest rate observed was low in comparison to rates reported for temperate and tropical
ecosystems. The low grazing rates in that study were not a consequence of low grazer
abundance. Microzooplankton grazers (ciliates and heterotrophic dinoflagellates) varied
considerably in abundance over the course of the experiments and were at times
comparable in abundance to those observed in locations such as the North Atlantic during
the spring bloom, as well as the Arabian Sea and Equatorial Pacific. The authors also
2
compiled a literature review of grazing rates from 19 published studies measured over a
wide range of temperatures and from a variety of marine systems and found significantly
lower grazing rates at temperatures below 2˚C compared to rates measured at 10˚C or
above. Similarly, two studies that examined ingestion rates of microzooplankton on
fluorescently labeled phytoplankton in the Atlantic and Indian sectors of the Southern
Ocean reported low uptake rates at ambient temperature (Becquevort 1997, Becquevort et
al. 2000). Maximal ingestion rates in those studies were ≤2 P. antarctica grazer
-1
h
-1
, and
≤0.01 P. antarctica grazer
-1
h
-1
.
We believe it is possible that low temperature acts as a strong constraint on the
growth of phototrophic and heterotrophic protists, as well as on the grazing activities of
heterotrophic protists in Antarctic marine ecosystems. We have combined a meta-
analysis of literature data with experiments in both field and laboratory settings to
investigate the extent to which low temperature acts as a constraining force on different
parts of the Antarctic microbial food web. This approach can be described as follows:

1. A literature review of growth rates of phototrophic protists, bacterivorous protists
and herbivorous protists in culture. This review sought to re-examine classic
reviews by Eppley (1972) and Goldman and Carpenter (1974) on the effect of
temperature on growth rates of phytoplankton in culture to see if the relationships
they derived were applicable to protists from polar and temperate regions growing
at extreme low temperature. This review also sought to describe the effect of
3
temperature on maximal growth rates of heterotrophic protists in a similar fashion
to Eppley’s relationship based on phytoplankton maximal growth rates.

2. A series of laboratory experiments examining the effects of temperature and prey
type on growth rate, nutrient regeneration rate, gross growth efficiency and
nutrient regeneration efficiency of an Antarctic strain of Paraphysomonas
imperforata. The experiments were designed to establish baseline information
about the physiological ecology of Antarctic bacterivorous protists. The
experiments were also designed to examine whether the relationship with
temperature for heterotrophic protists in general derived in the literature review
could be applied at the genus level for temperate and polar species within the
genus Paraphysomonas.

3. A combination of laboratory and field experiments measuring ingestion rates of
Antarctic herbivorous microzooplankton. These experiments were designed to
test the rate at which Antarctic microzooplankton in the Ross Sea during late
austral summer ingested phytoplankton, and estimate the daily grazing impact of
microzooplankton assemblages on phytoplankton standing stocks. These
experiments were also intended to compare specific ingestion rates of Antarctic
ciliates in the field and in culture with specific ingestion rates published for
temperate ciliates measured over a range of temperatures.

4
Chapter 1: Counting heterotrophic nanoplanktonic protists in cultures
and aquatic communities by flow cytometry
Introduction
Nanoplanktonic (2-20 μm) heterotrophic protists play integral roles within aquatic
environments. These organisms comprise a significant portion of the total living biomass
of planktonic ecosystems (Stoecker et al. 1994, Caron et al. 1995) and serve as an
important source of mortality for bacteria, microalgae and other heterotrophic protists
(Sanders et al. 1992, Sherr & Sherr 1994). As consumers, heterotrophic nanoplankton
function in the transfer of carbon and energy to higher trophic levels and as important
recyclers of organic matter and macronutrients (Azam et al. 1983, Caron & Goldman
1990). Therefore, the accurate enumeration of heterotrophic nanoplankton is essential for
understanding trophic dynamics and energy flow in aquatic ecosystems.
Traditionally, heterotrophic nanoplanktonic protists have been quantified using
microscopical techniques. Typically, assemblages in natural water samples are preserved
and stained with nucleic acid or protein stains, filtered and mounted onto slides, and
counted using epifluorescence microscopy (Sherr et al. 1993). Protists are first identified
based on the strong fluorescent signal of the stain, then heterotrophs and phototrophs
within the sample are distinguished and quantified based on the absence or presence of
autofluorescence due to photosynthetic pigments (Sherr et al. 1993). This process of
counting and classifying individual cells by epifluorescence microscopy is slow, tedious,
and subject to a number of potential sources of error. These sources of error can include
fixation artifacts, over- or understaining, incorrectly categorizing heterotrophs cells with
5
autofluorescent prey as phototrophs, and human error. Nanoplanktonic protists have
occasionally been counted live (Dale & Burkill 1982), but it is difficult to differentiate
small phototrophs from heterotrophs using this method. To address these issues, I sought
to develop a faster and more precise method of quantification, based on flow cytometry.
Flow cytometry has been employed in aquatic microbiology to quantify
autotrophic prokaryotes, small autotrophic eukaryotes, heterotrophic bacteria and viruses
(Olson et al. 1983, Yentsch et al. 1983, del Giorgio et al. 1996, Marie et al. 1997, Marie
et al. 1999). Prokaryotic and eukaryotic phototrophs can be detected and quantified
based on the autofluorescence of their photosynthetic pigments. Also, heterotrophic
bacteria and viruses can be detected and quantified using a variety of nucleic acid stains
to distinguish them from detrital particles. The speed and accuracy with which these
populations can be counted by this method has made the flow cytometer a valuable tool
for ecological studies in aquatic sciences (Olson et al. 1991, Porter 1999, Campbell
2001).
Unfortunately, an effective, accurate technique for the enumeration of
heterotrophic unicellular eukaryotes in natural water samples using flow cytometry has
not yet emerged. Most heterotrophic eukaryotes have little or no autofluorescence,
except some heterotrophic dinoflagellates which fluoresce apple-green with blue light
excitation (Carpenter et al. 1991). Therefore, detection of most heterotrophic
nanoplanktonic protists is not possible by cytometry without staining. Common
fluorescent compounds used to stain prokaryotes and eukaryotes do not differentiate
these populations on a flow cytometer. For example, one technique published recently
6
by Rifa et al (2002) used the nucleic acid stain SYTO-13
®
to quantify heterotrophic
eukaryotes in cultures and in field samples. However, both the prokaryotic and
eukaryotic assemblages were stained, and eukaryotes were indistinguishable from
prokaryotes on cytograms when the abundance of bacteria greatly exceeded the
abundance of protists. This drawback makes SYTO-13
®
problematic for use in growth
experiments, because starting bacterial concentrations are often several orders of
magnitude greater than the protistan populations. This complication may also preclude
the use of SYTO-13
®
for many natural samples where the ratio of bacteria to
heterotrophic protists may exceed 1000 (Sanders et al. 1992).
A new cytometric method was developed that employs a fluorogenic compound
that selectively stains eukaryotic cells that possess acidic vacuoles in order to avoid
problems distinguishing prokaryotes from eukaryotes by cytometry. The method
employs a pH-specific compound, LysoTracker Green
®
, (Molecular Probes, Eugene,
Oregon, USA) to allow the discrimination of eukaryotes from prokaryotes by flow
cytometry. The stain concentrates within acidic organelles of eukaryotes such as
intracellular food vacuoles, lysosomes and chloroplasts. Prokaryotes do not possess
acidic organelles and thus display low fluorescence. Phototrophic eukaryotes are
distinguished from heterotrophic eukaryotes by the autofluorescence of photosynthetic
pigments. This method was developed using cultures of heterotrophic protists and a
standard benchtop flow cytometer in order to demonstrate the efficacy of LysoTracker
Green
®
for enumerating these species. The stain was then applied to natural samples of
plankton.
7
Methods
Cultures. The LysoTracker Green
®
compound was tested on one strain of cyanobacteria
and eleven species of protists. The cyanobacterium Synechococcus sp. (CCMP 837), the
prasinophyte Micromonas pusilla (CCMP 494) and the phototrophic dinoflagellate
Heterocapsa triquetra (CCMP 448) were obtained from the Provasoli-Guillard National
Center for the Culture of Marine Phytoplankton, Boothbay Harbor, ME.
Paraphysomonas vestita was obtained from Dr. Robert W. Sanders, Temple University,
Philadelphia, PA. Paraphysomonas butcheri and Bodo caudatus were obtained from Dr.
Delma Bratvold, College of Charleston, Charleston SC. Paraphysomonas bandaiensis
was obtained from Dr. John Waterbury, Woods Hole Oceanographic Institution, Woods
Hole, MA. Paraphysomonas imperforata, Cafeteria roenbergensis, Pteridomonas sp., an
unidentified chrysophyte and a scuticociliate (Uronema sp.) were isolated from a variety
of aquatic environments and cultured in the laboratory of D.A. Caron. All protists, other
than P. vestita were marine. All heterotrophic protists were cultured on bacteria
endogenous in the cultures.

Natural samples. Eight coastal locations from Connecticut to Maine, USA were sampled
over the course of three days. 150 ml samples were collected in 75 cm
2
tissue culture
flasks from a single location within Milford, CT; Noank, CT; East Greenwich, RI;
Cohasset, MA; Portsmouth, NH; Portland, ME and two locations within Boothbay
Harbor, ME. Eighteen sites along the California coast, USA, between Carlsbad and
8
Malibu were also sampled. One liter samples were collected in polycarbonate flasks
from Agua Hedionda and Carlsbad State Beach, Carlsbad; Buena Vista Lagoon,
Oceanside; Dana Cove Park, Dana Point; Newport Beach; Newport Beach Harbor;
Huntington City Beach; Huntington Beach Harbor; Seal Beach; Bluff Park, Long Beach;
Los Angeles Harbor; Cabrillo Beach, Los Angeles; Redondo County Beach, Redondo;
Dockweiler State Beach, El Segundo; Santa Monica State Beach, Santa Monica; Leo
Carrillo Beach and Topanga County Beach, Malibu. Samples were kept in the dark, on
ice, prior to analysis, then allowed to come to room temperature before they were
analyzed on the flow cytometer and preserved for epifluorescence microscopy.

Microscopical counts. Transmitted light microscopy was used to count protists in
culture. Five ml samples were removed from experimental flasks and preserved in 0.1 ml
acid Lugol’s solution (2% final concentration). Subsamples were removed and 0.1 ml
aliquots were counted in a Palmer Maloney counting chamber. At least 200 heterotrophic
protists were counted per slide.
Heterotrophic protists in natural samples were counted using epifluorescence
microscopy (Sherr et al. 1993). A subsample was preserved for microscopical analysis at
the same time as the live sample was analyzed on the flow cytometer. Samples were
preserved with 0.3% glutaraldehyde and stored in the dark overnight at 5
o
C. Samples of
15-45 ml were placed in a glass filtering tower, stained for five minutes with 5-25 μg⋅ml
-1

DAPI (final stain concentration) and filtered onto 25 mm, 0.8 µm black polycarbonate
filters. Filters were immediately placed on slides, mounted with a drop of immersion oil,
9
covered with a coverslip, sealed with wax, and stored in the dark at –20
o
C until they
were counted. Duplicate slides were prepared for each sample. Slides were counted by
comparing two digital images for each field of view examined; one obtained using blue
light excitation and the other using UV light excitation. Protists were distinguished based
on the signal of DAPI with UV light excitation, then classified as either phototrophic or
heterotrophic based on the presence or absence of photosynthetic pigments using blue
light excitation. Multiple fields of view were examined until at least 100 heterotrophic
protists were counted per slide.

Staining protists with LysoTracker Green
®
. The LysoTracker Green
®
stain
(Molecular Probes, Eugene, Oregon, USA) consists of a fluorochrome attached to a weak
base, which causes the compound to accumulate in acidic organelles. The stain has a
peak excitation at 504 nm, but can be excited by the 488 nm argon-ion laser found in
some commercially available benchtop flow cytometers. Peak emission occurs at 511
nm, so the photomultiplier tube of the FL1 channel of the FACScan and FACScalibur
flow cytometers (Becton Dickinson, San Jose, California, USA) collects the strongest
signal, (515 to 545 nm, green fluorescence). There is no significant detectable emission
at wavelengths detected by the FL3 or FL4 channels (>653 nm).

Optimizing flow cytometer settings and staining procedures with cultured protists.
A FACScalibur flow cytometer (Becton Dickinson, San Jose, California, USA), equipped
with a 15 mW air-cooled 488 nm argon-ion laser was used to count heterotrophic protists.
10
The salinity of the sheath fluid (the fluid surrounding the sample stream within a flow
cytometer,) was matched to that of the sample to avoid distortion of the forward scatter
signal (Cucci & Sieracki 2001). Organisms stained with LysoTracker Green
®
were
recorded by the FL1 (green) fluorescence detector. Uniprotistan cultures of heterotrophic
protists in bacterized cultures were distinguished by their high fluorescence and high
forward scatter on a plot of green fluorescence (FL1) vs. relative size (forward scatter, or
FSC). Polygon gates were drawn around populations of protists, and events falling
within these polygons were counted using the software program CellQuest (Becton
Dickinson, San Jose, California, USA).
The number of events on the flow cytometer was converted to cell abundance
(protists ⋅ml
-1
) using one of two methods. A known concentration of green fluorescent 2.5
µm polystyrene beads (Molecular Probes, Eugene, Oregon, USA) was added to samples
of cultured protists and served as an internal standard for enumeration of the protistan
population. The beads were diluted 1:1000 to create a working stock that was
enumerated at the beginning of each experiment using a hemacytometer. Samples were
acquired on the HI flow rate setting (approximately 50 μl ⋅min
-1
) until at least 200 beads
and 200 protists were recorded. Alternatively, the sample tube was weighed on an
analytical balance before and after each analysis on the flow cytometer to determine the
volume examined over the course of the analysis. The total event count was converted to
cell abundance (protists ⋅ml
-1
) after each time point using the processed sample volume.
The first method was employed during growth experiments with cultured heterotrophs,
while the second method was employed during the experiments optimizing stain
11
conditions as well as in natural water samples. Both methods yielded comparable results,
but the second method required much less effort and therefore less time to prepare.
Stain concentration and appropriate staining time were determined for the
LysoTracker Green
®
stain based on experiments with a small heterotrophic flagellate
(Bodo caudatus; 4-6 µm) and a medium-sized ciliate (Uronema sp.; 10-20 µm) because
they represented the approximate range of sizes of heterotrophic protists in natural
samples that might be expected to be encountered and effectively counted by flow
cytometry. Eleven different incubation times and twelve different stain concentrations
were examined to determine appropriate stain conditions for both species.
A 1 mM stock of LysoTracker Green
®
was diluted 1:10 with 0.2 μm filtered
sterile seawater, and this working stock was added to two live cultures of heterotrophic
protists at a final stain concentration of 75 nM. This stain concentration was chosen
based on a recommendation from Molecular Probes (Molecular Probes, Eugene OR,
Technical Bulletin #07525). Subsamples were analyzed on the flow cytometer
immediately for 30 seconds on the HI setting, then again after 3, 5, 7, 10, 15, 20, 30, 45,
60, and 90 minutes. The samples were kept at room temperature between cytometric
analyses and kept in the dark to prevent the stain from fading. Samples of each culture
were also removed immediately prior to staining with LysoTracker Green
®
and preserved
with acid Lugol’s solution (2% final concentration) for direct determinations of cell
abundances by light microscopy.
Stain concentration was examined for its efficacy for staining heterotrophic
protists. A serial dilution of a 1 mM stock of LysoTracker Green
®
was used to prepare
12
final stain concentrations of 4, 7, 10, 16, 25, 38, 58, 89, 137, 211, 325 and 500 nM.
Cultures were stained for ten minutes, based on the results from the incubation time
series, then analyzed on the flow cytometer. Cell abundances were determined as
described above.
Three species of heterotrophic protists and one species of heterotrophic bacterium
were examined for the effect of preservative on the fluorescence of stained cultures:
Uronema sp., Pteridomonas sp. (~5 μm), Paraphysomonas bandaiensis (~5 μm) and
Halomonas halodurans. Three treatments were compared: 1) unpreserved and unstained,
2) unpreserved and stained with LysoTracker Green
®
, and 3) preserved and stained with
LysoTracker Green
®
. Cultures were grown to mid-logarithmic growth phase before
analysis. Unstained samples were not treated prior to analysis on the flow cytometer.
Stained samples were incubated for ten minutes in the dark with 75 nM LysoTracker
Green
®
(final concentration). Samples were preserved with 1% glutaraldehyde for at
least one hour before staining in the “Dead” treatment.
Counting cultured heterotrophic protists by flow cytometry. Five species of
bacterivorous protists were used to compare cytometric counts to microscopical counts:
Uronema sp., Paraphysomonas butcheri (4-6 μm), Bodo caudatus, Pteridomonas sp. and
Paraphysomonas bandaiensis. The marine bacterium Halomonas halodurans was used
as food for the protists. Bacteria were grown to late stationary phase in 0.1% yeast
extract, harvested by centrifugation (4000 rpm for 25 min), rinsed three times and
resuspended in 0.2 μm filtered, sterile seawater. Bacteria were added at a concentration
13
of approximately 10
7
cells ⋅ml
-1
to flasks of 0.2 μm filtered, sterile seawater. Duplicate
flasks were inoculated for each of the five protists at starting concentrations of
approximately 10
3
protists ⋅ml
-1
. Flasks were sampled throughout the exponential growth
phases of the cultures. Samples were analyzed by flow cytometry using LysoTracker
Green
®
to stain the cells and by light microscopy using a Palmer-Maloney counting
chamber as described above.
Three bacterivorous species of protists were used to test the accuracy of
LysoTracker Green
®
for counting cultures during all phases of growth, including the
stationary growth phase when ingestion rates should be low: Paraphysomonas vestita (6-
8 μm) Paraphysomonas imperforata (5-6 μm) and Cafeteria roenbergensis (3-5 μm).
Duplicate flasks of 0.2 μm filtered sterile seawater were prepared with Halomonas
halodurans as described above for the two marine protistan species (P. imperforata and
C. roenbergensis). Duplicate flasks of 0.2 μm filtered sterile fresh water were prepared
with an unidentified bacterium that served as prey for the freshwater protist (P. vestita).
Samples were removed every 5-12 hours for cytometric and microscopical analysis as
described above, until the populations reached the late stationary growth phase. Growth
rates were calculated from the slopes of the linear portions of plots of ln cell abundance
vs. time. Growth rates were calculated for each species based on cytometric counts and
microscopical counts, and the two rates were compared statistically using an ANCOVA
test.
Eukaryotic organisms could accumulate the LysoTracker Green
®
stain within
acidic organelles including chloroplasts. Therefore, phototrophic eukaryotes may
14
interfere with the detection of eukaryotic heterotrophs in natural planktonic communities
if the former emit a high green fluorescent signal from the uptake of LysoTracker Green
®

into chloroplasts. Thus, they need to be eliminated from cytograms used to count
heterotrophic protists.
Three phototrophic species were examined for uptake of the LysoTracker Green
®

stain, including two eukaryotes, Heterocapsa triquetra (20x25 μm) and Micromonas
pusilla (1-2 μm), and one prokaryote, Synechococcus sp. Preserved, stained cultures
were used as controls. The cultures were grown in sterile seawater F/2 medium (Guillard
& Ryther 1962, Guillard 1975), and cells were harvested in late exponential growth
phase. A 1 mM stock of LysoTracker Green
®
was diluted 1:100 with 0.2 μm filtered
sterile seawater and added to cultures at a final concentration of 75 nM. All cultures, live
and dead, were stained with LysoTracker Green
®
for 10 minutes in the dark before
cytometric analysis. Samples to be analyzed as dead cells were preserved using
paraformaldehyde (0.5% final concentration) for one hour prior to treatment. All samples
were analyzed on a FACScan flow cytometer. The population of phototrophs was gated
based on their red autofluorescent signal (FL3), and the mean green fluorescence (FL1)
was determined for all autofluorescent cells.
Heterotrophic protists with ingested fluorescent food particles will themselves
have a fluorescent signal that can be detected by a flow cytometer. Therefore,
heterotrophic protists with ingested algae will emit red fluorescence and thus may fall
within the “phototroph” region on a cytogram of red fluorescence (FL3) vs. relative size
(FSC). The extent to which cytometric counts of heterotrophic grazers may have been
15
affected by the fluorescence of ingested algal prey was investigated using an unidentified
chrysophyte and Uronema sp. The unidentified chrysophyte (2-3 μm) was fed to
Uronema sp., to examine the effect of algal ingestion on the chlorophyll fluorescence of
heterotrophic grazers. The ciliate was fed Halomonas halodurans for four days prior to
the addition of algae in order to eliminate algal fluorescence in the protistan food
vacuoles at the beginning of the experiment. The algae and protists were analyzed
separately on a FACScalibur flow cytometer prior to mixing the cultures in order to
establish baseline red fluorescence values for prey and predators. The samples were
acquired on the cytometer until at least 200 events of the population of interest were
recorded. The algae were then added to the culture of heterotrophic protists, and the
heterotrophs were allowed to feed for 30 minutes before subsamples of the mixtures were
analyzed on the flow cytometer.

Application of LysoTracker Green
®
to natural water samples. A FACScan flow
cytometer was used to analyze natural water samples from the east coast, and a
FACScalibur flow cytometer was used for samples from the west coast. A 1:10 dilution
of the 1 mM LysoTracker Green
®
stock was freshly prepared in 0.2 µm filtered seawater,
and was used as a working stock. An aliquot of natural sample (10 ml) was removed and
stained live at room temperature in the dark for 10 minutes with 7.5 µl of the working
stock of LysoTracker Green
®
(75 nM final concentration). The sample was acquired on
the HI flow rate setting for 10 minutes. The software was configured to record particles
with a detectable green fluorescence (FL1), and the threshold was raised until the event
16
rate dropped below 1000 events ⋅sec
-1
. An event rate of 1000 events ⋅sec
-1
represents the
upper limit of the processing speed by the CellQuest software.
A sequential analysis of the resulting light scatter and fluorescence data was
employed to determine the number of heterotrophic eukaryotes in the analyzed sample.
Eukaryotic phototrophs were first distinguished from eukaryotic heterotrophs on a
cytogram of chlorophyll fluorescence (FL3) vs. forward scatter (FSC), based on the high
relative chlorophyll fluorescence of the phototrophs (Olson et al. 1983, Yentsch et al.
1983). Detritus was then separated from cellular material on a cytogram of side scatter
(SSC) vs. green fluorescence (FL1), based on the high refractive index of the detritus
(Spinrad & Brown 1986, Ackleson & Spinrad 1988). Green fluorescent polystyrene
beads (2.5 μm) were then used to establish a size reference on a cytogram of green
fluorescence (FL1) vs. FSC. The regions of eukaryotic phototrophs, detritus, and beads
were removed from the cytogram of green fluorescence (FL1) vs. size, and all remaining
events that had an FSC signal greater than the previously marked 2.5 μm beads were
counted as heterotrophic protists.
Results
Staining conditions. Cell abundances determined by flow cytometric analyses showed
decreasing trends with staining time for the two species of heterotrophic protists
examined (Fig. 1A,C). At each sampling time, a clear population of protists was
distinguishable from background fluorescence on a cytogram of green fluorescence (FL1)
vs. relative size (FSC) (data not shown). The ratio of cytometric counts to microscopical
17
18
counts ranged from 0.86-1.14, averaging over all incubation times tested for the ciliate,
with an average ratio of 0.97. An exponential curve fit to the cytometric data for the
ciliate had the same cell abundance as the microscopical count at approximately 15
minutes. The ratio of cytometric counts to microscopical counts ranged from 0.68-1.32
for the flagellate over all incubation times tested, with an average ratio of 0.86. An
exponential curve fit to the cytometric data for the flagellate had the same cell abundance
as the microscopical count within the first 5 minutes. A stain time of 10 minutes was
chosen for all further analyses.
Stain concentration had no noticeable effect on cytometric counts for either
protistan species (Fig. 1B,D). The ratio of cytometric counts to microscopical counts for
Uronema sp. ranged from 0.87-1.52 between 4 nM and 325 nM final stain concentration,
with an average ratio of 1.06. The ratio of cytometric counts to microscopical counts
ranged from 0.90 to 1.08 for the same range of stain concentrations for Bodo caudatus,
with an average ratio of 0.99. Within this range, cytometric counts were not consistently
higher or lower than microscopical counts with increasing stain concentration. A final
stain concentration of 75 nM was used for all subsequent analyses based on these results
and the recommendation of the LysoTracker Green
®
manufacturer.

Analysis of cultured protists. Comparison of unstained, live-stained and preserved-
stained cultures of heterotrophic protists showed a marked increase in green fluorescence
for the live cells over preserved samples (Fig. 2). The magnitude of the increase in
fluorescence appeared to be related to the size of the species. Paraphysomonas
19
20
bandaiensis (~5 μm; 2A-C) had a mean fluorescence of 18.7 when stained live, but only
5.27 when stained after preservation. Pteridomonas sp. (~5 μm; 2D-F) had a mean
fluorescence of 107 when stained live, compared to a mean fluorescence of 4.56 when
stained after preservation. Uronema sp. (10x20 μm; 2G-I) had a mean green
fluorescence of 919 when stained live, compared to a mean fluorescence of 21.4 when
stained after preservation. Unstained protists had greatly reduced green fluorescent
signals relative to live stained cells. The heterotrophic bacterium (2J-L) did not show an
increase in mean green fluorescence between live and preserved samples, suggesting the
bacteria did not accumulate LysoTracker Green
®
.
Uptake of substantial amounts of the LysoTracker Green
®
stain by heterotrophic
protists enabled discrimination of the protists from large numbers of bacteria that were
also present in the cultures. Cultures of heterotrophic protists stained with LysoTracker
Green
®
were identified by their very high fluorescent signal and large relative size on a
cytogram of green fluorescence (FL1) vs. FSC (Fig. 3). The protistan populations were
distinct from bacteria on cytograms at all ranges of protistan and bacterial abundances
tested. As the protistan cultures grew, the number of events registered within the
previously identified ‘protistan’ region on the flow cytometer increased accordingly (Fig.
3B-D).
A comparison of microscopical counts to cytometric counts showed a close
agreement between the two methods of enumeration for five species of heterotrophic
protists examined (Fig. 4). All species were in exponential growth phase during the
sampling period. A regression of the flow cytometric vs. microscopical counts for all
21
22
23
species yielded a slope of 0.95 (r
2
= 0.98). The ratio of cytometric counts to
microscopical counts was 0.84 averaged over all species and growth phases.
Three small flagellates were examined over the full growth cycle of these species
in order to examine the effect of growth phase on the ratio of cytometric counts to
microscopical counts (Fig. 5). The cytometric and microscopical counts closely mirrored
each other throughout the entire growth cycle for Paraphysomonas vestita and Cafeteria
roenbergensis (Fig. 5A and B). The growth rate of P. vestita was 1.2 d
-1
based on
cytometric counts, and 1.1 d
-1
based on microscopical counts (Fig. 5A). The growth rate
of C. roenbergensis was 4.7 d
-1
based on cytometric counts, and 4.3 d
-1
based on
microscopical counts (Fig. 5B). Cytometric and microscopical counts of
Paraphysomonas imperforata also matched closely during the lag, log and early
stationary growth phases. During the late stationary growth phase, however, the
cytometric counts of P. imperforata remained relatively constant, while microscopical
counts decreased (last two samples in Fig. 5C). P. imperforata had a growth rate of 2.7
d
-1
based on cytometric counts, and 3.1 d
-1
based on microscopical counts (Fig. 5C). No
statistically significant differences were detected between growth rates calculated based
on cytometric or microscopical counts for any of the species when tested at α = 0.01. A
comparison of microscopical and cytometric counts for the three protists in all phases of
population growth yielded a regression of slope 0.99 (r
2
= 0.96) (Fig. 5D). The ratio of
cytometric counts to microscopical counts was 1.14 averaged over all species and growth
phases.

24
25

Live phototrophic eukaryotes had higher green fluorescence relative to preserved
cultures, but prokaryotic phototrophs showed no increase in green fluorescence between
the live and preserved samples (Table 1). After staining with LysoTracker Green
®
, the
live cultures of Heterocapsa triquetra emitted a much stronger green fluorescence than
the dead cultures. The fluorescence ratio of live:dead cultures was 128. The eukaryotic
alga, Micromonas pusilla, also showed an increase in fluorescence between live and dead
cultures, (ratio of 4.3). The live Synechococcus sp. cultures showed no difference in
green fluorescent signal relative to dead cultures.
Some heterotrophic grazers with ingested phototrophic prey may potentially be
counted as phototrophs in the cytometric analysis of a natural water sample. The
ingestion of algae by heterotrophic protists had a noticeable effect on red (chlorophyll)
fluorescence of the heterotrophs (Fig. 6). The ciliate, Uronema sp., had a very low red
fluorescent signal when fed bacteria (Fig. 6B). The chrysophyte prey had a strong red
fluorescent signal relative to its size, and formed a distinct region within the cytogram of
chlorophyll fluorescence (FL3) vs. FSC (gated areas in Fig.6A). After the heterotrophic
protists fed for 30 minutes on the algae, the red fluorescent signal of the heterotrophs
increased substantially and the culture displayed a fairly wide range of chlorophyll
fluorescence (Fig. 6C). This range of red fluorescent signals of the heterotrophic protists
partially overlapped the algal region on the cytogram of FL3 vs. FSC.

26

Table 1-1. Uptake of LysoTracker Green
®
by two phototrophic eukaryotes and one phototrophic
prokaryote. Samples were preserved with 0.5% paraformaldehyde in the “Dead” treatment.

Green Fluorescence (FL1)

Phototrophic Culture
MEAN SD CV (%)
RATIO
LIVE:DEAD

LIVE 437 126 29 128
Heterocapsa
triquetra
CCMP 448
DEAD 3.4 1.3 37
LIVE 9.5 5.2 54 4.3
Micromonas
pusilla
CCMP 494 DEAD 2.2 1.0 46
LIVE 2.0 1.0 51 1
Synechococcus
sp.
CCMP 837
DEAD 2.0 1.0 50

27
28
Analysis of natural water samples. The accurate cytometric enumeration of
heterotrophic protists within mixed microbial communities was accomplished by
distinguishing them from phototrophic eukaryotes and detritus on the cytogram of green
fluorescence (FL1) vs. relative size (FSC) (Fig. 7). This discrimination was
accomplished by sequential processing of light scatter and fluorescence data acquired for
each sample. Phototrophic eukaryotes had to be eliminated from the counts of
heterotrophic eukaryotes because phototrophic eukaryotes also accumulated the
LysoTracker Green
®
stain (Table 1). Phototrophic eukaryotes were detected based on
their high chlorophyll fluorescence relative to size, using a cytogram of FL3 vs. FSC (red
events in Fig. 7A). Detrital particles were identified based on their high side scatter and
low green fluorescence, using a cytogram of FL1 vs. SSC (yellow events in Fig. 7B).
Polystyrene beads (2.5 μm; gated green events in Fig. 7C) were used to establish a
minimum size for protists on the cytogram of green fluorescence (FL1) vs. forward
scatter (FSC) (vertical line in Fig. 7D). Subsequently, the phototrophic eukaryotes,
detritus and beads were removed from the cytogram of green fluorescence (FL1) vs. FSC
using logical gates, and all particles greater than the minimum size (to the right of the
vertical line in Fig 7D) were counted as eukaryotic heterotrophic nanoplanktonic protists
(blue events in Fig. 7D).
A comparison of microscopical counts and cytometric counts (determined as
described in the previous paragraph) of 26 natural water samples showed close agreement
between the two methods of quantification (Fig. 8). A regression of the data resulted in a
line of slope 1.16 (r
2
= 0.95). The ratio of cytometric counts to microscopical counts was
29
30
31
1.09 averaged for all 26 samples. The locations sampled included a wide range of coastal
environments, from relatively isolated beaches with a strong oceanic influence (Leo
Carrillo Beach, Malibu, CA) to highly eutrophic harbors (Newport Beach Harbor and Los
Angeles Harbor) and a town dock near a treated sewage outfall (East Greenwich, RI).

Discussion
Cytometric analysis of live protists. LysoTracker Green
®
can only be used with live
samples because preservation destroys membrane potential resulting in the rapid loss of
the fluorescent stain (Figure 2, Table 1). There are benefits as well as drawbacks to the
requirement for conducting this procedure with live cells. There is an obvious
inconvenience associated with the inability to analyze preserved samples, and there will
undoubtedly be situations where samples will be collected in the field that are too far
from the laboratory to return them to the lab for live analysis. If the samples are to be
transported to a lab prior to analysis, measures must be taken to insure their integrity
during transport. Samples should be kept in the dark and on ice (but not frozen) during
transport, and processed as soon as possible after returning to the lab. In addition, the
salinity of the sheath fluid should be matched approximately to the salinity of the sample
to avoid distortion of the forward scatter signal (Cucci & Sieracki 2001)
32
On the positive side, this method was developed using commercially available
benchtop flow cytometers (FACSCalibur and FACScan, Becton Dickinson, San Jose,
CA) that are relatively portable for use aboard ships and at field stations with electrical
power. Another advantage of acquiring cytometric information on live samples is the
avoidance of sample preservation that might cause lysis of some species of protists, and
the egestion of food vacuole contents (Sieracki et al. 1987). Moreover, the stain is not
strongly accumulated by prokaryotes, which do not possess intracellular vacuoles. This
ability to distinguish between prokaryotic and eukaryotic cells is a primary advantage of
LysoTracker Green
®
for detecting and counting heterotrophic protists.

Staining time and stain concentration. Bodo caudatus appeared to show more
dramatic decreases in observed cell abundance with increasing staining time than
Uronema sp. (Fig. 1A,C), although both cultures showed a trend of decreasing cytometric
counts with increasing time of staining. These results were not a consequence of
changing physiological state of the protists because both cultures remained in the
exponential growth phase throughout the analyses. A probable explanation for this trend
is the nature of the stain molecule itself. The stain molecule consists of a fluorophore
linked to a weak base that accumulates in intracellular locations with a low pH (such as
food vacuoles). I speculate that this accumulation could eventually raise the pH of the
acidic organelles in which it accumulates, thereby reducing the affinity of the stain for
these organelles over time. Thus, the organelles themselves may have the potential to
become neutralized, resulting in diffusion of the stain out of the vacuoles and loss of
33
fluorescence. The exponential curve fit to the cytometric counts for the B. caudatus
culture indicated a steeper initial decline in cytometric counts. A more rapid decrease in
apparent cell abundance for B. caudatus would be consistent with more rapid
neutralization of acidic vacuoles in this smaller species.
I examined whether LysoTracker Green
®
actually caused cell death as a result of
the presumptive neutralization of acidic vacuoles. A culture of Uronema sp. was stained
with LysoTracker Green
®
, cytometrically sorted into a bacterial culture, and observed for
a week. Some of the sorted cells were dead immediately after sorting (approximately
15% of the culture), but most were still alive and actively swimming. The culture
continued to grow for at least a week after staining and sorting, indicating that most of
the cells were not adversely affected by the combination of LysoTracker Green
®
staining
and cytometric sorting.
Surprisingly, cytometric counts of Uronema sp. and Bodo caudatus exceeded
microscopical counts for both species during the first several minutes of staining (Fig 1A,
C). I speculate that the initial cytometric counts were higher than the microscopical
counts due to nonspecific staining of small acidic pockets within detrital particles or
perhaps in bacterial aggregates, which were neutralized within the first few minutes of
staining. The exponential curve fit to the data indicated that the cytometric counts were
equivalent to the microscopical counts after approximately 15 minutes of staining for the
ciliate, and within the first 5 minutes for the flagellate. Based on my results with these
two species, I chose a staining time of 10 minutes for subsequent analyses of protistan
cultures and natural water samples as a compromise between
34
ensuring that all eukaryotes would emit a strong fluorescent signal on the one hand, and
minimizing nonspecific staining on the other.
Stain concentration did not have a consistent effect on cytometric counts when
stained for 10 minutes (Fig 1B, D). Cytometric counts of Bodo caudatus generated from
all stain concentrations compared well to the microscopical counts of the culture (Fig.
1D). Only the highest stain concentration resulted in a cytometric count of the ciliate that
greatly exceeded the microscopical count (Fig. 1B). I chose 75 nM as a final working
concentration based on my results and the recommendation of the manufacturer
(Molecular Probes, Eugene, Oregon, Technical Bulletin #07525).

Application of Lysotracker Green
®
to cultured heterotrophic protists. Flow
cytometry has been used successfully to enumerate heterotrophic protists in culture.
Lindström et al. (2002) observed a close correlation between microscopical and
cytometric counts for two ciliates fed Cryptomonas sp. and stained with TO-PRO-1
®
.
Bratvold et al. (2000) enumerated heterotrophic flagellates in cultures using flow
cytometry on cytograms of relative size (FSC) and relative refractive index (SSC) to
distinguish protists from bacteria. The latter investigation selected their study species
based on substantial size differences between protists and bacteria that allowed
unequivocal separation of the two populations. This strategy is effective for some
protistan species but can greatly constrain the protist/prey combinations that can be
examined. For example, Bratvold et al. (2000) reported that two of the four protistan
species considered for her study had cytometric patterns that overlapped with those of the
35
bacteria and therefore were not employed in her experiments. My method expands the
number of small, bacterivorous protistan species that can be effectively differentiated and
counted separately from their bacterial prey using flow cytometry because bacteria do not
exhibit a substantial LysoTracker Green
®
(FL1) fluorescent signal.
Heterotrophic bacteria showed no strong increase in mean green fluorescence
when stained live or preserved with LysoTracker Green
®
(Fig. 2J-L), indicating that
heterotrophic bacteria did not take up the stain to any appreciable degree. This result is
consistent with microscopical observations, which showed no visually detectable
fluorescence (Zeiss standard microscope equipped with a 450-490 nm excitation filter, a
510 nm dichroic beam splitter, and a 520 nm barrier filter). Live heterotrophic protists,
by contrast, showed a clear increase in mean green fluorescence over preserved or
unstained samples (Fig. 2A-I). The increase in fluorescence for live cells was related to
the size of the cells. The largest species, (Uronema sp.), had the largest increase in mean
fluorescence between dead and live samples while the smaller flagellate species
(Paraphysomonas bandaiensis and Pteridomonas sp.) showed smaller increases in mean
fluorescence between dead and live samples. The stained, preserved heterotrophic
protistan cultures still had some green fluorescence relative to unstained controls, and the
larger preserved cells again showed a higher signal than the smaller preserved cells.
Since the dead and live cells within the uniprotistan cultures had distinct fluorescent
signals, it was possible to distinguish them on a cytogram (Fig. 2A-I). However, since
the green fluorescent signal of the live and preserved cells varied based on the size of the
36
protistan cells, the method may not be able to distinguish live small protists from large
dead ones in a mixed protistan assemblage.
The use of LysoTracker Green
®
and relative size provided information on some
cultures of heterotrophic protists beyond simple enumeration. Eight of the nine protistan
species examined in this study formed tight clusters of events within the cytogram of
green fluorescence (FL1) vs. relative size (FSC). The mean green fluorescence for these
populations dropped slightly when food became limiting and the cultures entered
stationary growth phase (presumably due to a reduced number of food vacuoles), but the
population as a whole remained distinct and unified. Paraphysomonas imperforata
formed a tight cluster of events during lag and exponential growth, but the population
split into two somewhat distinct subpopulations on cytograms during early stationary
growth phase. These subpopulations had the same relative green fluorescence (FL1), but
distinctly different relative sizes (FSC). The culture again formed a tight cluster of
events within the cytogram after approximately 20 hours in stationary phase. These
findings corroborate the microscopical observations of Goldman and Caron (1985), in
which a subpopulation of P. imperforata apparently resorted to cannibalism during the
stationary growth phase when bacterial abundance was low in the culture. The authors
reported a shift to a bimodal size distribution during the stationary growth phase of the
flagellate as I observed. They described an initial wide range of cell sizes (5.5 to 10 μm)
shifting to a narrow range of small cells (3.5 to 6 μm) in early stationary phase (as prey
became limiting). The species then formed two distinct subpopulations in late stationary
37
phase that was presumably a result of feeding by some of the individuals on smaller
protistan cells within the population.
The cytometric counts of five species of heterotrophic protists in exponential
growth were highly correlated to microscopical counts of these cultures (Fig. 4). Further
analysis of three species of heterotrophic protists indicated excellent correlation between
cytometric and microscopical counts during active protistan growth (Fig. 5). Growth
rates of these three heterotrophic protistan cultures were calculated from cytometric and
microscopical counts while the populations were in exponential growth. These paired
measurements yielded rates that were in close agreement for each species (Fig. 5A-C).
The rates were compared using an ANCOVA test ( α = 0.01). No significant differences
were detected between the growth rates calculated based on cytometric counts or
microscopical counts. These results indicate that my flow cytometric technique has wide
applicability for experimental studies to examine the growth rates of cultured protists.
Populations in exponential growth were actively feeding, and would be expected
to have both lysosomes and food vacuoles (for which LysoTracker Green
®
should have
high affinity). Cells in the stationary growth phase, however, had few prey available, and
presumably the number of acidic vacuoles would have been reduced during this growth
phase. I examined whether populations in stationary phase were accurately quantified by
my method. Cytometric counts were similar to microscopical counts during the
stationary growth phase for Paraphysomonas vestita and Cafeteria roenbergensis (Fig.
5A,B). Microscopical counts of Paraphysomonas imperforata, however, were somewhat
less than cytometric counts as the culture of this protist entered late stationary growth
38
phase (two data points in Fig. 5C). This result was unexpected because I hypothesized
that cells in late stationary growth would stain less intensely and thus yield lower counts
than microscopical estimates. I speculate that this result might be explained by the
substantial amounts of aggregated detritus formed by this species as it feeds (Caron et al.
1985, Goldman & Caron 1985). It is possible that the detritus was colonized by bacteria
remaining in the culture and formed microzones of low pH that accumulated LysoTracker
Green
®
. Thus the cytometric counts may have overestimated the number of heterotrophic
protists in these samples. Conversely, detrital material may have obscured heterotrophic
protists in these samples and thus caused an underestimation in the microscopical counts.
Neither P. vestita nor C. roenbergensis formed significant amounts of detrital aggregates,
and neither culture showed a discrepancy between cytometric and microscopical counts
during late stationary growth. A comparison of cytometric and microscopical counts
from all stages of the growth cycle resulted in a regression of slope 0.99 (r
2
= 0.96) (Fig.
5D). These results indicated that LysoTracker Green
®
should provide accurate counts of
heterotrophic protists for most situations.

Uptake of LysoTracker Green
®
by phototrophic prokaryotes and eukaryotic algae.
LysoTracker Green
®
caused both heterotrophic and phototrophic eukaryotes to emit a
green fluorescent signal (Table 1, Figure 2). LysoTracker Green
®
stains all acidic
vacuoles, so chloroplasts should accumulate stain as well as lysosomes and food
vacuoles. Thus, phototrophic eukaryotes must be distinguished from heterotrophs during
cytometric analyses using gating based on the red autofluorescence of chlorophyll a.
39
Live cultures of two phototrophic eukaryotes (Heterocapsa triquetra and Micromonas
pusilla) demonstrated increased fluorescence relative to preserved controls (Table 1).
Preserved cells were used as controls because membrane potentials are lost upon cell
death, the acidic pH of vacuoles is neutralized, and LysoTracker Green
®
fluorescence
dissipates rapidly (based on microscopical examination). Cell abundances of the
phototrophic eukaryotes before and after staining with LysoTracker Green
®
did not vary,
suggesting uniform staining of the chloroplasts. Both cultures of eukaryotic algae
showed increased mean green fluorescence in live cells relative to preserved cells
indicating active uptake of the stain by the eukaryotic algae. The magnitude of the
increase in fluorescence differed markedly for the two species. H. triquetra is a much
larger species than M. pusilla, with many more chloroplasts, which explains the higher
fluorescent ratio between live and dead cultures of H. triquetra. In contrast,
Synechococcus sp., a prokaryote, stained poorly with LysoTracker Green
®
(Table 1). This
result indicated that no accommodation was necessary to account for the photosynthetic
prokaryote community within the cytometric analyses.

Application of LysoTracker Green
®
to natural water samples. The efficacy of
counting heterotrophic nanoplankton in natural samples by flow cytometry was
predicated on the differentiation of these cells from detrital particles and other groups of
co-occurring microorganisms including archaea, heterotrophic bacteria, phototrophic
prokaryotes, and phototrophic eukaryotes. These populations and particles were
40
sequentially eliminated from cytograms of natural samples stained with LysoTracker
Green
®
to obtain counts of heterotrophic protists.
Separation of stained phototrophic eukaryotes from heterotrophic eukaryotes was
possible on the flow cytometer by gating phototrophs based on their high red
autofluorescent signal, in the FL3 channel (Fig. 7A). The CellQuest software allowed for
their subsequent removal from cytograms of green fluorescence (FL1) vs. relative size
(FSC) (Fig. 7D). Detrital particles emitted a low level of green fluorescence, most likely
due to the accumulation of LysoTracker Green
®
in microzones of low pH associated with
bacteria. Conveniently, most detritus can be identified by its high side scatter properties
and relatively low green fluorescence (Spinrad & Brown 1986, Ackleson & Spinrad
1988). Based on these characters, detritus could be identified in cytograms of green
fluorescence (FL1) vs. side scatter (SSC) (Fig. 7B), gated, and subsequently eliminated
from cytograms of green fluorescence (FL1) vs. FSC (Fig. 7D).
The effectiveness with which detritus was identified in cytograms was somewhat
dependent on the total amount of detritus present in a natural water sample. Large
amounts of detritus in highly eutrophic samples were more difficult to accurately gate
because the range of side scatter from the detrital particles somewhat overlapped that of
live cells. Most coastal sites did not pose a problem but problems were observed for a
few highly eutrophic harbors (Newport Beach Harbor and Los Angeles Harbor).
Samples from oceanic waters, while not specifically tested, should not pose a problem
due to the normally low levels of detritus present in these ecosystems.
41
The minimal size of protists in the cytograms was established by gating 2.5 μm
fluorescent beads that were added to all natural water samples. Beads were used to set a
lower limit on the size of events accepted as ‘protists’ on a cytogram of green
fluorescence (FL1) vs. relative size (FSC) (Fig 7D). A rectangular gate was placed
through the center of the bead population and extended to include all events with a
greater FSC signal in order to gate protists. The beads were then removed from the
cytogram of green fluorescence (FL1) vs. FSC so that they were not counted as protists
(Fig 7C). In some samples, the heterotrophic protists could overlap with bacteria in size,
and a size gate smaller than 2.5 μm could be used. Since LysoTracker
®
appears to stain
small eukaryotes much better than prokaryotes it should allow adequate discrimination of
protists from bacteria.
Cytometric counts of events remaining after (1) elimination of phototrophic cells,
(2) elimination of detrital particles and (3) using beads to establish a minimum size, were
recorded as heterotrophic nanoplanktonic protists in natural water samples (Fig. 7D).
Cytometric counts of heterotrophic nanoplankton determined in this manner agreed
closely with counts based on DAPI staining and epifluorescence microscopy (Fig. 8).
Twenty-six different locations sampled on the East Coast and West Coast of the United
States and analyzed to test the applicability of this method provided samples from a wide
range of coastal environments. The regression of the resulting data yielded a line of slope
1.16 (r
2
= 0.95). The average ratio of cytometric counts to microscopical counts (1.09)
indicated a very close agreement between the two counting methods.
42
Cytometric counts of heterotrophic nanoplankton could have been expected to
provide an overestimation of the abundance of heterotrophic protists compared to
epifluorescence microscopy due to misidentification of detrital particles as heterotrophic
nanoplankton in the cytogram. The incomplete removal of detrital particles from the
region gated to count heterotrophic nanoplankton (Fig. 7B,D) would have artificially
increased cytometric counts. As noted above, this only appeared to be a problem in a few
extremely eutrophic environments with high concentrations of particulate material
(Newport Beach Harbor and Los Angeles Harbor). However, these samples were also
problematic to count using epifluorescence microscopy. The presence of large amounts
of detritus in the water samples may have obscured some cells on slide preparations,
resulting in underestimation of the abundances of heterotrophic nanoplankton. The
combination of a potential underestimation of heterotrophic nanoplankton by
epifluorescence microscopy and potential overestimation by flow cytometry most likely
caused the slight discrepancies observed between these two counting methods in my
study (ratio of cytometric:microscopical counts of 1.09; Fig. 8). Nevertheless, I found a
close relationship between the cytometric quantification of heterotrophic nanoplankton
and the commonly employed microscopical method of enumeration.
As previously noted, eukaryotic phototrophs accumulated LysoTracker Green
®

within their chloroplasts, and emitted a strong green fluorescent signal (Table 1). These
species were separated from eukaryotic heterotrophs on the flow cytometer based on the
strong red autofluorescence present in the phototrophs. It has been shown that the
fluorescent particles within heterotrophic protists can be detected by the flow cytometer,
43
giving the heterotrophs a fluorescent signal (Gerritsen et al. 1987, Cucci et al. 1989,
Keller et al. 1994, Weisse & Kirchoff 1997). One potential complication of my
methodological approach is that heterotrophic grazers with ingested algae also would
exhibit red fluorescence and may fall within the ‘phototroph’ region on a cytogram of red
fluorescence (FL3) vs. relative size (FSC). Therefore, the abundance of heterotrophic
nanoplankton would be underestimated if heterotrophs with ingested phototrophs were
counted as algae.
To examine the possibility of this artifact, I measured the change in red
fluorescence after the addition of algae to a culture of herbivorous, heterotrophic protists
(Fig. 6). Regions were gated encompassing the phototrophs on cytograms of the algal
culture alone (Fig. 6A), then observations were made with respect to how many
heterotrophs exhibited substantial chlorophyll fluorescence after feeding for 30 minutes
on the algae (Fig. 6C). The heterotrophs displayed a wide range of red fluorescent
signals, which was expected because each individual heterotroph might vary in the
number of ingested algae at any one time and the degree of digestion of the prey. I
observed that some of the heterotrophs did fall within the ‘phototroph’ region after
feeding on algae, but most were located outside the gated region (Fig. 6C). This result
indicated that the cytometric quantification of heterotrophic nanoplankton may
underestimate the total number of heterotrophic protists in a natural water sample if a
substantial portion of the phagotrophic assemblage is actively involved in herbivory.
Many of these herbivorous cells, however, would still be included in cytometric counts of
44
heterotrophic nanoplankton due to the low red autofluorescence of these cells relative to
their size.
It should be noted that this bias (i.e. mistaken classification of herbivorous protists
as phototrophic protists) can also be a problem with the established method of counting
heterotrophic protists by epifluorescence microscopy. The microscopical method
differentiates phototrophic nanoplankton (PNAN) from heterotrophic nanoplankton
(HNAN) based on the presence or absence of red autofluorescence under blue light
excitation. Some differentiation between heterotrophic protists with ingested algae and
phototrophic protists is possible using microscopy, but this task can be difficult for some
species of protists.
Similarly, ciliates and heterotrophic dinoflagellates that retain functional
chloroplasts obtained from ingested algal prey (Stoecker 1998) might be eliminated from
cytograms based on their ‘acquired’ red fluorescence from the chloroplasts. Elimination
of these cells could result in underestimation of the abundance of heterotrophic
nanoplankton by cytometry. However, most of these latter cells are larger than the cells
that can be effectively counted by flow cytometry. In any event, my cytometric counts
compared very well to microscopical counts of natural samples (Fig. 8), implying that
either herbivorous and mixotrophic protists did not contribute significantly to the samples
analyzed in this study, or that my method of enumeration does not exclude any more
heterotrophs than the commonly used microscopical method.

45
Conclusions
LysoTracker Green
®
is a very promising tool for the semi-automated enumeration
of nanoplanktonic heterotrophic protists by flow cytometry. Heterotrophic protists in
cultures and in natural water samples were accurately quantified using a standard
benchtop flow cytometer. The flow cytometer dramatically decreased the time for
sample processing relative to traditional microscopical methods and yielded results for
natural samples that were very comparable to counts obtained using epifluorescence
microscopy.

46
Chapter 2: Does low temperature constrain the growth rates of
heterotrophic protists? Evidence and implications for algal blooms in
cold waters
Introduction
Phytoplankton blooms are a common occurrence in many aquatic ecosystems.
These phenomena can positively or negatively affect food web structure and carbon flow
in marine ecosystems. For example, spring blooms in temperate environments are often
characterized by phytoplankton species that are subsequently grazed by larger
zooplankton, resulting in the efficient transfer of energy to higher trophic levels and away
from energetic losses within the microbial loop. Conversely, blooms of toxic or noxious
species of phytoplankton can disrupt energy transfer in planktonic food webs, and/or
result in illness or death of mammals, birds and commercially important fish and
shellfish.
The formation of a phytoplankton bloom indicates a fundamental imbalance
between growth and removal of phytoplankton. Assuming that grazing dominates other
loss factors, this imbalance may be accomplished through the stimulation of
phytoplankton growth relative to extant grazing pressure, the inhibition of herbivory in
the presence of phytoplankton growth, or some combination of the two. Most research
has emphasized the importance of the stimulation of the intrinsic growth rate of
phytoplankton as the primary underlying cause for massive accumulations of
phytoplankton, and deemphasized the importance of removal processes (Verity &
Smetacek 1996). This predilection is rooted in the preponderance of studies that have
47
examined the response of phytoplankton growth to the availability of nutrients and light
(Riley 1942, Sverdrup 1953, Platt et al. 1991). Increases in phytoplankton standing stock
can only occur if the algal population is growing. Nonetheless, reduced grazing pressure
due to constraints on herbivory (e.g., due to low zooplankton abundance, low individual
feeding rates, or the production of toxic or inhibitory compounds by phytoplankton) will
yield a higher net population growth rate for a given intrinsic growth rate of the
phytoplankton assemblage, and thus can play a role as an explanation for some
phytoplankton blooms (Riley 1942, Sverdrup 1953, Hansen 1989, Smayda 1997, Liu &
Buskey 2000, Tagliabue & Arrigo 2003).
One classical explanation for the initiation of non-toxic (i.e., ‘edible’)
phytoplankton blooms (e.g., the spring bloom of many temperate and polar ecosystems)
is the temporal offset that occurs between the onset of rapid phytoplankton growth in
early spring and the subsequent development of a zooplankton assemblage that is
sufficient to affect phytoplankton standing stock. Historically, the occurrence of these
blooms has been reconciled by the fact that planktonic metazoan development is slow
relative to maximal phytoplankton growth rates, particularly at very low temperature
(Walsh & McRoy 1986, Huntley & Lopez 1992, Napp et al. 2000). This explanation has
received less attention in recent years due to the recognition of the importance of
herbivorous microzooplankton, which are eukaryotic consumers of phytoplankton in the
size range 20-200 µm, as defined by Sieburth et al (1978). Herbivorous
microzooplankton have demonstrated the potential for rapid growth rates and thus rapid
response rates to increases in phytoplankton production. In fact, the absence of
48
phytoplankton blooms in regions that would be expected to manifest these phenomena
has been attributed to high microzooplankton standing stocks at those locations and times
(Frost 1987, Parsons & Lalli 1988, Frost 1991).
If the latter scenario is a general principle of marine ecosystems, then one might
ask, why are blooms of non-toxic phytoplankton such a common event in the ocean?
Given the ubiquity of herbivorous microzooplankton, and their potentially rapid instrinsic
growth rates, what environmental factors constrain the growth and grazing rates of
herbivorous microzooplankton (relative to phytoplankton growth rates) that would allow
phytoplankton blooms to develop? I hypothesize that the extreme low temperature
characteristic of high latitude environments provides a fundamental constraint on the
metabolic rates of microzooplankton relative to the effect of temperature on
phytoplankton growth rates. Given favorable conditions for phytoplankton growth, this
imbalance can result in a substantial net phytoplankton growth rate, thus contributing to
massive bloom formation in these environments. My hypothesis is supported by bloom
dynamics and an extensive analysis of growth rate data from the literature.
Massive annual algal blooms are common in high latitude marine ecosystems.
For example, the Ross Sea, the Bering Sea, and the high latitude North Atlantic are
characterized by seasonal algal blooms that are spatially extensive with high maximal
chlorophyll concentrations. While bloom composition and specific timing vary with ice
conditions and other annual variables, the overall dynamics of the blooms in these
environments are generally similar. Chlorophyll concentrations and algal cell
abundances increase dramatically during late spring in conjunction with rapidly
49
increasing light levels, daylengths, and ice retreat (El-Sayed et al. 1983, Smith & Nelson
1985, Lochte et al. 1993, Stramska et al. 1995, Smith et al. 2000).
Although herbivorous microzooplankton can be relatively abundant in these
ecosystems, field studies indicate that grazing pressure is insufficient in spring to prevent
the formation of large phytoplankton blooms during late spring and summer (Burkill et
al. 1993, Verity et al. 1993, Stoecker et al. 1994, Gifford et al. 1995, Caron et al. 2000).
This mismatch between production and consumption appears to be temperature-related.
In the Ross Sea where extreme low temperatures ( ≤0.5˚C) persist year-round, observed
rates of herbivory (d
-1
) were consistently low (Caron et al. 2000). Rates of
microzooplankton herbivory in the Bering Sea were positively correlated (albeit weakly)
with temperature (Olson & Strom 2002), while in the North Atlantic at 59˚N,
microzooplankton herbivory had relatively little effect on the phytoplankton assemblage
during spring yet removed most of the potential daily chlorophyll production during
summer (Gifford et al. 1995). Analysis of a much larger data set on the relationship
between phytoplankton mortality and temperature has indicated a positive relationship
between these parameters (Caron et al. 2000). This relationship may in part be due to
differences in phytoplankton community composition at different temperatures, but these
observations may also indicate the potential for a fundamental difference in the effect of
temperature on the growth of phototrophic and heterotrophic protists. I examine here this
difference as a factor in the formation of phytoplankton blooms at high latitude.
The relationship between growth rate and temperature for temperate and tropical
species of phytoplankton is well established. Eppley (1972) reviewed published growth
50
rates for marine phytoplankton available up to that time, and demonstrated that the upper
limit for phytoplankton growth rate was exponentially and positively correlated with
temperature. Goldman and Carpenter (1974) derived a similar relationship between
temperature and phytoplankton growth rate based on data derived from continuous
culture experiments using a variety of algal species. The Goldman-Carpenter curve had a
lower y-intercept but a similar slope to Eppley’s curve.
The relationships described by Eppley (1972) and Goldman and Carpenter (1974)
were based almost entirely on data derived from temperate and tropical phototrophic
protists. Thus the applicability of these data to extrapolate to potential maximal growth
rates of polar phytoplankton species is questionable. In addition, a large number of
studies to date have examined the relationship between temperature and growth rate for a
wide spectrum of heterotrophic protists, but relatively few studies have examined this
relationship for more than a few species at one time (Fenchel 1968, Finlay 1977, Baldock
et al. 1980, Muller & Geller 1993). To date, no comprehensive review has examined the
effects of temperature on the growth rates of heterotrophic protists.
Here I address these issues by expanding the analysis of phytoplankton growth
rate using information available since the publication of Eppley’s review more than thirty
years ago, to examine (1) its usage as a descriptor of phytoplankton maximal growth rate
and (2) its applicability to describe maximal growth rates of phototrophic protists in
permanently cold environments. In addition, I examine the relationship between
temperature and growth rate for heterotrophic protists, and specifically compare
temperature dependence of phototrophic and heterotrophic protists in order to examine
51
the contribution of extreme low temperature to phytoplankton blooms in high latitude
environments.
Methods
Protistan growth rates were compiled and organized into groups first based on the
local environment from which cultures were isolated (temperate/tropical vs. polar).
Temperate and tropical protists were grouped together since the limited data available for
protists from tropical regions did not provide enough information for a meta-analysis.
Protists were further grouped according to mode of nutrition (phototroph vs. heterotroph),
and then heterotrophs were divided into groups according to prey type (bacterivore vs.
herbivore). Growth rates measured on cultured protists were included, but rates
measured on mixed plankton assemblages were not considered. The total data set
consisted of 3,374 growth rates, representing a wide taxonomic range of protists isolated
from water samples obtained throughout the world, including both limnetic and marine
systems. The list of references is included as Appendix 1. The data set itself is included
as Appendix 2 (phototrophic protists) and Appendix 3 (heterotrophic protists). The data
set contained 2,867 growth rates of protists from temperate and tropical regions, and 483
from polar regions. There were 2,042 growth rates of phototrophic protists (1,590
temperate and 452 polar) from 58 published reports, and 1,308 growth rates of
heterotrophic protists (1,277 temperate and 31 polar; 597 bacterivores and 711
herbivores) from 53 publications.
All growth rates were converted to intrinsic growth rate (d
-1
) using the equations:
µ=(ln 2) x (t
g
)
-1

52
(1) x (t
g
)
-1
=doublings d
-1
where µ = intrinsic growth rate (d
-1
) and t
g
= generation time (days). Numerical values
for growth rates published in graphical form were acquired using the freeware program
Data Thief (Kees Huyser, http://www.datathief.org).
Results and Discussion
The entire data set of protistan growth rates compiled from the literature spanned
the temperature range –3 to 40˚C. This range was chosen because my primary interest
was the response of growth rate to temperatures that occur within typical marine and
limnetic systems. Growth rates have been measured at temperatures below –3˚C, but this
requires highly saline conditions found only in unusual environments such as brine
channels. Eppley’s (1972) published curve covered the temperature range 0-45˚C, and
was calculated using algal growth rates in doublings

d
-1
. I extrapolated the curve to –3˚C
using the equation reported in his paper (doublings

d
-1
; log
10
µ
max
= 0.0275T - 0.070; T =
temperature in degrees Celsius), converted to intrinsic growth rate (d
-1
).
Growth rates reported for cultures of phototrophic protists from polar, temperate
and tropical regions were nearly all less than or equal to Eppley’s predicted maximum
over the entire temperature range (solid line, Fig. 1). Thus, the overall effect of
temperature on maximal growth rates of phototrophic protists appears to be consistent
with Eppley’s original findings. Out of a total of 2,048 compiled growth rates of
phototrophic protists, 157 (~8%) were greater than the maximum predicted by the Eppley
curve. Brush et al. (2002) noted published growth rates in excess of the Eppley curve.
They suggested that the use of Eppley’s maximal growth rate equation resulted in an
53
0
1
2
3
4
5
6
7
-3 0 5 10 15 20 25 30 35 40
Temperature (
o
C)
Continuous Light


<24 Hour Daylength
Figure 2-1. Summary of the relationship between published growth rates of phototrophic
protists and temperature. Solid line represents the Eppley (1972) curve of maximal predicted
growth rates of phototrophic protists. Dashed line represents the curve from Brush et al. (2002)
54
underestimation of primary production in mathematical models of this process. Brush et
al. (2002) argued for an alternative to the Eppley curve based on their new data set, and
recommended increasing the y-intercept of the Eppley curve by roughly 60%. The new
predictor (henceforth called the Brush curve) is depicted as the dashed line in Fig. 1.
While the Eppley curve may underestimate the absolute maximal growth rate that
can be attained by the fastest growing phototrophic protists, this curve is still a valid
representation of maximal growth rate for >90% of all the growth rates in my compiled
dataset. Moreover, the extrapolation of Eppley’s curve below 0˚C appears to be a valid
representation of growth rates of polar phototrophic protists at extreme low temperature.
While the exact placement of Eppley’s line depicting maximal growth rate may be
debated, it is clear that the shape of the curve is still appropriate for most of the data
available before and since 1972.
The data sets of Eppley (1972) and Brush et al. (2002) were based primarily on
growth rates of cultured protists measured under continuous illumination, since these
authors were interested in obtaining the maximal possible growth rate for any
phototrophic protist at each temperature. This approach may be appropriate for extreme
high latitude ecosystems where protists will experience annual periods of continuous
illumination, but at lower latitudes the annual maximal photoperiod will be reduced
substantially from 24 h. Therefore, I divided my data set into growth rates measured
under continuous vs. <24 hour illumination to look for differences in maximal growth
rates between these two groups (open and closed symbols, respectively, in Fig. 1). No
55
discernable differences were apparent between phototrophic protists grown in continuous
or <24 hour illumination.
Growth rates of heterotrophic protists in culture were compared to maximal
growth rates of cultured phototrophic protists over the temperature range –3 to 40˚C (Fig.
2; maximal growth rates of phototrophs (Eppley curve from Fig. 1) are represented by the
solid line in Fig. 2A and B). Heterotrophic protists were divided according to prey type,
separating bacterivores from herbivores. Bacterivores were capable of extremely rapid
maximal growth rates at high temperatures (Fig. 2A), such as 8.3 d
-1
at 25˚C reported for
a strain of Uronema marinum (Martinez 1980) and 6 d
-1
at 20˚C reported for an
unidentified species within the genus Paraphysomonas (Caron et al. 1991). In general,
the fastest growth rates of bacterivores were far in excess of Eppley’s curve at similar
temperatures. As the Eppley curve represents the theoretical maximal growth rates that
can be achieved by phototrophic protists at any temperature, the data in Fig. 2A indicate
that bacterivores, in the presence of high prey abundances, have the potential for much
higher growth rates than phototrophic protists. At temperatures less than 18˚C maximal
published growth rates of bacterivores declined sharply and below 5˚C the growth rates
of these species were less than or equal to the maximal observed growth rates of
phototrophic protists. Q
10
values were calculated for bacterivorous protists using the
maximal reported growth rate at each temperature using the standard equation Q
10
=(µ
1
x
µ
2
-1
)
(10/(t1 - t2))
where µ is specific growth rate (d
-1
) and t is temperature ˚C). The Q
10
value
for maximal growth rates of bacterivorous protists between 5 and 25˚C was 2.4,
compared to a value of 1.88 for the Eppley (1972) curve. Between the temperature
56
0
1
2
3
4
5
6
7
8
9
-3 0 5 10 15 20 25 30 35 40
Temperature (
o
C)
0
1
2
3
4
5
6
7
8
9
-3 0 5 10 15 20 25 30 35 40
A
B
Figure 2-2. Summary of the relationship between growth rates and temperature
for bacterivores (A) and herbivores (B). Solid line represents the Eppley curve
for phototrophic protists from Figure 1.
57
ranges 0-10˚C and 10-20˚C, the Q
10
value for maximal growth rates of bacterivorous
protists was even higher: 2.68 and 2.97, respectively. The differences in Q
10
values
between phototrophic and bacterivorous protists imply a fundamentally different
relationship between temperature and growth rate for these two protistan groups.
Growth rates of herbivorous protozoa have been reported over the temperature
range 5-30˚C. I am unaware of published growth rates of cultured herbivorous protozoa
below 5˚C, or of growth rates of cultured herbivorous protozoa from polar environments
measured at any temperature. In general, growth rates of herbivores were lower than
those of bacterivorous protozoa (Fig. 2B). The maximal observed growth rate for a
herbivorous protist at any temperature was 4.04 d
-1
, compared to the maximal rate of 8.32
d
-1
observed for a bacterivorous protist. The Q
10
value derived from the maximal growth
rates of herbivores between 10 and 20˚C (Fig. 2B) was 3.75, which is larger than the Q
10

obtained by Eppley for phototrophs and the Q
10
observed in this review for bacterivores
(Fig. 2A). The maximal growth rates of herbivorous protozoa were in excess of those of
their prey at temperatures common to temperate and tropical regions (20-30˚C, Fig. 2B).
However, as temperature decreased, the maximal growth rates of herbivorous protists
decreased more rapidly than those of their algal prey. At the lowest temperature for
which data are available (5˚C), the highest reported growth rate for a herbivorous
protozoan was 0.3 d
-1
, which is less than half the value of 0.81 d
-1
predicted by the Eppley
curve for phototrophic protists at the same temperature.
Differences between the maximal growth rates of bacterivorous and herbivorous
modes of nutrition observed in the multi-taxa data sets of Fig. 2 were also apparent for a
58
narrower data set examining omnivorous species of the genus Paraphysomonas grown
primarily on phototrophic protists or bacteria (Fig. 3). Isolates of Paraphysomonas spp.
fed bacteria grew far in excess of the Eppley curve at temperatures representative of
temperate and tropical regions (20-28˚C; Fig. 3A) while Paraphysomonas spp. fed algal
prey grew at maximal rates only marginally above the Eppley curve (Fig. 3B). Maximal
growth rates of the flagellate fed bacteria decreased rapidly below 18˚C (Fig. 3A).
Maximal growth rates of bacterivorously grown Paraphysomonas appeared to cross the
Eppley curve at polar temperatures (-1˚C). However, Paraphysomonas spp. fed primarily
phototrophic protists (including both axenic and nonaxenic cultures of prey) exhibited
maximal growth rates that appeared to cross the Eppley curve at approximately 18˚C
(Fig. 3B). The Q
10
values for these species were similar between the herbivorous and
bacterivorous Paraphysomonas (2.57 between 14 and 26˚C for a herbivorous diet vs.
2.31 between 10 and 20˚C for a bacterivorous diet). The Q
10
values were higher for the
bacterivorous Paraphysomonas between 0 and10˚C than between 10 and 20˚C (3.97 vs.
2.31), but no information is available for herbivory by Paraphysomonas spp. below 14˚C,
so it is unclear whether this trend is true for both trophic modes.
I examined the possibility that the observed differences in maximal growth rates
between herbivorous and bacterivorous protists might be explained by differences in the
average size of herbivores and bacterivores. Size has previously been correlated to
zooplankton growth rate (Hansen et al. 1997), and since herbivorous protists are feeding
on much larger prey, they could themselves have been larger on average. Growth rates of
bacterivorous and herbivorous protists were compared on the basis of cell volume (Fig.
59
0
2
4
6
8
10
12
-3 0 5 10 15 20 25 30 35 40
Temperature (
o
C)
0
2
4
6
8
10
12
-3 0 5 10 15 20 25 30 35 40
A
B
Figure 2-3. Effect of temperature on the growth rate of bacterivorous (A) and
herbivorous (B) Paraphysomonas spp. Solid line represents the Eppley curve.
60
4). Volumes were calculated based on standard geometric shapes if cell measurements
but not cell volumes were published. Cell volume or measurements for the same species,
published elsewhere, were used if neither volumes nor measurements were published
with the growth rate data. The herbivorous dinoflagellate Noctiluca sp. was removed
from the data set prior to analysis because its unusual morphology made its cell volume a
clear outlier.
The cell volumes of species feeding herbivorously or bacterivorously overlapped
considerably and thus the more rapid growth rates of the bacterivorous protists were not
due to smaller average cell volumes than those of herbivorous protists (Fig. 4). The
largest cell volumes reported included both herbivorous and bacterivorous species. The
smallest cell volumes were strictly bacterivorous. The average cell volume of the
bacterivores was ~ 4,000 µm
3
. If the cell volumes <100 µm
3
were removed (thus
including only the range in which the herbivorous and bacterivorous protistan cell
volumes overlapped), the average cell volume was ~13,000 µm
3
.

The average cell
volume of the herbivores was ~14,000 µm
3
. Rapid growth rates (>4 d
-1
) were achieved
solely by bacterivores, and were attained by a wide range of sizes of bacterivorous
protists (55 to 54,000 µm
3
). The average growth rates of bacterivorous and herbivorous
protists were compared over the entire range of volumes using a modified ANCOVA test
(Wilcox 2003). This test has the advantage of no assumptions of normality,
homoscedasticity, linearity or that the regression lines are parallel. This test used a
running interval smooth to approximate the regression lines for each set of points, then
typical values for growth rate were compared (given some value for volume). Individual
61
0
1
2
3
4
5
6
7
8
9
0 1 2 3 4 5 6 7
Log Cell Volume (µm
3
)

Herbivores
 Bacterivores
Figure 2-4. Growth rates of herbivorous and bacterivorous protists as a function of cell volume.
62
tests were performed using a modified one-step M-estimator as a measure of location, the
distribution of the data set was approximated using a percentile bootstrap method and
familywise error rate was controlled by adjusting α values. The test was performed using
the freeware statistical program R (http://www.r-project.org/). The modified
ANCOVA found significant differences between the growth rates of bacterivores and
herbivores across the entire range of volume, at α = 0.05.
The relationships between maximal growth rates and temperature were compared
for phototrophic protists, bacterivorous protists, herbivorous protists and copepods (Fig.
5). Copepod growth rates were obtained from Huntley and Lopez (1992). The natural log
of all growth rates for each category was calculated and plotted against temperature. The
Eppley and Brush curves were converted to natural log and plotted against growth rates
of phototrophic protists (Fig. 5A). An upper envelope enclosing the maximal growth
rates of bacterivorous and herbivorous protists and copepods was generated, similar to
the Eppley and Brush curves for phototrophic protists (Fig. 5B-D). This envelope was
generated for each subset as follows: the maximal two growth rates for each temperature
were first placed in a separate group. Since many temperatures only had one or two
growth rates total, a fair amount of scatter was observed when this group was plotted
against temperature. Unusually low values were removed from the subset until a
regression of the remaining data, plotted against the entire data set, represented a
reasonable upper limit (visually determined against the data in Fig. 5B-D). A series of
figures illustrating the process by which I generated the upper envelope for herbivorous
protists, in addition to two figures demonstrating the robust nature of the regression lines,
63
-5
-4
-3
-2
-1
0
1
2
3
-5
-4
-3
-2
-1
0
1
2
3
-3 0 5 10 15 20 25 30 35 40
-5
-4
-3
-2
-1
0
1
2
3
-3 0 5 10 15 20 25 30 35 40
Temperature (
o
C)
B
C D
-5
-4
-3
-2
-1
0
1
2
3
A
E: y = 0.06x - 0.5
B: y = 0.06x - 0.03
y = 0.11x - 0.03
y = 0.13x - 1.5 y = 0.12x - 3.0
Figure 2-5. Growth rates of phototrophic protists (A), bacterivorous protists (B), herbivorous
protists (C) and copepods (D) as a function of temperature. Solid line in (A) represents the Eppley
curve, and dashed line represents the Brush curve. Solid larger symbols indicate the maximal
values within each data set used to calculate the regression equations, open smaller symbols
indicate values not used in calculation of the regression equation.
64
has been included as Appendix 4. The regressions obtained in this manner were y =
0.11x - 0.32 for bacterivorous protists (Fig. 5B), y = 0.13x - 1.5 for herbivorous protists
(Fig. 5C) and y = 0.12x – 3.0 for copepods (Fig. 5D).
The Eppley and Brush curves describing the effects of temperature on maximal
algal growth rates were then compared to the regressions generated in Figs. 5C and 5D
for algal grazers (herbivorous protists and copepods; Fig. 6). This comparison indicated
that the maximal growth rates of algal grazers decreases much more rapidly with
decreasing temperature than the maximal growth rates of their algal prey. Furthermore,
the slopes of the regressions of the herbivores (herbivorous protists and copepods) were
quite similar to each other.
Maximal physiological rates (e.g. clearance rates, growth rates and ingestion
rates) have been hypothesized to vary among different taxonomic groups of protists
(Hansen et al. 1997). These authors specifically noted that ciliates had much higher
maximal growth rates than those of dinoflagellates. It is possible that the steep decline in
maximal growth rate of herbivorous protists observed in this study could be accentuated
if the data set were biased at extreme low and high temperatures towards different
taxonomic groups of protists. For example, if the maximal growth rates published at high
temperatures (20-25˚C) were dominated by ciliates, but no studies of ciliates were
published at low temperature (0-5˚C), then more slowly-growing taxa such as
dinoflagellates would have contributed to the maximal growth rate estimation at low
temperature. This bias could have skewed the regression of temperature vs. maximal
growth rate at low temperature and could have resulted in a larger slope for the overall
65
-3
-2.5
-2
-1.5
-1
-0.5
0
0.5
1
1.5
2
-3 0 5 10 15 20 25 30 35 40
Temperature (
o
C)
Figure 2-6. Comparison of maximal growth rate predictor lines for phototrophic protists,
herbivorous protists and copepods generated in Figure 2-5. The thick solid line represents the
Eppley curve, the thin solid line represents the Brush curve, the dashed line represents maximal
growth rates of herbivorous protists and the dotted line represents maximal growth rates of
copepods.
66
regression. I have included the specific data points used to form the regression equation
for herbivorous protists as Appendix 5 to address this issue. This subset of the herbivore
data set was dominated by ciliates across the entire range of temperatures, with a few
nanoflagellate growth rates in the middle of the range. Since the data set used to generate
the regression was not comprised of different taxonomic groups at different temperatures,
I am confident my regression is an unbiased reflection of the relationship between
temperature and maximal growth rate for herbivorous protists in general.
I speculate that the more severe effect of decreasing temperature on the maximal
potential growth rates of herbivorous protists (relative to its effect on maximal growth
rates of their algal prey) may have major implications for microbial food web dynamics
in cold-water ecosystems. I infer from my analysis that, all other conditions being
optimal, the growth rates of herbivorous protists in cold waters will be lower than rates
for phototrophic protists. This effect may result in low grazing pressure on polar
phototrophic protists in spring when phytoplankton growth rates are rapid due to
increasing photoperiod and nutrient availability. In northern temperate systems, grazing
pressure on phototrophic protists may also be constrained (relative to phytoplankton
growth rates) in early spring when water temperatures are low. This situation appears
analogous to the poor temporal coupling between bacterial production and primary
production in some cold marine ecosystems (Pomeroy & Deibel 1986, Ducklow et al.
2001). The underlying physiological basis for this difference in maximal growth rates of
phototrophs and heterotrophs is not clear. I speculate that the catabolic processes
67
associated with heterotrophy would appear to be a likely source of the differences
because phototrophs and heterotrophs may contain similar suites of anabolic pathways.
Based on this reasoning, slow growth rates of herbivores relative to growth rates
of phototrophic protists may allow phototrophs to temporarily escape top-down control,
and contribute to the initiation of massive phytoplankton blooms. It is unfortunate that
little to no data are available for growth and grazing rates of cultured herbivorous protists
at the extremely cold (<5˚C) temperatures at which the disparity between maximal
growth rate of herbivorous and phototrophic protists may be the highest and thus the
potential for phytoplankton bloom formation the greatest.
It is, of course, important to recognize that growth rates of heterotrophic protists
do not translate directly into grazing pressure in marine systems. For example, it is
possible that slow growth rates that are the result of rapid ingestion rates but low growth
efficiencies, could still yield high rates of prey removal. However, an increasing amount
of field and laboratory information supports the idea that temperature significantly
constrains grazing rates. Burkill et al. (1995) measured low rates of herbivory in the
Bellingshausen Sea, Antarctica. Caron et al. (2000) measured rates of herbivory during
four cruises spanning three seasons in the polynya of the Ross Sea, Antarctica. Only 13
of 51 experiments yielded detectable rates of herbivory using the dilution method, and the
highest rate observed was low in comparison to rates reported for temperate and tropical
ecosystems. The low grazing rates in that study were not a consequence of low grazer
abundance. Microzooplankton grazers (ciliates and heterotrophic dinoflagellates) varied
considerably in abundance over the course of the experiments and were at times
68
comparable in abundance to those observed in locations such as the North Atlantic during
the spring bloom, as well as the Arabian Sea and Equatorial Pacific. The authors also
compiled a literature review of grazing rates from 19 published studies measured over a
wide range of temperatures and from a variety of marine systems and found significantly
lower grazing rates at temperatures below 2˚C compared to rates measured at 10˚C or
above. Similarly, two studies that examined ingestion rates of microzooplankton on
fluorescently labeled phytoplankton in the Atlantic and Indian sectors of the Southern
Ocean reported low uptake rates at ambient temperature (Becquevort 1997, Becquevort et
al. 2000). Maximal ingestion rates in those studies were ≤2 P. antarctica grazer
-1
h
-1
, and
≤0.01 P. antarctica grazer
-1
h
-1
. Measurements of specific ingestion rates of individual
ciliate taxa in natural plankton assemblages within the Ross Sea, Antarctica, as well as
specific ingestion rates of an Antarctic Strombidium sp. culture in the laboratory support
the idea of very low specific ingestion rates at natural prey abundances at ambient
Antarctic temperatures (Rose 2006).
Some of the largest annual phytoplankton blooms in the world occur in cold
waters. At the same time, neither polar nor temperate phototrophic protists in culture
have demonstrated rapid growth at low temperature, based on the maximal growth rate
information collected in this review. My analysis suggests that the formation of these
massive blooms may, in part, be related to the different effects of temperature on the
growth rates of phototrophic and heterotrophic protists in these systems. The growth
rates of heterotrophic protists in general appear to be more strongly constrained by
decreasing temperature than the growth rates of phototrophic protists. Also, my data
69
indicate that the growth rates of herbivorous protozoa are considerably lower than rates
for bacterivorous protozoa for the same temperature, regardless of cell size, and even
among members of the same genus. Maximal growth rates of herbivorous protozoa were
similar to the maximal growth rates of phototrophic protists at temperate temperatures
(20˚C), but decreased sharply with decreasing temperature to approximately half the
maximal growth rates of phototrophic protists at the lowest temperature for which data
are available (5˚C). Published reports of grazing and ingestion rates of microzooplankton
in cold waters support my overall conclusion that temperature also exhibits a strong
negative effect on herbivory. Slow growth and grazing by herbivores in cold waters may
result in a reduction in top-down control of phytoplankton by micrograzers at a critical
period when phytoplankton growth rates are stimulated. Reduced top-down control in
cold waters due to low temperatures could be a strong contributing factor in the formation
of algal blooms in these ecosystems.
70
Chapter 3: Effects of temperature and prey type on growth rate and
gross growth efficiency of an Antarctic bacterivorous protist

Introduction
Heterotrophic protists are key players in aquatic ecosystems. These organisms
contribute significantly to the total living biomass within these systems, serve as the
major top-down control on bacterial assemblages, and are an important source of
mortality for microalgae and other heterotrophic protists (Sanders et al. 1992, Sherr &
Sherr 1994). Heterotrophic protists also function as important remineralizers of organic
matter and nutrients in aquatic systems (Azam et al. 1983, Caron & Goldman 1990). In
accordance with these important ecological roles, heterotrophic protists have been the
subject of considerable study both in the field and laboratory. Cultures of these species
have been examined to determine the effects of a wide range of physical and chemical
parameters on growth rate and other physiological and biogeochemical processes.
Temperature is recognized as a fundamental determinant of physiological rates of
heterotrophic protists including growth rates. In general, growth rates increase with
increasing temperature up to an optimum and then decrease beyond the physiological
optimum for the species (Lee & Fenchel 1972, Martinez 1980, Baldock & Berger 1984,
Caron et al. 1986, Muller & Geller 1993, Weisse et al. 2001). This relationship is
generally expressed using a Q
10
value, which represents the ratio of change in growth rate
with a 10˚C change in temperature. Q
10
values are calculated using the equation Q
10
=
71
(µ
1
/µ
2
)
10/(t1-t2)
where µ
1
and µ
2
are growth rates determined at temperatures t
1
and t
2
,
respectively.
Most laboratory studies involving cultured heterotrophic protists have focused on
species isolated from temperate environments and have examined protistan physiology at
relatively high water temperatures (>15˚C). Thus, the extrapolation of physiological rate
information obtained from temperate cultures to infer rate processes of protists in
permanently cold environments may be inappropriate. To date, three published studies
have examined growth rates and/or gross growth efficiencies (GGE, defined as the
amount of protistan carbon or biovolume produced relative to the amount of prey
consumed) of cultured heterotrophic protists from permanently cold environments. All
three studies reported low growth rates of polar isolates at appropriate environmental
temperatures. However, these studies reported conflicting results when growth rates of
polar isolates were compared to growth rates of temperate congeners at low
environmental temperature.
Lee and Fenchel (Lee & Fenchel 1972) reported a growth rate of 0.16 d
-1
for the
Antarctic ciliate Euplotes antarcticus at –2˚C. The growth rate of this species increased
exponentially with increasing temperature although growth was not sustained above
10˚C. The Q
10
values from –1.8-2˚C and 2-5˚C were 4.8 and 4.6, respectively. The
growth rates of this Antarctic species were comparable to those of temperate and tropical
congeners when measured at similar temperatures (5 and 10˚C). In contrast, an Arctic
strain of P. imperforata examined by Choi and Peters (1992) grew more rapidly at –1.5
and 6˚C than a subarctic conspecific examined in the same study, but more slowly than
72
the subarctic conspecific at 15˚C. The authors interpreted these results as indicative of
adaptation by the Arctic P. imperforata to low environmental temperature. Mayes et al
(1997) reported growth rates for five species of Antarctic naked marine amoebae over the
temperature range –2 to 4˚C. The observed rates were significantly less than growth rates
predicted for ambient Antarctic temperatures by the extrapolation of available growth
rate information for temperate amoebae. In contrast to the conclusion of Choi and Peters,
Mayes et al hypothesized that the disparity between the observed growth rates of the
Antarctic amoebae and rates predicted from the extrapolation of growth from temperate
amoebae was due to the energetic costs of inhabiting cold environments.
In general, the effects of temperature on protistan GGE are not well understood.
Some studies have reported increases in GGE with decreasing temperature
(Rassoulzadegan 1982, Verity 1985, Choi & Peters 1992), while others have reported
decreases with decreasing temperature, or very low GGE at extreme low temperature
(Laybourn & Stewart 1975, Rogerson 1981, Mayes et al. 1997). Still others have
reported no effect of temperature on GGE (Caron et al. 1986, Ishigaki & Sleigh 2001).
Choi and Peters (1992) reported high volume-based GGEs (60 and 71%) for an Arctic
and a subarctic P. imperforata measured at –1.5˚C. In contrast, Mayes et al (1997)
reported exceptionally low GGEs for Antarctic amoebae over the temperature range –2 to
4˚C, (GGE = 0.8-7.2%).
Food quality is also recognized as an important factor controlling protistan
physiological processes including growth rate and GGE. These parameters may respond
to a wide range of characteristics including size, swimming speed, presence of large
73
amounts of undigestible compounds in prey biomass, or limiting quantities of elements or
compounds essential to heterotrophic protistan metabolism (Sherr et al. 1983, Gonzalez
et al. 1990, Caron et al. 1991, Matz et al. 2002, Mohapatra & Fukami 2004).
This study is the first to document growth characteristics of a cultured marine
Antarctic heterotrophic nanoflagellate. P. imperforata was selected as a study organism
due to its wide geographic distribution, its potential importance as a source of bacterial
mortality in aquatic ecosystems, and published laboratory studies detailing the growth
and grazing rates, growth efficiencies and nutrient remineralization efficiencies of this
and closely related species (Fenchel 1982, Sherr et al. 1983, Goldman & Caron 1985,
Caron et al. 1986, Choi & Peters 1992, Eccleston-Parry & Leadbeater 1995). Growth
rates observed for the Antarctic P. imperforata were consistent with maximum published
values for temperate congeners between 0 and 15˚C. Neither temperature nor bacterial
prey type significantly affected GGE of this flagellate, in contrast to considerable
variability in this parameter published for conspecific and congeneric flagellates. We
conclude that temperature is not a major factor controlling GGE of Paraphysomonas
species (i.e. much of the variability in published values for growth efficiency appears to
be due to specific methodological approaches). In addition, the relationship between
temperature and the maximal growth rate of all Paraphysomonas species is predictable.
In contrast to previous conclusions, our results indicate that there is no clear evidence for
strong adaptation of Paraphysomonas species/strains to environmental temperature.

74
Methods
Cultures. P. imperforata (clone RS-4-2) was isolated from an enriched water sample
obtained from the Ross Sea, Antarctica (76˚01.38’ S, 165˚24.46’ W). Water was
collected and the natural bacterial flora was enriched with sterilized yeast extract and rice
grains. Enrichment cultures were maintained at temperatures 0-2˚C at all times to avoid
selection against heat-sensitive protists. P. imperforata was isolated by dilution
extinction, and clonal (uniprotistan) cultures were identified based on the structure of
scales on the surface of the flagellate according to Preisig and Hibberd (1982) using
transmission electron microscopy (Fig 1). Two unidentified bacterial strains were
isolated from an enriched water sample also obtained from the Ross Sea, Antarctica
(76˚30’ S, 169˚33’ E). Enriched water was plated onto marine agar (DIFCO
Laboratories, Sparks, MD) and single colonies were picked and streaked repeatedly on
marine agar plates to ensure purity. A third bacterial strain, the temperate heterotrophic
marine bacterium Halomonas halodurans (Pseudomonas halodurans) was also used as
prey (originally obtained from Russel Cuhel, Great Lakes Water Institute).

Growth experiments. Two growth experiments were conducted with the Antarctic P.
imperforata. The first experiment employed three strains of bacteria (H. halodurans and
the two unidentified Antarctic bacteria) fed to P. imperforata at 0˚C, and a single strain
of Antarctic bacteria (Strain A) fed to the flagellate at 5 and 10˚C. A second experiment
was conducted using a single strain of Antarctic bacteria (Strain A) to measure growth at
0, 5, 10, 15 and 20˚C. P. imperforata cultures in both experiments were slowly
75
A
B
Figure 3-1. Transmission electron micrograph of Paraphysomonas imperforata (negative stain
using 1% uranyl acetate). (A) Whole cell view showing flagellation and surface scales, 5,000x.
Marker bar = 2um. (B) High magnification showing scale morphology, 27,000x. Marker bar = 0.5
um.
76
acclimated to experimental temperatures above 2˚C by gradually raising the culture
temperature 3-5˚C, and maintaining the culture at that temperature for a period of several
weeks before further increasing the temperature. Acclimatization therefore was
conducted over a period of several months prior to the experiments.
Bacteria in both experiments were grown to late stationary growth phase on
marine broth (DIFCO Laboratories, Sparks, MD), harvested by centrifugation (3220 x g
for 20 minutes), rinsed, resuspended and recentrifuged three times with sterile seawater to
remove residual organic medium. The two strains of Antarctic bacteria were grown at
5˚C, and H. halodurans was grown at 20˚C. Rinsed bacteria were inoculated into 600 ml
sterile 0.2 µm filtered seawater in 1.25 L polycarbonate bottles at a starting concentration
of ~10
7
cells ml
-1
and brought to experimental temperatures before beginning both
experiments. Protists were grown to high abundance prior to the experiments (at their
respective experimental temperatures) and added to the bacterized seawater at 1-5x10
3

cells ml
-1
to minimize bacteria and dissolved organics carried over with the protistan
inoculum. Protist-free bacterized seawater was used as a control for each temperature
and each prey type to monitor changes in prey populations not associated with protistan
grazing activities. All experimental treatments were performed in triplicate, and controls
were performed in duplicate. Experiments were conducted in incubators set to the
appropriate temperatures. All incubations were conducted in continuous darkness.
Sampling was performed on ice in a cold room for cultures incubated at 0˚C, in a 5˚C
cold room for cultures incubated at 5 and 10˚C, and in a 15˚C walk-in incubator for the
cultures at 15 and 20˚C in order to avoid temperature shock during the experiments.
77

Measurements and calculations. Bacterial abundances were measured by flow
cytometry on samples preserved in 1% formaldehyde and frozen at –20˚C until analysis
(FACScalibur flow cytometer, BD Biosciences, San Jose, CA). Samples were stained in
the dark for 15 minutes with 2.5 µM SYTO 13 (Molecular Probes, Eugene, OR) and
analyzed on the flow cytometer according to the protocol of del Giorgio et al (del Giorgio
et al. 1996). Event counts were converted to cell abundances by weighing sample tubes
on an analytical balance before and after each analysis on the flow cytometer to
determine the volume analyzed.
Protistan abundances were measured by light microscopy of samples preserved in
2% acid Lugols solution (final concentration) and kept in the dark at room temperature
until analysis. Protists were enumerated using a Palmer-Maloney counting chamber, and
at least 200 cells were counted per slide. Protistan growth rates were calculated based on
a regression of natural log cell abundance vs. time while cultures were in logarithmic
growth phase (linear portion of the growth curve).
Protistan GGEs in the first experiment were calculated in two ways: based on
changes in particulate carbon in the cultures, and based on bacterial volume consumed
relative to protistan volume produced. In the second experiment, protistan GGEs were
calculated based on volume only. For carbon-based estimates of GGE, 20 ml of sample
was filtered onto a precombusted GF/F filter (Gelman
®
) and frozen at –20˚C until
analysis for particulate organic carbon (analysis conducted by the MSI Analytical Lab,
Santa Barbara, CA). Gross growth efficiency was calculated by comparing the
78
particulate carbon concentration in each culture at the first sampling time showing the
maximum P. imperforata abundance (end of log phase growth) to initial particulate
carbon concentration, and assuming the initial particulate carbon from P. imperforata
was negligible.
Volume-based estimates of GGE were completed on samples preserved with 1%
glutaraldehyde (final concentration) and stored at 5˚C until analysis. Digital photographs
of protists and bacteria were taken using phase contrast microscopy and an inverted
microscope (Leica DM IRB) at 1000x magnification. Measurements of protistan
diameter and bacterial length and width were performed on the digital photographs using
the software program Openlab 3.5.1. Volumes of protists and bacteria were calculated
using standard equations for spheres and cylinders, respectively. The volumes of at least
25 protists and 50 bacteria from each sample were averaged for use in GGE calculations.
The initial total bacterial volume (abundance x cell volume) was compared to the total
bacterial volume at the time of maximum P. imperforata abundance to estimate the
volume of bacteria consumed. This value was compared to total protistan volume
produced to calculate volume-based gross GGE.

Statistics. P. imperforata growth rates and GGEs were analyzed with a percentile
bootstrap method for comparing means of multiple groups, using the statistical software
package R (Liu & Singh 1997, Wilcox 2003). The percentile bootstrap method was
selected over the commonly used ANOVA since it has the advantage of no assumptions
of equality of variance or normal distribution. If a significant p value was obtained for
79
this global comparison, a multiple comparison test was performed to identify significant
differences among pairs of growth rates or GGEs within experiments. The multiple
comparison test used here consists of a series of percentile bootstrap tests of each pair, in
conjunction with a sequentially rejective method derived by Hochberg to control
familywise error rate (Hochberg 1988, Wilcox 2003).
Results
Experiment #1: Growth rates of P. imperforata increased significantly with increasing
temperature (all p values <0.01), (Fig 2a-c; f). Growth rates increased from 0.61 d
-1
at
0˚C to 1.6 d
-1
at 5˚C and 2.6 d
-1
at 10˚C. Q
10
values determined from these growth rates
were 6.9 between 0 and 5˚C, 2.6 between 5 and 10˚C, and 4.3 between 0 and 10˚C.
Growth rates of P. imperforata at 0˚C were not significantly affected by the prey types
examined (p=0.62), (Fig 2c-e).
Gross growth efficiency of the flagellate was relatively recalcitrant to different
culture temperatures and prey types, although volume-based GGEs were consistently
lower than those based on particulate carbon (means=36% and 46%, respectively). GGE
was not significantly affected by temperature in the first experiment, regardless of
whether efficiencies were calculated based on estimates of carbon or volume (p=0.30 and
p=0.92, respectively; range 36-45%), (Fig 3). Gross growth efficiencies based on carbon
were not significantly different among the three prey types (p=0.33), and only one pair of
GGEs calculated based on volume measurements were significantly different among the
three prey types (Strain B and H. halodurans; p<0.01). Volume-based GGEs ranged
from 28% when P. imperforata was grown at 0˚C on Antarctic bacterial strain B, to 42%
80
6
8
10
12
14
16
18
20
0 1 2 3 4 5
Growth Rate = 2.6 d
-1
R
2
= 0.99
6
8
10
12
14
16
18
20
0 1 2 3 4 5 6 7
Growth Rate = 1.6 d
-1
R
2
= 0.99
6
8
10
12
14
16
18
20
0 2 4 6 8 10 12 14 16
Growth Rate = 0.61 d
-1
R
2
= 0.99
6
8
10
12
14
16
18
20
0 2 4 6 8 10 12 14 16
Growth Rate = 0.53 d
-1
R
2
= 0.99
6
8
10
12
14
16
18
20
0 2 4 6 8 10 12 14 16
Growth Rate = 0.52 d
-1
R
2
= 0.98
6
8
10
12
14
16
18
20
0 2 4 6 8 10 12 14 16
Time (days)
A
B
C D
E F
Figure 3-2. Growth dynamics of an Antarctic strain of P. imperforata (filled symbols) and removal of
bacterial prey (open symbols) at three temperatures and on three strains of bacteria (A-E): Antarctic bacterial
strain A as food at 10
o
C (A), 5
o
C (B) and 0
o
C (C); Antarctic bacterial strain B at 0
o
C (D);

H. halodurans at
0
o
C (E). Composite of information from (A)-(C) comparing the growth of P. imperforata at 10
o
C (■), 5
o
C
(▲), and 0
o
C (●) (F)
81
0
10
20
30
40
50
60
70
80
90
100
0
10
20
30
40
50
60
70
80
90
100
Volume Carbon
Bacteria A
10
o
C
Bacteria B
0
o
C
Bacteria A
5
o
C
Bacteria A
0
o
C
H. halodurans
0
o
C
Figure 3-3. Gross growth efficiencies of the Antarctic strain of P. imperforata based on
measurements of cell volume (solid) and carbon (hatched). Protists were growth at three
temperatures, and on three strains of bacteria, as indicated. Error bars represent one standard
deviation.
82
when the flagellate was grown at 0˚C on Halomonas halodurans. Carbon-based GGEs
ranged from 42% when P. imperforata was grown at 0˚C on Antarctic bacterial strain A
to 54% when the flagellate was grown at 0˚C on H. halodurans.

Experiment #2: Growth rates of P. imperforata in experiment #2 were examined when
the flagellate was fed Antarctic bacterial strain A over a wider temperature range than in
experiment #1. Flagellate growth rates increased significantly with increasing
temperature up to 15˚C, but did not increase further at 20˚C (p<0.01 for all pairs except
for 15˚C vs. 20˚C, which were not significantly different; Fig 4A). The Q
10
values for
growth rates calculated at 5˚ intervals between 0˚ and 20˚C in experiment #2 were 15,
2.2, 3.6 and 0.93. Gross growth efficiency was not consistently affected by increasing
temperature (Fig 4B). Only two pairs of GGEs were statistically different (0˚C vs. 10˚C
and 0˚C vs. 15˚C, p<0.01 for both; all other p values were >0.05). Thus, GGE observed
at 0˚C was not significantly different from the GGE observed at 5 or 20˚C. Overall,
GGEs in experiment #2 ranged from 31% at 0˚C to 53% at 15˚C based on volume.

Comparison with published data: A literature review was compiled of growth rates
and GGEs for Paraphysomonas spp. in order to compare with the information obtained in
this study. 131 growth rates from 14 studies (Fig 5A, open symbols) and 55 GGEs from
four studies (Fig 5B, open symbols) were compared to data reported in this study (Fig 5A
and B, closed symbols). Temperature had a strong effect on the maximal growth rates
reported for Paraphysomonas spp.. Growth rates obtained in this study were similar to
83
Temperature (
o
C)
0
1
2
3
4
5
0 5 10 15 20
0
20
40
60
80
100
0 5 10 15 20
A
B
Figure 3-4. Relationship between temperature and growth rate (A) or gross
growth efficiency (B) for an Antarctic strain of P. imperforata. Error bars
represent one standard deviation.
84
0
1
2
3
4
5
6
-5 0 5 10 15 20 25 30
0
20
40
60
80
100
-5 0 5 10 15 20 25 30
Temperature (
o
C)
A
B
Figure 3-5. Relationship between temperature and growth rate (A) or gross growth efficiency
(B) for strains of Paraphysomonas spp. observed in this study ( ), and in published reports for
temperate and polar strains ( ). Solid lines in (B) indicate mean values and dashed lines indicate
median values for each 5
o
C temperature interval.
85
rates reported for an arctic strain of P. imperforata at low temperatures ( ≤6˚C). Growth
rates obtained at 5, 10 and 15˚C in this study were higher than those reported for
temperate Paraphysomonas spp., while growth rates at the highest temperature examined
in this study (20˚C) were less than maximal rates observed for Paraphysomonas species
in other studies but within the range of rates reported for temperate species.
Gross growth efficiencies summarized for two Paraphysomonas species (P.
bandaiensis and P. imperforata) spanned a very wide range (Fig 5B; 2-80%). Values
obtained for P. imperforata in this study were considerably more constrained (28-54%)
and in general agreement with the overall mean and/or median values for GGEs across
the temperature range –1.5-26˚C (overall mean = 34%; overall median = 32%).
Averaged over the temperature intervals <5˚, ≥5˚<10˚, ≥10<15˚C, ≥15<20˚, ≥20˚, the
range of median and mean GGEs (25-44% for median and 29-50% for mean GGE) were
quite similar to the narrow range of GGEs obtained in the present study.
Discussion
Studies detailing the effects of temperature on the growth rates of heterotrophic
protists are numerous. A wide range of values for this parameter has been published,
particularly for taxa that are believed to be of cosmopolitan distribution and considerable
ecological significance such as species of the nanoflagellate genus Paraphysomonas (Fig
5A) insert Finlay reference and other Paraphysomonas papers, re-cite refs from p5.
Collectively, these data indicate that temperature sets an upper limit on the growth rates
of Paraphysomonas spp. (maximal growth rates are directly related to temperature).
While it is probable that long-term exposure of conspecifics to different climatic regions
86
has resulted in the evolution of unique biochemical and physiological adaptations to
allow existence in different temperature regimes, there is little direct evidence that these
adaptations can result in rapid growth of protists at low temperature.
Choi and Peters (Choi & Peters 1992) compared the growth rates of arctic and
subarctic strains of P. imperforata at three temperatures between –1.5 and 15
o
C. These
authors reported that the arctic P. imperforata was capable of faster growth at –1.5˚C
than its subarctic conspecific (0.77 and 0.53, respectively). The authors attributed these
differences to adaptation of the arctic strain to extreme low temperature. These minor
intraspecific variations aside, there appears to be a generalized temperature constraint on
growth rate of these species (Fig 5A). This relationship is not unlike the classical
relationship between temperature and growth rate reported for phytoplankton by Eppley
(Eppley 1972) and recently reviewed by Rose and Caron (submitted).
The Antarctic P. imperforata examined in this study was capable of long-term
growth at all temperatures examined (Fig 5A). Growth rate increased with increasing
temperature up to 15˚C, but no further increase was observed at 20˚C (Fig 4A). Q
10

values were relatively high at low temperatures and decreased as temperature increased
within the range examined. Q
10
values considerably higher than 2 are typically observed
at temperatures below optimal, and may be indicative of increased energy barriers to the
biochemical processes involved in determining growth rates (Hochachka & Somero
2002). The Q
10
values obtained in this study between 0 and 5˚C (6.9, experiment 1; 15,
experiment 2) were greater than all other values observed between 5 and 20˚C
(experiment 1: 2.6; experiment 2: 2.2, 3.6 and 0.93). The high Q
10
values observed
87
between 0 and 5˚C imply that our Antarctic P. imperforata was growing outside its
optimal physiological range at ambient Antarctic temperatures. The Antarctic strain was
acclimated to experimental temperatures slowly over a period of several months, thus is
unlikely that the high Q
10
values were a result of an overly rapid rate of acclimation.
We believe that the response of growth rate of our isolate of P. imperforata to
temperature is representative of this species in Antarctic marine ecosystems. The
isolation and temperature acclimation of the strain of P. imperforata used in this study
was done with utmost care to prevent heat shock and selection against heat-sensitive
clones. Accordingly, the growth rates observed in this study were among the highest
rates reported over the temperature range –1.5-15˚C. The growth rates of this isolate
were comparable to maximal rates for arctic and subarctic conspecifics of P. imperforata
at low temperature (-1.5 to 6˚C; (Choi & Peters 1992)) and are among the highest rates
reported at 10 and 15˚C for any species within the genus Paraphysomonas (Fig 5A),
although it is worth noting that available data are limited within this temperature range
relative to higher temperatures. In contrast with very high growth rates observed from 0-
15˚C for the Antarctic P. imperforata, our isolate has a lower maximal growth rate at
20˚C relative to the maximal growth rates reported for other Paraphysomonas spp. (solid
circles vs. open triangles in Fig 5A). We speculate that the inability of our Antarctic
strain of P. imperforata to grow as rapidly as some conspecific and congeneric strains at
20˚C may reflect a physiological compromise that enables its growth at extreme low
temperature.
88
Prey type had no significant effect on growth rate of the Antarctic P. imperforata
in this study (Fig 3). H. halodurans, a temperate marine bacterium grown at 20˚C in this
experiment, was apparently as good of a food source for the Antarctic P. imperforata
growing at 0˚C as were the two strains of Antarctic bacteria grown at 5˚C. Growth
temperature can affect the biochemical composition of bacteria, in particular the types,
classes and species of lipids present (Hazel 1995, Hochachka & Somero 2002). It is
therefore possible that bacteria isolated from the permanently cold waters of the Antarctic
could differ biochemically from temperate marine bacteria or Antarctic bacteria grown at
warmer temperatures. If such differences existed in our study, they did not affect growth
rates of our Antarctic strain of P. imperforata at low temperature.
While temperature exerts a significant effect on maximal growth rates of protists,
its effect on GGE is less predictable. Experimental data indicate that GGE may increase,
decrease, or remain unchanged with changing temperature. Choi and Peters (Choi &
Peters 1992) reported very high GGEs (60-70%) for both strains of P. imperforata at low
temperature, due to an apparent increase in cell volume with decreasing temperature. In
contrast, low GGEs (0.8-7.2%) have been reported for several species of Antarctic
amoebae grown between -2 and +4˚C (Mayes et al. 1997). The latter authors
hypothesized that the extremely low GGEs for the Antarctic amoebae were due to
energetic costs from inhabiting a permanently cold environment.
Gross growth efficiencies measured in this study yielded a relatively narrow range
of values which did not vary consistently with temperature. Overall averages were 39%
based on volume and 46% based on carbon. Such minor differences between these
89
methods for calculating GGE have been noted previously and are a consequence of the
different assumptions inherent in the two approaches (Caron et al. 1986).
Some of the treatments in experiment 1 were duplicated in experiment 2 in this
study and allow comparisons between the same treatments examined at different times.
Individual GGEs compared at 0 and 5˚C between the two experiments yielded no
significant differences (p=0.48 and 0.37, respectively), and differences between GGEs
measured at 10˚C for the two experiments were only slightly significantly different
(p=0.01). While the range of reported values of GGE for two species of
Paraphysomonas was quite large from a single study (2-80%; (Selph et al. 2003)), the
mean and median values for each 5˚C increment of temperature were constrained to a
relatively narrow range (29-50% for mean, 25-44% for median; solid and dashed lines
respectively in Fig 5B). Selph et al (Selph et al. 2003) reported extremely low GGEs at
18˚C, which brought the mean and median estimates down in the ≥15-20˚C range,
otherwise the range of mean and median values would have been even narrower: 33-49%
and 40-51%, respectively. The mean and median values of GGE thus show no apparent
relationship with temperature.
Given the narrow range of GGEs measured in this study, and the agreement of our values
with mean GGEs reported for two species of Paraphysomonas, it is not clear why the
range of GGEs in the literature was so wide (2-80%, Fig 5B). Factors such as poor food
quality or rapid acclimation of cultures to experimental temperatures might explain
reports of low values, but do not satisfactorily explain some high GGEs that have been
reported. Some differences in GGE are to be expected due to inherent differences in the
90
calculation of efficiencies. Carbon-based GGE assumes that protists make an
insignificant contribution to particulate carbon at the beginning of an experiment (due to
relatively high bacterial abundances). This assumption allows the calculation of carbon
content in a single bacterium, which is then used as a constant in the calculation of GGE.
The use of a constant assumes that average carbon content per bacterium does not change
over the course of an experiment. This method also assumes that all particulate carbon
can be attributed to either protistan or bacterial cells, and that egested particulate carbon
does not make a contribute significantly to total carbon. If the average carbon content of
bacteria decreases over the course of an experiment (due to starvation), carbon-based
GGE could underestimate actual GGE. On the other hand, if egested particulate carbon
comprises a significant portion of the total, carbon-based GGE could overestimate actual
protistan GGE.
Volume-based GGE takes into account changes in bacterial volume at the
beginning and end of an experiment and excludes egested particulate carbon from the
calculation. Volume-based GGEs can underestimate actual GGE, though, due to greater
loss of protistan cell volume than bacterial cell volume from cell shrinkage upon
preservation and preparation for microscopy. Shrinkage due to preservation can be
prevented through the analysis of live samples. Estimates of cell volume (and GGEs)
have been reported based on measurements of live cultures of protists and bacteria by
flow cytometry (Choi & Peters 1992), but the accurate conversion of measurements of
forward scatter to cell size using reference beads is difficult, since the surface of the
reference beads will scatter light differently than the surface of a cell.
91
Given the vagaries of these different calculations of GGE, and inherent variability
in efficiency due to food quality, the variability observed in this parameter is not
surprising (Fig 5B). Despite this variability, our analysis of mean and median values
indicated little to no effect of temperature on GGE.
Conclusions
This study provides some of the first information on growth of a cold-adapted
nanoflagellate at extreme low temperature (0˚C). Our Antarctic strain of P. imperforata
grew slowly but efficiently at this temperature but grew much more rapidly (with the
same efficiency) at higher temperatures. The relationship between temperature and GGE
of heterotrophic protists has been debated in the past. Gross growth efficiency values
compiled here indicate there was no relationship between temperature and GGE for two
species of Paraphysomonas. In addition, the wide variance in GGE reported in the
literature demonstrates that the introduction of artifact into calculations of GGE due to
methodological issues can be significant.

92
Chapter 4: Effect of temperature and prey type on nutrient
regeneration by an Antarctic bacterivorous protist

Introduction
Remineralization of major nutrients (nitrogen/phosphorus) via the grazing
activities of phagotrophic protists is an important source of nutrients fueling
phytoplankton growth in the ocean (Strom 2000, Sherr & Sherr 2002). Dissolved
inorganic and organic nutrients and particulate material not incorporated into protistan
biomass are excreted after feeding on bacteria, phytoplankton or other heterotrophic
protists, and recycled back to primary producers. Protistan trophic activities can be
important contributors to nutrient remineralization given that heterotrophic bacteria may
at times compete with phytoplankton for dissolved inorganic nutrients, and thus may be a
sink rather than a source for these substances (Thingstad et al. 1993, Cotner et al. 1997,
Caron et al. 2000). Therefore, characterizing the mechanism of protistan nutrient
remineralization and the factors controlling this process are fundamental aspects for
understanding nutrient dynamics in marine ecosystems.
Heterotrophic protists excrete a variety of forms of dissolved nitrogen and
phosphorus, but ammonium and phosphate (soluble reactive phosphorus, SRP) are the
dominant forms released by most species (Goldman et al. 1985, Andersen et al. 1986,
Caron & Goldman 1990, Dolan 1997). Concentrations of dissolved organic phosphorus
(largely uncharacterized) and a variety of dissolved organic nitrogen compounds (e.g.
amino acids, urea, uric acid, and small quantities of hypoxanthine, dihydrouracil, adenine
93
and guanine have been detected) are significant but secondary in absolute amount to the
inorganic forms (Soldo & Wagtendonk 1961, Leboy et al. 1964, Soldo et al. 1978,
Nagata & Kirchman 1991, 1992). Similarly, the release of undigested particulate
material by protists constitutes a significant albeit minor porportion of the total amount of
ingested prey biomass (Caron et al. 1985).
Laboratory experiments with heterotrophic protists have demonstrated their
ability to remineralize a substantial portion of the phosphorus and nitrogen ingested as
prey, especially when feeding on bacteria (Sherr et al. 1983, Goldman et al. 1985,
Andersen et al. 1986, Caron et al. 1986, Caron et al. 1990, Dolan 1997). The latter
outcome is based on the fact that the C:N and C:P ratios of bacteria are generally much
lower than those of heterotrophic protistan biomass. Thus, bacterial prey are rich in
nitrogen and phosphorus relative to the nutritional needs of their consumers, and
bacterivorous protists maintain appropriate intracellular C:N:P ratios by excreting the
excess nitrogen and phosphorus (Caron 1990). Laboratory experiments have
demonstrated the maintenance of intracellular stoichiometric balance in heterotrophic
protists through excretion of excess N or P in P- or N-limited bacteria and diatoms
(Goldman et al. 1985, Andersen et al. 1986).
While the percentage of nitrogen and phosphorus excreted by phagotrophic
protists appears to be dependent on elemental stoichiometry of the consumer and prey, in
general, weight-specific excretion rates of zooplankton have been shown to be inversely
related to organismal size (Hargrave & Geen 1968, Ikeda et al. 1982). The relatively
small size of heterotrophic protists implies a potential for extremely high weight-specific
94
excretion rates in comparison to metazoa. Metazoan-based equations have been
extrapolated to the size range of heterotrophic protists, and compared to excretion rate
information from laboratory studies of cultures of heterotrophic protists (Caron &
Goldman 1990, Dolan 1997). These analyses have found higher maximal phosphorus
excretion rates but comparable maximal nitrogen excretion rates compared to rates
predicted by the metazoan-based equations. These results indicate that are protists
capable of some of the highest weight-specific excretion rates, and that heterotrophic
protists may play a particularly important role in phosphorus remineralization in some
aquatic ecosystems.
Most experimental studies of nutrient remineralization by heterotrophic protists
carried out to date have been performed with temperate isolates cultured at relatively
warm environmental temperatures (15-25˚C). One exception was the study by Sherr et al
(1983) that examined ammonium excretion by a temperate strain of Monas sp. at four
temperatures (range = 3-30˚C. These authors observed lower gross growth efficiencies
(based on carbon) and higher ammonium excretion rates at the two temperature extremes
compared to efficiencies and rates observed at intermediate temperatures of 18 and
23.5˚C. However, they noted that the lowest and highest temperatures were clearly
outside the optimal physiological temperature range for the isolate. To my knowledge,
no other study has examined protistan nutrient remineralization of protistan strains
isolated at ambient polar temperatures and studied that these low temperatures.
Therefore, it is not clear whether the extrapolation of excretion rates obtained for
temperate heterotrophic protists is appropriate to describe the excretion rates of
95
heterotrophic protists from permanently cold environments growing at environmentally
pertinent temperatures.
This study is the first to examine the effect of temperature (including
environmentally appropriate temperatures) and prey type on nutrient excretion rates and
remineralization efficiencies of a polar heterotrophic nanoflagellate. Paraphysomonas
imperforata was selected as the study organism because it has a global distribution, it is
an important bacterial consumer in aquatic ecosystems, and temperate conspecific strains
of this species have been extensively studied. My results indicated no effect of prey type
on either nutrient remineralization rates or total nutrient remineralization. Higher rates of
nutrient remineralization by the nanoflagellate were observed at higher temperatures, but
total nutrient remineralization was unaffected by growth over a range of temperatures.
Total nutrient remineralization in these experiments was comparable to values published
for temperate conspecifics at much higher temperatures.
Methods
Cultures. P. imperforata (clone RS-4-2) was isolated from an enriched water sample
obtained from the Ross Sea, Antarctica (76˚01.383’ S, 165˚24.459’ W). Water was
collected and the natural bacterial flora was enriched with sterilized yeast extract and rice
grains. Cultures were maintained at temperatures below 2˚C at all times to avoid
selection against heat-sensitive protists. P. imperforata was isolated by dilution
extinction, and clonal (uniprotistan) cultures were identified based on the structure of
scales on the surface of the flagellate according to Preisig and Hibberd (Preisig &
Hibberd 1982) using transmission electron microscopy. Two unidentified bacterial
96
strains were isolated from an enriched water sample also obtained from the Ross Sea,
Antarctica (76˚30 S, 169˚33 E). Enriched water was plated onto marine agar (DIFCO
Laboratories, Sparks, MD) and single colonies were picked and streaked repeatedly on
marine agar plates to ensure purity. A temperate bacterial strain, Halomonas halodurans,
was also used in the nutrient regeneration experiments.

Nutrient regeneration experiments. Subcultures of the Antarctic P. imperforata were
acclimated to three temperatures (0, 5 and 10˚C) for several months prior to the
beginning of experiments. Subcultures of the Antarctic P. imperforata were transferred
at high protistan and low bacterial density into pure cultures of the three bacterial strains
for several transfers prior to the beginning of experiments to eliminate other strains of
bacterial prey from the cultures.
All bacterial strains were grown to late stationary growth phase on marine broth
(DIFCO Laboratories, Sparks, MD), harvested by centrifugation, rinsed and resuspended
several times in sterile seawater to remove residual organics from the marine broth. The
two strains of Antarctic bacteria were grown at 5˚C and Halomonas halodurans was
grown at 20˚C. Bacteria were then added to 600 ml sterile seawater at a starting
concentration of approximately 10
7
cells ml
-1
and flasks containing bacterized seawater
were brought to the appropriate experimental temperatures in separate temperature-
controlled incubators.
Protists were grown to high abundance and inoculated into each of the
experimental treatments at low starting abundance (~10
3
cells ml
-1
), to minimize the
97
organic material and bacteria carried over from the inoculum. Protist-free controls of
bacterized seawater were used in duplicate at each temperature and each prey type to
monitor changes in nutrient concentration not associated with protistan grazing activity.
Experimental treatments were performed in triplicate. Experimental and control
treatments were kept in continuous darkness. The 5˚C and 10˚C treatments were sampled
in a 5˚C cold room and the 0˚C treatment was sampled on ice in the cold room to avoid
heat-shocking experimental organisms.

Sample analysis. Samples were periodically removed and fixed with 2% lugols solution
for enumeration of protists by light microscopy using a Palmer Maloney counting
chamber. Samples were also fixed with 1% formalin for enumeration of bacteria by flow
cytometry according to the protocol of del Giorgio et al (1996).
Samples of 20 ml volume were collected onto pre-combusted GF/F glass fiber
filters (Gelman) for determination of particulate carbon and nitrogen. Samples were kept
frozen at –20˚C until analysis by the MSI Analytical Lab, Santa Barbara, CA. The
filtrate was collected and kept frozen at –20˚C for analysis of NH
4
concentration
according to Parsons et al (1984). Samples of 10 ml volume were collected onto pre-
combusted GF/F glass fiber filters for analysis of particulate phosphorus according to
Menzel and Corwin (1965). The filtrate was also collected for analysis of soluble
reactive phosphorus according to Parsons et al (1984) and total dissolved phosphorus
according to Menzel and Corwin (1965). Both filters and filtrate were kept frozen at –
20˚C until analysis. Dissolved organic phosphorus concentrations were calculated for
98
each sample by subtracting the soluble reactive phosphorus from the total dissolved
phosphorus concentration.
Results
Particulate carbon (PC) concentrations in all five treatments decreased during the
period when P. imperforata was actively growing and bacteria were being actively
consumed (Fig 1). PC concentrations decreased more rapidly at higher temperature (Fig
1C) but the loss rate was unaffected by prey type (Fig 1D). The concentrations of PC at
the time of peak abundance of P. imperforata were similar regardless of incubation
temperature or prey type (Table 1).
Particulate nitrogen (PN) concentrations also decreased during bacterial grazing
by P. imperforata in all five treatments (Fig 2, solid symbols). Increases in ammonium
concentrations in each treatment lagged behind increases in protistan abundance (Fig 2,
open symbols). The rates of PN loss and ammonium regeneration increased with
increasing temperature but were unaffected by prey type (Fig 2F). The total amount of
ammonium regenerated was similar at all temperatures and prey types. Final ammonium
concentrations in all treatments ranged from 118 to 148 µM.
Changes in particulate and dissolved phosphorus constituents mirrored those of
nitrogen constituents (Fig 3). After an initial lag, particulate phosphorus (PP)
concentrations decreased (Fig 3, solid symbols) and dissolved phosphorus concentrations
increased (Fig 3, open symbols). The dissolved phosphorus constituents primarily
consisted of soluble reactive phosphorus (SRP, open circles), with dissolved organic
phosphorus (DOP, open triangles) comprising 10-20% of the total dissolved phosphorus
99
6
7
8
9
10
11
12
13
14
6
7
8
9
10
11
12
13
14
A B
0
200
400
600
800
1000
1200
0 2 4 6 8 10 12 14 16
0
200
400
600
800
1000
1200
0 2 4 6 8 10 12 14 16
C D
Time (days)
Figure 4-1. Changes in protistan abundance (A,B) and particulate carbon concentrations (C,D) in cultures of
Paraphysomonas imperforata growth at three temperatures (A,C) and on three different strains of bacteria at
a single temperature (B,D). Symbols indicate particulate carbon concentration (C,D) and lines indicate
protistan abundance (A,B). (A,C) 0
o
C, Antarctic bacterial strain A (square, solid line); 5
o
C, Antarctic
bacterial strain A (circle, dashed line); 10
o
C, Antarctic bacterial strain A (triangle, dotted line). (B,D) 0
o
C,
Antarctic bacterial strain A (square, solid line); 0
o
C Antarctic bacterial strain B (diamond, dashed line); 0
o
C,
Halomonas halodurans (inverted triangle, dotted line)
100
0
50
100
150
200
250
0 2 4 6 8 10 12 14 16
0
50
100
150
200
250
0 1 2 3 4 5 6 7
0
50
100
150
200
250
0 1 2 3 4 5
0
50
100
150
200
250
0 2 4 6 8 10 12 14 16
0
50
100
150
200
250
0 2 4 6 8 10 12 14 16
0
25
50
75
100
125
150
0 2 4 6 8 10 12 14 16
Time (days)
D
E
F
A B
C
F
Figure 4-2. Changes in particulate nitrogen (solid symbols) and ammonium (open symbols) during
bacterial grazing by Paraphysomonas imperforata on Antarctic bacterial strain A at 0
o
C (A), 5
o
C (B) and
10
o
C (C), and at 0
o
C on Antarctic bacterial strain B (D) and Halomonas halodurans (E). Changes in
ammonium concentration in all treatments (F). Note different ranges for x and y axes in (A-F).
101
0
3
6
9
12
15
18
0 2 4 6 8 10 12 14 16
0
3
6
9
12
15
18
0 1 2 3 4 5 6 7
0
3
6
9
12
15
18
0 1 2 3 4 5
0
3
6
9
12
15
18
0 2 4 6 8 10 12 14 16
0 2 4 6 8 10 12 14 16
0
3
6
9
12
15
18
0 2 4 6 8 10 12 14 16
Time (days)
A B
C D
E F
Figure 4-3. Changes in particulate phosphorus (solid symbols) and dissolved phosphorus (open symbols),
including dissolved organic phosphorus (triangle), soluble reactive phosphorus (circle) and total dissolved
phosphorus (square) (A-E). Grazing by Paraphysomonas imperforata on Antarctic bacterial strain A at 0
o
C
(A), 5
o
C (B) and 10
o
C (C), and at 0
o
C on Antarctic bacterial strain B (D) and Halomonas halodurans (E).
Changes in phophorus concentration in all treatments (F). Note different ranges for x and y axes in (A-F).
102
(TDP, open squares) in each treatment by the end of protistan log phase growth. Similar
to nitrogen dynamics, the rates of PP loss and TDP release increased with increasing
temperature but were unaffected by prey type (Fig 3f). The total amount of TDP released
was not affected by either temperature or prey type. Final TDP concentrations in four of
the five treatments were 6-8 µM. One treatment, P. imperforata grazing on H.
halodurans at 0˚C, had higher final TDP concentrations (12 µM), but also had higher
initial TDP concentrations (3.3 µM vs. 0.4 - 1.0 µM in the other treatments).
Absolute changes in the dissolved and particulate fractions of nitrogen and
phosphorus were compared at the time that cultures were inoculated and the time of peak
protistan abundance (Table 1). Initial PN varied from 208-247 µM (average 218 µM)
and decreased to 43-128 µM (average 89 µM) by the time of peak protistan abundance.
Ammonium concentrations increased from 2.2-6.0 µM (average 4.5 µM) to 82.9-101 µM
(average 88.4 µM). Initial PP varied from 8.6-15.8 µm (average 10.7 µM) and decreased
to 3.5-10.3 µM (average 7.1 µM) by the time of peak protistan abundance. Soluble
reactive phosphorus concentrations increased from 0.1-1.9 µM (average 0.6 µM) to 2.1-
6.7 µM (average 3.8 µM).
The rates of nutrient regeneration at different temperatures were normalized by
plotting changes in concentrations of regenerated nutrients against natural log cell
abundance (Fig 4). The patterns of ammonium (Fig 4a) and SRP (Fig 4b) regeneration
vs. natural log cell abundance were similar regardless of temperature or prey type.
Ammonium and SRP showed similar changes in concentration as protistan abundance
increased.
103

Table 4-1. Changes in concentration of particulate carbon and particulate and dissolved nitrogen
and phosphorus before and after bacterial grazing by Paraphysomonas imperforata.

0˚C
Antarctic
bacterial
strain A
5˚C
Antarctic
bacterial
strain A
10˚C
Antarctic
bacterial
strain A
0˚C
Antarctic
bacterial
strain B
0˚C
Halomonas
halodurans

Average
Initial
PC
(µM)
977 952 935 979 1075 984
Final
PC
(µM)
416 217 430 436 571 414
Initial
PN
(µM)
213

208

205

216

247

218
Initial
NH
4

(µM)
4.7

2.2

4.3

5.2

6.0

4.5
Final
PN
(µM)
89.6

43

92.1

92.1

128

89.0

Final
NH
4

(µM)
86.1

101

84.7

87.5

82.9

88.4
Initial
PP
(µM)
8.6

9.6

9.3

10.4

15.8

10.7
Initial
SRP
(µM)
0.1

0.2

0.2

0.4

1.9

0.6
Final
PP
(µM)
6.8

3.5

7.2

7.5

10.3

7.1
Final
SRP
(µM)
2.1

4.6

2.4

3.1

6.7

3.8
104
0
0.5
1
1.5
2
2.5
6 7 8 9 10 11 12 13 14
LN Cell Abundance (cells ml
-1
)
Figure 4-4. Regeneration of ammonium normalized to the natural logarithm abundance of
Paraphysomonas imperforata in the different treatments. 0
o
C, Antarctic bacterial strain A
(square); 5
o
C, Antarctic bacterial strain A (circle); 10
o
C, Antarctic bacterial strain A
(triangle); 0
o
C, Antarctic bacterial strain B (diamond); 0
o
C, Halomonas halodurans
(inverted triangle).
105
Changes in the relative percentages of dissolved and particulate fractions of
nitrogen and phosphorus constituents were compared at the initial time point and the time
of peak protistan abundance (Fig 5). Initially ≥98% of nitrogen was in particulate form,
and 89-99% of phosphorus was in particulate form in all treatments. By the time of peak
protistan abundance, an average of 38% of the initial total nitrogen was in the form of
ammonium across all treatments (range = 30-47%). An average of 29% of the initial total
phosphorus was in the soluble reactive form across all treatments at the time of peak
protistan abundance. A wide range of soluble reactive phosphorus was observed (23-
45%) due to one high value (for 5˚C, Antarctic bacterial strain A). If that value is
removed from the range, the range is 23-27%, and the average is 25%.
The percentages of initial particulate nitrogen and phosphorus that were
regenerated as ammonium or dissolved phosphorus at the time of peak protistan
abundance were compared to results published for temperate conspecifics (Table 2).
These values were calculated by subtracting the initial ammonium or soluble reactive
phosphorus concentration from the final concentration, then dividing by the initial
particulate nitrogen or phosphorus concentration. Values ranged from 29-47% of
particulate nitrogen regenerated as ammonium, with an average of 37%. The percent of
particulate phosphorus regenerated as soluble reactive phosphorus ranged from 23-45%
with an average of 30%. There were no apparent trends in percent regeneration of
nutrients with temperature or prey type. The values for percent regeneration obtained in
this study were compared to published values for a temperate strain of Paraphysomonas
imperforata (Goldman et al. 1985, Andersen et al. 1986, Caron et al. 1986). The percent
106
0
20
40
60
80
100
0
20
40
60
80
100
0
20
40
60
80
100
0
20
40
60
80
100
Initial
PN
Initial
NH
4
+
Final
PN
Final
NH
4
+
Initial
PP
Initial
SRP
Final
PP
Final
SRP
A
B
Figure 4-5. Changes in particulate and dissolved fractions of initial total nitrogen (A) and phosphorus
(B). 0
o
C, Antarctic bacterial strain A (solid bars); 5
o
C, Antarctic bacterial strain A (horizontal stripes);
10
o
C, Antarctic bacterial strain A (diagonal stripes); 0
o
C, Antarctic bacterial strain B (dotted bars); 0
o
C,
Halomonas halodurans (white bars). Bars correspond to particulate or dissolved nutrient concentrations
as a percentage of the initial total nutrient concentration (particulate plus dissolved).
107
Table 4-2. Comparison of percent of initial particulate nutrients regenerated as ammonium or
soluble phosphorus at the time of peak protistan abundance for strains of bacterivorous
Paraphysomonas imperforata examined at different temperatures.

Source Temp
(˚C)
% PN regenerated
as NH
4
+
% PP regenerated
as SRP
Caron et al
(1986)
14

27
18 44
22 51
26 47
Goldman et
al (1985)
20 35
Andersen et
al (1986)
20 36
This study 0 38 23
5 47 45
10 37 25

0 36 26

0 29 30
Average
(SD)
39
(8)
31
(9)
108
of particulate nitrogen regenerated as ammonium ranged from 27-51% for the entire data
set (including published values and ones presented here), with an average of 39%. Only
one additional value for percent of particulate phosphorus regenerated as soluble reactive
phosphorus was available, but it was similar to the average observed here (36% vs. 30%
in this study).
Discussion
Heterotrophic protists play an important role as major nutrient remineralizers in
aquatic ecosystems (Caron & Goldman 1990, Dolan 1997). However, data for rates and
efficiencies of remineralization by heterotrophic protists are limited, and observations
have been conducted primarily on cultures isolated from temperate regions. Growth rates
and gross growth efficiencies of heterotrophic protists from permanently cold regions
have previously been compared to those of temperate heterotrophic protists (Lee &
Fenchel 1972, Choi & Peters 1992, Mayes et al. 1997, Rose et al. in prep). In general, it
is recognized that protists from all environments grow slowly at low temperature (<5˚C)
relative to growth rates of protists at higher temperatures (Rose & Caron submitted).
However, there is conflicting information regarding the response of growth efficiency to
changes in temperature. Some studies have reported increases in gross growth efficiency
with increases in temperature (Rassoulzadegan 1982, Verity 1985, Choi & Peters 1992),
others have reported decreases in gross growth efficiency with increases in temperature
or very low gross growth efficiency at low temperature (Laybourn & Stewart 1975,
Rogerson 1981, Mayes et al. 1997), and others have reported no change in gross growth
109
efficiency with increasing temperature (Caron et al. 1986, Ishigaki & Sleigh 2001, Rose
et al. in prep).
Much less information is available for temperature effects on rates and
efficiencies of nutrient regeneration. One study examining the effect of temperature on
nutrient remineralization found increased rates of ammonium regeneration at higher
temperatures for a temperate strain of the heterotrophic protist P. imperforata, but total
amount of ammonium regenerated was unaffected (Caron et al. 1986). Another study
reported increased excretion rates when the heterotrophic protist Monas sp. was outside
the optimal temperatures for growth, and also reported increased excretion rates at higher
temperatures (Sherr et al. 1983). We are unaware of any studies reporting baseline
nutrient regeneration information for heterotrophic protists from permanently cold
environments.
Particulate carbon decreased as protistan grazing reduced the bacterial
populations in experimental flasks (Fig 1, Table 1). The rate of loss of particulate
carbon was higher at higher temperatures (Fig. 1a, c). The rate of decrease of particulate
carbon did not change with different prey types and there was no apparent difference
between grazing on Antarctic vs. temperate strains of bacteria (Fig 1b, d). The total
number of protists produced and the total amount of particulate carbon consumed was not
affected by either temperature or prey type (Table 1).
The Antarctic P. imperforata had faster rates of nitrogen regeneration at higher
temperatures, but no apparent difference in rates of nitrogen regeneration when fed
different strains of bacterial prey at a single temperature (Fig. 2). Particulate nitrogen
110
decreased steadily in all treatments from the initial time point, and loss rates were faster
at higher temperatures (Fig 2a-e, filled symbols). Ammonium concentration increased in
all treatments after an initial lag and continued to increase into the stationary growth
phase of the protist (Fig 2a-e, open symbols). Ammonium regeneration occurred more
rapidly at higher temperatures, but rates were not affected by prey type (Fig. 2f). The
two strains of Antarctic bacteria grown at 5˚C did not affect rates or magnitude of
nutrient regeneration by the Antarctic P. imperforata relative to H. halodurans grown at
room temperature.
As observed for particulate carbon, particulate nitrogen and ammonium
concentrations, changes in the particulate and dissolved constituents of phosphorus were
dependent on temperature but not prey type (Fig. 3). Decreases in particulate phosphorus
and increases in dissolved phosphorus were only observed as the protistan cultures
approached the end of log phase growth (Fig. 3a-e). Regeneration of soluble reactive
phosphorus occurred more rapidly at higher temperature, but was not affected by prey
type.
Soluble reactive phosphorus concentrations in the P. imperforata culture feeding
on H. halodurans increased at cell abundances of the protists that were lower than those
observed in the other treatments (Fig 4). Initial particulate phosphorus concentrations
were also high within this treatment relative to particulate carbon and particulate nitrogen
concentrations, suggesting that H. halodurans was phosphorus-rich relative to the two
Antarctic bacterial strains used in this experiment. The excess of particulate phosphorus
in the H. halodurans treatment led to a greater remineralization of soluble reactive
111
phosphorus because the flagellate could not use the excess phosphorus for growth (6.7
µM final SRP vs. average 3.0 µM final SRP for the other treatments, Table 1).
The overall dynamics of carbon, nitrogen and phosphorus were similar for all
treatments in this study. Changes in absolute abundances of particulate and dissolved
nutrients between the initial time point and the time of peak protistan abundance were
similar across all treatments (Table 1). Slight differences in these values were due to the
inability to sample at the exact same point in growth phase of the flagellate across all
treatments. The treatment at 5˚C containing Antarctic bacterial strain A had higher
dissolved nutrients and lower particulate nutrients (both nitrogen and phosphorus) than
the other treatments. The sampling times for this treatment were just before and just after
the protistan culture entered stationary growth phase, while the other treatments had
sampling times that were almost exactly at the beginning of stationary growth phase. The
sampling time after the 5˚C culture entered stationary growth phase was closer to the
projected time of peak protistan abundance, so this point was used for nutrient
calculations. Sampling later in stationary phase relative to the other treatments meant
that the protists had more time to generate dissolved nutrients and consume the
particulate nutrients (in the form of the bacterial prey). When the values of Table 1 are
recalculated using the earlier time point, they are as follows: final particulate nitrogen =
113 µM, final particulate phosphorus = 8.7 µM, final NH
4
+
= 70 µM, final SRP = 0.6
µM. It is unlikely that the differences in particulate and regenerated nutrients in this
treatment are due to effects of the temperature or the bacterial type. No trend was
112
apparent when this treatment was compared to other temperatures or bacterial types in
total particulate nutrient loss or dissolved nutrient regeneration.
The similarity in the patterns of nutrient dynamics became obvious when the
effects of temperature on rate processes were eliminated. Rates of nutrient regeneration
were normalized by plotting ammonium and SRP concentrations against the natural log
cell abundance for each treatment (Fig 4). Both nutrients showed similar patterns of
regeneration as protistan cell abundance increased, with lags in dissolved nutrient
production early in log phase growth, and sharp increases as protistan cells reached
stationary phase growth. Slight differences in the buildup of SRP and ammonium were
apparent (Fig 4). Sharp increases in SRP were observed when cell abundance exceeded
5x10
5
cells ml
-1
, while ammonium concentrations increased rapidly when cell abundance
exceeded 10
5
cells ml
-1
.
Differences in total initial nutrients across treatments were removed by plotting
total regeneration of dissolved nutrients as a percentage of total initial nutrient
concentration (Fig 5). This normalization removes the effect of higher initial nutrient
concentrations in the 0˚C H. halodurans treatment to demonstrate that the amount of N
and P remineralized was similar in this treatment to the other bacterial types as well as
the higher temperatures. As observed in Table 1, total regeneration of nutrients was
similar across all treatments.
Total regeneration of nutrients by the Antarctic strain of P. imperforata observed
in this study was similar to nutrient regeneration values published for temperate
conspecifics (Goldman et al. 1985, Andersen et al. 1986, Caron et al. 1986) (Table 2).
113
Since initial nutrient concentrations differed among studies, calculating the percentage of
initial particulate nutrient concentrations normalized the values for dissolved nutrients
generated by protistan grazing. These studies and the present study include observations
measured across a temperature range of 0-26˚C, however, percent ammonium
regeneration was remarkably consistent. The total average percent ammonium
regenerated across all studies was 39%, the average for this study was 37%, and the
average for the studies examining temperate P. imperforata was 41%. Only one study
examined regeneration of soluble reactive phosphorus, however, the percent regenerated
phosphorus observed in that study (36%) was similar to the average observed here (30%).
Conclusions
Temperature affected the rate of nutrient regeneration by an Antarctic strain of P.
imperforata, but did not affect the total quantity of nutrients regenerated. Prey type had
no effect on rates of nutrient regeneration or total quantity of nutrients regenerated. Total
nutrient regeneration by the Antarctic P. imperforata was also similar to values published
for a temperate conspecific examined at much higher temperatures. Rates of nutrient
regeneration were greatly reduced at low temperature, suggesting that in permanently
cold systems, nutrient recycling by heterotrophic protists occurs slowly. However, since
total regeneration of nutrients was similar at all temperatures examined in this study, the
efficiency with which heterotrophic protists convert nutrients contained in particulate
prey to their remineralized forms is similar across ecosystems with extremely different
temperature regimes.
114
Chapter 5: Low temperature constraints on grazing by Antarctic
microzooplankton in culture and in natural assemblages
Introduction
Herbivorous protists play important roles as consumers within aquatic foodwebs.
These organisms have been reported to graze a high percentage of phytoplankton
production on a daily basis in many regions (Chavez et al. 1991, Verity et al. 1996,
Lessard & Murrell 1998, Strom et al. 2001). Rates of herbivory by protists have been
demonstrated to be as high or higher as rates of herbivory by metazooplankton at
numerous times and locations in the ocean (Gifford et al. 1995). Herbivorous protists
have also been identified as key players in top-down control of phytoplankton in aquatic
systems. The high grazing rates of microzooplankton have been used to explain the
absence of algal blooms in some aquatic systems at times and locations during which
these phenomenon would be expected to occur (Frost 1987, Parsons & Lalli 1988, Frost
1991).
Microzooplankton also have been implicated as the dominant herbivores in
Antarctic marine ecosystems, as opposed to copepods (Atkinson 1995, 1996, Lonsdale et
al. 2000). The permanently cold coastal waters surrounding the continent of Antarctica
have been reported to have periodically high abundances of microzooplankton (Garrison
et al. 1986, Garrison 1991, Gowing & Garrison 1992, Stoecker et al. 1993, Caron et al.
2000). Ciliates and dinoflagellates are especially abundant in mid- to late summer (Caron
et al. 2000). Abundances have been reported on the order of 10
4
cells l
-1
for ciliates and
115
dinoflagellates in the Ross Sea (Caron et al. 2000), 10
3
heterotrophic dinoflagellates l
-1
in
the Weddell Sea (Garrison & Buck 1989), 10
4
ciliates l
-1
in the Weddell Sea (Buck &
Garrison 1983) and 10
5
ciliates l
-1
in Prydz Bay (Davidson & Marchant 1992). The high
potential abundances of herbivorous protists in coastal waters surrounding Antarctica
have led some researchers to suggest grazing may control phytoplankton blooms in this
ecosystem (Klaas 1997).
The Ross Sea, Antarctica is the location of one of the world’s largest annual
phytoplankton blooms. Chlorophyll values of 14 µg l
-1
were observed in the southern
central polynya during the US Southern Ocean JGOFS cruises, and abundances of the
dominant phytoplankton in this region, Phaeocystis antarctica, were reported to be as
high as 10
6
cells l
-1
(Smith et al. 2000, Smith et al. 2003). This massive phytoplankton
bloom does not appear to be heavily grazed. Caron et al (2000) measured rates of
herbivory in 51 dilution experiments at four stations in the Ross Sea during austral
spring, summer and fall. Only 13 of these experiments yielded significant grazing rates,
and the significant rates were universally low when compared to rates of herbivory
reported for temperate systems. In addition, sediment trap data and surface sediment
samples from the Ross Sea have indicated a large vertical flux at the end of the
phytoplankton bloom, suggesting that a large portion of phytoplankton production is
transported to the sediment rather than recycled in the water column by grazing activities
(Smith & Dunbar 1998, DiTullio et al. 2000).
There is thus a striking contrast between the high microzooplankton abundance in
late summer and early fall in the Ross Sea and the apparent weak top-down control on
116
primary production in this system. Recent evidence suggests that low temperature plays
a role in inhibiting microzooplankton growth in cold oceans relative to phytoplankton
growth rate. A review of protistan growth rates by Rose and Caron (submitted) suggests
that growth rates of heterotrophs are extremely low at cold temperatures relative to
growth rates of phototrophic protists. Moreover, growth rates of herbivorous
microzooplankton appear to be suppressed by low temperature to a greater extent than
their phytoplankton prey. Additionally, Caron et al (2000) reported a direct relationship
of temperature on grazing rate based on a literature review of published reports of
measurements of herbivory.
We hypothesized that the low ambient temperatures of the Ross Sea constrain
individual grazing rates by herbivorous microzooplankton. We also hypothesize that low
ingestion rates are the reason why rates of herbivory are low even when micrograzers are
abundant. This study combined laboratory and field experiments to examine ingestion
rates by Antarctic microzooplankton. Short-term ingestion rate experiments were used
to obtain estimates of ingestion rates of the whole microzooplankton assemblages as well
as rates of the numerically dominant genera during austral summer in the Ross Sea.
Ingestion rates of natural assemblages of microzooplankton based on tracer-level
additions of fluorescently labeled algae were consistently low, and estimates of grazing
impact on standing stocks of phytoplankton suggested grazers did not exhibit strong top-
down control on phytoplankton at any station. Specific ingestion rates of individual
genera of Antarctic ciliates in field populations were low when compared to values for
temperate ciliates reported in the literature. The effects on ingestion rate of low
117
temperature, prey abundance and physiological state were further examined using a
laboratory culture of an Antarctic strain of Strombidium. Specific ingestion rates of the
ciliate were highly dependent on physiological state of the ciliate. Low temperature
imparted a strong overall constraint on growth rate of the ciliate. Ingestion rates for a
starved culture were comparable to rates reported for temperate ciliates. Ingestion rates
for the same culture in balanced growth were much lower, and instead were comparable
to rates measured for the field populations of Antarctic ciliates. Our results indicate that
low ingestion rates by Antarctic microzooplankton may contribute to the weak top-down
control by grazers on phytoplankton assemblages in the Ross Sea that has been noted in
previously published studies.
Methods
Cultures. Strombidium sp. (isolate I-256Ciliate) was isolated from an ice core that was
collected from the pack ice region of the Ross Sea, Antarctica. The core was collected at
175˚53’ E, 68˚07’ S using a hand powered Sipre corer. Two sections (20 cm between 27
and 47 cm from the surface, plus the bottom 15 cm) were slowly melted into 1L sterile
seawater in the dark at 0˚C over the course of several days. The melted core was
enriched with F/2 to enrich the natural phytoplankton assemblage. Once ciliate
abundance increased, individual cells were micropipetted, rinsed and placed into sterile
seawater, to which a few drops of cultured Antarctic algae were added. A ciliate culture
obtained in this manner was used for the experiments described in this study. The ciliate
was identified as Strombidium sp. based on morphological features and 18S rDNA
sequence similarity when blasted against the GenBank data set [Accession No. 0000].
118
The cultures of Antarctic algae used as prey in these experiments were isolated
from water samples collected over a series of research cruises to the Ross Sea, Antarctica.
Cultures were identified to closest phylogenetic taxon based on 18S rDNA sequence
similarity [Accession Nos. 0000, etc]. Pyramimonas sp. (isolate RS-11) was isolated by
dilution extinction from an enriched water sample collected at 73˚30’S, 176˚50’W on
11/15/97, and was also identified using light microscopy according to (Daugbjerg 2001).
Thalassiosira sp.1 (isolate SL-64/78Cheetos) and Thalassiosira sp. 2 (isolate SL-
64/78Frag) were isolated by dilution extinction from an enriched meltwater sample
collected at 68˚59’S, 164˚59’W on 1/5/99. Polarella glacialis (isolate RS-6) was isolated
by dilution extinction from an enriched water sample collected at 76˚37’S, 169˚33’E on
11/21/97, and was also identified using light microscopy according to (Montresor &
Procaccini 1999). Chlamydomonas sp. (isolate I-155Chlamydomonas) was isolated by
dilution extinction from an ice core collected at 71˚59’S, 150˚02’W on 1/16/99. The ice
core was collected, melted and enriched as described above. Mallomonas sp. (isolate I-
76) was isolated by dilution extinction from an ice core collected at 68˚05’S, 164˚58’W
on 1/4/99. All water and ice samples were collected, enriched and maintained at
temperatures <2˚C to prevent selection against heat-sensitive protists. All cultures were
maintained on F/2 medium (Guillard 1975) at 0˚C on a 12:12 L:D cycle prior to
beginning experiments.

Field sample collection and cell counts. Sample collection and short-term ingestion
rate experiments were conducted in the Ross Sea, Antarctica aboard the RVIB Nathaniel
119
B. Palmer. Four experiments were carried out within the Ross Sea polynya during austral
summer (late December and January). The stations were located at 77˚00’S, 180˚00’E
(Experiment 1), 74˚30’S, 173˚30’E (Experiments 2 and 4, separated in time by
approximately 2 weeks) and 75˚00’S, 167˚00’E (Experiment 3). Water for all analyses
and experiments was collected using Niskin bottles from 5m depth. Short-term ingestion
rate experiments were conducted using cultures of Antarctic algae and natural plankton
assemblages. In addition, samples were preserved with 10% acid lugols solution and
stored in the dark for enumeration of microzooplankton back in the laboratory. One
hundred milliliters were settled for at least 18 hours in the laboratory into settling
chambers before counting using an inverted microscope at 200x magnification (Utermohl
1958). Ciliates were enumerated and abundant taxa were identified to genus level when
possible. Heterotrophic dinoflagellates such as Protoperidinium and Gyrodinium were
also enumerated based on morphology but other dinoflagellate genera were not recorded
since the lugols fixative made it impossible to distinguish phototrophic from
heterotrophic taxa.

Field ingestion rate experiments. Short-term ingestion rate experiments were
conducted using cultures of Antarctic algae fluorescently stained with Cell Tracker Green
(Molecular Probes, Eugene, OR). Cell Tracker Green was chosen to stain the algae in
these experiments over the more commonly used DTAF because algal cultures could be
stained live and retained their stain inside and outside of microzooplankton food vacuoles
upon preservation with ice cold 1% glutaraldehyde. Algal cultures were stained in the
120
dark for two hours immediately prior to the short-term ingestion rate experiments using 1
µM, 0.2 µm filtered Cell Tracker Green according to the protocol of Li et al (1996).
Short-term ingestion rate experiments were conducted using a wide range of
abundance of added fluorescently labeled algae in order to estimate actual grazing rates
of the natural microzooplankton assemblages as well as maximal grazing rates of the
same assemblages. These experiments consisted of a series of treatments to which
increasing concentrations of fluorescently stained, cultured Antarctic Thalassiosira sp.
(clone SL-64/78Cheetos) were added to whole seawater. Abundances of added stained
Thalassiosira sp. ranged from 10
6
-10
8
cells L
-1
. We were limited in the maximal
abundance of Thalassiosira sp. that we could add to each mixed plankton assemblage.
Cell Tracker Green could not be added to whole seawater at less than a 1:10 dilution
without the risk of fluorescently staining algal and microzooplankton cells present in the
mixed plankton assemblage. In addition, an actively growing algal culture was required
in order to achieve bright, even staining of all cultured cells with Cell Tracker Green.
Since Thalassiosira sp. consistently entered stationary growth phase at an abundance of
5x10
8
- 10
9
cells L
-1
, the maximum abundance of Thalassiosira sp. that we could have in
our experiments was 5x10
7
- 10
8
cells L
-1
.
Experiments were carried out in the dark in a 0˚C cold room, in 150 cm
2

polystyrene tissue culture flasks. An aliquot of the fluorescently stained algal culture was
filtered (0.2 µm) and the filtrate was added to whole seawater as a control treatment to
check for uptake of the fluorescent stain by the mixed natural plankton assemblage. This
problem never occurred in any of the treatments or experiments. Experimental and
121
control treatments were performed in triplicate. Samples of 125 ml were removed
initially and after 30 minutes in the algal addition treatments with ≥10
7
cells L
-1
, and were
removed initially and after 120 minutes in the algal addition treatments with ≤5x10
6
cells
L
-1
. Ingestion of cells by the microzooplankton was assumed to be linear over this time
period based on the results of ingestion rate experiments conducted with Strombidium sp.
in the laboratory (see Results section). Samples were preserved with 1% ice-cold
glutaraldehyde (final concentration) and allowed to settle for at least 24 hours in the dark
in 250 ml amber glass jars. Samples were then concentrated as follows: after settling, the
top 200 ml were pumped off under gentle vacuum through a 1 ml pipet. The remaining
50 ml were mixed, poured into a 50 ml centrifuge tube and centrifuged in a clinical
centrifuge at top speed for 10 minutes. The top 45 ml were pumped off as before and
again, the supernatant was examined periodically to ensure that no microzooplankton
were lost in this process. The remaining 5 ml were examined in 1 ml aliquots under light
microscopy using a Sedgwick Rafter counting chamber. The first 30 microzooplankton
encountered were examined for ingested stained algal cells to determine community
grazing rates. Grazing rates of individual genera of microzooplankton were determined
by examining either the first 30 cells encountered (for common genera) or the number of
cells encountered in the entire 1 ml aliquot (for rarer genera). Rates reported here for
individual genera are only those for which at least 15 cells were examined per
experiment. Ingestion rates could not be determined for Protoperidinium sp. because the
natural apple-green fluorescence of the dinoflagellate interfered with visualization of
ingested fluorescently labeled algae. However, since Protoperidinium sp. did not
122
numerically dominate the microzooplankton assemblage during any of the experiments,
we believe this did not significantly affect our estimations of ingestion rate by the total
microzooplankton assemblage.

Laboratory ingestion rates of cultured Strombidium sp. Based on results from the
field experiments that showed variability in ingestion rate within individual ciliate genera
between experiments, we conducted a series of experiments examining the relationship
between physiological state and ingestion rate using a culture of the Antarctic ciliate
Strombidium sp. Thalassiosira sp. 1 was selected for use as prey in these experiments,
because Strombidium sp. grew well on this alga in the growth rate experiment and it
stained well using the vital stain Cell Tracker Green. Thalassiosira sp. was grown to the
end of log phase growth on F/2 medium to ensure prey were provided in excess.
Strombidium sp. was grown to high abundance on Thalassiosira sp. until Thalassiosira
sp. were at very low abundance. Strombidium sp. was starved on this low abundance of
Thalassiosira sp. for two weeks prior to the beginning of the experiment. Starvation of
Strombidium sp. resulted in greatly reduced cell volume.
A dense sample of the Thalassiosira sp. was stained in the dark at 0˚C for two
hours with 1 µM Cell Tracker Green according to the protocol of Li et al (1996). This
stained culture was added to an unstained sample of dense Thalassiosira sp. culture at
~15% of the unstained cell abundance. It was confirmed beforehand that a stain
concentration of 0.15 µM did not stain the Thalassiosira or Strombidium cells after the
addition of stained Thalassiosira sp. cells. Starved Strombidium sp. were added at ~300
123
cells ml
-1
to cultures of dense, unstained Thalassiosira and separate aliquots of the same
culture of starved Strombidium sp. were added to triplicate mixed cultures of stained and
unstained Thalassiosira sp. at the same initial abundance. A sample of the Strombidium
and mixed Thalassiosira sp. culture was immediately removed from each replicate and
preserved with 1% ice cold glutaraldehyde (final concentration). Additional samples
were removed and preserved with 1% ice cold glutaraldehyde after 15, 30, 60 and 120
minutes. One milliliter subsamples were examined using epifluorescence microscopy
and a Sedgwick Rafter slide. The number of ingested stained algal cells was counted
within 30 ciliates per sample for each of the replicates at each time point. The total
number of ingested cells was then calculated from the ratio of unstained:stained cells in
each replicate and the number of ingested stained cells.
The Strombidium cultures containing dense unstained Thalassiosira were kept in
a lighted incubator at 0˚C for two days, then an aliquot was removed from each for a
second ingestion rate experiment. Stained Thalassiosira sp. cells were added to the
Strombidium culture at ~15% the abundance of unstained Thalassiosira sp. and
subsamples were removed after 0, 15, 30, 60 and 120 minutes, preserved with 1% ice
cold glutaraldehyde and counted under epifluorescence microscopy as described above.
This experiment was repeated a final time after the Strombidium culture fed on the dense
unstained Thalassiosira sp. for four days.

Laboratory growth rates of cultured Strombidium sp. Algae used for prey in the
growth rate experiments were grown to early stationary growth phase on F/2 medium to
124
ensure prey were provided to the ciliate in excess. Strombidium sp. were acclimated to
growth on the following six algal species: Thalassiosira sp. 1, Thalassiosira sp. 2,
Pyramimonas sp., Polarella glacialis, Chlamydomonas sp. and Mallomonas sp. At the
beginning of the growth experiment, Strombidium sp. were inoculated at approximately
40 cells ml
-1
into triplicate algal cultures of each prey species and were grown for 26
days, sampling approximately every 3-4 days. Subsamples were removed and preserved
with 2% acid lugols solution and kept at room temperature in the dark until analysis. One
milliliter samples were placed in a Sedgwick Rafter counting chamber and the entire
volume counted using light microscopy. Growth rates were calculated based on the slope
of the graph of natural log cell abundance vs. time when cells were in logarithmic growth
phase.

Calculations. Estimates of ingestion rates (algal cells ingested ciliate
-1
h
-1
) of
Strombidium, Eutintinnus and Codonellopsis were converted into specific ingestion rate
(h
-1
). Specific ingestion rate normalizes ingestion rates to the size of the herbivorous
microzooplankton by dividing the ingestion rate of algal cells in grams carbon per unit
time by the biomass of the microzooplankton in grams carbon (Hansen et al. 1997). The
dimensions of ciliates and algae in culture and in natural plankton assemblages, preserved
in 10% lugols solution, were determined using digital photographs of the protists taken
using phase contrast microscopy and an inverted microscope (Leica DM IRB) at 200x
(ciliates) and 400x (algae) magnification. Measurements of the length and width of
preserved cells were performed on the digital photographs using the software program
125
Openlab 3.5.1. Linear dimensions were converted to volume based on standard
equations. Biovolumes of the cells were converted into grams C cell
-1
assuming a
conversion factor of (0.2*wet weight in grams) = dry weight and (0.4*dry weight) =
grams carbon (Beers & Stewart 1971, Beers et al. 1975, Gifford & Caron 2000). Algal
cell volumes were converted into grams carbon based on the equation log carbon = –
0.314+0.712*(log volume) from Strathmann (1967). Specific ingestion rate was
calculated as (pg C ingested ciliate
-1
h
-1
)/(pg C ciliate
-1
).
Results
Field experiments. Total phytoplankton abundance was high at the three stations at the
time of all four experiments, ranging from 3.0-6.2 x10
7
cells L
-1
(Table 1). The
phytoplankton assemblage was numerically dominated at all stations by a small pennate
diatom present in excess of 10
7
cells L
-1
(D. Karentz, pers. comm.) The diatom was
identified as Nitzschia sp. by scanning electron microscopy (P. Miller, pers. comm.). The
second most common taxon was Phaeocystis antarctica, which was present at
abundances ranging from 5x10
5
to 5x10
6
cells L
-1
. Phytoplankton abundance generally
increased from Experiments 1-3 but decreased at the time of Experiment 4.
Phytoplankton abundance was nearly 50% lower at the time of Experiment 4 relative to
2, which were both located at 74˚30’S, 173˚30’E but separated in time by approximately
two weeks.
Total microzooplankton abundance generally increased from Experiments 1-4,
from 6,600 cells L
-1
at station 1 to 19,000 cells L
-1
at station 4 (Table 1).
Microzooplankton abundance was dominated at station 1 by aloricate ciliates, which were
126

Table 5-1. Abundance of phytoplankton, total microzooplankton and subgroups of
microzooplankton in late austral summer at four stations in the Ross Sea, Antarctica.

Station
#
Phyto-
plankton
abundance
(cells L
-1
)
Total
Microzoo-
plankton
(cells L
-1
)
Eutin-
tinnus sp.
(cells L
-1
)
Codonel-
lopsis sp.
(cells L
-1
)
Aloricate
Ciliates
(cells L
-1
)
Protoperi
-dinium
sp.
(cells L
-1
)

1

4.6x10
7

6.6x10
3

180

250

5500

600

2

5.5x10
7

5.1x10
3

2000

190

2600

290

3

6.2x10
7

1.3x10
4

8000

530

3200

810

4

3.0x10
7

1.9x10
4

12000

1200

5100

560

127
present at 5,500 cells L
-1
. Aloricate ciliates were present at similar abundances among all
four stations, ranging from 2,600 to 5,500 cells L
-1
. Total microzooplankton abundance
was dominated by loricate ciliates at stations 2 and 3 (Experiments 2, 3 and 4). Most
loricate ciliates at these stations belonged to the genera Eutintinnus or Codonellopsis.
The abundance of both genera increased dramatically over the three stations, from 250 to
1,200 cells L
-1
for Codonellopsis and from 180 to 12,000 cells L
-1
for Eutintinnus.
Morphologically identifiable heterotrophic dinoflagellates were dominated by
Protoperidinium sp. (Gyrodinium sp. were observed only extremely rarely). The
abundance of Protoperidinium sp. ranged from 290 to 810 cells L
-1
across all three
stations, and nearly doubled in abundance during the two week interim between
Experiments 2 and 4 (from 290 to 560 cells L
-1
).
Ingestion rates of the total microzooplankton assemblage increased with
increasing abundances of added stained Thalassiosira sp. at all four stations (Fig 1). The
number of ingested cells microzooplankton
-1
after 30 minutes appeared to approach a
maximum at the highest abundance of added Thalassiosira sp. during Experiments 1, 3
and 4, but we were not able to obtain definitive maximum ingestion rates at any station.
We were unable to add higher abundances of Thalassiosira sp. due to limitations imposed
by the Cell Tracker Green stain, as described in the Methods section. The highest
number of average ingested algal cells microzooplankton
-1
was 10.8 h
-1
at station 4, when
10
8
stained Thalassiosira sp. L
-1
were added to the mixed plankton assemblage. The
average ingested prey microzooplankton
-1
ranged from 2.7 to 8.1 h
-1
across all stations
when Thalassiosira sp. was added at 5x10
7
cells L
-1
. The average ingested prey
128
0
2
4
6
8
10
12
0
2
4
6
8
10
12
0
2
4
6
8
10
12
0 20 40 60 80 100
0
2
4
6
8
10
12
0 20 40 60 80 100
Thalassiosira sp. abundance (x10
6
cells L
-1
)
A
B
C
D
Figure 5-1. Ingestion rates with different abundances of fluorescently labeled algae. Average number of ingested
Thalassiosira cells microzooplankton
-1
at four stations in late austral summer in the Ross Sea, Antarctica. Station 1
(A), Station 2 (B), Station 3 (C) and Station 4 (D).
129
microzooplankton
-1
was more similar among stations when fewer prey were added,
ranging from 0.07 to 0.1 cells h
-1
at all four stations when Thalassiosira sp. was added at
10
6
cells L
-1
.
Ingestion rates of two individual ciliate genera were calculated at Experiments 2
and 4 (Fig 2). Rates of Eutintinnus sp. during Experiment 2 varied from 0.07 to 2.3 cells
h
-1
when Thalassiosira sp. was added at four abundances ranging from 10
6
to 5x10
7
cells
L
-1
. Rates during Experiment 4 ranged from 0.06 to 7.2 cells h
-1
when Thalassiosira sp.
was added at five abundances ranging from 10
6
to 10
8
cells L
-1
(Fig 2A, B). Maximum
ingestion rates of Eutintinnus sp. were observed during Experiment 4, with both 5x10
7

and 10
8
cells L
-1
showing an average ingestion rate of ~7 cells h
-1
. Ingestion rates for
Codonellopsis sp. were generally much higher than those observed for Eutintinnus sp.
(Fig 2C, D). Ingestion rates during Experiment 2 ranged from 0.12 to 10.7 cells h
-1
,
when Thalassiosira sp. was added between 10
6
and 5x10
7
cells L
-1
. Ingestion rates
during Experiment 4 ranged from 0.13 to 40.3 cells h
-1
when Thalassiosira sp. was added
from 10
6
to 10
8
cells L
-1
. Ingestion rates showed no signs of leveling off at either 5x10
7

or 10
8
Thalassiosira sp. L
-1
.
There was a general trend of increasing ingestion rate at high abundances of
added prey for all four experiments. This trend was consistent for both the total
microzooplankton as well as the two most common individual genera. At 5x10
7

Thalassiosira sp. L
-1
, total microzooplankton ingestion rates increased from 2.7 ingested
prey microzooplankton
-1
h
-1
(Experiment 1) to 3.8 (Experiment 2), to 5.1 (Experiment 3)
to 10.8 (Experiment 4). The two most common genera, Eutintinnus and Codonellopsis,
130
0
1
2
3
4
5
6
7
8
0
1
2
3
4
5
6
7
8
0
10
20
30
40
50
0 20 40 60 80 100
0
10
20
30
40
50
0 20 40 60 80 100
Thalassiosira sp. abundance (x10
6
cells L
-1
)
A B
C D
Figure 5-2. Ingestion rates with different abundances of fluorescently labeled algae. Average number of ingestion
Thalassiosira cells per Eutintinnus (A,B) or per Codonellopsis (C,D) at Station 2 (A,C) and Station 4 (B,D).
131
were only examined during Experiments 2 and 4 (which had the same location but a two
week difference in time), but they demonstrated higher potential ingestion rates during
Experiment 4 than Experiment 2. Ingestion rates at 5x10
7
Thalassiosira L
-1
were 3 cells
Eutintinnus
-1
h
-1
during Experiment 2 vs. 7 cells Eutintinnus
-1
h
-1
during Experiment 4.
Ingestion rates at 5x10
7
Thalassiosira L
-1
were 10.7 cells Codonellopsis
-1
h
-1
during
Experiment 2 vs. 40.3 cells Codonellopsis
-1
h
-1
during Experiment 4.
Specific ingestion rates of Eutintinnus sp. and Codonellopsis sp. increased at
higher abundances of Thalassiosira sp. but were low at even the highest abundance of
Thalassiosira sp. (Table 2). Specific ingestion rates were similar among both genera
between Experiments 2 and 4 when Thalassiosira were added at 10
6
cells L
-1
. However,
specific ingestion rates were consistently higher within each genus during Experiment 4
than during Experiment 2 when Thalassiosira sp. were added at 5x10
6
, 10
7
and 5x10
7

cells L
-1
.

Laboratory experiments. Ingestion rates of the Antarctic Strombidium sp. varied
greatly depending on whether cultures were in balanced growth or starved (Table 2, Fig
3). When the ciliates were fed a high number of Thalassiosira sp. following a period of
starvation (~ 2 weeks), the average number of ingested algal cells was 12 ciliate
-1
after 15
minutes, and began to reach a maximum of approximately 18 ingested cells ciliate
-1
by 60
minutes. Digestion of stained cells had begun by 120 minutes in many of the ciliates and
individual cells could not always be reliably enumerated in food vacuoles within each
ciliate. Therefore, the results from this time point were not included. Ingestion rates of
132

Table 5-2. Ingestion rates and specific ingestion rates of Antarctic ciliates in culture and in
natural plankton assemblages.

Ciliate Station Prey
Abundance
(cells l
-1
)
Ingestion Rate
(cells ingested
ciliate
-1
h
-1
)
Specific
Ingestion
Rate (h
-1
)
Eutintinnus sp. 2 5x10
7
2.30 0.0043
10
7
0.68 0.0013
5x10
6
0.25 0.0005
10
6
0.07 0.0001
4 10
8
7.23 0.0135
5x10
7
7.05 0.0131
10
7
2.43 0.0045
5x10
6
0.69 0.0013
10
6
0.06 0.0001
Codonellopsis sp. 2 5x10
7
10.7 0.0067
10
7
2.12 0.0013
5x10
6
0.51 0.0003
10
6
0.12 0.0001
4 10
8
40.3 0.0252
5x10
7
21.1 0.0132
10
7
5.41 0.0034
5x10
6
1.03 0.0006
10
6
0.13 0.0001
Strombidium sp.
starved
N/A 7x10
8
32.9 0.187
Strombidium sp.
fed 2 days
N/A 7x10
8
8.94 0.0510
Strombidium sp.
fed 4 days
N/A 7x10
8
6.23 0.0354
133
0
2
4
6
8
10
12
14
16
18
0 30 60 90 120 150 180
Time (hours)
Figure 5-3. Ingestion rates of the cultured Antarctic Strombidium sp. feeding on the Antarctic
alga Thalassiosira sp. 1 after starvation (squares), after two days of feeding on a dense culture of
Thalassiosira sp. 1 (circles) and after four days of feeding (triangles).
134
the Antarctic Strombidium sp. were considerably lower after the ciliate culture fed on a
high density of Thalassiosira sp. for two days (Fig 3, circles). Fifteen minutes after the
addition of stained cells, the average total number of ingested algal cells was 2.3 ciliate
-1
.
The number of ingested cells appeared to be still increasing at 120 minutes. The number
of ingested algal cells ciliate
-1
after 120 minutes was 10.8. Ingestion rates of the
Antarctic Strombidium sp. continued to decline after the ciliate culture fed on a high
density of Thalassiosira sp. for four days (Fig 3, triangles). Fifteen minutes after the
addition of stained cells, the average total number of ingested algal cells was 1.8 ciliate
-1
.
The number of ingested cells ciliate
-1
began to approach a maximum after 60 minutes,
and the number of ingested algal cells was the same at 120 and 180 minutes (7.7).
Ingestion rates (calculated based on the linear portion of the curve of algal cells ingested
vs. time) from these three experiments were 32.9, 8.94 and 6.23 algal cells ingested
ciliate
-1
h
-1
for the starved, fed two days and fed four days treatments, respectively.
Specific ingestion rates were also different for the cultures in balanced growth vs.
starved (Table 2). The specific ingestion rate of the starved Strombidium sp. was 0.187 h
-
1
but dropped to 0.0510 h
-1
after two days of feeding on a high abundance of prey and
0.0354 h
-1
after four days. The specific ingestion rate of the starved ciliate at 0˚C was
comparable to specific ingestion rates reported for temperate ciliates at appropriate
temperatures for those environments (0.187 for Strombidium was within the range 0.1-
0.481 of temperate ciliates, Fig 4). The specific ingestion rates of the Strombidium sp. in
balanced growth were more similar to the maximal specific ingestion rates of the
Antarctic Eutintinnus sp. and Codonellopsis sp. observed in the natural plankton
135
0
0.1
0.2
0.3
0.4
0.5
-5 0 5 10 15 20 25
Temperature (
o
C)
Figure 5-4. Comparison of specific ingestion rates measured in this study to values for temperate
ciliates reported in Hansen et al (1997). Cultures of Antarctic Strombidium sp. (circles), field
populations of Eutintinnus sp. (triangles), field populations of Codonellopsis sp. (diamonds) and
literature values for cultures of temperate ciliates (squares).
136
assemblages (0.0510 and 0.0354 for Strombidium vs. 0.0135 and 0.0252 for Eutintinnus
and Codonellopsis, respectively, Table 2).
Growth rates of the Antarctic Strombidium sp. were low, and varied with the type
of algae used as prey (Fig 5). The growth rates were calculated from the slope of the line
of natural log cell abundance vs. time (expressed in d
-1
) when the culture was in
logarithmic phase growth (see example, Fig 5a). Growth rates varied from 0.21 d
-1

(when Pyramimonas sp. was used as food) to -0.03 d
-1
(P. glacialis) (Fig 5b). Positive
growth of the ciliate was supported by Pyramimonas sp., Thalassiosira sp. 1 and sp 2.
and Mallomonas sp. Net cell loss was observed when ciliates were fed Chlamydomonas
sp. and P. glacialis. Growth rates were similar when the Strombidium sp. was fed the
two strains of Thalassiosira sp (0.16 and 0.13 d
-1
).
Discussion
Ingestion rates of the total microzooplankton assemblage were similar among all
four stations when Thalassiosira sp. were added at low abundance ( ≤5x10
6
cells L
-1
) (Fig
1). Rates ranged from 0.07-0.21 Thalassiosira sp. ingested microzooplankton
-1
h
-1
when
Thalassiosira sp. were added at 10
6
cells L
-1
and from 0.3-0.8 Thalassiosira sp. ingested
microzooplankton
-1
h
-1
when prey were added at 5x10
6
cells L
-1
. These two abundances
of added Thalassiosira sp. were ≤10% of the natural phytoplankton abundance. These
ingestion rates on these labeled cells were used to calculate ingestion rates on the total
phytoplankton assemblage using the total phytoplankton abundance at each station. This
calculation yielded ingestion rates ranging from 2-10 ingested algal cells
microzooplankton
-1
h
-1
.
137
3
4
5
6
7
8
0 5 10 15 20 25 30
-0.05
0
0.05
0.1
0.15
0.2
0.25
Growth Rate = 0.21 d
-1
r
2
= 0.99
a b c d e
f
0.21
0.16
0.13
0.04
-0.005 -0.03
A
B
Time (days)
Figure 5-5. Sample growth rate of the Antarctic Strombidium sp. feeding on Pyramimonas sp. (A).
A comparison of growth rates of the Antarctic Strombidium sp. feeding on six species of Antarctic
algae (B): Pyramimonas sp. (a), Thalassiosira sp. 1 (b), Thalassiosira sp. 2 (c), Mallomonas sp. (d),
Chlamydomonas sp. (e), Polarella glacialis (f).
138
The information obtained for microzooplankton counts, chlorophyll concentration in µg
L
-1
, the ingestion rates obtained in these experiments and assuming an average
carbon:chlorophyll ratio for all four experiments were used to calculate the percent
standing stock of phytoplankton grazed each day by the microzooplankton assemblage.
Chlorophyll values were 4.79, 4.28, 3.27 and 3.00 µg chl L
-1
at stations 1-4, respectively
(C. Sobrino, personal comm.) The total phytoplankton ingestion rate was calculated by
multiplying the average number of ingested fluorescently labeled cells microzooplankton
-
1
h
-1
by the ratio of labeled phytoplankton: total phytoplankton at each station.
Microzooplankton abundance ranged from 5.1-19x10
3
cells
-1
L
-1
(Table 1). The total
phytoplankton standing stock was estimated using a carbon:chlorophyll ratio of 111
obtained from the Ross Sea in January (Smith et al. 1996). The total phytoplankton
carbon grazed per day was estimated by multiplying the ingestion rate for the total
phytoplankton assemblage (in g C d
-1
) by the total number of microzooplankton. The
percentage of phytoplankton standing stock grazed per day was estimated by dividing the
total phytoplankton carbon grazed per day by the phytoplankton standing stock.
These calculations yielded estimates of 3, 1, 7 and 3% of the phytoplankton
standing stock grazed by the microzooplankton assemblage per day in the four
experiments, respectively. The carbon:chlorophyll ratio used was based on total
measurements of particulate carbon, which assumed that all particulate carbon was in
phytoplankton biomass. In the Ross Sea in January, a substantial proportion of protistan
biomass is heterotrophic (Dennett et al. 2001). However, even if the carbon:chlorophyll
ratio overestimated phytoplankton carbon by 50%, the microzooplankton assemblage still
139
only grazed between 2 and 15% of the phytoplankton standing stock per day. Rates of
phytoplankton production are low in the Antarctic relative to rates measured in temperate
systems at higher temperatures. Thus, grazing rates of 7-15% of the standing stock of
phytoplankton could represent significant portions of total daily primary production in
late austral summer in the Ross Sea. However, the grazing rates between 1 and 3%
estimated for three of the four experiments were much less likely to significantly impact
phytoplankton assemblages, suggesting that even though microzooplankton were
abundant at all stations, they generally did not exert strong grazing pressure on the total
phytoplankton assemblage.
Estimates of ingestion rate are based on several assumptions that must be
considered when interpreting experimental data. First, it is assumed that the fluorescently
labeled algae are consumed at the same rate as other phytoplankton in a natural
assemblage. A common consideration when using the stain DTAF to fluorescently label
prey is the potential for grazing selection against heat-killed prey by microzooplankton
(Landry et al. 1991). We instead chose to use Cell Tracker Green to label prey so that
algal cells were stained live and added live to mixed microzooplankton assemblages. The
presence of live, labeled prey should have reduced feeding selectivity by
microzooplankton against dead prey cells. The second major assumption in short-term
ingestion rate experiments is that the fluorescently labeled algae added are representative
of the algae present in the mixed plankton assemblage, so ingestion rates calculated based
on the fluorescently labeled prey are similar to ingestion rates on the natural algal
assemblage. If labeled prey were a better food source than the phytoplankton available in
140
the mixed plankton community, our estimates of ingested algal prey microzooplankton
-1

would be higher than the actual ingestion rates on the dominant phytoplankton taxa.
Conversely, if the labeled prey were of poor food quality relative to naturally-occurring
phytoplankton, our ingestion rate calculations would underestimate actual ingestion rates
on the dominant phytoplankton taxa. The third major assumption in short-term ingestion
rate experiments is that ingestion of fluorescently labeled prey by microzooplankton
occurs linearly throughout the duration of the experiment. Based on the laboratory
experiments with the cultured Antarctic ciliate, we believe the microzooplankton in the
natural plankton assemblages were ingesting cells linearly throughout ingestion rate
experiments in the field.
One disadvantage of the Cell Tracker Green stain is that not every algal taxa will
fluoresce or retain fluorescence upon preservation. We were unable to stain any of
several cultured strains of Phaeocystis antarctica with the Cell Tracker Green, so instead
used Thalassiosira sp. to approximate grazing on small phytoplankton present in the
mixed assemblage. The numerically dominant phytoplankton in the mixed assemblage
was the pennate diatom Nitzschia sp. The very long, very narrow morphology of this
particular species (~ 50-100 x 3-5 µm) difficult for microzooplankton to ingest and thus
of poor food quality. The second most dominant phytoplankton species at all four
stations was Phaeocystis antarctica, which is primarily in colonial form during mid- to
late austral summer (Smith et al. 2003). The colonial forms of Phaeocystis spp. are
grazed inefficiently, if at all by micro- and metazooplankton (Haberman et al. 2003, Tang
2003). It is therefore quite likely that the Thalassiosira sp. that were added to the mixed
141
microzooplankton assemblage were a better food source than the dominant phytoplankton
taxa and may have been grazed preferentially during the measurements of ingestion rate.
If the labeled algae were grazed preferentially, then even alga added at ‘tracer’
abundances would overestimate ingestion rates on natural phytoplankton.
The results of this study are consistent with low phytoplankton grazing rates
reported for the Ross Sea obtained using the dilution method (Caron et al. 2000). Those
authors reported detectable grazing rates in only 13 out of 51 dilution experiments during
austral spring, summer and fall in the Ross Sea polynya. The highest grazing rate
reported for experiments in this study was 0.26 d
-1
. The authors suggested that low
temperature and poor phytoplankton food quality were the likely causes of low grazing
rates. Our grazing rates were measured using an algal species which does not have the
same morphological features such as large size and colony formation that have been
hypothesized to make common Antarctic diatom species and Phaeocystis antarctica
unpalatable to microzooplankton grazers. In addition, by using the vital stain CMFDA to
fluorescently label algal prey the problem of underestimation of grazing rates due to
selective avoidance of dead prey cells should have been avoided. The low grazing rates
that we observed in this study were thus more likely due to inhibition of the
microzooplankton by physical or chemical factors such as low temperature than by poor
food quality of the fluorescently labeled algae. There is evidence that grazing rates of
microzooplankton are inversely related to temperature (Caron et al. 2000). There is also
recent evidence to suggest that growth rates of heterotrophic protists and copepods are
more strongly affected by low environmental temperature than the growth rates of
142
phototrophic protists (Rose & Caron submitted). It is likely that the extreme low
temperature of the Ross Sea, Antarctica significantly constrained microzooplankton
grazing during this study.
Ingestion rates of Codonellopsis sp. and Eutintinnus sp. were similar among
experiments within each genus when Thalassiosira sp. were added at low abundance
( ≤5x10
6
cells L
-1
) (Fig 2). Eutintinnus sp. ingestion rates were 0.06-0.7 Thalassiosira sp.
ingested ciliate
-1
h
-1
when Thalassiosira sp. were added at 10
6
and 5x10
6
cells L
-1
at
stations 2 and 4. Codonellopsis sp. ingestion rates were 0.12-1.03 Thalassiosira sp.
ingested ciliate
-1
h
-1
when Thalassiosira sp. were added at 10
6
and 5x10
6
cells L
-1
during
Experiments 2 and 4. Specific ingestion rates for the two ciliate genera were similar
between experiments and between genera, ranging from 0.0001-0.0013 h
-1
when
Thalassiosira sp. were added at 10
6
and 5x10
6
cells L
-1
. Differences in ingestion rate
between the two ciliates were likely due to size differences, as the Eutintinnus sp. was
127x26 µm on average, while the Codonellopsis sp. was 238x55 µm on average.
Ingestion rates of the total microzooplankton assemblage, Eutintinnus sp. and
Codonellopsis sp. were much higher in treatments with higher abundances of added
Thalassiosira sp. Ingestion rates at the highest level of added Thalassiosira were also
variable between stations both for the total microzooplankton assemblage as well as the
individual genera. These highest ingestion rates measured cannot be technically
considered maximum ingestion rates since ingestion rates were still increasing at the
highest abundance of added Thalassiosira sp. However, at the highest abundance of
algae added at all four stations (5x10
7
cells L
-1
), ingestion rates increased consistently
143
among Experiments 1-4. Increases in ingestion rates by the total microzooplankton
assemblage among Experiments 1-4 could potentially be explained by changes in
microzooplankton assemblage composition. Individual microzooplankton taxa have
different ingestion rates (e.g. Fig 2, Table 2), so the composition of the microzooplankton
assemblage can play a large role in determining the ingestion rate of the entire
assemblage. As illustrated in Table 1, the total abundance of microzooplankton varied by
nearly 400% over the four experiments and different components of the
microzooplankton assemblage dominated numerically at the time of the different
experiments.
Both ingestion rates and specific ingestion rates of each of the two numerically
dominant ciliate genera increased between experiments 2 and 4. Ingestion rates increased
from 2.3 to 7.2 algal cells Eutintinnus
-1
h
-1
and from 10.7 to 40.3 algal cells
Codonellopsis
-1
h
-1
between experiments 2 and 4. Specific ingestion rates of the
Eutintinnus sp. increased from 0.0043 to 0.0135 h
-1
and rates of the Codonellopsis sp.
increased from 0.0067 to 0.0252 between stations 2 and 4. We believe that the increases
are more likely reflective of changes in physiological state of the individual
microzooplankton taxa between experiments 2 and 4, based on the following reasoning:
The increases in ingestion rates between stations were consistent between genera (and
also consistent with increases observed in ingestion rates for the total microzooplankton
assemblage). In addition, increasing ingestion rate was also accompanied by a nearly
50% decrease in total phytoplankton abundance and a 400% increase in total
microzooplankton abundance between stations 2 and 4. These results are consistent with
144
a scenario in which the numerically dominant members of the microzooplankton
assemblage fed and grew on the phytoplankton bloom, but became increasingly starved
as phytoplankton abundance declined at this station.
The laboratory experiments with a culture of Antarctic ciliate, Strombidium sp.
also support the hypothesis that ingestion rates can be affected by physiological state of
the grazer. When Strombidium sp. was starved for two weeks before the addition of a
high density of prey, ingestion rates were much higher than when the ciliate culture was
in balanced growth (Fig 3). The specific ingestion rates of the starved ciliate were so
high that they fell within the range of maximal specific ingestion rates reported for
temperate ciliates at much higher temperatures (Fig 4). In contrast, the specific ingestion
rates of the ciliate culture in balanced growth were 19-27% of the rates of the starved
ciliates, were nearly an order of magnitude lower than specific ingestion rates reported
for the temperate ciliates, and were similar to the specific ingestion rates calculated for
the Eutintinnus sp. and Codonellopsis sp. in the natural plankton assemblages at station 4.
It is important to note that the high ingestion rates observed for the cultured
Antarctic Strombidium sp. did not translate into rapid growth rates (Fig 5). Growth rates
of the ciliate were consistently low. The Strombidium sp. grew relatively well on the
Thalassiosira sp. that was used in the ingestion rate experiments, however, growth rates
were low when compared to rates reported for temperate Strombidium sp. feeding on
algal prey (Jonsson 1986, Montagnes 1996). The very high ingestion rates of the starved
ciliate when taken out of context seem to indicate temperature compensation by the
Antarctic ciliate, since these rates are roughly equivalent to rates reported for temperate
145
ciliates at much higher temperatures. However, the subsequent comparison of ingestion
and growth rates measured during balanced growth clearly indicate that the Antarctic
ciliate was ingesting and processing algal cells much more slowly. These results
highlight the need for caution when interpreting ingestion rate data, especially from
natural assemblages whose physiological state is unknown.
Conclusions
Ingestion rates of the natural microzooplankton assemblages in the Ross Sea,
Antarctica were low at all stations, whether fluorescently labeled prey were added at
tracer levels or at high abundance. The estimated grazing pressure on the phytoplankton
assemblage, calculated using chlorophyll measurements, total abundances of
microzooplankton and measurements of grazing rates, was also low at all four stations.
Specific ingestion rates of two numerically dominant microzooplankton taxa were also
low, regardless of abundance of added prey, when compared to rates for temperate
microzooplankton. These results indicate that even though microzooplankton were
present in high abundance in the Ross Sea, they did not exert significant grazing pressure
on the phytoplankton assemblage. Reduced top-down control by the microzooplankton
may contribute to the formation and persistence of annual phytoplankton blooms in this
ecosystem.

146
Dissertation Conclusions
A literature review of protistan growth rates indicated that maximal growth rates
of heterotrophs (including bacterivorous protists, herbivorous protists and copepods)
were more strongly affected by decreases in temperature than maximal growth rates of
phototrophic protists. At temperatures below ~15˚C, maximal growth rates of
herbivorous protists were consistently lower than maximal growth rates attainable by
phototrophic protists. At the extreme low temperatures common to polar environments,
the maximal growth rates of herbivorous protists as predicted by the regression were
approximately one third the maximal growth rates of phototrophic protists as predicted by
the Eppley curve. These results have important implications for plankton dynamics in
cold ecosystems. If the maximal growth rates of phototrophic protists outpace the
maximal growth rates of herbivorous protists at low temperature, this may help to explain
how phototrophic protists escape top-down control to form the massive annual algal
blooms common to high latitude ecosystems.
While growth rates of heterotrophic protists reported in the literature span a wide
range of temperatures and include strains isolated from temperate, tropical and polar
environments, the majority of growth rates published were measured on temperate strains
at temperatures between 10-25˚C. Relatively few growth rates of polar heterotrophic
protists or of temperate protists measured at extreme low temperature (< 5˚C) have been
reported, and even less information has been published about other physiological
parameters such as growth efficiency, ingestion rate, remineralization rate or
remineralization efficiency. I examined the effect of temperature on a cultured Antarctic
147
bacterivorous protist, Paraphysomonas imperforata, to establish baseline information
about the effect of temperature and prey type on growth rate, gross growth efficiency as
well as nutrient regeneration rates and efficiencies of the Antarctic P. imperforata. I also
compared this data with physiological information published for temperate congeners and
conspecifics of this globally-distributed heterotrophic protist.
Growth and nutrient regeneration rates of the Antarctic P. imperforata increased
rapidly with increased temperature. Q
10
values for growth rates of the bacterivorous
protist between 0 and 5˚C were 6.9 and 15 during two separate experiments.
Regeneration of nutrients also occurred more rapidly as experimental temperatures
increased from 0 to 5˚C and from 5 to 10˚C. Classical Q
10
values describing increases in
physiological rate with increased temperature have a value of approximately 2 when
organisms are not physiologically stressed by temperature, but values much higher than 2
have been reported for organisms growing below their optimal temperature range (values
between 0 and 1 are reported for organisms growing above their optimal temperature
range, as physiological functioning decreases rapidly with increased temperature). The
high Q
10
values observed at the lowest temperatures in this study suggest that the
Antarctic bacterivorous protist was physiologically stressed at the low (but
environmentally pertinent) temperatures used in the experiment.
Gross growth and nutrient regeneration efficiencies were not affected by temperature
in the experiments. Gross growth efficiencies of heterotrophic protists have been
reported in the literature to increase, decrease and stay constant with increased
temperature. Published gross growth efficiencies for species within the single genus
148
Paraphysomonas also span a huge range, from 2-80%, and this range has been reported
across a 20
o
C temperature range. The median of these literature values (grouped into 5
o
C
temperature bins) were similar to the gross growth efficiencies for the Antarctic P.
imperforata reported here. The gross growth efficiencies observed here span a much
narrower range (28-53% for all experiments). The relatively large range of published
growth efficiencies is likely artifactual, due to the difficulties of measuring gross growth
efficiency even in controlled laboratory settings.
Both growth rates and gross growth efficiencies of the Antarctic P. imperforata could
be predicted by the extrapolation of data published for temperate congeners. These
results suggest that the P. imperforata from the Antarctic exhibited a similar
physiological ecology (at the whole-cell level) as other morphologically-related
Paraphysomonas spp. from temperate and tropical regions. When growth rates of the
Antarctic P. imperforata were compared to published values for other species within the
genus Paraphysomonas, the Antarctic P. imperforata had similar growth rates to those
already published at the lowest and highest temperatures (0, 5 and 20˚C) and the highest
growth rate reported at 10 and 15˚C. In addition, the trend of rapidly increased maximal
growth rates with increasing temperature observed in the literature review was mirrored
in this minireview of growth rates of Paraphysomonas spp. A comparison of the gross
growth efficiencies of the Antarctic P. imperforata to literature values for
Paraphysomonas sp. revealed a wide range of published gross growth efficiencies.
However, mean and median values for gross growth efficiencies grouped by 5˚C
temperature bins did not steadily increase or decrease with increased temperature, which
149
was consistent with our observation of no effect of temperature on gross growth
efficiency for the Antarctic P. imperforata.
While few studies have examined growth rates of temperate and polar bacterivorous
protists at extreme low temperature, even fewer studies have examined the physiological
rates of cultured herbivorous protists at low temperature. I was unable to obtain any
published growth rates of cultured herbivorous protists measured at temperatures below
5˚C. Very few papers have been published examining growth and grazing by natural
assemblages of herbivorous protists from the Antarctic. By combining laboratory
measurements of growth and ingestion rates of a cultured Antarctic ciliate with field
measurements of ingestion rates of natural assemblages of herbivorous protists, I sought
to establish baseline information for Antarctic herbivorous protists as well as determine
whether low predicted growth rates was due in part to low ingestion rates by these
grazers.
Ingestion rates by the natural assemblages of herbivorous protists were universally
low during four experiments in austral summer in the Ross Sea, Antarctica.
Microzooplankton abundance was high in all experiments, but ingestion rates were so
low that the estimated grazing impact on phytoplankton standing stocks at during the four
experiments was 3, 1, 7 and 3% per day, respectively. While rates of phytoplankton
production are low in the Ross Sea relative to more temperate ecosystems, it is still likely
that only the protistan grazers in Experiment 3 (7% of phytoplankton standing stock
grazed per day) had a significant impact on the phytoplankton assemblages.
150
When high abundances of fluorescently labeled algae were added to the natural
plankton assemblages, ingestion rates of the total herbivorous microzooplankton
assemblage increased but never approached a maximum. These results indicate that a
combination of high microzooplankton abundance and either low natural prey abundance
or poor food quality prevented us from saturating ingestion rates of the microzooplankton
species present. All four experiments were carried out during a late summer
phytoplankton bloom in the Ross Sea, which was dominated by a mixture of Phaeocystis
antarctica and a small pennate diatom. Prey abundance at all four experimental sites was
high (>10
7
cells L
-1
), suggesting that food limitation of microzooplankton was not due to
low predator-prey encounter rates. The colonial morphology common to Phaeocystis
spp. has been hypothesized to be deter grazers, and the very long, narrow morphology of
the pennate diatom (~50-100 um x 3-5 μm) may have instead rendered both dominant
phytoplankton taxa unpalatable to the microzooplankton assemblage.
Ingestion rates of two numerically-dominant ciliate genera were measured in the Ross
Sea in late austral summer during two experiments conducted at the same location but
separated in time by two weeks. Maximum ingestion rates were only observed for one of
the two genera, Eutintinnus sp., during one of the two experiments. These results suggest
that the natural food limitation (likely due to poor prey quality as explained above)
observed in these experiments was occurring not only for the microzooplankton
assemblage as a whole, but also the two dominant genera within the assemblage.
Specific ingestion rates calculated for these two ciliate genera using the volumes of
the ciliates and of the fluorescently labeled algae were at least an order of magnitude
151
lower than rates published for temperate ciliates measured at higher temperatures. The
specific ingestion rates of the two Antarctic ciliates should logically have been lower than
maximal specific ingestion rates published for temperate ciliates, because maximal
ingestion rates of the two Antarctic ciliates were not obtained in either experiment.
Therefore higher specific ingestion rates of the Antarctic ciliates were theoretically
possible. However, the order of magnitude difference between specific ingestion rates of
the Antarctic and temperate ciliate genera suggest that the low ingestion rates observed
for the Antarctic ciliates were not simply caused by non-saturating prey abundance in the
experiments. It is possible that low environmental temperature also contributed to the
relatively low specific ingestion rates observed for the total microzooplankton
assemblage as well as the individual microzooplankton genera.
An increased number of microzooplankton and decreased phytoplankton abundance
paralleled an increase in specific ingestion rate observed in the two ingestion rate
experiments conducted in the same location within the Ross Sea but separated in time by
two weeks. Specific ingestion rates calculated for the individual genera varied from
0.004 vs. 0.013 h
-1
for Eutintinnus sp. and 0.007 vs. 0.013 h
-1
for Codonellopsis sp. when
prey were added at 5x10
7
cells L
-1
. At the same time, microzooplankton abundance
quadrupled and total phytoplankton abundance decreased by nearly 50%. These data
support the hypothesis that the inability to saturate microzooplankton ingestion rate was
due to food limitation of the microzooplankton assemblage in these experiments.
The hypothesis of food limitation affecting specific ingestion rate was further
supported by laboratory experiments with a culture of Antarctic Strombidium sp. When
152
ciliates were starved for two weeks, then fed a high abundance of prey, the specific
ingestion rate measured was comparable to rates published for temperate ciliates at much
higher temperature (0.2 vs. a range of 0.1-0.4 h
-1
). In contrast, specific ingestion rates
measured on the same ciliate culture in balanced growth were an order of magnitude
lower, and were instead comparable to rates measured for the Antarctic ciliates in the
field. These data highlight the need for caution in interpretation of field estimates of
ingestion rate, because the physiological state of a natural microzooplankton assemblage
is generally unknown.
In summary, low temperature exerts a strong constraint on growth and grazing by
heterotrophic protists and this is a fundamental determinant of plankton processes and
dynamics in polar ecosystems. Published values for maximal growth rates of
heterotrophic protists were more strongly affected by decreases in temperature than
maximal growth rates of phototrophic protists. The same pattern was observed in growth
rates of a cultured Antarctic strain of the bacterivorous protist Paraphysomonas
imperforata, as well as when the growth rates of this Antarctic protist were compared to
rates published for temperate and polar congeners. The Antarctic P. imperforata grew
slowly at low temperature but gross growth efficiency was unchanged. Neither gross
growth nor nutrient regeneration efficiencies were affected by increased temperature.
Ingestion rates of Antarctic microzooplankton in natural assemblages were consistently
low, as were ingestion rates of a cultured Antarctic ciliate in balanced growth. These low
ingestion rates resulted in generally low estimates of grazing impact on phytoplankton
standing stocks in the Ross Sea during austral summer, even though total
153
microzooplankton abundance was relatively high. The constraints imposed by low
temperature on growth and grazing of heterotrophic protists may contribute to the
formation of the massive, spatially extensive annual algal blooms common in high
latitude environments.
154
Bibliography

Ackleson SG, Spinrad RW (1988) Size and refractive index of individual marine
particulates: a flow cytometric approach. Applied Optics 27:1270

Andersen OK, Goldman JC, Caron DA, Dennett MR (1986) Nutrient cycling in a
microflagellate food chain: III. Phosphorus dynamics. Marine Ecology Progress
Series 31:47-55

Atkinson A (1995) Omnivory and feeding selectivity in five copepod species during
spring in the Bellinghausen Sea, Antarctica. ICES Journal of Marine Science
52:385-396

Atkinson A (1996) Subantarctic copepods in an oceanic, low chlorophyll environment:
ciliate predation, food selectivity and impact on prey populations. Marine Ecology
Progress Series 130:85-96

Azam F, Fenchel T, Field JG, Gray JS, Meyer-Reil LA, Thingstad F (1983) The
ecological role of water-column microbes in the sea. Marine Ecology Progress
Series 10:257-263

Baldock BM, Baker JH, Sleigh MA (1980) Laboratory growth rates of six species of
freshwater Gymnamoebia. Oecologia 47:156-159

Baldock BM, Berger J (1984) The effects of low temperatures on the growth of four
fresh-water amoebae (Protozoa: Gymnamoebia). Transactions of the American
Microscopical Society 103:233-239

Becquevort S (1997) Nanoprotozooplankton in the Atlantic sector of the Southern Ocean
during early spring: biomass and feeding activities. Deep-Sea Research II 44:355-
373

Becquevort S, Menon P, Lancelot C (2000) Differences of the protozoan biomass and
grazing during spring and summer in the Indian sector of the Southern Ocean.
Polar Biology 23:309-320

Beers JR, Reid FMH, Stewart GL (1975) Microplankton of the North Pacific Central
Gyre. Population structure and abundance, June 1973. Int Revue Ges Hydrobiol
60:607-638

Beers JR, Stewart GL (1971) Micro-zooplankters in the plankton communities of the
upper waters of the easter tropical Pacific. Deep-Sea Research I 18:861-883

155
Bratvold D, Srienc F, Taub SR (2000) Analysis of the distribution of ingested bacteria in
nanoflagellates and estimation of grazing rates with flow cytometry. Aquatic
Microbial Ecology 21:1-12
Buck KR, Garrison DL (1983) Protists from the water column of the Weddell Sea ice
edge. Deep-Sea Research I 30:1261-1277

Burkill PH, Edwards ES, John AWG, Sleigh MA (1993) Microzooplankton and their
herbivorous activity in the northeastern Atlantic Ocean. Deep-Sea Research II
40:479-493

Campbell L (2001) Flow cytometric analysis of autotrophic picoplankton. In: Methods in
Microbiology, Vol 30. Academic Press, p 317-341

Caron DA, Dam HG, Kremer P, Lessard EJ, Madin LP, Malone TC, Napp JM, Peele ER,
Roman MR, Youngbluth MJ (1995) The contribution of microorganisms to
particulate carbon and nitrogen in surface waters of the Sargasso Sea near
Bermuda. Deep-Sea Research I 42:943-972

Caron DA, Dennett MR, Lonsdale DJ, Moran DM, Shalapyonok L (2000)
Microzooplankton herbivory in the Ross Sea, Antarctica. Deep-Sea Research II
47:3249-3272

Caron DA, Goldman JC (1990) Protozoan nutrient regeneration. In: Capriulo GM (ed)
Ecology of marine protozoa. Oxford University Press, New York, p 283-306

Caron DA, Goldman JC, Andersen OK, Dennett MR (1985) Nutrient cycling in a
microflagellate food chain: II. Population dynamics and carbon cycling. Marine
Ecology Progress Series 24:243-254

Caron DA, Goldman JC, Dennett MR (1986) Effect of temperature on growth, respiration
and nutrient regeneration by an omnivorous microflagellate. Applied and
Environmental Microbiology 52:1340-1347

Caron DA, Lim EL, Miceli G, Waterbury JB, Valois FW (1991) Grazing and utilization
of chroococcoid cyanobacteria and heterotrophic bacteria by protozoa in
laboratory cultures and a coastal plankton community. Marine Ecology Progress
Series 76:205-217

Caron DA, Lim EL, Sanders RW, Dennett MR, Berninger UG (2000) Responses of
bacterioplankton and phytoplankton to organic carbon and inorganic nutrient
additions in contrasting oceanic ecosystems. Aquatic Microbial Ecology 22:175-
184

156
Carpenter EJ, Chang J, Shapiro LP (1991) Green and blue fluorescing dinoflagellates in
Bahamian waters. Marine Biology 108:145-149

Chavez F, Buck KR, Coale KH, Martin JH, DiTullio GR, Welschmeyer NA, Jacobson
AC, Barber RT (1991) Growth rates, grazing, sinking, and iron limitation of
equatorial Pacific phytoplankton. Limnology and Oceanography 36:1816-1833
Choi JW, Peters F (1992) Effects of temperature on two psychrophilic ecotypes of a
heterotrophic nanoflagellate, Paraphysomonas imperforata. Applied and
Environmental Microbiology 58:593-599

Cotner JB, Ammerman JW, Peele ER, Bentzen E (1997) Phosphorus-limited
bacterioplankton growth in the Sargasso Sea. Aquatic Microbial Ecology 13:141-
149

Cucci TL, Shumway SE, Brown WS, Newell CR (1989) Using phytoplankton and flow
cytometry to analyze grazing by marine organisms. Cytometry 10:659-669

Cucci TL, Sieracki ME (2001) Effects of mismatched refractive indices in aquatic flow
cytometry. Cytometry 44:173-178

Dale T, Burkill PH (1982) "Live counting" - a quick and simple technique for
enumerating pelagic ciliates. Annales de L'institut Océanographique, Paris
58(S):267-276

Daugbjerg N (2001) Pyramimonas tychotreta, sp. nov. (Prasinophyceae), a new marine
species from Antarctica: light and electron microscopy of the motile stage and
notes on growth rates. Journal of Phycology 36:160-171

Davidson AT, Marchant HJ (1992) Protist abundance and carbon concentration during a
Phaeocystis-dominated bloom at an Antarctic coastal site. Polar Biology 12:387-
395

del Giorgio PA, Bird DF, Prairie YT, Planas D (1996) Flow cytometric determination of
bacterial abundance in lake plankton with the green nucleic acid stain SYTO 13.
Limnology and Oceanography 41:783-789

Dennett MR, Mathot S, Caron DA, Smith WO, Jr, Lonsdale DJ (2001) Abundance and
distribution of phototrophic and heterotrophic nano- and microplankton in the
southern Ross Sea. Deep-Sea Research II 48:4019-4037

DiTullio GR, Grebmeier J, Arrigo KR (2000) Rapid and early export of Phaeocystis
antarctica blooms in the Ross Sea, Antarctica. Nature 404:595-598

157
Dolan JR (1997) Phosphorus and ammonia excretion by planktonic protists. Marine
Geology 139:109-122

Ducklow H, Carlson C, Church M, Kirchman DL, Smith D, Steward G (2001) The
seasonal development of the bacterioplankton bloom in the Ross Sea, Antarctica,
1994-1997. Deep-Sea Research II 48:4199-4221

Eccleston-Parry JD, Leadbeater BS (1995) Regeneration of phosphorus and nitrogen by
four species of heterotrophic nanoflagellates feeding on three nutritional states of
a single bacterial strain. Applied and Environmental Microbiology 61:1033-1038

Eppley RW (1972) Temperature and phytoplankton growth in the sea. Fishery Bulletin
70:1063-1085

Fenchel T (1982) Ecology of heterotrophic microflagellates. II. Bioenergetics and
growth. Marine Ecology Progress Series 8:225-231

Finlay B (1977) The dependence of reproductive rate on cell size and temperature in
freshwater ciliated protozoa. Oecologia 30:75-81

Frost BW (1987) Grazing control of phytoplankton stock in the open subarctic Pacific
Ocean: a model assessing the role of mesozooplankton, particularly the large
calanoid copepods, Neocalanus spp. Marine Ecology Progress Series 39:49-68

Frost BW (1991) The role of grazing in nutrient-rich areas of the open sea. Limnology
and Oceanography 36:1616-1630

Garrison DL (1991) An overview of the abundance and role of protozooplankton in
Antarctic waters. Journal of Marine Systems 2:317-331

Garrison DL, Buck KR (1989) Protozooplankton in the Weddell Sea, Antarctica:
abundance and distribution in the ice-edge zone. Polar Biology 9:341-351

Garrison DL, Sullivan CW, Ackley SF (1986) Sea ice microbial communities in
Antarctica. BioScience 36:243-250

Gerritsen J, Sanders RW, Bradley SW, Porter KG (1987) Individual feeding variability of
protozoan and crustacean zooplankton analyzed with flow cytometry. Limnology
and Oceanography 32:691-699

Gifford DJ, Caron DA (2000) Sampling, preservation, enumeration and biomass of
marine zooplankton. In: Harris R, Wiebe P, Lenz J, Skjoldal HR, Huntley M (eds)
ICES Zooplankton Methodolody Manual. Academic Press, San Diego, p 193-221

158
Gifford DJ, Fessenden LM, Garrahan PR, Martin E (1995) Grazing by microzooplankton
and mesozooplankton in the high-latitude North Atlantic Ocean: spring versus
summer dynamics. Journal of Geophysical Research 100:6665-6675

Goldman JC, Caron DA (1985) Experimental studies on an omnivorous microflagellate:
implications for grazing and nutrient regeneration in the marine microbial food
chain. Deep-Sea Research I 32:899-915
Goldman JC, Caron DA, Andersen OK, Dennett MR (1985) Nutrient cycling in a
microflagellate food chain: I. Nitrogen dynamics. Marine Ecology Progress Series
24:231-242

Goldman JC, Carpenter EJ (1974) A kinetic approach to the effect of temperature on
algal growth. Limnology and Oceanography 19:756-766

Gonzalez JM, Iriberri J, Egea L, Barcina I (1990) Differential rates of digestion of
bacteria by fresh-water and marine phagotrophic protozoa. Applied and
Environmental Microbiology 56:1851-1857

Gowing MM, Garrison DL (1992) Abundance and feeding ecology of larger
protozooplankton in the ice edge zone of the Weddell and Scotia Seas during the
austral winter. Deep-Sea Research I 39:893-919

Guillard RRL (1975) Culture of phytoplankton for feeding marine invertebrates. In:
Smith WL, Chanley MH (eds) Culture of Marine Invertebrate Animals. Plenum
Press, New York

Guillard RRL, Ryther JH (1962) Studies of marine planktonic diatoms. I. Cyclotella nana
Hustedt and Detonula confervacea Cleve. Canadian Journal of Microbiology
8:229-239

Haberman KL, Quetin LB, Ross RM (2003) Diet of the Antarctic krill (Euphausia
superba Dana): I. Comparisons of grazing on Phaeocystis antarctica (Karsten)
and Thalassiosira antarctica (Comber). Journal of Experimental Marine Biology
and Ecology 283:79-95

Hansen PJ, Bjornsen PK, Hansen BW (1997) Zooplankton grazing and growth: Scaling
within the 2-2,000um body size range. Limnology and Oceanography 42:687-704

Hargrave BT, Geen GH (1968) Phosphorus excretion by zooplankton. Limnology and
Oceanography 13:332-342

Hazel JR (1995) Thermal adaptation in biological membranes: is homeoviscous
adaptation the explanation? Annual Review of Physiology 57:19-42

159
Hochachka P, Somero G (2002) Biochemical Adaptation, Vol. Oxford University Press,
New York

Hochberg Y (1988) A sharper Bonferroni procedure for multiple tests of significance.
Biometrika 75:800-802

Huntley ME, Lopez MDG (1992) Temperature-dependent production of marine
copepods: a global synthesis. The American Naturalist 140:201-242
Ikeda T, Hing Fay E, Hutchinson SA, Boto GM (1982) Ammonia and inorganic
phosphate excretion by zooplankton from inshore waters of the Great Barrier
Reef, Queensland. I. Relationship between excretion rates and body size.
Australian Journal of Marine and Freshwater Research 33

Ishigaki T, Sleigh MA (2001) Grazing characteristics and growth efficiencies at two
different temperatures for three nanoflagellates fed with Vibrio bacteria at three
different concentrations. Microbial Ecology 41:264-271

Jonsson PR (1986) Particle size selection, feeding rates and growth dynamics of marine
planktonic oligotrichous ciliates (Ciliophora:Oligotrichina). Marine Ecology
Progress Series 33:265-277

Keller MD, Shapiro LP, Haugen EM, Cucci TL, Sherr EB, Sherr BF (1994) Phagotrophy
of fluorescently labeled bacteria by an oceanic phytoplankter. Microbial Ecology
28:39-52

Klaas C (1997) Microzooplankton distribution and their potential grazing impact in the
Antarctic Circumpolar Current. Deep-Sea Research II 44:375-393

Landry MR, Lehner-Fournier JM, Sundstrom JA, Fagerness VL, Selph KE (1991)
Discrimination between living and heat-killed prey by a marine zooflagellate,
Paraphysomonas vestita (Stokes). Journal of Experimental Marine Biology and
Ecology 146:139-151

Laybourn J, Stewart JM (1975) Studies on consumption and growth in the ciliate
Colpidium campylum Stokes. Journal of Animal Ecology 44:165-174

Leboy PS, Cline SG, Conner RL (1964) Phosphate, purines and pyrimidines as excretory
products of Tetrahymena. Journal of Protozoology 11:217-222

Lee CC, Fenchel T (1972) Studies on ciliates associated with sea ice from Antarctica. II.
Temperature responses and tolerances in ciliates from Antarctic, Temperate and
Tropical habitats. Archiv fur Protistenk 114:237-244

160
Lessard EJ, Murrell MC (1998) Microzooplankton herbivory and phytoplankton growth
in the northwestern Sargasso Sea. Aquatic Microbial Ecology 16:173-188

Li A, Stoecker DK, Coats DW, Adam EJ (1996) Ingestion of fluorescently labeled and
phycoerythrin-containing prey by mixotrophic dinoflagellates. Aquatic Microbial
Ecology 10:139-147

Lindstrom ES, Weisse T, Stadler P (2002) Enumeration of small ciliates in culture by
flow cytometry and nucleic acid staining. Journal of Microbiological Methods
49:173-182
Liu H, Buskey EJ (2000) The exopolymer secretions (EPS) layer surrounding
Aureoumbra lagunensis cells affects growth, grazing, and behavior of protozoa.
Limnology and Oceanography 45:1187-1191

Liu RG, Singh K (1997) Notions of limiting P values based on data depth and bootstrap.
Journal of the American Statistical Association 92:266-277

Lochte K, Ducklow HW, Fasham MJR, Stienen C (1993) Plankton succession and carbon
cycling at 47
o
N 20
o
W during the JGOFS North Atlantic Bloom Experiment.
Deep-Sea Research II 40:91-114

Lonsdale DJ, Caron DA, Dennett MR, Schaffner RA (2000) Predation by Oithona spp on
protozooplankton in the Ross Sea, Antarctica. Deep-Sea Research II 47:3273-
3283

Marie D, Brussaard CPD, Thyrhaug R, Bratbak G, Vaulot D (1999) Enumeration of
marine viruses in culture and natural samples by flow cytometry. Applied and
Environmental Microbiology 65:45-52

Marie D, Partensky F, Jacquet S, Vaulot D (1997) Enumeration and cell cycle analysis of
natural populations of marine picoplankton by flow cytometry using the nucleic
acid stain SYBR Green I. Applied and Environmental Microbiology 63:186-193

Martinez EA (1980) Sensitivity of marine ciliates (Protozoa, Ciliophora) to high thermal
stress. Estuarine and Coastal Marine Science 10:369-381

Matz C, Boenigk J, Arndt H, Jurgens K (2002) Role of bacterial phenotypic traits in
selective feeding of the heterotrophic nanoflagellate Spumella sp. Aquatic
Microbial Ecology 27:137-148

Mayes DF, Rogerson A, Marchant H, Laybourn-Parry J (1997) Growth and consumption
rates of bacterivorous Antarctic naked marine amoebae. Marine Ecology Progress
Series 160:101-108

161
Menzel D, Corwin N (1965) The measurement of total phosphorus in seawater based on
the liberation of organically bound fractions by persulfate oxidation. Limnology
and Oceanography 10:280-282

Mohapatra BR, Fukami K (2004) Comparison of the numerical grazing response of two
marine heterotrophic nanoflagellates fed with different bacteria. Journal of Sea
Research 52:99-107

Montagnes DJS (1996) Growth responses of planktonic ciliates in the genera
Strobilidium and Strombidium. Marine Ecology Progress Series 130:241-254

Montresor M, Procaccini G (1999) Polarella glacialis, gen. nov., sp. nov. (Dinophyceae):
Suessiaceae are still alive! Journal of Phycology 35:186-197

Muller H, Geller W (1993) Maximum growth rates of aquatic ciliated protozoa: the
dependence on body size and temperature reconsidered. Archiv fur Hydrobiologie
126:315-327

Nagata T, Kirchman DL (1991) Release of dissolved free and combined amino-acids by
bacterivorous marine flagellates. Limnology and Oceanography 36:433-443

Nagata T, Kirchman DL (1992) Release of macromolecular organic-complexes by
heterotrophic marine flagellates. Marine Ecology-Progress Series 83:233-240

Napp JM, Kendall AW, Schumacher JD (2000) A synthesis of biological and physical
processes affecting the feeding environment of larval walleye pollock (Theragra
chalcogramma) in the eastern Bering Sea. Fisheries Oceanography 9:147-162

Olson MB, Strom SL (2002) Phytoplankton growth, microzooplankton herbivory and
community structure in the southeast Bering Sea: insight into the formation and
temporal persistence of an Emiliana huxleyi bloom. Deep-Sea Research II
49:5969-5990

Olson RJ, Frankel SL, Chisholm SW, Shapiro HM (1983) An inexpensive flow
cytometer for the analysis of fluorescence signals in phytoplankton: chlorophyll
and DNA distributions. Journal of Experimental Marine Biology and Ecology
68:129-144

Olson RJ, Zettler ER, Chisholm SW, Dusenberry JA (1991) Advances in oceanography
through flow cytometry. In: P.F. Kemp BFS, E.B. Sherr and J.J. Cole (ed)
Handbook of Methods in Aquatic Microbial Ecology. Lewis Publishers, Boca
Raton, p 175-186

162
Parsons TR, Lalli CM (1988) Comparative oceanic ecology of the plankton communities
of the subarctic Atlantic and subarctic Pacific Oceans. Oceanography and Marine
Biology Annual Review 26:317-359

Parsons TR, Maita Y, Lalli CM (1984) A manual of chemical and biological methods for
seawater analysis, Vol. Pergamon, New York

Platt T, Bird DF, Sathyendranath S (1991) Critical depth and marine primary production.
Proceedings of the Royal Society of London Series B 246:205-217

Pomeroy LR, Deibel D (1986) Temperature regulation of bacterial activity during the
spring bloom in Newfoundland coastal waters. Science 233:359-361

Porter J (1999) Flow cytometry and enviromental microbiology. In: Robinson JP (ed)
Current Protocols in Cytometry. J. Wiley and Sons, New York

Preisig HR, Hibberd DJ (1982) Ultrastructure and taxonomy of Paraphysomonas
(Chrysophyceae) and related genera I. Nordic Journal of Botany 2:397-420

Rassoulzadegan F (1982) Dependence of grazing rate, gross growth efficiency and food
size range on temperature in a pelagic oligotrichous ciliate Lohmanniella spiralis
Leeg., fed on naturally occurring particulate matter. Annales de L'institut
Océanographique 58:177-184

Riley GA (1942) The relationship of vertical turbulence and spring diatom flowerings.
Journal of Marine Research 5:67-87

Rogerson A (1981) The ecological energetics of Amoeba proteus (Protozoa).
Hydrobiologia 85:117-128

Rose JM (2006) Physiological and ecological consequences of environmental
temperature on Antarctic protists. University of Southern California

Rose JM, Caron DA (submitted) Does low temperature constrain the growth rates of
heterotrophic protists? Evidence and implications for algal blooms in cold waters.
Limnology and Oceanography

Rose JM, Vora NM, Gast RJ, Caron DA (in prep) Effects of temperature and prey type
on growth rate and gross growth efficiency of an Antarctic bacterivorous protist.

Sanders RW, Caron DA, Berninger UG (1992) Relationships between bacteria and
heterotrophic nanoplankton in marine and fresh water: an interecosystem
comparison. Marine Ecology Progress Series 86:1-14

163
Selph KE, Landry MR, Laws EA (2003) Heterotrophic nanoflagellate enhancement of
bacterial growth through nutrient remineralization in chemostat culture. Aquatic
Microbial Ecology 32:23-37

Sherr BF, Caron DA, Sherr EB (1993) Staining of heterotrophic protists for visualization
via epifluorescence microscopy. In: Kemp PF, Sherr BF, Sherr EB, Cole JJ (eds)
Handbook of Methods in Aquatic Microbial Ecology. Lewis Publishers, Boca
Raton, p 213-228

Sherr BF, Sherr EB (1994) Bacterivory and herbivory: key roles of phagotrophic protists
in pelagic food webs. Microbial Ecology 28:223-235

Sherr BF, Sherr EB, Berman T (1983) Grazing, growth and ammonium excretion rates of
a heterotrophic microflagellate fed with four species of bacteria. Applied and
Environmental Microbiology 45:1196-1201
Sherr EB, Sherr BF (2002) Significance of predation by protists in aquatic microbial food
webs. Antonie van Leeuwenhoek 81:293-308

Sieburth JM, Smetacek V, Lenz J (1978) Pelagic ecosystem structure: heterotrophic
compartments of the plankton and their relationship to plankton size fractions.
Limnology and Oceanography 23

Sieracki ME, Haas LW, Caron DA, Lessard EJ (1987) The effect of fixation on particle
retention by microflagellates: underestimation of grazing rates. Marine Ecology
Progress Series 38:251-258

Smayda TJ (1997) Harmful algal blooms: their ecophysiology and general relevance to
phytoplankton blooms in the sea. Limnology and Oceanography 42:1137-1153

Smith WO, Jr, Dennett MR, Mathot S, Caron DA (2003) The temporal dynamics of the
flagellated and colonial stages of Phaeocystis antarctica in the Ross Sea. Deep-
Sea Research II 50:605-617

Smith WO, Jr, Dunbar RB (1998) The relationship between new production and vertical
flux on the Ross Sea continental shelf. Journal of Marine Systems 17:445-457

Smith WO, Jr, Marra J, Hiscock MR, Barber RT (2000) The seasonal cycle of
phytoplankton biomass and primary productivity in the Ross Sea, Antarctica.
Deep-Sea Research II 47:3119-3140

Smith WO, Jr, Nelson DM, DiTullio GR, Leventer AR (1996) Temporal and spatial
patterns in the Ross Sea: Phytoplankton biomass, elemental composition,
productivity and growth rates. Journal of Geophysical Research 101:18455-18466

164
Soldo AT, Godoy GA, Larin F (1978) Purine-excretory nature of refractile bodies in the
marine ciliate Parauronema acutum. Journal of Protozoology 25:416-418

Soldo AT, Wagtendonk WJ (1961) Nitrogen metabolism in Paramecium aurelia. Journal
of Protozoology 8:41-55

Spinrad RW, Brown JF (1986) Relative refractive index of marine microorganisms: a
technique for flow cytometric estimation. Applied Optics 12:1930

Stoecker DK (1998) Conceptual models of mixotrophy in planktonic protists and some
ecological and evolutionary implications. European Journal of Protistology
34:281-290

Stoecker DK, Buck KR, Putt M (1993) Changes in the sea-ice brine community during
the spring-summer transition, McMurdo Sound, Antarctica. II. Phagotrophic
protists. Marine Ecology Progress Series 95:103-113
Stoecker DK, Sieracki ME, Verity PG, Michaels AE, Haugen E, Burkill PH, Edwards.
ES (1994) Nanoplankton and protozoan microzooplankton during the JGOFS N.
Atlantic Bloom Experiment: 1989 and 1990. Journal of the Marine Biological
Association of the United Kingdom 74:427-443

Stramska M, Dickey TD, Plueddemann A, Weller R, Langdon C, Marra J (1995)
Biooptical variability associated with phytoplankton dynamics in the North-
Atlantic Ocean during spring and summer of 1991. Journal of Geophysical
Research 100:6621-6632

Strathmann RR (1967) Estimating the organic carbon content of phytoplankton from cell
volume or plasma volume. Limnology and Oceanography 12:411-418

Strom SL (2000) Bacterivory: Interactions between bacteria and their grazers. In:
Kirchman DL (ed) Microbial ecology of the oceans. Wiley-Liss, Inc, p 351-386

Strom SL, Brainard MA, Holmes JL, Olson MB (2001) Phytoplankton blooms are
strongly impacted by microzooplankton grazing in coastal North Pacific waters.
Marine Biology 138:355-368

Sverdrup HU (1953) On the conditions for the vernal blooming of phytoplankton. Journal
du Conseil Permanent International pour l'Exploration de la Mer 18:287-295

Tagliabue A, Arrigo KR (2003) Anomalously low zooplankton abundance in the Ross
Sea: An alternative explanation. Limnology and Oceanography 48:686-699

165
Tang KW (2003) Grazing and colony size development in Phaeocystis globosa
(Prymnesiophyceae): the role of a chemical signal. Journal of Plankton Research
25:831-842

Thingstad TF, Skjoldal EF, Bohne RA (1993) Phosphorus Cycling and Algal-Bacterial
Competition in Sandsfjord, Western Norway. Marine Ecology-Progress Series
99:239-259

Utermohl H (1958) Zur Vervollkommung der quantitativen phytoplankton-methodik.
Mitt Int Ver Limnol 9:38

Verity PG (1985) Grazing, respiration, excretion, and growth rates of tintinnids.
Limnology and Oceanography 30:1268-1282

Verity PG, Smetacek V (1996) Organism life cycles, predation, and the structure of
marine pelagic ecosystems. Marine Ecology Progress Series 130:277-293

Verity PG, Stoecker DK, Sieracki ME, Nelson DM (1996) Microzooplankton grazing of
primary production at 140
o
W in the equatorial Pacific. Deep-Sea Research II
43:1227-1255
Walsh JJ, McRoy CP (1986) Ecosystem analysis in the southeastern Bering Sea.
Continental Shelf Research 5:259-288

Weisse T, Karstens N, Meyer VCL, Janke L, Lettner S, Teichgraber K (2001) Niche
separation in common prostome freshwater ciliates: the effect of food and
temperature. Aquatic Microbial Ecology 26:167-179

Weisse T, Kirchoff B (1997) Feeding of the heterotrophic freshwater dinoflagellate
Peridiniopsis berolinense on cryptophytes: analysis by flow cytometry and
electronic particle counting. Aquatic Microbial Ecology 12:153-164

Wilcox RR (2003) Applying contemporary statistical techniques, Vol. Academic Press,
New York

Yentsch CM, Horan PK, Muirhead K, Dortch Q, Haugen EM, Legendre L, Murphy LS,
Phinney D, Pomponi SA, Spinrad RW, Wood AM, Yentsch CS, Zahurenec BJ
(1983) Flow cytometry and sorting: a powerful technique with potential
applications in aquatic sciences. Limnology and Oceanography 28:1275-1280

166
Appendix 1: List of References Comprising Data Set for Chapter 2

Admiraal, W. 1977. Influence of light and temperature on the growth rate of estuarine
benthic diatoms in culture. Mar. Biol. 39: 1-9.
Aelion, C. M., and S. W. Chisholm. 1985. Effect of temperature on growth and ingestion
rates of Favella sp. J. Plankton Res. 7: 821-830.
Baars, J. W. M. 1981. Autecological investigations on marine diatoms. Hydrobiol. Bull. 15:
137-151.
Baars, J. W. M. 1982. Autecological investigation on marine diatoms: 3. Thalassiosira
nordenskioeldii and Chaetoceros diadema. Mar. Biol. 68: 343-350.
Baldock, B. M., and J. Berger. 1984. The effects of low temperatures on the growth of four
fresh-water amoebae (Protozoa: Gymnamoebia). Trans. Am. Microsc. Soc. 103: 233-
239.
Baldock, B. M., J. H. Baker, and M. A. Sleigh. 1980. Laboratory growth rates of six species
of freshwater Gymnamoebia. Oecologia 47: 156-159.
Boenigk, J., and G. Novarino. 2004. Effect of suspended clay on the feeding and growth of
bacterivorous flagellates and ciliates. Aquat. Microb. Ecol. 34: 181-192.
Brand, L., and R. R. L. Guillard. 1981. The effects of continuous light and light intensity on
the reproduction rates of twenty-two species of marine phytoplankton. J. Exp. Mar.
Biol. Ecol. 50: 119-132.
Brand, L. E., L. S. Murphy, R. R. L. Guillard, and H. T. Lee. 1981. Genetic variability and
differentiation in the temperature niche component of the diatom Thalassiosira
pseudonana. Mar. Biol. 62: 103-110.
167

Bravo, I., and D. M. Anderson. 1994. The effects of temperature, growth medium and
darkness on excystment and growth of the toxic dinoflagellate Gymnodinium
catenatum from northwest Spain. J. Plankton Res. 16: 513-525.
Buma, A. G. J., A. A. M. Noordeloos, and J. Larsen. 1993. Strategies and kinetics of
photoacclimation in three Antarctic nanophytoflagellates. J. Phycol. 29: 407-417.
Buskey, E. J., and C. J. Hyatt. 1995. Effects of the Texas (USA) 'brown tide' alga on
planktonic grazers. Mar. Ecol. Prog. Ser. 126: 285-292.
Butler, H., and A. Rogerson. 1996. Growth potential, production efficiency and annual
production of marine benthic naked amoebae (gymnamoebae) inhabiting sediments of
the Clyde Sea area, Scotland. Aquat. Microb. Ecol. 10: 123-129.
Capriulo, G. M., R. A. Schreiner, and B. L. Dexter. 1988. Differential growth of Euplotes
vannus fed fragmented versus unfragmented chains of Skeletonema costatum. Mar.
Ecol. Prog. Ser. 47: 205-209.
Caron, D. A. 1990. Growth of two species of bacterivorous nanoflagellates in batch and
continuous culture, and implications for their planktonic existence. Mar. Microb.
Food Webs 4: 143-159.
Caron, D. A., J. C. Goldman, O. K. Andersen, and M. R. Dennett. 1985. Nutrient cycling in a
microflagellate food chain: II. Population dynamics and carbon cycling. Mar. Ecol.
Prog. Ser. 24: 243-254.
Caron, D. A., J. C. Goldman, and M. R. Dennett. 1986. Effect of temperature on growth,
respiration and nutrient regeneration by an omnivorous microflagellate. Appl.
168
Environ. Microbiol. 52: 1340-1347.
Caron, D. A., E. L. Lim, G. Miceli, J. B. Waterbury, and F. W. Valois. 1991. Grazing and
utilization of chroococcoid cyanobacteria and heterotrophic bacteria by protozoa in
laboratory cultures and a coastal plankton community. Mar. Ecol. Prog. Ser. 76: 205-
217.
Choi, J. W., and F. Peters. 1992. Effects of temperature on two psychrophilic ecotypes of a
heterotrophic nanoflagellate, Paraphysomonas imperforata. Appl. Environ.
Microbiol. 58: 593-599.
Cloern, J. E. 1977. Effects of light intensity and temperature on Cryptomonas ovata
(Cryptophyceae) growth and nutrient uptake rates. J. Phycol. 13: 389-395.
Daugbjerg, N., H. J. Marchant, and H. A. Thomsen. 2000. Life history stages of
Pyramimonas tychotreta (Prasinophyceae, Chlorophyta), a marine flagellate from the
Ross Sea, Antarctica. Phycol. Res. 48: 199-209.
Davis, C. O., P. J. Harrison, and R. C. Dugdale. 1973. Continuous culture of marine diatoms
under silicate limitation. I. Synchronized life cycle of Skeletonema costatum. J.
Phycol. 9: 175-180.
Delaney, M. P. 2003. Effects of temperature and turbulence on the predator-prey interactions
between a heterotrophic flagellate and a marine bacterium. Microb. Ecol. 45: 218-
225.
Dini, F., and D. Nyberg. 1999. Growth rates of marine ciliates on diverse organisms reveal
ecological specializations within morphospecies. Microb. Ecol. 37: 13-22.
Durbin, E. G. 1974. Studies on the autecology of the marine diatom Thalassiosira
169
nordenskioldii Cleve. I. The influence of daylength, light intensity and temperature on
growth. J. Phycol. 10: 220-225.
Eccleston-Parry, J. D., and B. S. Leadbeater. 1994. The effect of long-term low bacterial
density on the growth kinetics of three marine heterotrophic nanoflagellates. J. Exp.
Mar. Biol. Ecol. 177: 219-233.
Eccleston-Parry, J. D. 1995. Regeneration of phosphorus and nitrogen by four species of
heterotrophic nanoflagellates feeding on three nutritional states of a single bacterial
strain. Appl. Environ. Microbiol. 61: 1033-1038.
Fawley, M. W. 1984. Effects of light intensity and temperature interactions on growth
characteristics of Phaeodactylum tricornutum (Bacillariophyceae). J. Phycol. 20: 67-
72.
Fenchel, T. 1968. The ecology of marine microbenthos: III. The reproductive potential of
ciliates. Ophelia 5: 123-136.
Fenchel, T. 1982. Ecology of heterotrophic microflagellates. II. Bioenergetics and growth.
Mar. Ecol. Prog. Ser. 8: 225-231.
Fenchel, T., and P. R. Jonsson. 1988. The functional biology of Strombidium sulcatum, a
marine oligotrich ciliate (Ciliophora, Oligotrichina). Mar. Ecol. Prog. Ser. 48: 1-15.
Fiala, M., and L. Oriol. 1990. Light-temperature interactions on the growth of antarctic
diatoms. Polar Biol. 10: 629-636.
Finlay, B. 1977. The dependence of reproductive rate on cell size and temperature in
freshwater ciliated protozoa. Oecologia 30: 75-81.
170
Fu, Y., C. O'Kelly, M. E. Sieracki, and D. L. Distel. 2003. Protistan grazing analysis by flow
cytometry using prey labeled by in vivo expression of fluorescent proteins. Appl.
Environ. Microbiol. 69: 6848-6855.
Furnas, M. 1978. Influence of temperature and cell size on the division rate and chemical
content of the diatom Chaetoceros curvisetum Cleve. J. Exp. Mar. Biol. Ecol. 34: 97-
109.
Gallagher, J. C. 1982. Physiological variation and electrophoretic banding patterns of
genetically different seasonal populations of Skeletonema costatum
(Bacillariophyceae). J. Phycol. 18: 148-162
Gilstad, M., and E. Sakshaug. 1990. Growth rates of ten diatom species from the Barents Sea
at different irradiances and day lengths. Mar. Ecol. Prog. Ser. 64: 169-173.
Goldman, J. C., and D. A. Caron. 1985. Experimental studies on an omnivorous
microflagellate: implications for grazing and nutrient regeneration in the marine
microbial food chain. Deep-Sea Res. I 32: 899-915.
Goldman, J. C., and E. J. Carpenter. 1974. A kinetic approach to the effect of temperature on
algal growth. Limnol. Oceanogr. 19: 756-766.
Goldman, J. C., and J. J. McCarthy. 1978. Steady state growth and ammonium uptake of a
fast-growing marine diatom. Limnol. Oceanogr. 23: 695-703.
Gransden, S. G., and A. J. Lewitus. 2003. Grazing of two euplotid ciliates on the
heterotrophic dinoflagellates Pfiesteria piscicida and Cryptoperidiniopsis sp. Aquat.
Microb. Ecol. 33: 303-308.
Grant, W. S., and R. A. Horner. 1976. Growth responses to salinity variation in four arctic ice
171
diatoms. J. Phycol. 12: 180-185.
Guillard, R. R. L., and J. H. Ryther. 1962. Studies of marine planktonic diatoms. I. Cyclotella
nana Hustedt and Detonula confervacea Cleve. Can. J. Microbiol. 8: 229-239.
Hobson, L. A. 1974. Effects of interactions of irradiance, daylength and temperature on
division rates of three species of marine unicellular algae. J. Fish. Res. Bd. Can. 31:
391-395.
Hoogenhout, H., and J. Amesz. 1965. Growth rates of photosynthetic microorganisms in
laboratory cultures. Arch. Mikrobiol. 50: 10-25.
Ignatiades, L., and T. J. Smayda. 1970. Autecological studies on the marine diatom
Rhizosolenia fragilissima Bergon . 1. The influence of light, temperature and salinity.
J. Phycol. 6: 332-339.
Jack, J. D., and J. J. Gilbert. 1993. The effect of suspended clay on ciliate population growth
rates. Freshw. Biol. 29: 385-394.
Jacques, G. 1983. Some ecophysiological aspects of the Antarctic phytoplankton. Polar Biol.
2: 27-33.
Jakobsen, H. H., and P. J. Hansen. 1997. Prey size selection, grazing and growth response of
the small heterotrophic dinoflagellate Gymnodinium sp. and the ciliate Balanion
comatum -- a comparative study. Mar. Ecol. Prog. Ser. 158: 75-86.
Jensen, M. O., and O. Moestrup. 1997. Autecology of the toxic dinoflagellate Alexandrum
ostenfeldii: life history and growth at different temperatures and salinities. Eur. J.
Phycol. 32: 9-18.
John, E. H., and K. Davidson. 2001. Prey selectivity and the influence of prey
172
carbon:nitrogen ratio on microflagellate grazing. J. Exp. Mar. Biol. Ecol. 260: 93-
111.
Jonsson, P. R. 1986. Particle size selection, feeding rates and growth dynamics of marine
planktonic oligotrichous ciliates (Ciliophora:Oligotrichina). Mar. Ecol. Prog. Ser. 33:
265-277.
Jorgensen, E. G. 1968. The adaptation of plankton algae II. Aspects of the temperature
adaptation of Skeletonema costatum. Physiol. Plant. 21: 423-427.
Kamiyama, T. 1997. Growth and grazing responses of tintinnid ciliates feeding on the toxic
dinoflagellate Heterocapsa circularisquama. Mar. Biol. 128: 509-515.
Krawiec, R. W. 1982. Autecology and clonal variability of the marine centric diatom
Thalassiosira rotula (Bacillariophyceae) in response to light, temperature and salinity.
Mar. Biol. 69: 79-89.
Langdon, C. 1987. On the causes of interspecific differences in the growth-irradiance
relationship for phytoplankton. Part I. A comparative study of the growth-irradiance
relationship of three marine phytoplankton species: Skeletonema costatum,
Olisthodiscus luteus and Gonyaulax tamarensis. J. Plankton Res. 9: 459-482.
Langdon, C. 1988. On the causes of interspecific differences in the growth-irradiance
relationship for phytoplankton. II. A general review. J. Plankton Res. 10: 1291-1312.
Laybourn, J., and J. M. Stewart. 1974. Effect of food consumption and temperature on
reproduction in the ciliate Colpidium campylum. J. Zool., Lond. 174: 277-283.
Laybourn, J., and L. Whymant. 1980. The effect of diet and temperature on reproductive rate
in Arcella vulgaris Ehrenberg (Sarcodina: Testacida). Oecologia 45: 282-284.
173
Lee, C. C., and T. Fenchel. 1972. Studies on ciliates associated with sea ice from Antarctica.
II. Temperature responses and tolerances in ciliates from Antarctic, Temperate and
Tropical habitats. Archiv fur Protistenk 114: 237-244.
Li, W. K. W., and I. Morris. 1982. Temperature adaptation in Phaeodactylum tricornutum
Bohlin: photosynthetic rate compensation and capacity. J. Exp. Mar. Biol. Ecol. 58:
135-150.
Liu, H., and E. J. Buskey. 2000. The exopolymer secretions (EPS) layer surrounding
Aureoumbra lagunensis cells affects growth, grazing, and behavior of protozoa.
Limnol. Oceanogr. 45: 1187-1191.
Lundholm, N., R. Pocklington, and O. Moestrup. 1997. Studies on the marine planktonic
diatom Pseudo-nitzschia. 2. Autecology of P. pseudodelicatissima based on isolates
from Danish coastal waters. Phycologia 36: 381-388.
Martinez, E. A. 1980. Sensitivity of marine ciliates (Protozoa, Ciliophora) to high thermal
stress. Estuar. Coast. Mar. Sci. 10: 369-381.
Mayes, D. F., A. Rogerson, H. Marchant, and J. Laybourn-Parry. 1997. Growth and
consumption rates of bacterivorous Antarctic naked marine amoebae. Mar. Ecol.
Prog. Ser. 160: 101-108.
Montagnes, D. J. S. 1996. Growth responses of planktonic ciliates in the genera Strobilidium
and Strombidium. Mar. Ecol. Prog. Ser. 130: 241-254.
Morgan, K. C., and J. Kalff. 1979. Effect of light and temperature interactions on growth of
Cryptomonas erosa (Cryptophyceae). J. Phycol. 15: 127-134.
Mortain-Bertrand. 1989. Effects of light fluctuations on the growth and productivity of
174
Antarctic diatoms in cultures. Polar Biol. 9: 245-252.
Morton, S. L., D. R. Norris, and J. W. Bomber. 1992. Effect of temperature, salinity and light
intensity on the growth and seasonality of toxic dinoflagellates associated with
ciguatera. J. Exp. Mar. Biol. Ecol. 157: 79-90.
Muller, H., and W. Geller. 1993. Maximum growth rates of aquatic ciliated protozoa: the
dependence on body size and temperature reconsidered. Arch. Hydrobiol. 126: 315-
327.
Nakamura, Y. 1998. Growth and grazing of a large heterotrophic dinoflagellate, Noctiluca
scintillans, in laboratory cultures. J. Plankton Res. 20: 1711-1720.
Naustvoll, L.-J. 2000. Prey size spectra in naked heterotrophic dinoflagellates. Phycologia
39: 448-455.
Nilawati, J., B. M. Greenberg, and R. E. H. Smith. 1997. Influence of ultraviolet radiation on
growth and photosynthesis of two cold ocean diatoms. J. Phycol. 33: 215-224.
Ohman, M. D., and R. A. Snyder. 1991. Growth kinetics of the omnivorous oligotrich ciliate
Strombidium sp. Limnol. Oceanogr. 36: 922-935.
Paasche E. 1973. Silicon and the ecology of marine plankton diatoms. I. Thalassiosira
psuedonana (Cyclotella nana) grown in a chemostat with silicate as limiting nutrient.
Mar. Biol. 19:117-126.
Parsons, T. R., K. Stephens, and J. D. H. Strickland. 1961. On the chemical composition of
eleven species of marine phytoplankters. J. Fish. Res. Bd. Can. 18: 1001-1016.
Pedersen, M. F., and P. J. Hansen. 2003. Effects of high pH on the growth and survival of six
marine heterotrophic protists. Mar. Ecol. Prog. Ser. 260: 33-41.
175
Pérez-Uz, B. 1995. Growth rate variability in geographically diverse clones of Uronema
(Ciliophora: Scuticociliatida). FEMS Microbiol. Ecol. 16: 193-204.
Perez-Uz, B. 1996. Bacterial preferences and growth kinetic variation in Uronema marinum
and Uronema nigricans (Ciliophora: Scuticociliatida). Microb. Ecol. 31: 189-198.
Reay, D. S., D. B. Nedwell, J. Priddle, and J. C. Ellis-Evans. 1999. Temperature dependence
of inorganic nitrogen uptake: reduced affinity for nitrate at suboptimal temperatures
in both algae and bacteria. Appl. Environ. Microbiol. 65: 2577-2584.
Sakshaug, E., and K. Andresen. 1986. Effect of light regime upon growth rate and chemical
composition of a clone of Skeletonema costatum from the Trondheimsfjord, Norway.
J. Plankton Res. 8: 619-637.
Sanders, R. W., D. A. Caron, J. M. Davidson, M. R. Dennett, and D. M. Moran. 2001.
Nutrient acquisition and population growth of a mixotrophic alga in axenic and
bacterized cultures. Microb. Ecol. 442: 513-523.
Schlesinger, D. A., L. A. Molot, and B. J. Shuter. 1981. Specific growth rates of freshwater
algae in relation to cell size and light intensity. Can. J. Fish. Aquat. Sci. 38: 1052-
1058.
Seaburg, K. G., B. C. Parker, R. A. W. Jr., and G. M. S. Jr. 1981. Temperature-growth
responses of algal isolates from Antarctic oases. J. Phycol. 17: 353-360.
Selph, K. E., M. R. Landry, and E. A. Laws. 2003. Heterotrophic nanoflagellate enhancement
of bacterial growth through nutrient remineralization in chemostat culture. Aquat.
Microb. Ecol. 32: 23-37.
176
Sherr, B. F., E. B. Sherr, and T. Berman. 1983. Grazing, growth and ammonium excretion
rates of a heterotrophic microflagellate fed with four species of bacteria. Appl.
Environ. Microbiol. 45: 1196-1201.
Smayda, T. J. 1969. Experimental observations on the influence of temperature, light and
salinity on cell division of the marine diatom, Detonula confervacea (Cleve) Gran. J.
Phycol. 5: 150-157.
Smith, R. E. H., L. C. Stapleford, and R. S. Ridings. 1994. The acclimated response of
growth, photosynthesis, composition and carbon balance to temperature in the
psychrophilic ice diatom Nitzschia seriata. J. Phycol. 30: 8-16.
Sommer, U. 1989. Maximal growth rates of antarctic phytoplankton: only weak dependence
on cell size. Limnol. Oceanogr. 34: 1109-1112.
Stapleford, L. S., and R. E. H. Smith. 1996. The interactive effects of temperature and silicon
limitation on the psychrophilic ice diatom Pseudonitszchia seriata. Polar Biol. 16:
589-594.
Strom, S. L., and T. A. Morello. 1998. Comparative growth rates and yields of ciliates and
heterotrophic dinoflagellates. J. Plankton Res. 20: 571-584.
Strzepek, R. F., and N. M. Price. 2000. Influence of irradiance and temperature on the iron
content of the marine diatom Thalassiosira weissflogii (Bacillariophyceae). Mar.
Ecol. Prog. Ser. 206: 107-117.
Suzuki, Y., and M. Takahashi. 1995. Growth responses of several diatom species isolated
from various environments to temperature. J. Phycol. 31: 880-888.
Thomas, W. H. 1966. Effects of temperature and illuminance on cell division rates of three
177
species of tropical oceanic phytoplankton. J. Phycol. 2: 17-22.
Thompson, P. A., M.-x. Guo, and P. J. Harrison. 1992. Effects of variation in temperature. I.
On the biochemical composition of eight species of marine phytoplankton. J. Phycol.
28: 481-488.
Thompson, P. A., M.-x. Guo, P. J. Harrison, and J. N. C. Whyte. 1992. Effects of variation in
temperature. II. On the fatty acid composition of eight species of marine
phytoplankton. J. Phycol. 28: 488-497.
Timmermans, K. R. and others 2001. Growth rates of large and small Southern Ocean
diatoms in relation to availability of iron in natural seawater. Limnol. Oceanogr. 46:
260-266.
Valbonesi, A., and P. Luporini. 1993. Biology of Euplotes focardii, an Antarctic ciliate. Polar
Biol. 13: 489-493.
Verity, P. G. 1982. Effects of temperature, irradiance, and daylength on the marine diatom
Leptocylindrus danicus Cleve. IV. Growth. J. Exp. Mar. Biol. Ecol. 60: 209-222.
Walton, B. M., M. A. Gates, A. Kloos, and J. Fisher. 1995. Intraspecific variability in the
thermal dependence of locomotion, population growth, and mating in the ciliated
protist Euplotes vannus. Physiol. Zool. 68: 98-113.
Watras, C. J., S. W. Chisholm, and D. M. Anderson. 1982. Regulation of growth in an
estuarine clone of Gonyaulax tamarensis Lebour: salinity-dependent temperature
responses. J. Exp. Mar. Biol. Ecol. 62: 25-37.
Weisse, T., and A. Frahm. 2002. Direct and indirect impact of two common rotifer species
(Keratella spp.) on two abundant ciliate species (Urotricha furcata, Balanion
178
planctonicum). Freshw. Biol. 47: 53-64.
Weisse, T., and D. J. S. Montagnes. 1998. Effect of temperature on inter- and intraspecific
isolates of Urotricha (Prostomatida, Ciliophora). Aquat. Microb. Ecol. 15: 285-291.
Weisse, T., N. Karstens, V. C. L. Meyer, L. Janke, S. Lettner, and K. Teichgraber. 2001.
Niche separation in common prostome freshwater ciliates: the effect of food and
temperature. Aquat. Microb. Ecol. 26: 167-179.
Weisse, T., P. Stadler, E. S. Lindstrom, S. A. Kimmance, and D. J. S. Montagnes. 2002.
Interactive effect of temperature and food concentration on growth rate: a test case
using the small freshwater ciliate Urotricha farcta. Limnol. Oceanogr. 47: 1447-
1455.
Yoder, J. A. 1979. Effect of temperature on light-limited growth and chemical composition
of Skeletonema costatum (Bacillariophyceae). J. Phycol. 15: 362-370.
Zubkov, M. V., and M. A. Sleigh. 2000. Comparison of growth efficiencies of protozoa
growing on bacteria deposited on surfaces and in suspension. J. Eukaryot. Microbiol.
47: 62-69.
179
Appendix 2: Growth Rates of Phototrophic Protists Within Chapter 2
Data Set

Organism Temperature (
o
C) Growth rate (d
-1
) Study
Amphiprora sp. 4 0.30 Admiraal (1977)
Amphiprora sp. 8 0.47 Admiraal (1977)
Amphiprora sp. 12 0.63 Admiraal (1977)
Amphiprora sp. 16 1.01 Admiraal (1977)
Amphiprora sp. 20 1.24 Admiraal (1977)
Amphiprora sp. 25 1.57 Admiraal (1977)
Nitzschia dissipata 4 0.21 Admiraal (1977)
Nitzschia dissipata 8 0.49 Admiraal (1977)
Nitzschia dissipata 12 0.76 Admiraal (1977)
Nitzschia dissipata 16 1.00 Admiraal (1977)
Nitzschia dissipata 20 1.55 Admiraal (1977)
Nitzschia dissipata 25 1.75 Admiraal (1977)
Navicula arenaria 4 0.15 Admiraal (1977)
Navicula arenaria 8 0.52 Admiraal (1977)
Navicula arenaria 12 0.73 Admiraal (1977)
Navicula arenaria 16 1.00 Admiraal (1977)
Navicula arenaria 20 0.91 Admiraal (1977)
Navicula arenaria 25 0.40 Admiraal (1977)
Nitzschia sigma 4 0.21 Admiraal (1977)
Nitzschia sigma 8 0.16 Admiraal (1977)
Nitzschia sigma 12 0.25 Admiraal (1977)
Nitzschia sigma 16 0.29 Admiraal (1977)
Nitzschia sigma 20 0.33 Admiraal (1977)
Nitzschia sigma 25 0.43 Admiraal (1977)
Heterocapsa triquetra 8 0.35 Aelion and Chisholm (1985)
Heterocapsa triquetra 8 0.46 Aelion and Chisholm (1985)
Heterocapsa triquetra 11.4 0.52 Aelion and Chisholm (1985)
Heterocapsa triquetra 11.4 0.56 Aelion and Chisholm (1985)
Heterocapsa triquetra 16.4 0.59 Aelion and Chisholm (1985)
Heterocapsa triquetra 16.4 0.61 Aelion and Chisholm (1985)
Heterocapsa triquetra 16.4 0.64 Aelion and Chisholm (1985)
Heterocapsa triquetra 16.4 0.79 Aelion and Chisholm (1985)
Heterocapsa triquetra 16.4 0.85 Aelion and Chisholm (1985)
Heterocapsa triquetra 18.8 0.68 Aelion and Chisholm (1985)
Heterocapsa triquetra 18.8 0.71 Aelion and Chisholm (1985)
Heterocapsa triquetra 18.8 0.80 Aelion and Chisholm (1985)
Heterocapsa triquetra 18.8 0.85 Aelion and Chisholm (1985)
Heterocapsa triquetra 21.1 0.76 Aelion and Chisholm (1985)
Heterocapsa triquetra 21.1 0.82 Aelion and Chisholm (1985)
Chaetoceros diadema -1.5 0.27 Baars (1981)
Coscinodiscus concinnus -1.5 0.10 Baars (1981)
Thalassiosira nordenskioeldii -1.5 0.25 Baars (1981)
Thalassiosira nordenskioeldii -1.5 0.22 Baars (1981)
Thalassiosira nordenskioeldii -1.5 0.12 Baars (1981)
Biddulphia aurita -1.5 0.21 Baars (1981)
Chaetoceros diadema -1.5 0.30 Baars (1981)
180
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Coscinodiscus concinnus -1.5 0.10 Baars (1981)
Rhizosolenia setigera -1.5 0.14 Baars (1981)
Thalassiosira nordenskioeldii -1.5 0.17 Baars (1981)
Coscinodiscus concinnus -1 0.12 Baars (1981)
Thalassiosira nordenskioeldii -1 0.35 Baars (1981)
Thalassiosira rotula -1 0.05 Baars (1981)
Thalassiosira nordenskioeldii 0 0.20 Baars (1981)
Thalassiosira nordenskioeldii 0 0.17 Baars (1981)
Thalassiosira nordenskioeldii 0 0.47 Baars (1981)
Thalassiosira nordenskioeldii 0 0.30 Baars (1981)
Thalassiosira nordenskioeldii 0 0.44 Baars (1981)
Thalassiosira nordenskioeldii 0 0.30 Baars (1981)
Biddulphia aurita 0 0.36 Baars (1981)
Chaetoceros diadema 0 0.43 Baars (1981)
Chaetoceros teres 0 0.33 Baars (1981)
Coscinodiscus concinnus 0 0.17 Baars (1981)
Ditylum brightwellii 0 0.15 Baars (1981)
Melosira nummuloides 0 0.12 Baars (1981)
Rhizosolenia setigera 0 0.28 Baars (1981)
Skeletonema costatum 0 0.35 Baars (1981)
Thalassiosira decipiens 0 0.11 Baars (1981)
Thalassiosira levanderi 0 0.35 Baars (1981)
Thalassiosira nordenskioeldii 0 0.74 Baars (1981)
Thalassiosira nordenskioeldii 0 0.36 Baars (1981)
Thalassiosira nordenskioeldii 0 0.20 Baars (1981)
Thalassiosira rotula 0 0.17 Baars (1981)
Thalassiosira rotula 0 0.12 Baars (1981)
Thalassiosira rotula 0 0.09 Baars (1981)
Biddulphia aurita 0 0.31 Baars (1981)
Biddulphia aurita 0 0.26 Baars (1981)
Chaetoceros diadema 0 0.45 Baars (1981)
Coscinodiscus concinnus 0 0.15 Baars (1981)
Rhizosolenia setigera 0 0.27 Baars (1981)
Thalassiosira nordenskioeldii 0 0.19 Baars (1981)
Biddulphia aurita 3 0.57 Baars (1981)
Biddulphia aurita 3 0.39 Baars (1981)
Coscinodiscus concinnus 3 0.32 Baars (1981)
Thalassiosira nordenskioeldii 3 0.46 Baars (1981)
Thalassiosira rotula 3 0.51 Baars (1981)
Thalassiosira decipiens 6 0.27 Baars (1981)
Thalassiosira decipiens 6 0.27 Baars (1981)
Thalassiosira levanderi 6 0.44 Baars (1981)
Thalassiosira nordenskioeldii 6 0.37 Baars (1981)
Thalassiosira rotula 6 0.27 Baars (1981)
Thalassiosira rotula 6 0.27 Baars (1981)
Actinoptychus senarius 6 0.13 Baars (1981)
Biddulphia aurita 6 0.44 Baars (1981)
Coscinodiscus concinnus 6 0.35 Baars (1981)
181
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira eccentrica 6 0.11 Baars (1981)
Thalassiosira nordenskioeldii 6 0.65 Baars (1981)
Thalassiosira nordenskioeldii 6 0.43 Baars (1981)
Thalassiosira rotula 6 0.54 Baars (1981)
Actinoptychus senarius 6 0.09 Baars (1981)
Biddulphia aurita 6 0.44 Baars (1981)
Coscinodiscus concinnus 6 0.34 Baars (1981)
Thalassiosira nordenskioeldii 6 0.34 Baars (1981)
Thalassiosira rotula 6 0.60 Baars (1981)
Thalassiosira decipiens 6 0.41 Baars (1981)
Thalassiosira decipiens 6 0.37 Baars (1981)
Thalassiosira levanderi 6 0.54 Baars (1981)
Thalassiosira nordenskioeldii 6 0.64 Baars (1981)
Thalassiosira rotula 6 0.40 Baars (1981)
Thalassiosira rotula 6 0.40 Baars (1981)
Biddulphia aurita 6 0.48 Baars (1981)
Biddulphia regia 6 0.33 Baars (1981)
Coscinodiscus concinnus 6 0.41 Baars (1981)
Ditylum brightwellii 6 0.56 Baars (1981)
Guinarida flaccida 6 0.27 Baars (1981)
Lauderia annulata 6 0.55 Baars (1981)
Melosira moniliformis 6 0.48 Baars (1981)
Rhizosolenia setigera 6 0.64 Baars (1981)
Thalassiosira nordenskioeldii 6 0.50 Baars (1981)
Thalassiosira rotula 6 0.74 Baars (1981)
Biddulphia aurita 6 0.61 Baars (1981)
Biddulphia regia 6 0.35 Baars (1981)
Chaetoceros debilis 6 0.50 Baars (1981)
Coscinodiscus concinnus 6 0.46 Baars (1981)
Coscinodiscus granii 6 0.14 Baars (1981)
Ditylum brightwellii 6 0.44 Baars (1981)
Guinarida flaccida 6 0.24 Baars (1981)
Melosira nummuloides 6 0.43 Baars (1981)
Thalassiosira decipiens 6 0.44 Baars (1981)
Thalassiosira eccentrica 6 0.13 Baars (1981)
Thalassiosira levanderi 6 0.77 Baars (1981)
Thalassiosira nordenskioeldii 6 0.75 Baars (1981)
Thalassiosira nordenskioeldii 6 0.69 Baars (1981)
Thalassiosira nordenskioeldii 6 0.62 Baars (1981)
Thalassiosira nordenskioeldii 6 0.53 Baars (1981)
Thalassiosira rotula 6 0.58 Baars (1981)
Thalassiosira rotula 6 0.58 Baars (1981)
Biddulphia aurita 6 0.71 Baars (1981)
Chaetoceros didymus 6 0.31 Baars (1981)
Coscinodiscus concinnus 6 0.39 Baars (1981)
Thalassiosira rotula 6 0.74 Baars (1981)
Biddulphia aurita 6 0.40 Baars (1981)
Chaetoceros decipiens 6 0.56 Baars (1981)
182
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Chaetoceros densus 6 0.31 Baars (1981)
Chaetoceros diadema 6 0.74 Baars (1981)
Chaetoceros diadema 6 0.68 Baars (1981)
Chaetoceros teres 6 0.67 Baars (1981)
Coscinodiscus concinnus 6 0.40 Baars (1981)
Eucampia zoodiacus 6 0.35 Baars (1981)
Rhizosolenia setigera 6 0.49 Baars (1981)
Thalassiosira decipiens 6 0.49 Baars (1981)
Thalassiosira levanderi 6 0.70 Baars (1981)
Thalassiosira nordenskioeldii 6 0.73 Baars (1981)
Thalassiosira rotula 6 0.60 Baars (1981)
Thalassiosira rotula 6 0.59 Baars (1981)
Thalassiosira nordenskioeldii 6 1.12 Baars (1981)
Thalassiosira rotula 6 0.71 Baars (1981)
Thalassiosira rotula 6 0.70 Baars (1981)
Biddulphia aurita 6 0.62 Baars (1981)
Lauderia annulata 6 0.46 Baars (1981)
Thalassiosira eccentrica 6 0.20 Baars (1981)
Thalassiosira rotula 6 0.84 Baars (1981)
Chaetoceros decipiens 6 0.74 Baars (1981)
Chaetoceros diadema 6 0.95 Baars (1981)
Thalassiosira nordenskioeldii 6 0.83 Baars (1981)
Thalassiosira rotula 6 0.78 Baars (1981)
Thalassiosira rotula 6 0.74 Baars (1981)
Chaetoceros decipiens 6 0.51 Baars (1981)
Chaetoceros diadema 6 0.87 Baars (1981)
Coscinodiscus concinnus 6 0.38 Baars (1981)
Thalassiosira decipiens 6 0.30 Baars (1981)
Thalassiosira nordenskioeldii 6 0.84 Baars (1981)
Thalassiosira rotula 6 0.54 Baars (1981)
Thalassiosira rotula 6 0.33 Baars (1981)
Thalassiosira rotula 8 0.71 Baars (1981)
Thalassiosira rotula 8 0.83 Baars (1981)
Coscinodiscus granii 10 0.31 Baars (1981)
Thalassiosira rotula 10 0.96 Baars (1981)
Thalassiosira rotula 10 0.43 Baars (1981)
Coscinodiscus concinnus 12 0.44 Baars (1981)
Thalassiosira eccentrica 12 0.39 Baars (1981)
Thalassiosira hendeyi 12 0.28 Baars (1981)
Coscinodiscus concinnus 12 0.59 Baars (1981)
Biddulphia aurita 12 0.14 Baars (1981)
Chaetoceros teres 12 0.17 Baars (1981)
Coscinodiscus concinnus 12 0.09 Baars (1981)
Thalassiosira polychorda 12 0.11 Baars (1981)
Chaetoceros teres 12 0.32 Baars (1981)
Coscinodiscus centralis 12 0.16 Baars (1981)
Coscinodiscus concinnus 12 0.14 Baars (1981)
Thalassiosira eccentrica 12 0.24 Baars (1981)
183
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Biddulphia aurita 12 0.56 Baars (1981)
Coscinodiscus concinnus 12 0.25 Baars (1981)
Chaetoceros debilis 12 0.80 Baars (1981)
Chaetoceros teres 12 0.53 Baars (1981)
Coscinodiscus centralis 12 0.41 Baars (1981)
Coscinodiscus concinnus 12 0.30 Baars (1981)
Thalassiosira eccentrica 12 0.51 Baars (1981)
Thalassiosira polychorda 12 0.44 Baars (1981)
Biddulphia aurita 12 0.67 Baars (1981)
Coscinodiscus centralis 12 0.53 Baars (1981)
Coscinodiscus concinnus 12 0.48 Baars (1981)
Thalassiosira eccentrica 12 0.52 Baars (1981)
Thalassiosira polychorda 12 0.68 Baars (1981)
Actinocyclus octonarius 12 0.26 Baars (1981)
Actinoptychus senarius 12 0.35 Baars (1981)
Asterionella glacialis 12 1.15 Baars (1981)
Asterionella kariana 12 0.21 Baars (1981)
Bacillaria paxillifer 12 0.56 Baars (1981)
Bellerochea malleus 12 0.26 Baars (1981)
Biddulphia aurita 12 0.78 Baars (1981)
Biddulphia aurita 12 0.70 Baars (1981)
Biddulphia regia 12 0.68 Baars (1981)
Biddulphia sinensis 12 0.42 Baars (1981)
Chaetoceros debilis 12 1.14 Baars (1981)
Chaetoceros didymus 12 1.07 Baars (1981)
Chaetoceros tortissimus 12 0.98 Baars (1981)
Coscinodiscus centralis 12 0.61 Baars (1981)
Coscinodiscus concinnus 12 0.56 Baars (1981)
Coscinodiscus granii 12 0.45 Baars (1981)
Coscinodiscus jonesianus var. commutata 12 0.23 Baars (1981)
Coscinodiscus pavillardii 12 0.53 Baars (1981)
Ditylum brightwellii 12 1.12 Baars (1981)
Ditylum brightwellii 12 0.94 Baars (1981)
Eucampia zoodiacus 12 0.95 Baars (1981)
Guinardia flaccida 12 0.57 Baars (1981)
Lauderia annulata 12 1.13 Baars (1981)
Leptocylindrus danicus 12 0.91 Baars (1981)
Lithodesmium undulatum 12 0.69 Baars (1981)
Melosira monoliformis 12 0.82 Baars (1981)
Navicula comoides 12 0.46 Baars (1981)
Navicula sp. 12 1.13 Baars (1981)
Rhizosolenia robusta 12 0.31 Baars (1981)
Rhizosolenia setigera 12 0.83 Baars (1981)
Rhizosolenia stolterfothii 12 0.97 Baars (1981)
Sceletonema costatum 12 1.20 Baars (1981)
Stephanopyxis palmeriana 12 0.54 Baars (1981)
Synedra tabulata 12 0.37 Baars (1981)
Thalassiosira decipiens 12 0.89 Baars (1981)
184
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira eccentrica 12 0.54 Baars (1981)
Thalassiosira hendeyi 12 0.38 Baars (1981)
Thalassiosira levanderi 12 1.13 Baars (1981)
Thalassiosira nordenskioeldii 12 0.98 Baars (1981)
Thalassiosira nordenskioeldii 12 0.91 Baars (1981)
Thalassiosira nordenskioeldii 12 0.75 Baars (1981)
Thalassiosira rotula 12 0.81 Baars (1981)
Thalassiosira rotula 12 0.83 Baars (1981)
Triceratium alternans 12 0.74 Baars (1981)
Biddulphia aurita 12 0.73 Baars (1981)
Chaetoceros teres 12 0.98 Baars (1981)
Thalassiosira eccentrica 12 0.56 Baars (1981)
Thalassiosira polychorda 12 0.79 Baars (1981)
Chaetoceros teres 12 1.19 Baars (1981)
Thalassiosira eccentrica 12 0.54 Baars (1981)
Biddulphia aurita 12 0.89 Baars (1981)
Coscinodiscus centralis 12 0.74 Baars (1981)
Coscinodiscus concinnus 12 0.50 Baars (1981)
Thalassiosira polychorda 12 1.09 Baars (1981)
Chaetoceros teres 12 1.05 Baars (1981)
Thalassiosira eccentrica 12 0.56 Baars (1981)
Biddulphia aurita 12 0.93 Baars (1981)
Coscinodiscus concinnus 12 0.51 Baars (1981)
Thalassiosira polychorda 12 1.15 Baars (1981)
Chaetoceros teres 12 1.21 Baars (1981)
Coscinodiscus concinnus 12 0.50 Baars (1981)
Thalassiosira eccentrica 12 0.55 Baars (1981)
Thalassiosira hendeyi 12 0.28 Baars (1981)
Coscinodiscus concinnus 12 0.52 Baars (1981)
Thalassiosira polychorda 12 1.16 Baars (1981)
Biddulphia aurita 12 0.96 Baars (1981)
Chaetoceros teres 12 1.17 Baars (1981)
Thalassiosira eccentrica 12 0.55 Baars (1981)
Chaetoceros teres 12 1.08 Baars (1981)
Coscinodiscus centralis 12 0.74 Baars (1981)
Thalassiosira eccentrica 12 0.56 Baars (1981)
Coscinodiscus concinnus 16 0.25 Baars (1981)
Thalassiosira eccentrica 16 0.27 Baars (1981)
Thalassiosira hendeyi 16 0.26 Baars (1981)
Biddulphia aurita 16 0.50 Baars (1981)
Chaetoceros didymus 16 0.82 Baars (1981)
Coscinodiscus concinnus 16 0.48 Baars (1981)
Lauderia annulata 16 0.90 Baars (1981)
Thalassiosira eccentrica 16 0.71 Baars (1981)
Thalassiosira eccentrica 16 0.68 Baars (1981)
Thalassiosira hendeyi 16 0.52 Baars (1981)
Thalassiosira nordenskioeldii 16 0.49 Baars (1981)
Thalassiosira rotula 16 1.02 Baars (1981)
185
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Actinoptychus senarius 16 0.38 Baars (1981)
Bacillaria paxillifer 16 0.55 Baars (1981)
Biddulphia aurita 16 0.49 Baars (1981)
Coscinodiscus radiatus 16 0.21 Baars (1981)
Rhizosolenia alata f. indica 16 0.31 Baars (1981)
Rhizosolenia robusta 16 0.40 Baars (1981)
Rhizosolenia stolterfothii 16 0.58 Baars (1981)
Stephanopyxis palmeriana 16 0.42 Baars (1981)
Actinoptychus senarius 16 0.32 Baars (1981)
Bacillaria paxillifer 16 0.41 Baars (1981)
Biddulphia aurita 16 0.52 Baars (1981)
Coscinodiscus radiatus 16 0.26 Baars (1981)
Rhizosolenia robusta 16 0.57 Baars (1981)
Rhizosolenia stolterfothii 16 0.50 Baars (1981)
Stephanopyxis palmeriana 16 0.38 Baars (1981)
Biddulphia aurita 16 0.58 Baars (1981)
Coscinodiscus concinnus 16 0.43 Baars (1981)
Lauderia annulata 16 1.71 Baars (1981)
Thalassiosira eccentrica 16 1.00 Baars (1981)
Thalassiosira hendeyi 16 0.52 Baars (1981)
Thalassiosira rotula 16 2.13 Baars (1981)
Cerataulina bergonii 18 0.71 Baars (1981)
Thalassiosira decipiens 18 1.13 Baars (1981)
Thalassiosira decipiens 18 1.03 Baars (1981)
Thalassiosira nordenskioeldii 18 0.70 Baars (1981)
Thalassiosira rotula 18 1.31 Baars (1981)
Thalassiosira rotula 18 1.41 Baars (1981)
Actinocyclus octonarius 20 0.14 Baars (1981)
Asterionella kariana 20 0.29 Baars (1981)
Biddulphia aurita 20 0.10 Baars (1981)
Biddulphia regia 20 0.13 Baars (1981)
Biddulphia sinensis 20 0.27 Baars (1981)
Chaetoceros debilis 20 0.38 Baars (1981)
Coscinodiscus granii 20 0.16 Baars (1981)
Coscinodiscus jonesianus var. commutata 20 0.14 Baars (1981)
Coscinodiscus radiatus 20 0.12 Baars (1981)
Ditylum brightwellii 20 0.33 Baars (1981)
Guinardia flaccida 20 0.24 Baars (1981)
Lauderia annulata 20 0.28 Baars (1981)
Rhizosolenia alata f. indica 20 0.25 Baars (1981)
Rhizosolenia imbricata 20 0.24 Baars (1981)
Rhizosolenia robusta 20 0.22 Baars (1981)
Rhizosolenia setigera 20 0.22 Baars (1981)
Rhizosolenia stolterfothii 20 0.24 Baars (1981)
Sceletonema costatum 20 0.51 Baars (1981)
Stephanopyxis palmeriana 20 0.21 Baars (1981)
Thalassiosira eccentrica 20 0.26 Baars (1981)
Thalassiosira rotula 20 0.24 Baars (1981)
186
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Actinocyclus octonarius 20 0.54 Baars (1981)
Actinoptychus senarius 20 0.57 Baars (1981)
Asterionella kariana 20 0.36 Baars (1981)
Biddulphia regia 20 0.66 Baars (1981)
Biddulphia sinensis 20 0.62 Baars (1981)
Chaetoceros debilis 20 0.98 Baars (1981)
Coscinodiscus granii 20 0.60 Baars (1981)
Coscinodiscus pavillardii 20 0.68 Baars (1981)
Coscinodiscus radiatus 20 0.61 Baars (1981)
Ditylum brightwellii 20 1.05 Baars (1981)
Guinardia flaccida 20 0.66 Baars (1981)
Lauderia annulata 20 1.00 Baars (1981)
Navicula comoides 20 0.52 Baars (1981)
Navicula sp. 20 1.03 Baars (1981)
Rhizosolenia alata f. indica 20 0.70 Baars (1981)
Rhizosolenia imbricata 20 0.68 Baars (1981)
Rhizosolenia robusta 20 0.64 Baars (1981)
Rhizosolenia setigera 20 0.63 Baars (1981)
Rhizosolenia stolterfothii 20 0.71 Baars (1981)
Stephanopyxis palmeriana 20 0.79 Baars (1981)
Synedra tabulata 20 0.41 Baars (1981)
Thalassiosira decipiens 20 0.97 Baars (1981)
Thalassiosira eccentrica 20 0.89 Baars (1981)
Thalassiosira hendeyi 20 0.71 Baars (1981)
Thalassiosira rotula 20 0.99 Baars (1981)
Actinocyclus octonarius 26 0.40 Baars (1981)
Biddulphia regia 26 0.24 Baars (1981)
Biddulphia sinensis 26 0.48 Baars (1981)
Coscinodiscus granii 26 0.33 Baars (1981)
Coscinodiscus jonesianus var. commutata 26 0.34 Baars (1981)
Coscinodiscus radiatus 26 0.28 Baars (1981)
Ditylum brightwellii 26 0.64 Baars (1981)
Ditylum brightwellii 26 0.62 Baars (1981)
Lauderia annulata 26 0.52 Baars (1981)
Rhizosolenia alata f. indica 26 0.63 Baars (1981)
Rhizosolenia robusta 26 0.37 Baars (1981)
Stephanopyxis palmeriana 26 0.43 Baars (1981)
Thalassiosira eccentrica 26 0.48 Baars (1981)
Thalassiosira rotula 26 0.59 Baars (1981)
Actinocyclus octonarius 30 0.79 Baars (1981)
Actinoptychus senarius 30 0.73 Baars (1981)
Biddulphia sinensis 30 0.99 Baars (1981)
Coscinodiscus granii 30 0.70 Baars (1981)
Coscinodiscus jonesianus var. commutata 30 0.74 Baars (1981)
Coscinodiscus pavillardii 30 0.88 Baars (1981)
Coscinodiscus radiatus 30 0.83 Baars (1981)
Ditylum brightwellii 30 1.08 Baars (1981)
Rhizosolenia imbricata 30 1.08 Baars (1981)
187
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira eccentrica 30 0.81 Baars (1981)
Thalassiosira nordenskioeldii -1.5 0.25 Baars (1982)
Thalassiosira nordenskioeldii -1.5 0.22 Baars (1982)
Thalassiosira nordenskioeldii -1.5 0.12 Baars (1982)
Thalassiosira nordenskioeldii -1.5 0.17 Baars (1982)
Thalassiosira nordenskioeldii -1 0.35 Baars (1982)
Thalassiosira nordenskioeldii 0 0.24 Baars (1982)
Thalassiosira nordenskioeldii 0 0.17 Baars (1982)
Thalassiosira nordenskioeldii 0 0.47 Baars (1982)
Thalassiosira nordenskioeldii 0 0.30 Baars (1982)
Thalassiosira nordenskioeldii 0 0.44 Baars (1982)
Thalassiosira nordenskioeldii 0 0.30 Baars (1982)
Thalassiosira nordenskioeldii 0 0.51 Baars (1982)
Thalassiosira nordenskioeldii 0 0.36 Baars (1982)
Thalassiosira nordenskioeldii 0 0.20 Baars (1982)
Thalassiosira nordenskioeldii 0 0.19 Baars (1982)
Thalassiosira nordenskioeldii 3 0.46 Baars (1982)
Thalassiosira nordenskioeldii 6 0.37 Baars (1982)
Thalassiosira nordenskioeldii 6 0.65 Baars (1982)
Thalassiosira nordenskioeldii 6 0.43 Baars (1982)
Thalassiosira nordenskioeldii 6 0.34 Baars (1982)
Thalassiosira nordenskioeldii 6 0.64 Baars (1982)
Thalassiosira nordenskioeldii 6 0.50 Baars (1982)
Thalassiosira nordenskioeldii 6 0.75 Baars (1982)
Thalassiosira nordenskioeldii 6 0.69 Baars (1982)
Thalassiosira nordenskioeldii 6 0.62 Baars (1982)
Thalassiosira nordenskioeldii 6 0.53 Baars (1982)
Thalassiosira nordenskioeldii 6 0.73 Baars (1982)
Thalassiosira nordenskioeldii 6 1.12 Baars (1982)
Thalassiosira nordenskioeldii 6 0.84 Baars (1982)
Thalassiosira nordenskioeldii 12 0.98 Baars (1982)
Thalassiosira nordenskioeldii 12 0.91 Baars (1982)
Thalassiosira nordenskioeldii 12 0.75 Baars (1982)
Thalassiosira nordenskioeldii 16 0.49 Baars (1982)
Thalassiosira nordenskioeldii 18 0.70 Baars (1982)
Chaetoceros diadema -1.5 0.27 Baars (1982)
Chaetoceros diadema -1.5 0.30 Baars (1982)
Chaetoceros diadema 0 0.43 Baars (1982)
Chaetoceros diadema 0 0.45 Baars (1982)
Chaetoceros diadema 6 0.74 Baars (1982)
Chaetoceros diadema 6 0.68 Baars (1982)
Chaetoceros diadema 6 0.95 Baars (1982)
Chaetoceros diadema 6 0.87 Baars (1982)
Asterionella glacialis 24 1.39 Brand and Guillard (1981)
Asterionella glacialis 24 0.83 Brand and Guillard (1981)
Asterionella glacialis 24 1.66 Brand and Guillard (1981)
Asterionella glacialis 24 2.01 Brand and Guillard (1981)
Asterionella glacialis 24 3.26 Brand and Guillard (1981)
188
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Asterionella glacialis 24 2.56 Brand and Guillard (1981)
Asterionella glacialis 24 3.60 Brand and Guillard (1981)
Asterionella glacialis 24 2.84 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 0.40 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 0.62 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 1.25 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 1.04 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 2.08 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 2.50 Brand and Guillard (1981)
Bacteriastrum delicatulum 24 0.83 Brand and Guillard (1981)
Biddulphia sp. 24 0.69 Brand and Guillard (1981)
Biddulphia sp. 24 1.18 Brand and Guillard (1981)
Biddulphia sp. 24 1.73 Brand and Guillard (1981)
Biddulphia sp. 24 1.73 Brand and Guillard (1981)
Corethron criophilum 24 0.76 Brand and Guillard (1981)
Corethron criophilum 24 1.25 Brand and Guillard (1981)
Corethron criophilum 24 1.32 Brand and Guillard (1981)
Corethron criophilum 24 2.22 Brand and Guillard (1981)
Corethron criophilum 24 2.08 Brand and Guillard (1981)
Corethron criophilum 24 1.94 Brand and Guillard (1981)
Corethron criophilum 24 1.66 Brand and Guillard (1981)
Coscinodiscus sp. 24 0.55 Brand and Guillard (1981)
Coscinodiscus sp. 24 0.83 Brand and Guillard (1981)
Coscinodiscus sp. 24 0.97 Brand and Guillard (1981)
Coscinodiscus sp. 24 1.11 Brand and Guillard (1981)
Coscinodiscus sp. 24 0.97 Brand and Guillard (1981)
Ditylum brightwellii 24 0.55 Brand and Guillard (1981)
Ditylum brightwellii 24 0.28 Brand and Guillard (1981)
Ditylum brightwellii 24 0.97 Brand and Guillard (1981)
Ditylum brightwellii 24 0.76 Brand and Guillard (1981)
Ditylum brightwellii 24 1.32 Brand and Guillard (1981)
Ditylum brightwellii 24 0.83 Brand and Guillard (1981)
Ditylum brightwellii 24 1.11 Brand and Guillard (1981)
Hemiaulus hauckii 24 0.26 Brand and Guillard (1981)
Hemiaulus hauckii 24 0.67 Brand and Guillard (1981)
Hemiaulus hauckii 24 1.39 Brand and Guillard (1981)
Hemiaulus hauckii 24 0.76 Brand and Guillard (1981)
Hemiaulus hauckii 24 2.70 Brand and Guillard (1981)
Hemiaulus hauckii 24 0.33 Brand and Guillard (1981)
Hemiaulus hauckii 24 2.50 Brand and Guillard (1981)
Streptotheca tamesis 24 0.55 Brand and Guillard (1981)
Streptotheca tamesis 24 0.97 Brand and Guillard (1981)
Streptotheca tamesis 24 1.52 Brand and Guillard (1981)
Streptotheca tamesis 24 1.52 Brand and Guillard (1981)
Thalassiosira pseudonana 24 0.76 Brand and Guillard (1981)
Thalassiosira pseudonana 24 0.97 Brand and Guillard (1981)
Thalassiosira pseudonana 24 1.32 Brand and Guillard (1981)
Thalassiosira pseudonana 24 1.73 Brand and Guillard (1981)
189
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira pseudonana 24 2.08 Brand and Guillard (1981)
Thalassiosira pseudonana 24 3.40 Brand and Guillard (1981)
Thalassiosira pseudonana 24 3.26 Brand and Guillard (1981)
Thalassiosira pseudonana 24 2.63 Brand and Guillard (1981)
Thalassiosira sp. 24 0.76 Brand and Guillard (1981)
Thalassiosira sp. 24 1.87 Brand and Guillard (1981)
Thalassiosira sp. 24 1.04 Brand and Guillard (1981)
Thalassiosira sp. 24 1.87 Brand and Guillard (1981)
Ceratium platycorne 24 0.48 Brand and Guillard (1981)
Ceratium platycorne 24 0.24 Brand and Guillard (1981)
Ceratium platycorne 24 0.17 Brand and Guillard (1981)
Ceratium platycorne 24 0.33 Brand and Guillard (1981)
Ceratium platycorne 24 0.25 Brand and Guillard (1981)
Ceratium candelabrum 24 0.11 Brand and Guillard (1981)
Ceratium candelabrum 24 0.09 Brand and Guillard (1981)
Ceratium ranipes 24 0.33 Brand and Guillard (1981)
Ceratium ranipes 24 0.35 Brand and Guillard (1981)
Dissodinium lunula 24 0.19 Brand and Guillard (1981)
Dissodinium lunula 24 0.23 Brand and Guillard (1981)
Gonyaulax polyedra 24 0.62 Brand and Guillard (1981)
Gonyaulax polyedra 24 0.42 Brand and Guillard (1981)
Gonyaulax polyedra 24 0.69 Brand and Guillard (1981)
Gonyaulax polyedra 24 0.49 Brand and Guillard (1981)
Prorocentrum micans 24 0.62 Brand and Guillard (1981)
Prorocentrum micans 24 0.35 Brand and Guillard (1981)
Prorocentrum micans 24 0.49 Brand and Guillard (1981)
Prorocentrum micans 24 0.76 Brand and Guillard (1981)
Prorocentrum micans 24 0.90 Brand and Guillard (1981)
Prorocentrum micans 24 0.83 Brand and Guillard (1981)
Cyclococcolinthinia leptopora 24 0.39 Brand and Guillard (1981)
Cyclococcolinthinia leptopora 24 0.68 Brand and Guillard (1981)
Cyclococcolinthinia leptopora 24 0.62 Brand and Guillard (1981)
Cyclococcolinthinia leptopora 24 0.58 Brand and Guillard (1981)
Emiliana huxleyi 24 0.76 Brand and Guillard (1981)
Emiliana huxleyi 24 0.83 Brand and Guillard (1981)
Emiliana huxleyi 24 1.32 Brand and Guillard (1981)
Emiliana huxleyi 24 1.59 Brand and Guillard (1981)
Emiliana huxleyi 24 1.73 Brand and Guillard (1981)
Emiliana huxleyi 24 1.46 Brand and Guillard (1981)
Emiliana huxleyi 24 1.94 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 0.76 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 1.04 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 1.32 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 1.66 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 1.11 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 1.59 Brand and Guillard (1981)
Gephyrocapsa oceanica 24 1.25 Brand and Guillard (1981)
Hymenomonas carterae 24 0.58 Brand and Guillard (1981)
190
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Hymenomonas carterae 24 0.76 Brand and Guillard (1981)
Hymenomonas carterae 24 1.11 Brand and Guillard (1981)
Hymenomonas carterae 24 1.32 Brand and Guillard (1981)
Hymenomonas carterae 24 1.66 Brand and Guillard (1981)
Hymenomonas carterae 24 1.59 Brand and Guillard (1981)
Hymenomonas carterae 24 1.73 Brand and Guillard (1981)
Hymenomonas carterae 24 1.66 Brand and Guillard (1981)
Thalassiosira pseudonana 16 0.62 Brand et al (1981)
Thalassiosira pseudonana 16 0.90 Brand et al (1981)
Thalassiosira pseudonana 20 1.25 Brand et al (1981)
Thalassiosira pseudonana 20 1.59 Brand et al (1981)
Thalassiosira pseudonana 20 1.80 Brand et al (1981)
Thalassiosira pseudonana 24 1.52 Brand et al (1981)
Thalassiosira pseudonana 24 1.94 Brand et al (1981)
Thalassiosira pseudonana 24 2.08 Brand et al (1981)
Thalassiosira pseudonana 24 2.50 Brand et al (1981)
Thalassiosira pseudonana 12 0.26 Brand et al (1981)
Thalassiosira pseudonana 12 0.35 Brand et al (1981)
Thalassiosira pseudonana 12 0.52 Brand et al (1981)
Thalassiosira pseudonana 16 1.04 Brand et al (1981)
Thalassiosira pseudonana 20 1.39 Brand et al (1981)
Thalassiosira pseudonana 20 1.59 Brand et al (1981)
Thalassiosira pseudonana 20 1.80 Brand et al (1981)
Thalassiosira pseudonana 20 2.01 Brand et al (1981)
Thalassiosira pseudonana 20 2.22 Brand et al (1981)
Thalassiosira pseudonana 24 2.08 Brand et al (1981)
Thalassiosira pseudonana 24 2.43 Brand et al (1981)
Thalassiosira pseudonana 24 2.63 Brand et al (1981)
Thalassiosira pseudonana 24 2.77 Brand et al (1981)
Thalassiosira pseudonana 24 3.05 Brand et al (1981)
Thalassiosira pseudonana 12 0.62 Brand et al (1981)
Thalassiosira pseudonana 12 0.97 Brand et al (1981)
Thalassiosira pseudonana 12 1.11 Brand et al (1981)
Thalassiosira pseudonana 16 1.25 Brand et al (1981)
Thalassiosira pseudonana 16 1.94 Brand et al (1981)
Thalassiosira pseudonana 20 1.80 Brand et al (1981)
Thalassiosira pseudonana 20 2.63 Brand et al (1981)
Thalassiosira pseudonana 24 2.50 Brand et al (1981)
Thalassiosira pseudonana 24 3.26 Brand et al (1981)
Gymnodinium catenatum 13 0.08 Bravo and Anderson (1994)
Gymnodinium catenatum 15 0.15 Bravo and Anderson (1994)
Gymnodinium catenatum 16.5 0.26 Bravo and Anderson (1994)
Gymnodinium catenatum 18 0.30 Bravo and Anderson (1994)
Gymnodinium catenatum 19.5 0.24 Bravo and Anderson (1994)
Gymnodinium catenatum 21 0.28 Bravo and Anderson (1994)
Gymnodinium catenatum 22 0.37 Bravo and Anderson (1994)
Gymnodinium catenatum 24 0.36 Bravo and Anderson (1994)
Gymnodinium catenatum 25 0.32 Bravo and Anderson (1994)
191
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Gymnodinium catenatum 26 0.35 Bravo and Anderson (1994)
Gymnodinium catenatum 27 0.35 Bravo and Anderson (1994)
UnID cryptophyte 1 0.84 Buma et. al (1993)
UnID cryptophyte 1 0.72 Buma et. al (1993)
Pyramimonas sp. 1 0.08 Buma et. al (1993)
Cryptomonas ovata 8 0.07 Cloern (1977)
Cryptomonas ovata 14 0.20 Cloern (1977)
Cryptomonas ovata 20 0.60 Cloern (1977)
Cryptomonas ovata 26 0.35 Cloern (1977)
Pyramimonas tychotreta 4.6 0.45 Daugbjerg (2000)
Pyramimonas tychotreta 7 0.28 Daugbjerg (2000)
Skeletonema costatum 18 3.02 Davis et al (1973)
Skeletonema costatum 18 1.80 Davis et al (1973)
Skeletonema costatum 18 1.56 Davis et al (1973)
Skeletonema costatum 18 1.34 Davis et al (1973)
Skeletonema costatum 18 1.46 Davis et al (1973)
Skeletonema costatum 18 1.44 Davis et al (1973)
Skeletonema costatum 18 1.44 Davis et al (1973)
Thalassiosira nordenskioldii 0 0.39 Durbin (1974)
Thalassiosira nordenskioldii 0 0.35 Durbin (1974)
Thalassiosira nordenskioldii 0 0.30 Durbin (1974)
Thalassiosira nordenskioldii 0 0.46 Durbin (1974)
Thalassiosira nordenskioldii 0 0.42 Durbin (1974)
Thalassiosira nordenskioldii 0 0.46 Durbin (1974)
Thalassiosira nordenskioldii 0 0.40 Durbin (1974)
Thalassiosira nordenskioldii 0 0.28 Durbin (1974)
Thalassiosira nordenskioldii 5 0.33 Durbin (1974)
Thalassiosira nordenskioldii 5 0.58 Durbin (1974)
Thalassiosira nordenskioldii 5 0.69 Durbin (1974)
Thalassiosira nordenskioldii 5 0.67 Durbin (1974)
Thalassiosira nordenskioldii 5 0.53 Durbin (1974)
Thalassiosira nordenskioldii 5 0.90 Durbin (1974)
Thalassiosira nordenskioldii 5 0.83 Durbin (1974)
Thalassiosira nordenskioldii 5 0.69 Durbin (1974)
Thalassiosira nordenskioldii 5 0.83 Durbin (1974)
Thalassiosira nordenskioldii 5 0.76 Durbin (1974)
Thalassiosira nordenskioldii 5 0.76 Durbin (1974)
Thalassiosira nordenskioldii 10 0.90 Durbin (1974)
Thalassiosira nordenskioldii 10 0.67 Durbin (1974)
Thalassiosira nordenskioldii 10 0.46 Durbin (1974)
Thalassiosira nordenskioldii 10 0.76 Durbin (1974)
Thalassiosira nordenskioldii 10 0.97 Durbin (1974)
Thalassiosira nordenskioldii 10 1.25 Durbin (1974)
Thalassiosira nordenskioldii 10 0.97 Durbin (1974)
Thalassiosira nordenskioldii 10 1.11 Durbin (1974)
Thalassiosira nordenskioldii 10 1.25 Durbin (1974)
Thalassiosira nordenskioldii 10 0.90 Durbin (1974)
Thalassiosira nordenskioldii 10 0.97 Durbin (1974)
192
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira nordenskioldii 10 1.11 Durbin (1974)
Thalassiosira nordenskioldii 15 0.83 Durbin (1974)
Thalassiosira nordenskioldii 15 0.69 Durbin (1974)
Thalassiosira nordenskioldii 15 0.42 Durbin (1974)
Thalassiosira nordenskioldii 15 0.90 Durbin (1974)
Thalassiosira nordenskioldii 15 1.04 Durbin (1974)
Thalassiosira nordenskioldii 15 1.11 Durbin (1974)
Thalassiosira nordenskioldii 15 0.90 Durbin (1974)
Thalassiosira nordenskioldii 15 1.11 Durbin (1974)
Thalassiosira nordenskioldii 15 1.18 Durbin (1974)
Phaeodactylum tricornutum 14 0.24 Fawley (1984)
Phaeodactylum tricornutum 14 0.94 Fawley (1984)
Phaeodactylum tricornutum 14 0.98 Fawley (1984)
Phaeodactylum tricornutum 14 1.20 Fawley (1984)
Phaeodactylum tricornutum 14 1.40 Fawley (1984)
Phaeodactylum tricornutum 16 0.32 Fawley (1984)
Phaeodactylum tricornutum 16 1.10 Fawley (1984)
Phaeodactylum tricornutum 16 1.30 Fawley (1984)
Phaeodactylum tricornutum 16 1.40 Fawley (1984)
Phaeodactylum tricornutum 16 1.80 Fawley (1984)
Phaeodactylum tricornutum 19 0.30 Fawley (1984)
Phaeodactylum tricornutum 19 1.10 Fawley (1984)
Phaeodactylum tricornutum 19 1.40 Fawley (1984)
Phaeodactylum tricornutum 19 1.70 Fawley (1984)
Phaeodactylum tricornutum 19 1.90 Fawley (1984)
Phaeodactylum tricornutum 21 0.28 Fawley (1984)
Phaeodactylum tricornutum 21 1.20 Fawley (1984)
Phaeodactylum tricornutum 21 1.40 Fawley (1984)
Phaeodactylum tricornutum 21 1.90 Fawley (1984)
Phaeodactylum tricornutum 21 2.10 Fawley (1984)
Phaeodactylum tricornutum 23 0.26 Fawley (1984)
Phaeodactylum tricornutum 23 1.10 Fawley (1984)
Phaeodactylum tricornutum 23 1.50 Fawley (1984)
Phaeodactylum tricornutum 23 2.00 Fawley (1984)
Phaeodactylum tricornutum 23 2.10 Fawley (1984)
Phaeodactylum tricornutum 25 0.24 Fawley (1984)
Phaeodactylum tricornutum 25 1.00 Fawley (1984)
Phaeodactylum tricornutum 25 1.20 Fawley (1984)
Phaeodactylum tricornutum 25 1.70 Fawley (1984)
Phaeodactylum tricornutum 25 1.80 Fawley (1984)
Stellarima microtrias 3 0.60 Fiala and Oriol (1990)
Nitzschia turgiduloides 5 0.43 Fiala and Oriol (1990)
Chaetoceros deflandrei 5 1.04 Fiala and Oriol (1990)
Corethron criophilum 4 0.38 Fiala and Oriol (1990)
Synedra sp. 5 0.56 Fiala and Oriol (1990)
Nitzschia turgiduloides 3 0.71 Fiala and Oriol (1990)
Nitzschia kerguelensis 4 0.78 Fiala and Oriol (1990)
Stellarima microtrias 4 0.85 Fiala and Oriol (1990)
193
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Nitzschia cylindrus 5 0.86 Fiala and Oriol (1990)
Chaetoceros deflandrei 5 0.88 Fiala and Oriol (1990)
Chaetoceros curvisetum 15 0.97 Furnas (1978)
Chaetoceros curvisetum 15 0.97 Furnas (1978)
Chaetoceros curvisetum 15 0.97 Furnas (1978)
Chaetoceros curvisetum 15 0.54 Furnas (1978)
Chaetoceros curvisetum 15 0.83 Furnas (1978)
Chaetoceros curvisetum 15 0.97 Furnas (1978)
Chaetoceros curvisetum 15 0.97 Furnas (1978)
Chaetoceros curvisetum 15 1.25 Furnas (1978)
Chaetoceros curvisetum 15 1.25 Furnas (1978)
Chaetoceros curvisetum 15 1.25 Furnas (1978)
Chaetoceros curvisetum 15 1.04 Furnas (1978)
Chaetoceros curvisetum 20 0.04 Furnas (1978)
Chaetoceros curvisetum 20 0.31 Furnas (1978)
Chaetoceros curvisetum 20 1.04 Furnas (1978)
Chaetoceros curvisetum 20 1.11 Furnas (1978)
Chaetoceros curvisetum 20 0.90 Furnas (1978)
Chaetoceros curvisetum 20 0.83 Furnas (1978)
Chaetoceros curvisetum 20 0.90 Furnas (1978)
Chaetoceros curvisetum 20 0.97 Furnas (1978)
Chaetoceros curvisetum 25 0.97 Furnas (1978)
Chaetoceros curvisetum 25 0.97 Furnas (1978)
Chaetoceros curvisetum 25 1.04 Furnas (1978)
Chaetoceros curvisetum 25 1.18 Furnas (1978)
Chaetoceros curvisetum 25 1.39 Furnas (1978)
Chaetoceros curvisetum 25 1.39 Furnas (1978)
Chaetoceros curvisetum 25 1.39 Furnas (1978)
Chaetoceros curvisetum 25 0.90 Furnas (1978)
Chaetoceros curvisetum 25 0.76 Furnas (1978)
Chaetoceros curvisetum 25 0.69 Furnas (1978)
Skeletonema costatum 20 1.39 Gallagher (1982)
Skeletonema costatum 20 1.46 Gallagher (1982)
Skeletonema costatum 20 1.66 Gallagher (1982)
Skeletonema costatum 20 2.15 Gallagher (1982)
Skeletonema costatum 20 0.07 Gallagher (1982)
Skeletonema costatum 20 0.26 Gallagher (1982)
Skeletonema costatum 20 1.25 Gallagher (1982)
Skeletonema costatum 20 0.27 Gallagher (1982)
Skeletonema costatum 20 0.90 Gallagher (1982)
Skeletonema costatum 20 0.97 Gallagher (1982)
Skeletonema costatum 20 0.07 Gallagher (1982)
Skeletonema costatum 20 1.32 Gallagher (1982)
Skeletonema costatum 20 1.59 Gallagher (1982)
Skeletonema costatum 20 0.69 Gallagher (1982)
Skeletonema costatum 20 1.73 Gallagher (1982)
Skeletonema costatum 20 0.42 Gallagher (1982)
Skeletonema costatum 20 1.46 Gallagher (1982)
194
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Skeletonema costatum 20 0.54 Gallagher (1982)
Skeletonema costatum 20 1.04 Gallagher (1982)
Skeletonema costatum 20 0.90 Gallagher (1982)
Skeletonema costatum 20 0.24 Gallagher (1982)
Skeletonema costatum 20 0.28 Gallagher (1982)
Skeletonema costatum 20 0.37 Gallagher (1982)
Skeletonema costatum 20 1.04 Gallagher (1982)
Skeletonema costatum 20 1.59 Gallagher (1982)
Skeletonema costatum 20 0.30 Gallagher (1982)
Skeletonema costatum 20 0.83 Gallagher (1982)
Skeletonema costatum 20 1.04 Gallagher (1982)
Skeletonema costatum 20 0.97 Gallagher (1982)
Skeletonema costatum 20 0.25 Gallagher (1982)
Skeletonema costatum 20 0.97 Gallagher (1982)
Skeletonema costatum 20 2.98 Gallagher (1982)
Skeletonema costatum 20 2.22 Gallagher (1982)
Skeletonema costatum 10 0.55 Gallagher (1982)
Skeletonema costatum 20 1.59 Gallagher (1982)
Skeletonema costatum 10 0.76 Gallagher (1982)
Skeletonema costatum 20 1.04 Gallagher (1982)
Skeletonema costatum 10 0.40 Gallagher (1982)
Skeletonema costatum 20 0.90 Gallagher (1982)
Skeletonema costatum 10 0.43 Gallagher (1982)
Skeletonema costatum 20 0.24 Gallagher (1982)
Skeletonema costatum 10 0.40 Gallagher (1982)
Skeletonema costatum 20 3.47 Gallagher (1982)
Skeletonema costatum 20 2.91 Gallagher (1982)
Skeletonema costatum 20 2.77 Gallagher (1982)
Skeletonema costatum 20 0.27 Gallagher (1982)
Skeletonema costatum 20 2.56 Gallagher (1982)
Skeletonema costatum 20 2.22 Gallagher (1982)
Thalassiosira antarctica -0.5 0.14 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.16 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.20 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.15 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.16 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.07 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.31 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.31 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.33 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.33 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.46 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.52 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.53 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.58 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.08 Gilstad and Sakshaug (1990)
195
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira antarctica -0.5 0.01 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.05 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.33 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.60 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.59 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.58 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.48 Gilstad and Sakshaug (1990)
Thalassiosira antarctica -0.5 0.51 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.11 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.12 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.13 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.20 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.13 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.02 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.10 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.13 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.31 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.34 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.31 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.33 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.29 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.27 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.39 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.44 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.43 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.13 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.20 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.37 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.56 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.46 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.50 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.58 Gilstad and Sakshaug (1990)
Thalassiosira nordenskioeldii -0.5 0.52 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.05 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.07 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.12 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.13 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.15 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.04 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.08 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.17 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.30 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.29 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.25 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.27 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.26 Gilstad and Sakshaug (1990)
196
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Porosira glacialis -0.5 0.25 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.31 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.36 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.19 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.25 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.16 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.10 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.42 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.40 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.24 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.32 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.38 Gilstad and Sakshaug (1990)
Porosira glacialis -0.5 0.35 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.10 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.05 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.06 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.01 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.11 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.02 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.08 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.12 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.19 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.12 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.13 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.11 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.29 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.05 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.35 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.44 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.42 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.34 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.30 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.23 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.36 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.41 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.36 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.39 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.50 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.46 Gilstad and Sakshaug (1990)
Nitzschia delicatissima -0.5 0.36 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.07 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.07 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.20 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.12 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.02 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.07 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.12 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.20 Gilstad and Sakshaug (1990)
197
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira bulbosa -0.5 0.31 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.38 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.34 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.32 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.32 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.43 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.45 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.46 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.44 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.12 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.24 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.42 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.59 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.59 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.60 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.46 Gilstad and Sakshaug (1990)
Thalassiosira bulbosa -0.5 0.54 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.12 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.01 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.05 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.07 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.11 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.16 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.15 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.19 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.23 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.19 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.29 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.33 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.34 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.03 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.19 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.38 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.42 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.40 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.30 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.43 Gilstad and Sakshaug (1990)
Thalassiosira bioculata -0.5 0.34 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.15 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.16 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.21 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.14 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.16 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.05 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.11 Gilstad and Sakshaug (1990)
198
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Chaetoceros furcellatus -0.5 0.14 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.23 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.37 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.37 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.41 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.49 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.30 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.28 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.33 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.33 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.24 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.46 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.10 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.15 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.38 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.42 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.43 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.39 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.42 Gilstad and Sakshaug (1990)
Chaetoceros furcellatus -0.5 0.31 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.15 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.16 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.12 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.14 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.19 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.04 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.11 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.09 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.22 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.12 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.18 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.21 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.25 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.16 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.41 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.39 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.41 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.39 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.37 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.22 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.36 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.37 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.30 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.31 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.21 Gilstad and Sakshaug (1990)
Amphiprora sp. -0.5 0.35 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.10 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.03 Gilstad and Sakshaug (1990)
199
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Nitzschia grunowii -0.5 0.16 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.15 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.17 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.03 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.14 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.17 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.23 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.34 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.31 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.33 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.33 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.34 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.44 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.43 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.45 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.41 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.28 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.25 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.27 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.36 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.55 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.55 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.56 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.34 Gilstad and Sakshaug (1990)
Nitzschia grunowii -0.5 0.14 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.07 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.09 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.13 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.15 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.19 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.04 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.07 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.09 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.15 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.30 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.38 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.34 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.37 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.26 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.33 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.43 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.42 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.34 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.45 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.02 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.20 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.27 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.37 Gilstad and Sakshaug (1990)
200
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Nitzschia vanhoeffenii -0.5 0.28 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.41 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.51 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.29 Gilstad and Sakshaug (1990)
Nitzschia vanhoeffenii -0.5 0.19 Gilstad and Sakshaug (1990)
Chlorella pyrenoidosa 19 1.36 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 19 1.45 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 25 1.95 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 25 2.14 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 28.5 1.84 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 28.5 2.22 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 35 3.94 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 35 4.32 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 39.2 4.26 Goldman and Carpenter (1974)
Chlorella pyrenoidosa 39.2 5.65 Goldman and Carpenter (1974)
Chlorella sp. 25 1.88 Goldman and Carpenter (1974)
Selenastrum capricornutum 24 1.85 Goldman and Carpenter (1974)
Selenastrum capricornutum 27 2.45 Goldman and Carpenter (1974)
Scenedesmus quadricauda 27 2.29 Goldman and Carpenter (1974)
Skeletonema costatum 19 1.27 Goldman and Carpenter (1974)
Thalassiosira pseudonana 13.5 0.48 Goldman and Carpenter (1974)
Thalassiosira pseudonana 18 1.14 Goldman and Carpenter (1974)
Thalassiosira pseudonana 24 1.46 Goldman and Carpenter (1974)
Thalassiosira pseudonana 20 2.77 Goldman and Carpenter (1974)
Thalassiosira pseudonana 16 1.34 Goldman and Carpenter (1974)
Nitzschia actinastroides 23 2.06 Goldman and Carpenter (1974)
Monochrysis lutheri 19 0.84 Goldman and Carpenter (1974)
Monochrysis lutheri 25 1.83 Goldman and Carpenter (1974)
Thalassiosira pseudonana 18 1.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.60 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.60 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.60 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.60 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.70 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.70 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.80 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.80 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 1.90 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.00 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.20 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.30 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.30 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.30 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.30 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.30 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.40 Goldman and McCarthy (1978)
201
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira pseudonana 18 2.40 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.50 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.60 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.60 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 2.80 Goldman and McCarthy (1978)
Thalassiosira pseudonana 18 3.00 Goldman and McCarthy (1978)
Melosira juergensii 5 0.30 Grant and Horner (1976)
Melosira juergensii 5 0.55 Grant and Horner (1976)
Melosira juergensii 5 0.90 Grant and Horner (1976)
Melosira juergensii 5 0.75 Grant and Horner (1976)
Melosira juergensii 5 0.80 Grant and Horner (1976)
Melosira juergensii 5 0.80 Grant and Horner (1976)
Melosira juergensii 5 0.70 Grant and Horner (1976)
Melosira juergensii 5 0.50 Grant and Horner (1976)
Melosira juergensii 5 0.45 Grant and Horner (1976)
Melosira juergensii 5 0.25 Grant and Horner (1976)
Porosira glacialis 5 0.45 Grant and Horner (1976)
Porosira glacialis 5 0.95 Grant and Horner (1976)
Porosira glacialis 5 0.85 Grant and Horner (1976)
Porosira glacialis 5 0.70 Grant and Horner (1976)
Porosira glacialis 5 0.80 Grant and Horner (1976)
Porosira glacialis 5 0.80 Grant and Horner (1976)
Porosira glacialis 5 0.60 Grant and Horner (1976)
Porosira glacialis 5 0.65 Grant and Horner (1976)
Porosira glacialis 5 0.75 Grant and Horner (1976)
Porosira glacialis 5 0.30 Grant and Horner (1976)
Porosira glacialis 5 0.15 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.55 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.80 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.75 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.75 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.75 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.75 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.75 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.80 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.75 Grant and Horner (1976)
Coscinodiscus lacustris 5 0.50 Grant and Horner (1976)
Navicula transitans 5 0.35 Grant and Horner (1976)
Navicula transitans 5 0.45 Grant and Horner (1976)
Navicula transitans 5 0.35 Grant and Horner (1976)
Navicula transitans 5 0.35 Grant and Horner (1976)
Navicula transitans 5 0.35 Grant and Horner (1976)
Navicula transitans 5 0.40 Grant and Horner (1976)
Navicula transitans 5 0.15 Grant and Horner (1976)
Cyclotella nana 10 0.90 Guillard and Ryther (1962)
202
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Cyclotella nana 10 0.90 Guillard and Ryther (1962)
Cyclotella nana 10 1.18 Guillard and Ryther (1962)
Cyclotella nana 10 1.46 Guillard and Ryther (1962)
Cyclotella nana 15 0.69 Guillard and Ryther (1962)
Cyclotella nana 15 1.39 Guillard and Ryther (1962)
Cyclotella nana 15 1.46 Guillard and Ryther (1962)
Cyclotella nana 15 1.46 Guillard and Ryther (1962)
Cyclotella nana 20 1.25 Guillard and Ryther (1962)
Cyclotella nana 20 1.46 Guillard and Ryther (1962)
Cyclotella nana 20 1.73 Guillard and Ryther (1962)
Cyclotella nana 20 1.80 Guillard and Ryther (1962)
Cyclotella nana 20 1.87 Guillard and Ryther (1962)
Detonula confervacea 10 0.97 Guillard and Ryther (1962)
Detonula confervacea 15 0.54 Guillard and Ryther (1962)
Detonula confervacea 10 0.34 Guillard and Ryther (1962)
Detonula confervacea 10 0.45 Guillard and Ryther (1962)
Detonula confervacea 10 0.53 Guillard and Ryther (1962)
Detonula confervacea 10 0.50 Guillard and Ryther (1962)
Cyclotella nana 20 0.14 Guillard and Ryther (1962)
Cyclotella nana 20 0.22 Guillard and Ryther (1962)
Cyclotella nana 20 0.67 Guillard and Ryther (1962)
Cyclotella nana 20 0.94 Guillard and Ryther (1962)
Cyclotella nana 20 1.25 Guillard and Ryther (1962)
Cyclotella nana 20 1.52 Guillard and Ryther (1962)
Cyclotella nana 20 1.52 Guillard and Ryther (1962)
Cyclotella nana 20 1.52 Guillard and Ryther (1962)
Cyclotella nana 20 1.25 Guillard and Ryther (1962)
Cyclotella nana 20 1.73 Guillard and Ryther (1962)
Cyclotella nana 20 1.87 Guillard and Ryther (1962)
Cyclotella nana 20 1.87 Guillard and Ryther (1962)
Cyclotella nana 20 1.87 Guillard and Ryther (1962)
Cyclotella nana 20 1.94 Guillard and Ryther (1962)
Cyclotella nana 20 1.52 Guillard and Ryther (1962)
Cyclotella nana 20 1.46 Guillard and Ryther (1962)
Cyclotella nana 20 1.52 Guillard and Ryther (1962)
Cyclotella nana 20 1.66 Guillard and Ryther (1962)
Cyclotella nana 20 1.73 Guillard and Ryther (1962)
Cyclotella nana 20 1.80 Guillard and Ryther (1962)
Cyclotella nana 20 1.94 Guillard and Ryther (1962)
Cyclotella nana 20 2.01 Guillard and Ryther (1962)
Cyclotella nana 20 2.01 Guillard and Ryther (1962)
Cyclotella nana 20 1.87 Guillard and Ryther (1962)
Cyclotella nana 20 1.94 Guillard and Ryther (1962)
Cyclotella nana 20 2.01 Guillard and Ryther (1962)
Cyclotella nana 20 1.94 Guillard and Ryther (1962)
Cyclotella nana 20 2.01 Guillard and Ryther (1962)
Cyclotella nana 20 2.08 Guillard and Ryther (1962)
Cyclotella nana 20 2.08 Guillard and Ryther (1962)
203
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Cyclotella nana 20 2.15 Guillard and Ryther (1962)
Cyclotella nana 20 2.36 Guillard and Ryther (1962)
Cyclotella nana 20 2.08 Guillard and Ryther (1962)
Cyclotella nana 20 2.15 Guillard and Ryther (1962)
Cyclotella nana 20 2.15 Guillard and Ryther (1962)
Cyclotella nana 20 1.94 Guillard and Ryther (1962)
Cyclotella nana 20 1.94 Guillard and Ryther (1962)
Cyclotella nana 20 2.29 Guillard and Ryther (1962)
Thalassiosira fluviatilis 15 0.70 Hobson (1974)
Thalassiosira fluviatilis 15 0.80 Hobson (1974)
Thalassiosira fluviatilis 15 1.10 Hobson (1974)
Thalassiosira fluviatilis 20 1.10 Hobson (1974)
Thalassiosira fluviatilis 20 1.30 Hobson (1974)
Thalassiosira fluviatilis 20 1.70 Hobson (1974)
Thalassiosira fluviatilis 25 1.20 Hobson (1974)
Thalassiosira fluviatilis 25 2.30 Hobson (1974)
Thalassiosira fluviatilis 25 3.60 Hobson (1974)
Isochrysis sp. 10 0.58 Hobson (1974)
Isochrysis sp. 10 0.93 Hobson (1974)
Isochrysis sp. 10 0.70 Hobson (1974)
Isochrysis sp. 15 0.87 Hobson (1974)
Isochrysis sp. 15 1.00 Hobson (1974)
Isochrysis sp. 15 1.30 Hobson (1974)
Isochrysis sp. 20 1.40 Hobson (1974)
Isochrysis sp. 20 1.60 Hobson (1974)
Isochrysis sp. 20 1.90 Hobson (1974)
Isochrysis sp. 25 1.60 Hobson (1974)
Isochrysis sp. 25 3.50 Hobson (1974)
Isochrysis sp. 25 1.20 Hobson (1974)
Chroomonas salina 10 0.54 Hobson (1974)
Chroomonas salina 10 0.97 Hobson (1974)
Chroomonas salina 10 0.62 Hobson (1974)
Chroomonas salina 15 0.87 Hobson (1974)
Chroomonas salina 15 1.20 Hobson (1974)
Chroomonas salina 15 1.40 Hobson (1974)
Chroomonas salina 20 0.87 Hobson (1974)
Chroomonas salina 20 1.30 Hobson (1974)
Chroomonas salina 20 1.70 Hobson (1974)
Chroomonas salina 25 1.40 Hobson (1974)
Chroomonas salina 25 2.20 Hobson (1974)
Chroomonas salina 25 2.30 Hobson (1974)
Porphyridium aerugineum 21 0.67 Hoogenhout and Amesz (1965)
Prorocentrum micans 20 0.30 Hoogenhout and Amesz (1965)
Asterionella formosa 18.5 0.91 Hoogenhout and Amesz (1965)
Asterionella japonica 18 0.62 Hoogenhout and Amesz (1965)
Asterionella japonica 25 0.82 Hoogenhout and Amesz (1965)
Navicula minima 25 0.67 Hoogenhout and Amesz (1965)
Nitzschia palea 25 1.01 Hoogenhout and Amesz (1965)
204
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Phaeodactylum tricornutum 19 1.30 Hoogenhout and Amesz (1965)
Tabellaria flocculosa var. flocculosa 20 0.67 Hoogenhout and Amesz (1965)
Botrydiopsis intercedens 25 0.72 Hoogenhout and Amesz (1965)
Bumilleriopsis brevis 25 1.39 Hoogenhout and Amesz (1965)
Polyedriella helvetica 25 0.53 Hoogenhout and Amesz (1965)
Tribonema aequale 25 0.33 Hoogenhout and Amesz (1965)
Tribonema minus 25 0.48 Hoogenhout and Amesz (1965)
Vischeria stellata 20 0.42 Hoogenhout and Amesz (1965)
Euglena gracilis 25 1.05 Hoogenhout and Amesz (1965)
Euglena gracilis 25 0.91 Hoogenhout and Amesz (1965)
Euglena gracilis var. bacillaris 25 0.72 Hoogenhout and Amesz (1965)
Ankistrodesmus braunii 25 1.11 Hoogenhout and Amesz (1965)
Chlorella ellipsoidea 25 1.73 Hoogenhout and Amesz (1965)
Chlorella ellipsoidea 25 1.59 Hoogenhout and Amesz (1965)
Chlorella pyrenoidosa 25 1.49 Hoogenhout and Amesz (1965)
Chlorella pyrenoidosa 25 1.49 Hoogenhout and Amesz (1965)
Chlorella pyrenoidosa 25 1.49 Hoogenhout and Amesz (1965)
Chlorella pyrenoidosa 25 1.44 Hoogenhout and Amesz (1965)
Chlorella vulgaris 25 1.20 Hoogenhout and Amesz (1965)
Scenedesmus obliquus 25 1.05 Hoogenhout and Amesz (1965)
Rhizosolenia fragilissima 9 0.33 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.64 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.43 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.41 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.62 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.79 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.43 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.73 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.83 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.46 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.64 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.38 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.50 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.63 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.69 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.35 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.49 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.64 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.57 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.41 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.69 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.55 Ignatiades and Smayda (1970)
205
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Rhizosolenia fragilissima 9 0.46 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.73 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.80 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.48 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.83 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.84 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.84 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.40 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.82 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.38 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.80 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.76 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.76 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.31 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.80 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.72 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.69 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.39 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.54 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.67 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.47 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.41 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.57 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.82 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.71 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.45 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.70 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.82 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 30 0.63 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.35 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.65 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.79 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.33 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.63 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.68 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.31 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.60 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.69 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.71 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.33 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.61 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.57 Ignatiades and Smayda (1970)
206
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Rhizosolenia fragilissima 25 0.39 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.39 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.61 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.80 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.71 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.43 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.64 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.81 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.80 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.46 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.58 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.75 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.78 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.41 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.54 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.74 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.55 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 9 0.40 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 12 0.50 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 18 0.72 Ignatiades and Smayda (1970)
Rhizosolenia fragilissima 25 0.75 Ignatiades and Smayda (1970)
Chaetoceros sp. 3 0.28 Jacques (1983)
Chaetoceros sp. 5 0.44 Jacques (1983)
Chaetoceros sp. 7 0.45 Jacques (1983)
Chaetoceros sp. 10 0.45 Jacques (1983)
Fragilariopsis kerguelensis 3 0.17 Jacques (1983)
Fragilariopsis kerguelensis 5 0.33 Jacques (1983)
Nitzschia turgiduloides 3 0.31 Jacques (1983)
Nitzschia turgiduloides 5 0.30 Jacques (1983)
Nitzschia turgiduloides 7 0.30 Jacques (1983)
Alexandrium ostenfeldii 11.3 0.06 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 11.4 0.06 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 16.3 0.15 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 16.4 0.16 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 20.3 0.22 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 20.6 0.21 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 23.5 0.16 Jensen and Moestrup (1997)
Alexandrium ostenfeldii 23.7 0.15 Jensen and Moestrup (1997)
Skeletonema costatum 20 1.60 Jorgensen (1968)
Skeletonema costatum 15 1.30 Jorgensen (1968)
Skeletonema costatum 10 1.10 Jorgensen (1968)
Skeletonema costatum 7 0.69 Jorgensen (1968)
Thalassiosira rotula 5 0.33 Krawiec (1982)
Thalassiosira rotula 5 0.38 Krawiec (1982)
Thalassiosira rotula 10 0.82 Krawiec (1982)
Thalassiosira rotula 10 1.20 Krawiec (1982)
Thalassiosira rotula 15 1.30 Krawiec (1982)
Thalassiosira rotula 15 1.50 Krawiec (1982)
207
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira rotula 20 1.70 Krawiec (1982)
Thalassiosira rotula 20 1.30 Krawiec (1982)
Thalassiosira rotula 25 1.60 Krawiec (1982)
Thalassiosira rotula 30 1.30 Krawiec (1982)
Thalassiosira rotula 5 0.52 Krawiec (1982)
Thalassiosira rotula 5 0.76 Krawiec (1982)
Thalassiosira rotula 10 1.00 Krawiec (1982)
Thalassiosira rotula 10 1.30 Krawiec (1982)
Thalassiosira rotula 15 1.50 Krawiec (1982)
Thalassiosira rotula 15 1.60 Krawiec (1982)
Thalassiosira rotula 20 1.70 Krawiec (1982)
Thalassiosira rotula 20 2.10 Krawiec (1982)
Thalassiosira rotula 25 1.90 Krawiec (1982)
Thalassiosira rotula 25 2.30 Krawiec (1982)
Thalassiosira rotula 30 0.88 Krawiec (1982)
Thalassiosira rotula 5 0.49 Krawiec (1982)
Thalassiosira rotula 5 0.59 Krawiec (1982)
Thalassiosira rotula 10 0.95 Krawiec (1982)
Thalassiosira rotula 10 1.20 Krawiec (1982)
Thalassiosira rotula 15 1.40 Krawiec (1982)
Thalassiosira rotula 15 1.50 Krawiec (1982)
Thalassiosira rotula 20 1.60 Krawiec (1982)
Thalassiosira rotula 20 1.90 Krawiec (1982)
Thalassiosira rotula 25 1.50 Krawiec (1982)
Thalassiosira rotula 25 2.30 Krawiec (1982)
Thalassiosira rotula 30 1.30 Krawiec (1982)
Thalassiosira rotula 5 0.58 Krawiec (1982)
Thalassiosira rotula 5 0.63 Krawiec (1982)
Thalassiosira rotula 10 0.90 Krawiec (1982)
Thalassiosira rotula 10 1.10 Krawiec (1982)
Thalassiosira rotula 15 1.40 Krawiec (1982)
Thalassiosira rotula 15 1.50 Krawiec (1982)
Thalassiosira rotula 20 1.60 Krawiec (1982)
Thalassiosira rotula 20 1.80 Krawiec (1982)
Thalassiosira rotula 25 1.60 Krawiec (1982)
Thalassiosira rotula 25 1.90 Krawiec (1982)
Thalassiosira rotula 5 0.61 Krawiec (1982)
Thalassiosira rotula 5 0.75 Krawiec (1982)
Thalassiosira rotula 10 1.10 Krawiec (1982)
Thalassiosira rotula 10 1.30 Krawiec (1982)
Thalassiosira rotula 15 1.40 Krawiec (1982)
Thalassiosira rotula 15 1.50 Krawiec (1982)
Thalassiosira rotula 20 1.70 Krawiec (1982)
Thalassiosira rotula 20 2.00 Krawiec (1982)
Thalassiosira rotula 25 1.80 Krawiec (1982)
Thalassiosira rotula 25 2.40 Krawiec (1982)
Thalassiosira rotula 10 0.45 Krawiec (1982)
Thalassiosira rotula 15 0.46 Krawiec (1982)
208
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Thalassiosira rotula 15 0.91 Krawiec (1982)
Thalassiosira rotula 20 0.42 Krawiec (1982)
Thalassiosira rotula 20 0.96 Krawiec (1982)
Thalassiosira rotula 5 0.12 Krawiec (1982)
Thalassiosira rotula 10 0.86 Krawiec (1982)
Thalassiosira rotula 15 1.30 Krawiec (1982)
Thalassiosira rotula 20 1.60 Krawiec (1982)
Thalassiosira rotula 20 1.80 Krawiec (1982)
Thalassiosira rotula 25 0.72 Krawiec (1982)
Thalassiosira rotula 25 0.63 Krawiec (1982)
Skeletonema costatum 15 0.04 Langdon (1987)
Skeletonema costatum 15 0.23 Langdon (1987)
Skeletonema costatum 15 0.86 Langdon (1987)
Skeletonema costatum 15 0.98 Langdon (1987)
Skeletonema costatum 15 1.34 Langdon (1987)
Skeletonema costatum 15 1.30 Langdon (1987)
Skeletonema costatum 15 1.64 Langdon (1987)
Skeletonema costatum 15 1.59 Langdon (1987)
Skeletonema costatum 15 1.67 Langdon (1987)
Skeletonema costatum 15 1.69 Langdon (1987)
Skeletonema costatum 15 1.62 Langdon (1987)
Olisthodiscus luteus 15 0.04 Langdon (1987)
Olisthodiscus luteus 15 0.12 Langdon (1987)
Olisthodiscus luteus 15 0.24 Langdon (1987)
Olisthodiscus luteus 15 0.42 Langdon (1987)
Olisthodiscus luteus 15 0.61 Langdon (1987)
Olisthodiscus luteus 15 0.58 Langdon (1987)
Olisthodiscus luteus 15 0.56 Langdon (1987)
Gonyaulax tamarensis 15 0.02 Langdon (1987)
Gonyaulax tamarensis 15 0.07 Langdon (1987)
Gonyaulax tamarensis 15 0.21 Langdon (1987)
Gonyaulax tamarensis 15 0.36 Langdon (1987)
Gonyaulax tamarensis 15 0.40 Langdon (1987)
Gonyaulax tamarensis 15 0.38 Langdon (1987)
Gonyaulax tamarensis 15 0.42 Langdon (1987)
Skeletonema costatum 5 0.58 Langdon (1988)
Skeletonema costatum 10 0.85 Langdon (1988)
Skeletonema costatum 15 1.11 Langdon (1988)
Skeletonema costatum 15 1.10 Langdon (1988)
Skeletonema costatum 15 1.64 Langdon (1988)
Skeletonema costatum 20 1.35 Langdon (1988)
Skeletonema costatum 25 1.72 Langdon (1988)
Olisthodiscus luteus 10 0.35 Langdon (1988)
Olisthodiscus luteus 15 0.56 Langdon (1988)
Olisthodiscus luteus 15 0.64 Langdon (1988)
Olisthodiscus luteus 15 0.58 Langdon (1988)
Olisthodiscus luteus 20 0.80 Langdon (1988)
Olisthodiscus luteus 25 0.84 Langdon (1988)
209
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Gonyaulax tamarensis 5 0.15 Langdon (1988)
Gonyaulax tamarensis 10 0.26 Langdon (1988)
Gonyaulax tamarensis 15 0.28 Langdon (1988)
Gonyaulax tamarensis 15 0.35 Langdon (1988)
Gonyaulax tamarensis 15 0.44 Langdon (1988)
Gonyaulax tamarensis 20 0.33 Langdon (1988)
Phaeodactylum tricornutum 5 0.24 Li and Morris (1982)
Phaeodactylum tricornutum 5 0.28 Li and Morris (1982)
Phaeodactylum tricornutum 9 0.37 Li and Morris (1982)
Phaeodactylum tricornutum 10 0.51 Li and Morris (1982)
Phaeodactylum tricornutum 10 0.53 Li and Morris (1982)
Phaeodactylum tricornutum 12 0.65 Li and Morris (1982)
Phaeodactylum tricornutum 15 0.95 Li and Morris (1982)
Phaeodactylum tricornutum 20 1.20 Li and Morris (1982)
Phaeodactylum tricornutum 25 1.10 Li and Morris (1982)
Phaeodactylum tricornutum 25 1.00 Li and Morris (1982)
Pseudo-nitzschia pseudodelicatissima 5 0.15 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 10 0.58 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 10 0.66 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 10 0.68 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 10 0.62 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 10 0.31 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 10 0.28 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 15 0.78 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 15 0.87 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 15 1.06 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 15 0.85 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 15 0.67 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 15 0.64 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 18 1.17 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 18 1.14 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 18 1.39 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 18 1.37 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 18 1.16 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 18 1.10 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 22 1.59 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 22 1.64 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 22 1.66 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 22 1.67 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 22 1.40 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 22 1.55 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 25 1.80 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 25 1.81 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 25 2.36 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 25 1.63 Lundholm et al (1997)
Pseudo-nitzschia pseudodelicatissima 25 1.60 Lundholm et al (1997)
Cryptomonas erosa 1 0.01 Morgan and Kalff (1979)
Cryptomonas erosa 4 0.06 Morgan and Kalff (1979)
210
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Cryptomonas erosa 4 0.14 Morgan and Kalff (1979)
Cryptomonas erosa 4 0.06 Morgan and Kalff (1979)
Cryptomonas erosa 4 0.06 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.12 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.21 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.28 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.39 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.48 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.33 Morgan and Kalff (1979)
Cryptomonas erosa 15 0.26 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.03 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.14 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.36 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.60 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.83 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.83 Morgan and Kalff (1979)
Cryptomonas erosa 20 0.76 Morgan and Kalff (1979)
Skeletonema costatum 18 1.40 Mortain-Bertrand (1989)
Skeletonema costatum 18 0.85 Mortain-Bertrand (1989)
Skeletonema costatum 18 0.85 Mortain-Bertrand (1989)
Skeletonema costatum 18 0.12 Mortain-Bertrand (1989)
Skeletonema costatum 3 0.12 Mortain-Bertrand (1989)
Corethron criophilum 3 0.23 Mortain-Bertrand (1989)
Corethron criophilum 3 0.14 Mortain-Bertrand (1989)
Corethron criophilum 3 0.15 Mortain-Bertrand (1989)
Nitzschia turgiduloides 3 0.42 Mortain-Bertrand (1989)
Nitzschia turgiduloides 3 0.37 Mortain-Bertrand (1989)
Nitzschia turgiduloides 3 0.37 Mortain-Bertrand (1989)
Stellarima microtrias 3 0.09 Mortain-Bertrand (1989)
Stellarima microtrias 3 0.12 Mortain-Bertrand (1989)
Stellarima microtrias 3 0.15 Mortain-Bertrand (1989)
Amphidinium klebsii 19 0.60 Morton et al (1992)
Amphidinium klebsii 21 0.14 Morton et al (1992)
Amphidinium klebsii 23 0.21 Morton et al (1992)
Amphidinium klebsii 25 0.25 Morton et al (1992)
Amphidinium klebsii 27 0.37 Morton et al (1992)
Amphidinium klebsii 29 0.34 Morton et al (1992)
Amphidinium klebsii 31 0.23 Morton et al (1992)
Amphidinium klebsii 33 0.19 Morton et al (1992)
Amphidinium klebsii 35 0.06 Morton et al (1992)
Amphidinium klebsii 23 0.05 Morton et al (1992)
Amphidinium klebsii 25 0.07 Morton et al (1992)
Amphidinium klebsii 27 0.10 Morton et al (1992)
Amphidinium klebsii 29 0.20 Morton et al (1992)
Amphidinium klebsii 31 0.14 Morton et al (1992)
Amphidinium klebsii 33 0.11 Morton et al (1992)
Amphidinium klebsii 19 0.04 Morton et al (1992)
Amphidinium klebsii 21 0.13 Morton et al (1992)
211
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Amphidinium klebsii 23 0.15 Morton et al (1992)
Amphidinium klebsii 25 0.21 Morton et al (1992)
Amphidinium klebsii 27 0.13 Morton et al (1992)
Amphidinium klebsii 29 0.07 Morton et al (1992)
Amphidinium klebsii 31 0.04 Morton et al (1992)
Amphidinium klebsii 21 0.05 Morton et al (1992)
Amphidinium klebsii 23 0.07 Morton et al (1992)
Amphidinium klebsii 25 0.10 Morton et al (1992)
Amphidinium klebsii 27 0.19 Morton et al (1992)
Amphidinium klebsii 29 0.14 Morton et al (1992)
Amphidinium klebsii 31 0.12 Morton et al (1992)
Amphidinium klebsii 21 0.05 Morton et al (1992)
Amphidinium klebsii 23 0.08 Morton et al (1992)
Amphidinium klebsii 25 0.19 Morton et al (1992)
Amphidinium klebsii 27 0.21 Morton et al (1992)
Amphidinium klebsii 29 0.17 Morton et al (1992)
Amphidinium klebsii 31 0.12 Morton et al (1992)
Amphidinium klebsii 33 0.04 Morton et al (1992)
Amphidinium klebsii 23 0.05 Morton et al (1992)
Amphidinium klebsii 25 0.11 Morton et al (1992)
Amphidinium klebsii 27 0.17 Morton et al (1992)
Amphidinium klebsii 29 0.15 Morton et al (1992)
Amphidinium klebsii 31 0.10 Morton et al (1992)
Amphidinium klebsii 33 0.07 Morton et al (1992)
Psuedonitzschia seriata 6 0.27 Nilawati et al (1997)
Nitzschia sp. 6 0.26 Nilawati et al (1997)
Thalassiosira pseudonana 20 0.48 Paasche (1973)
Thalassiosira pseudonana 20 1.00 Paasche (1973)
Thalassiosira pseudonana 20 1.60 Paasche (1973)
Thalassiosira pseudonana 20 2.00 Paasche (1973)
Thalassiosira pseudonana 20 2.40 Paasche (1973)
Thalassiosira pseudonana 20 0.50 Paasche (1973)
Thalassiosira pseudonana 20 1.00 Paasche (1973)
Thalassiosira pseudonana 20 1.70 Paasche (1973)
Thalassiosira pseudonana 20 2.10 Paasche (1973)
Thalassiosira pseudonana 20 2.70 Paasche (1973)
Thalassiosira pseudonana 20 2.50 Paasche (1973)
Tetraselmis maculata 18 0.65 Parsons et al (1961)
Dunaliella salina 18 0.35 Parsons et al (1961)
Monochrysis lutheri 18 0.27 Parsons et al (1961)
Syracosphaera carterae 18 0.25 Parsons et al (1961)
Chaetoceros sp. 18 0.30 Parsons et al (1961)
Skeletonema costatum 18 0.38 Parsons et al (1961)
Coscinodiscus sp. 18 0.14 Parsons et al (1961)
Phaeodactylum tricornutum 18 0.22 Parsons et al (1961)
Amphidinium carteri 18 0.57 Parsons et al (1961)
Exuviella sp. 18 0.15 Parsons et al (1961)
Agmenellum quadruplicatum 18 0.33 Parsons et al (1961)
212
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Chaetoceros sp. 1 0.53 Reay et al (1999)
Chaetoceros sp. 3 0.70 Reay et al (1999)
Chaetoceros sp. 5 0.79 Reay et al (1999)
Skeletonema costatum 15 0.04 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.15 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.26 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.32 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.36 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.62 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.23 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.30 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.60 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.69 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.97 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.97 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.26 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.47 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.62 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.83 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.04 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.11 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.32 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.39 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.46 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.69 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.90 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.18 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.76 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.83 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.97 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.32 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.66 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.55 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.69 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.90 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.32 Sakshaug and Andresen (1986)
Skeletonema costatum 15 1.39 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.44 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.60 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.69 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.76 Sakshaug and Andresen (1986)
Skeletonema costatum 15 0.83 Sakshaug and Andresen (1986)
Skeletonema costatum 18 2.08 Sakshaug and Holm-Hansen
(1977)
Skeletonema costatum 18 2.15 Sakshaug and Holm-Hansen
(1977)
Skeletonema costatum 18 2.08 Sakshaug and Holm-Hansen
(1977)
Skeletonema costatum 18 2.01 Sakshaug and Holm-Hansen
213
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
(1977)
Pavlova lutheri 18 1.32 Sakshaug and Holm-Hansen
(1977)
Pavlova lutheri 18 1.32 Sakshaug and Holm-Hansen
(1977)
Pavlova lutheri 18 0.83 Sakshaug and Holm-Hansen
(1977)
Ochromonas sp. 20 0.05 Sanders et al (2001)
Ochromonas sp. 20 0.04 Sanders et al (2001)
Ochromonas sp. 20 0.16 Sanders et al (2001)
Ochromonas sp. 20 0.27 Sanders et al (2001)
Ochromonas sp. 20 0.24 Sanders et al (2001)
Ochromonas sp. 20 0.36 Sanders et al (2001)
Ochromonas sp. 20 0.41 Sanders et al (2001)
Coelastrum sp. 16 0.96 Schlesinger et al (1981)
Coelastrum sp. 16 0.60 Schlesinger et al (1981)
Coelastrum sp. 16 0.37 Schlesinger et al (1981)
Coelastrum sp. 16 0.18 Schlesinger et al (1981)
Coelastrum sp. 26 1.43 Schlesinger et al (1981)
Coelastrum sp. 26 0.86 Schlesinger et al (1981)
Coelastrum sp. 26 0.61 Schlesinger et al (1981)
Coelastrum sp. 26 0.24 Schlesinger et al (1981)
Chlorella sp. 16 0.59 Schlesinger et al (1981)
Chlorella sp. 16 0.36 Schlesinger et al (1981)
Chlorella sp. 16 0.29 Schlesinger et al (1981)
Chlorella sp. 16 0.15 Schlesinger et al (1981)
Chlorella sp. 26 0.86 Schlesinger et al (1981)
Chlorella sp. 26 0.57 Schlesinger et al (1981)
Chlorella sp. 26 0.53 Schlesinger et al (1981)
Chlorella sp. 26 0.24 Schlesinger et al (1981)
Tetraedron sp. 16 0.40 Schlesinger et al (1981)
Tetraedron sp. 16 0.22 Schlesinger et al (1981)
Tetraedron sp. 16 0.19 Schlesinger et al (1981)
Tetraedron sp. 16 0.14 Schlesinger et al (1981)
Tetraedron sp. 26 0.83 Schlesinger et al (1981)
Tetraedron sp. 26 0.27 Schlesinger et al (1981)
Tetraedron sp. 26 0.24 Schlesinger et al (1981)
Tetraedron sp. 26 0.19 Schlesinger et al (1981)
Asterococcus sp. 16 0.53 Schlesinger et al (1981)
Asterococcus sp. 16 0.36 Schlesinger et al (1981)
Asterococcus sp. 16 0.29 Schlesinger et al (1981)
Asterococcus sp. 16 0.14 Schlesinger et al (1981)
Asterococcus sp. 26 0.30 Schlesinger et al (1981)
Asterococcus sp. 26 0.33 Schlesinger et al (1981)
Asterococcus sp. 26 0.29 Schlesinger et al (1981)
Asterococcus sp. 26 0.14 Schlesinger et al (1981)
Chlamydomonas globosa 5 0.77 Seaburg et al. (1981)
Chlamydomonas globosa 7.5 0.88 Seaburg et al. (1981)
Chlamydomonas globosa 10 1.02 Seaburg et al. (1981)
214
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Chlamydomonas globosa 12.5 1.22 Seaburg et al. (1981)
Chlamydomonas globosa 15 1.52 Seaburg et al. (1981)
Chlamydomonas globosa 18 1.82 Seaburg et al. (1981)
Chlamydomonas globosa 20 1.76 Seaburg et al. (1981)
Chlamydomonas intermedia -1 0.48 Seaburg et al. (1981)
Chlamydomonas intermedia 2 0.62 Seaburg et al. (1981)
Chlamydomonas intermedia 5 1.00 Seaburg et al. (1981)
Chlamydomonas intermedia 7.5 1.15 Seaburg et al. (1981)
Chlamydomonas intermedia 10 1.14 Seaburg et al. (1981)
Chlamydomonas intermedia 12.5 1.29 Seaburg et al. (1981)
Chlamydomonas intermedia 15 1.54 Seaburg et al. (1981)
Chlamydomonas intermedia 18 1.58 Seaburg et al. (1981)
Chlamydomonas subcaudata -1 0.36 Seaburg et al. (1981)
Chlamydomonas subcaudata 2 0.60 Seaburg et al. (1981)
Chlamydomonas subcaudata 5 0.81 Seaburg et al. (1981)
Chlamydomonas subcaudata 7.5 1.00 Seaburg et al. (1981)
Chlamydomonas subcaudata 10 1.10 Seaburg et al. (1981)
Chlamydomonas subcaudata 12.5 1.35 Seaburg et al. (1981)
Chlamydomonas subcaudata 15 1.06 Seaburg et al. (1981)
Chlamydomonas subcaudata 18 0.20 Seaburg et al. (1981)
Chlamydomonas alpina -1 0.08 Seaburg et al. (1981)
Chlamydomonas alpina 2 0.11 Seaburg et al. (1981)
Chlamydomonas alpina 5 0.34 Seaburg et al. (1981)
Chlamydomonas alpina 7.5 0.51 Seaburg et al. (1981)
Chlamydomonas alpina 10 0.84 Seaburg et al. (1981)
Chlamydomonas alpina 12.5 1.42 Seaburg et al. (1981)
Chlamydomonas alpina 15 1.04 Seaburg et al. (1981)
Chlamydomonas alpina 18 0.30 Seaburg et al. (1981)
Detonula confervacea 2 0.58 Smayda (1969)
Detonula confervacea 2 0.69 Smayda (1969)
Detonula confervacea 2 0.64 Smayda (1969)
Detonula confervacea 2 0.39 Smayda (1969)
Detonula confervacea 2 0.41 Smayda (1969)
Detonula confervacea 2 0.24 Smayda (1969)
Detonula confervacea 7 0.15 Smayda (1969)
Detonula confervacea 2 0.19 Smayda (1969)
Detonula confervacea 7 0.39 Smayda (1969)
Detonula confervacea 2 0.07 Smayda (1969)
Detonula confervacea 7 0.18 Smayda (1969)
Detonula confervacea 2 0.01 Smayda (1969)
Detonula confervacea 2 0.52 Smayda (1969)
Detonula confervacea 7 0.55 Smayda (1969)
Detonula confervacea 12 0.62 Smayda (1969)
Detonula confervacea 2 0.60 Smayda (1969)
Detonula confervacea 7 0.64 Smayda (1969)
Detonula confervacea 12 0.50 Smayda (1969)
Detonula confervacea 2 0.09 Smayda (1969)
Detonula confervacea 7 0.62 Smayda (1969)
215
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Detonula confervacea 12 0.75 Smayda (1969)
Detonula confervacea 2 0.11 Smayda (1969)
Detonula confervacea 7 0.72 Smayda (1969)
Detonula confervacea 12 0.17 Smayda (1969)
Detonula confervacea 2 0.58 Smayda (1969)
Detonula confervacea 2 0.81 Smayda (1969)
Detonula confervacea 7 0.95 Smayda (1969)
Detonula confervacea 12 0.68 Smayda (1969)
Detonula confervacea 2 0.70 Smayda (1969)
Detonula confervacea 7 0.95 Smayda (1969)
Detonula confervacea 12 0.67 Smayda (1969)
Detonula confervacea 2 0.53 Smayda (1969)
Detonula confervacea 7 0.83 Smayda (1969)
Detonula confervacea 12 1.03 Smayda (1969)
Detonula confervacea 2 0.28 Smayda (1969)
Detonula confervacea 7 0.82 Smayda (1969)
Detonula confervacea 12 0.52 Smayda (1969)
Detonula confervacea 2 0.69 Smayda (1969)
Detonula confervacea 2 0.83 Smayda (1969)
Detonula confervacea 7 0.94 Smayda (1969)
Detonula confervacea 12 0.73 Smayda (1969)
Detonula confervacea 2 0.76 Smayda (1969)
Detonula confervacea 7 1.04 Smayda (1969)
Detonula confervacea 12 0.66 Smayda (1969)
Detonula confervacea 2 0.56 Smayda (1969)
Detonula confervacea 7 1.02 Smayda (1969)
Detonula confervacea 12 1.07 Smayda (1969)
Detonula confervacea 2 0.30 Smayda (1969)
Detonula confervacea 7 0.98 Smayda (1969)
Detonula confervacea 12 1.03 Smayda (1969)
Detonula confervacea 2 0.64 Smayda (1969)
Detonula confervacea 2 0.84 Smayda (1969)
Detonula confervacea 7 0.93 Smayda (1969)
Detonula confervacea 12 0.82 Smayda (1969)
Detonula confervacea 2 0.76 Smayda (1969)
Detonula confervacea 7 1.05 Smayda (1969)
Detonula confervacea 12 0.69 Smayda (1969)
Detonula confervacea 2 0.49 Smayda (1969)
Detonula confervacea 7 0.99 Smayda (1969)
Detonula confervacea 12 0.96 Smayda (1969)
Detonula confervacea 2 0.28 Smayda (1969)
Detonula confervacea 7 0.97 Smayda (1969)
Detonula confervacea 12 1.02 Smayda (1969)
Detonula confervacea 2 0.39 Smayda (1969)
Detonula confervacea 2 0.84 Smayda (1969)
Detonula confervacea 7 0.84 Smayda (1969)
Detonula confervacea 2 0.73 Smayda (1969)
Detonula confervacea 7 1.10 Smayda (1969)
216
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Detonula confervacea 12 0.63 Smayda (1969)
Detonula confervacea 2 0.40 Smayda (1969)
Detonula confervacea 7 0.90 Smayda (1969)
Detonula confervacea 12 1.04 Smayda (1969)
Detonula confervacea 2 0.35 Smayda (1969)
Detonula confervacea 7 0.87 Smayda (1969)
Detonula confervacea 12 0.69 Smayda (1969)
Detonula confervacea 2 0.41 Smayda (1969)
Detonula confervacea 2 0.80 Smayda (1969)
Detonula confervacea 7 0.84 Smayda (1969)
Detonula confervacea 2 0.73 Smayda (1969)
Detonula confervacea 7 0.89 Smayda (1969)
Detonula confervacea 12 0.46 Smayda (1969)
Detonula confervacea 2 0.44 Smayda (1969)
Detonula confervacea 7 0.65 Smayda (1969)
Detonula confervacea 2 0.35 Smayda (1969)
Detonula confervacea 7 0.67 Smayda (1969)
Detonula confervacea 12 0.33 Smayda (1969)
Nitzschia seriata 0 0.30 Smith et al. (1994)
Nitzschia seriata -1.6 0.34 Smith et al. (1994)
Nitzschia seriata -0.5 0.26 Smith et al. (1994)
Nitzschia seriata 2 0.43 Smith et al. (1994)
Nitzschia seriata 4 0.44 Smith et al. (1994)
Nitzschia seriata 6 0.43 Smith et al. (1994)
Nitzschia seriata 6 0.40 Smith et al. (1994)
Nitzschia seriata 10 0.57 Smith et al. (1994)
Nitzschia seriata 12 0.36 Smith et al. (1994)
Corethron criophilum 0 0.40 Sommer (1989)
Rhizosolenia alata 0 0.45 Sommer (1989)
Rhizosolenia truncata 0 0.45 Sommer (1989)
Thalassiosira antarctica 0 0.47 Sommer (1989)
Biddulphia weissflogii 0 0.46 Sommer (1989)
Eucampia zodiacus 0 0.38 Sommer (1989)
Rhizosolenia hebetata 0 0.34 Sommer (1989)
Thalassiosira subtilis 0 0.46 Sommer (1989)
Thalassiothrix longissima 0 0.41 Sommer (1989)
Nitzschia kerguelensis 0 0.56 Sommer (1989)
Chaetoceros criophilum 0 0.32 Sommer (1989)
Pyramimonas sp. 0 0.53 Sommer (1989)
Nitzschia seriata 0 0.55 Sommer (1989)
Phaeocystis pouchetii 0 0.71 Sommer (1989)
Nitzschia cylindrus 0 0.69 Sommer (1989)
Pseudonitzschia seriata -1.5 0.30 Stapleford and Smith (1996)
Pseudonitzschia seriata -1 0.32 Stapleford and Smith (1996)
Pseudonitzschia seriata 0 0.35 Stapleford and Smith (1996)
Pseudonitzschia seriata 2 0.37 Stapleford and Smith (1996)
Pseudonitzschia seriata 4 0.35 Stapleford and Smith (1996)
Pseudonitzschia seriata 6 0.49 Stapleford and Smith (1996)
217
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Pseudonitzschia seriata 10 0.46 Stapleford and Smith (1996)
Thalassiosira weissflogii 13 0.18 Strzepek and Price (2000)
Thalassiosira weissflogii 13 0.36 Strzepek and Price (2000)
Thalassiosira weissflogii 16 0.54 Strzepek and Price (2000)
Thalassiosira weissflogii 16 0.67 Strzepek and Price (2000)
Thalassiosira weissflogii 18 0.93 Strzepek and Price (2000)
Thalassiosira weissflogii 18 1.05 Strzepek and Price (2000)
Thalassiosira weissflogii 21 1.46 Strzepek and Price (2000)
Thalassiosira weissflogii 21 1.53 Strzepek and Price (2000)
Thalassiosira weissflogii 24 1.62 Strzepek and Price (2000)
Thalassiosira weissflogii 24 1.84 Strzepek and Price (2000)
Thalassiosira weissflogii 27 1.88 Strzepek and Price (2000)
Thalassiosira weissflogii 27 1.92 Strzepek and Price (2000)
Thalassiosira weissflogii 29 2.00 Strzepek and Price (2000)
Thalassiosira weissflogii 29 2.37 Strzepek and Price (2000)
Chaetoceros sp. 0 0.40 Suzuki and Takahashi (1995)
Chaetoceros sp. 2 0.30 Suzuki and Takahashi (1995)
Chaetoceros sp. 5 0.60 Suzuki and Takahashi (1995)
Chaetoceros sp. 10 0.25 Suzuki and Takahashi (1995)
Nitzschia frigida 0 0.40 Suzuki and Takahashi (1995)
Nitzschia frigida -1.8 0.40 Suzuki and Takahashi (1995)
Nitzschia frigida 2 0.40 Suzuki and Takahashi (1995)
Nannochloris sp. 20 1.50 Thomas (1966)
Nannochloris sp. 24 2.10 Thomas (1966)
Nannochloris sp. 29 2.60 Thomas (1966)
Nannochloris sp. 33 3.00 Thomas (1966)
Nannochloris sp. 36 2.80 Thomas (1966)
Nannochloris sp. 15 0.46 Thomas (1966)
Nannochloris sp. 19.5 1.20 Thomas (1966)
Nannochloris sp. 23 1.50 Thomas (1966)
Nannochloris sp. 26.5 2.10 Thomas (1966)
Nannochloris sp. 30 3.00 Thomas (1966)
Nannochloris sp. 15.5 0.49 Thomas (1966)
Nannochloris sp. 19.5 1.20 Thomas (1966)
Nannochloris sp. 23 1.50 Thomas (1966)
Nannochloris sp. 26.5 1.60 Thomas (1966)
Nannochloris sp. 30 1.70 Thomas (1966)
Nannochloris sp. 33 1.90 Thomas (1966)
Nannochloris sp. 16 0.28 Thomas (1966)
Nannochloris sp. 20 1.00 Thomas (1966)
Nannochloris sp. 24 1.50 Thomas (1966)
Nannochloris sp. 27 1.80 Thomas (1966)
Nannochloris sp. 30 2.00 Thomas (1966)
Nannochloris sp. 33 2.00 Thomas (1966)
Chaetoceros sp. 20 2.70 Thomas (1966)
Chaetoceros sp. 25 3.60 Thomas (1966)
Chaetoceros sp. 29 4.40 Thomas (1966)
Chaetoceros sp. 33 3.90 Thomas (1966)
218
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Chaetoceros sp. 37 3.50 Thomas (1966)
Chaetoceros sp. 20 2.10 Thomas (1966)
Chaetoceros sp. 25 2.20 Thomas (1966)
Chaetoceros sp. 29 2.80 Thomas (1966)
Chaetoceros sp. 33 3.50 Thomas (1966)
Chaetoceros sp. 37 2.60 Thomas (1966)
Chaetoceros sp. 14.5 0.39 Thomas (1966)
Chaetoceros sp. 19.5 1.30 Thomas (1966)
Chaetoceros sp. 23 2.10 Thomas (1966)
Chaetoceros sp. 27 2.30 Thomas (1966)
Chaetoceros sp. 30 2.60 Thomas (1966)
Chaetoceros sp. 16 0.83 Thomas (1966)
Chaetoceros sp. 20 1.20 Thomas (1966)
Chaetoceros sp. 23.5 1.90 Thomas (1966)
Chaetoceros sp. 27 2.30 Thomas (1966)
Chaetoceros sp. 31 2.30 Thomas (1966)
Chaetoceros sp. 34 2.40 Thomas (1966)
Chaetoceros calcitrans 10 0.96 Thompson et al (1992)
Chaetoceros calcitrans 15 1.87 Thompson et al (1992)
Chaetoceros calcitrans 17.5 2.00 Thompson et al (1992)
Chaetoceros calcitrans 20 1.52 Thompson et al (1992)
Chaetoceros calcitrans 25 3.39 Thompson et al (1992)
Thalassiosira pseudonana 10 0.69 Thompson et al (1992)
Thalassiosira pseudonana 15 1.25 Thompson et al (1992)
Thalassiosira pseudonana 17.5 1.84 Thompson et al (1992)
Thalassiosira pseudonana 20 1.71 Thompson et al (1992)
Thalassiosira pseudonana 25 2.35 Thompson et al (1992)
Chaetoceros simplex 10 0.65 Thompson et al (1992)
Chaetoceros simplex 15 1.19 Thompson et al (1992)
Chaetoceros simplex 17.5 1.95 Thompson et al (1992)
Chaetoceros simplex 20 1.33 Thompson et al (1992)
Chaetoceros simplex 25 2.23 Thompson et al (1992)
Chaetoceros gracilis 10 0.74 Thompson et al (1992)
Chaetoceros gracilis 15 1.10 Thompson et al (1992)
Chaetoceros gracilis 17.5 1.54 Thompson et al (1992)
Chaetoceros gracilis 20 1.67 Thompson et al (1992)
Chaetoceros gracilis 25 1.01 Thompson et al (1992)
Phaeodactylum tricornutum 10 0.67 Thompson et al (1992)
Phaeodactylum tricornutum 15 1.37 Thompson et al (1992)
Phaeodactylum tricornutum 17.5 1.22 Thompson et al (1992)
Phaeodactylum tricornutum 20 1.47 Thompson et al (1992)
Phaeodactylum tricornutum 25 1.56 Thompson et al (1992)
Dunaliella tertiolecta 10 0.29 Thompson et al (1992)
Dunaliella tertiolecta 15 0.77 Thompson et al (1992)
Dunaliella tertiolecta 17.5 1.41 Thompson et al (1992)
Dunaliella tertiolecta 20 1.14 Thompson et al (1992)
Dunaliella tertiolecta 25 1.41 Thompson et al (1992)
Pavlova lutheri 10 0.62 Thompson et al (1992)
219
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Pavlova lutheri 15 1.09 Thompson et al (1992)
Pavlova lutheri 17.5 0.56 Thompson et al (1992)
Pavlova lutheri 20 1.79 Thompson et al (1992)
Pavlova lutheri 25 2.36 Thompson et al (1992)
Isochrysis galbana 10 0.63 Thompson et al (1992)
Isochrysis galbana 15 1.33 Thompson et al (1992)
Isochrysis galbana 17.5 0.43 Thompson et al (1992)
Isochrysis galbana 20 0.84 Thompson et al (1992)
Isochrysis galbana 25 1.38 Thompson et al (1992)
Thalassiosira pseudonana 10 0.64 Thompson et al (1992b)
Thalassiosira pseudonana 15 0.93 Thompson et al (1992b)
Thalassiosira pseudonana 15 1.26 Thompson et al (1992b)
Thalassiosira pseudonana 17.5 1.85 Thompson et al (1992b)
Thalassiosira pseudonana 20 1.77 Thompson et al (1992b)
Thalassiosira pseudonana 20 1.31 Thompson et al (1992b)
Thalassiosira pseudonana 25 2.37 Thompson et al (1992b)
Thalassiosira pseudonana 25 1.65 Thompson et al (1992b)
Chaetoceros gracilis 10 0.46 Thompson et al (1992b)
Chaetoceros gracilis 10 0.79 Thompson et al (1992b)
Chaetoceros gracilis 15 1.18 Thompson et al (1992b)
Chaetoceros gracilis 17.5 1.59 Thompson et al (1992b)
Chaetoceros gracilis 20 1.35 Thompson et al (1992b)
Chaetoceros gracilis 20 1.54 Thompson et al (1992b)
Chaetoceros gracilis 20 1.71 Thompson et al (1992b)
Chaetoceros gracilis 20 1.82 Thompson et al (1992b)
Chaetoceros gracilis 25 1.07 Thompson et al (1992b)
Chaetoceros gracilis 25 1.35 Thompson et al (1992b)
Chaetoceros gracilis 25 1.89 Thompson et al (1992b)
Chaetoceros simplex 10 0.36 Thompson et al (1992b)
Chaetoceros simplex 10 0.67 Thompson et al (1992b)
Chaetoceros simplex 15 0.74 Thompson et al (1992b)
Chaetoceros simplex 15 0.87 Thompson et al (1992b)
Chaetoceros simplex 15 1.23 Thompson et al (1992b)
Chaetoceros simplex 17.5 2.00 Thompson et al (1992b)
Chaetoceros simplex 20 1.38 Thompson et al (1992b)
Chaetoceros simplex 20 1.80 Thompson et al (1992b)
Chaetoceros simplex 20 2.00 Thompson et al (1992b)
Chaetoceros simplex 25 2.20 Thompson et al (1992b)
Chaetoceros simplex 25 2.40 Thompson et al (1992b)
Phaeodactylum tricornutum 10 0.65 Thompson et al (1992b)
Phaeodactylum tricornutum 15 0.89 Thompson et al (1992b)
Phaeodactylum tricornutum 15 1.40 Thompson et al (1992b)
Phaeodactylum tricornutum 17.5 1.10 Thompson et al (1992b)
Phaeodactylum tricornutum 17.5 1.30 Thompson et al (1992b)
Phaeodactylum tricornutum 20 1.30 Thompson et al (1992b)
Phaeodactylum tricornutum 20 1.50 Thompson et al (1992b)
Phaeodactylum tricornutum 25 1.60 Thompson et al (1992b)
Dunaliella tertiolecta 10 0.41 Thompson et al (1992b)
220
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Dunaliella tertiolecta 10 0.28 Thompson et al (1992b)
Dunaliella tertiolecta 15 0.70 Thompson et al (1992b)
Dunaliella tertiolecta 17.5 1.30 Thompson et al (1992b)
Dunaliella tertiolecta 17.5 1.40 Thompson et al (1992b)
Dunaliella tertiolecta 20 1.10 Thompson et al (1992b)
Dunaliella tertiolecta 25 1.40 Thompson et al (1992b)
Dunaliella tertiolecta 25 1.80 Thompson et al (1992b)
Pavlova lutheri 10 0.51 Thompson et al (1992b)
Pavlova lutheri 15 0.72 Thompson et al (1992b)
Pavlova lutheri 15 1.10 Thompson et al (1992b)
Pavlova lutheri 17.5 0.53 Thompson et al (1992b)
Pavlova lutheri 20 1.50 Thompson et al (1992b)
Pavlova lutheri 20 1.70 Thompson et al (1992b)
Pavlova lutheri 25 1.50 Thompson et al (1992b)
Pavlova lutheri 25 2.10 Thompson et al (1992b)
Pavlova lutheri 25 2.40 Thompson et al (1992b)
Isochrysis galbana 10 0.56 Thompson et al (1992b)
Isochrysis galbana 15 0.20 Thompson et al (1992b)
Isochrysis galbana 15 1.30 Thompson et al (1992b)
Isochrysis galbana 17.5 0.41 Thompson et al (1992b)
Isochrysis galbana 20 0.59 Thompson et al (1992b)
Isochrysis galbana 20 0.82 Thompson et al (1992b)
Isochrysis galbana 25 1.40 Thompson et al (1992b)
Isochrysis galbana 25 1.90 Thompson et al (1992b)
Chaetoceros calcitrans 10 0.97 Thompson et al (1992b)
Chaetoceros calcitrans 10 0.63 Thompson et al (1992b)
Chaetoceros calcitrans 15 1.10 Thompson et al (1992b)
Chaetoceros calcitrans 15 1.90 Thompson et al (1992b)
Chaetoceros calcitrans 17.5 2.00 Thompson et al (1992b)
Chaetoceros calcitrans 20 2.10 Thompson et al (1992b)
Chaetoceros calcitrans 20 1.80 Thompson et al (1992b)
Chaetoceros calcitrans 20 1.60 Thompson et al (1992b)
Chaetoceros calcitrans 25 3.00 Thompson et al (1992b)
Chaetoceros calcitrans 25 3.10 Thompson et al (1992b)
Chaetoceros calcitrans 25 3.40 Thompson et al (1992b)
Chaetoceros dichaeta 1.5 0.62 Timmermans et al (2001)
Chaetoceros dichaeta 1.5 Timmermans et al (2001)
Chaetoceros brevis 1.5 0.39 Timmermans et al (2001)
Leptocylindrus danicus 20 1.84 Verity (1982)
Leptocylindrus danicus 20 1.59 Verity (1982)
Leptocylindrus danicus 15 1.80 Verity (1982)
Leptocylindrus danicus 15 1.39 Verity (1982)
Leptocylindrus danicus 10 0.75 Verity (1982)
Leptocylindrus danicus 5 0.21 Verity (1982)
Gonyaulax tamarensis 8.5 0.12 Watras et al (1982)
Gonyaulax tamarensis 8.5 0.18 Watras et al (1982)
Gonyaulax tamarensis 8.5 0.20 Watras et al (1982)
Gonyaulax tamarensis 8.5 0.21 Watras et al (1982)
221
Appendix 2 (cont’d): Growth Rates of Phototrophic Protists Within
Chapter 2 Data Set
Gonyaulax tamarensis 8.5 0.22 Watras et al (1982)
Gonyaulax tamarensis 10.5 0.22 Watras et al (1982)
Gonyaulax tamarensis 10.5 0.24 Watras et al (1982)
Gonyaulax tamarensis 10.5 0.26 Watras et al (1982)
Gonyaulax tamarensis 13 0.28 Watras et al (1982)
Gonyaulax tamarensis 13 0.34 Watras et al (1982)
Gonyaulax tamarensis 13 0.38 Watras et al (1982)
Gonyaulax tamarensis 13 0.40 Watras et al (1982)
Gonyaulax tamarensis 13 0.42 Watras et al (1982)
Gonyaulax tamarensis 16 0.32 Watras et al (1982)
Gonyaulax tamarensis 16 0.36 Watras et al (1982)
Gonyaulax tamarensis 16 0.37 Watras et al (1982)
Gonyaulax tamarensis 16 0.42 Watras et al (1982)
Gonyaulax tamarensis 18 0.34 Watras et al (1982)
Gonyaulax tamarensis 18 0.36 Watras et al (1982)
Gonyaulax tamarensis 18 0.40 Watras et al (1982)
Gonyaulax tamarensis 18 0.44 Watras et al (1982)
Gonyaulax tamarensis 20 0.38 Watras et al (1982)
Gonyaulax tamarensis 20 0.44 Watras et al (1982)
Gonyaulax tamarensis 20 0.47 Watras et al (1982)
Gonyaulax tamarensis 20 0.48 Watras et al (1982)
Gonyaulax tamarensis 22.5 0.38 Watras et al (1982)
Gonyaulax tamarensis 22.5 0.40 Watras et al (1982)
Gonyaulax tamarensis 22.5 0.42 Watras et al (1982)
Gonyaulax tamarensis 22.5 0.45 Watras et al (1982)
Gonyaulax tamarensis 22.5 0.46 Watras et al (1982)
Gonyaulax tamarensis 24.5 0.20 Watras et al (1982)
Gonyaulax tamarensis 24.5 0.22 Watras et al (1982)
Skeletonema costatum 0 0.33 Yoder (1979)
Skeletonema costatum 5 0.67 Yoder (1979)
Skeletonema costatum 10 1.18 Yoder (1979)
Skeletonema costatum 16 2.22 Yoder (1979)
Skeletonema costatum 22 2.70 Yoder (1979)

222
Appendix 3: Growth Rates of Heterotrophic Protists Within Chapter 2
Data Set
Organism Temperature (
o
C) Growth rate (d
-1
) Study
Favella sp. 8 0.15 Aelion and Chisholm (1985)
Favella sp. 8 0.21 Aelion and Chisholm (1985)
Favella sp. 11.4 0.40 Aelion and Chisholm (1985)
Favella sp. 11.4 0.44 Aelion and Chisholm (1985)
Favella sp. 11.4 0.48 Aelion and Chisholm (1985)
Favella sp. 11.4 0.53 Aelion and Chisholm (1985)
Favella sp. 11.4 0.59 Aelion and Chisholm (1985)
Favella sp. 16.4 0.50 Aelion and Chisholm (1985)
Favella sp. 16.4 0.57 Aelion and Chisholm (1985)
Favella sp. 16.4 0.63 Aelion and Chisholm (1985)
Favella sp. 16.4 0.68 Aelion and Chisholm (1985)
Favella sp. 16.4 0.78 Aelion and Chisholm (1985)
Favella sp. 16.4 0.82 Aelion and Chisholm (1985)
Favella sp. 16.4 0.84 Aelion and Chisholm (1985)
Favella sp. 16.4 0.87 Aelion and Chisholm (1985)
Favella sp. 18.8 0.45 Aelion and Chisholm (1985)
Favella sp. 18.8 0.48 Aelion and Chisholm (1985)
Favella sp. 18.8 0.52 Aelion and Chisholm (1985)
Favella sp. 18.8 0.60 Aelion and Chisholm (1985)
Favella sp. 18.8 0.87 Aelion and Chisholm (1985)
Favella sp. 18.8 0.93 Aelion and Chisholm (1985)
Favella sp. 21.1 0.60 Aelion and Chisholm (1985)
Favella sp. 21.1 0.74 Aelion and Chisholm (1985)
Favella sp. 21.1 0.82 Aelion and Chisholm (1985)
Favella sp. 21.1 0.97 Aelion and Chisholm (1985)
Amoeba algonquinensis 5 0.12 Baldock and Berger (1984)
Amoeba algonquinensis 7.5 0.13 Baldock and Berger (1984)
Amoeba algonquinensis 10 0.23 Baldock and Berger (1984)
Amoeba algonquinensis 12 0.29 Baldock and Berger (1984)
Saccamoeba limax 5 0.24 Baldock and Berger (1984)
Saccamoeba limax 7.5 0.53 Baldock and Berger (1984)
Saccamoeba limax 10 0.62 Baldock and Berger (1984)
Saccamoeba limax 12 0.89 Baldock and Berger (1984)
Vannella sp. 5 0.09 Baldock and Berger (1984)
Vannella sp. 7.5 0.29 Baldock and Berger (1984)
Vannella sp. 10 0.36 Baldock and Berger (1984)
Vannella sp. 12 0.70 Baldock and Berger (1984)
Vannella sp. 7.5 0.14 Baldock and Berger (1984)
Vannella sp. 10 0.26 Baldock and Berger (1984)
Vannella sp. 12 0.36 Baldock and Berger (1984)
Acanthamoeba polyphaga 10 1.10 Baldock et al (1980)
Acanthamoeba polyphaga 15 3.20 Baldock et al (1980)
Acanthamoeba polyphaga 20 3.60 Baldock et al (1980)
Acanthamoeba polyphaga 25 3.00 Baldock et al (1980)
Cochliopodium minus 10 0.50 Baldock et al (1980)
223
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Cochliopodium minus 15 0.92 Baldock et al (1980)
Cochliopodium minus 20 1.60 Baldock et al (1980)
Cochliopodium minus 25 2.30 Baldock et al (1980)
Glaseria mira 10 1.00 Baldock et al (1980)
Glaseria mira 15 2.40 Baldock et al (1980)
Glaseria mira 20 2.40 Baldock et al (1980)
Glaseria mira 25 3.30 Baldock et al (1980)
Saccamoeba limax 10 0.82 Baldock et al (1980)
Saccamoeba limax 15 1.40 Baldock et al (1980)
Saccamoeba limax 20 2.20 Baldock et al (1980)
Saccamoeba limax 25 2.00 Baldock et al (1980)
Vanella sp. 10 0.56 Baldock et al (1980)
Vanella sp. 15 0.97 Baldock et al (1980)
Vanella sp. 20 1.50 Baldock et al (1980)
Vanella sp. 25 1.90 Baldock et al (1980)
Vexillifera bacillipedes 15 1.90 Baldock et al (1980)
Vexillifera bacillipedes 20 2.40 Baldock et al (1980)
Vexillifera bacillipedes 25 3.20 Baldock et al (1980)
Bodo saltans 20 4.56 Boenigk and Novarino (2004)
Bodo saltans 20 4.56 Boenigk and Novarino (2004)
Bodo saltans 20 4.80 Boenigk and Novarino (2004)
Spumella sp. 20 3.36 Boenigk and Novarino (2004)
Spumella sp. 20 4.80 Boenigk and Novarino (2004)
Spumella sp. 20 4.56 Boenigk and Novarino (2004)
Monosiga ovata 20 1.73 Boenigk and Novarino (2004)
Monosiga ovata 20 1.75 Boenigk and Novarino (2004)
Monosiga ovata 20 1.37 Boenigk and Novarino (2004)
Cyclidium sp. 20 2.88 Boenigk and Novarino (2004)
Cyclidium sp. 20 3.60 Boenigk and Novarino (2004)
Cyclidium sp. 20 3.84 Boenigk and Novarino (2004)
Tetryhymena pyriformis 20 3.84 Boenigk and Novarino (2004)
Tetryhymena pyriformis 20 3.60 Boenigk and Novarino (2004)
Tetryhymena pyriformis 20 3.36 Boenigk and Novarino (2004)
Halteria sp. 20 1.37 Boenigk and Novarino (2004)
Halteria sp. 20 1.34 Boenigk and Novarino (2004)
Halteria sp. 20 1.37 Boenigk and Novarino (2004)
Vorticella similis 20 2.26 Boenigk and Novarino (2004)
Vorticella similis 20 1.56 Boenigk and Novarino (2004)
Vorticella similis 20 2.04 Boenigk and Novarino (2004)
Entosiphon sulcatum 20 3.12 Boenigk and Novarino (2004)
Entosiphon sulcatum 20 2.40 Boenigk and Novarino (2004)
Entosiphon sulcatum 20 1.85 Boenigk and Novarino (2004)
Strombidinopsis sp. 20 0.59 Buskey and Hyatt (1995)
Strombidinopsis sp. 20 0.66 Buskey and Hyatt (1995)
Strombidinopsis sp. 20 0.95 Buskey and Hyatt (1995)
Strombidinopsis sp. 20 0.88 Buskey and Hyatt (1995)
224
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Fabrea salina 20 0.26 Buskey and Hyatt (1995)
Fabrea salina 20 0.39 Buskey and Hyatt (1995)
Fabrea salina 20 0.52 Buskey and Hyatt (1995)
Fabrea salina 20 0.51 Buskey and Hyatt (1995)
Fabrea salina 20 0.24 Buskey and Hyatt (1995)
Fabrea salina 20 0.26 Buskey and Hyatt (1995)
Fabrea salina 20 0.43 Buskey and Hyatt (1995)
Fabrea salina 20 0.54 Buskey and Hyatt (1995)
Fabrea salina 20 0.61 Buskey and Hyatt (1995)
Fabrea salina 20 0.71 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.39 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.50 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.61 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.56 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.58 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.22 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.27 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.37 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.40 Buskey and Hyatt (1995)
Noctiluca scintillans 20 0.38 Buskey and Hyatt (1995)
Euplotes sp. 20 0.07 Buskey and Hyatt (1995)
Euplotes sp. 20 0.21 Buskey and Hyatt (1995)
Euplotes sp. 20 0.39 Buskey and Hyatt (1995)
Euplotes sp. 20 0.41 Buskey and Hyatt (1995)
Euplotes sp. 20 0.45 Buskey and Hyatt (1995)
Euplotes sp. 20 0.48 Buskey and Hyatt (1995)
Euplotes sp. 20 0.09 Buskey and Hyatt (1995)
Euplotes sp. 20 0.19 Buskey and Hyatt (1995)
Euplotes sp. 20 0.28 Buskey and Hyatt (1995)
Euplotes sp. 20 0.50 Buskey and Hyatt (1995)
Euplotes sp. 20 0.35 Buskey and Hyatt (1995)
Euplotes sp. 20 0.27 Buskey and Hyatt (1995)
Euplotes sp. 20 0.29 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.05 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.23 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.45 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.66 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.62 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.73 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.59 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.12 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.20 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.24 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.36 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.42 Buskey and Hyatt (1995)
Oxyrrhis marina 20 0.54 Buskey and Hyatt (1995)
225
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Clydonella rosenfieldi 5 0.23 Butler and Rogerson (1996)
Clydonella rosenfieldi 10 0.29 Butler and Rogerson (1996)
Clydonella rosenfieldi 15 0.63 Butler and Rogerson (1996)
Clydonella rosenfieldi 20 1.34 Butler and Rogerson (1996)
Dactylamoeba sp. 10 0.16 Butler and Rogerson (1996)
Dactylamoeba sp. 15 0.20 Butler and Rogerson (1996)
Dactylamoeba sp. 20 0.49 Butler and Rogerson (1996)
Paraflabellula reniformis 5 0.26 Butler and Rogerson (1996)
Paraflabellula reniformis 10 0.60 Butler and Rogerson (1996)
Paraflabellula reniformis 15 0.84 Butler and Rogerson (1996)
Paraflabellula reniformis 20 0.96 Butler and Rogerson (1996)
Platyamoeba sp. 5 0.52 Butler and Rogerson (1996)
Platyamoeba sp. 10 0.69 Butler and Rogerson (1996)
Platyamoeba sp. 15 0.78 Butler and Rogerson (1996)
Platyamoeba sp. 20 1.49 Butler and Rogerson (1996)
Rhizamoeba sp. 5 0.13 Butler and Rogerson (1996)
Rhizamoeba sp. 10 0.25 Butler and Rogerson (1996)
Rhizamoeba sp. 15 0.31 Butler and Rogerson (1996)
Rhizamoeba sp. 20 0.55 Butler and Rogerson (1996)
Stereomyxa ramosa 5 0.16 Butler and Rogerson (1996)
Stereomyxa ramosa 10 0.19 Butler and Rogerson (1996)
Stereomyxa ramosa 15 0.38 Butler and Rogerson (1996)
Stereomyxa ramosa 20 0.37 Butler and Rogerson (1996)
Vahlkampfia baltica 5 0.23 Butler and Rogerson (1996)
Vahlkampfia baltica 10 0.26 Butler and Rogerson (1996)
Vahlkampfia baltica 15 0.71 Butler and Rogerson (1996)
Vahlkampfia baltica 20 1.05 Butler and Rogerson (1996)
Vahlkampfia damariscottae 5 0.26 Butler and Rogerson (1996)
Vahlkampfia damariscottae 10 0.80 Butler and Rogerson (1996)
Vahlkampfia damariscottae 15 0.77 Butler and Rogerson (1996)
Vahlkampfia damariscottae 20 1.32 Butler and Rogerson (1996)
Vannella caledonica 5 0.44 Butler and Rogerson (1996)
Vannella caledonica 10 0.60 Butler and Rogerson (1996)
Vannella caledonica 15 0.62 Butler and Rogerson (1996)
Vannella caledonica 20 0.70 Butler and Rogerson (1996)
Vannella sp. 5 0.31 Butler and Rogerson (1996)
Vannella sp. 10 0.41 Butler and Rogerson (1996)
Vannella sp. 15 0.76 Butler and Rogerson (1996)
Vannella sp. 20 1.09 Butler and Rogerson (1996)
Euplotes vannus 22 0.91 Capriulo et al (1988)
Euplotes vannus 22 0.82 Capriulo et al (1988)
Euplotes vannus 22 0.46 Capriulo et al (1988)
Euplotes vannus 22 0.17 Capriulo et al (1988)
Euplotes vannus 22 0.34 Capriulo et al (1988)
Euplotes vannus 22 0.06 Capriulo et al (1988)
Euplotes vannus 22 0.72 Capriulo et al (1988)
226
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Euplotes vannus 22 0.91 Capriulo et al (1988)
Euplotes vannus 22 1.22 Capriulo et al (1988)
Euplotes vannus 22 1.30 Capriulo et al (1988)
UnID chrysomonad 25 4.40 Caron (1990)
UNID kinetoplastid 25 1.00 Caron (1990)
Paraphysomonas imperforata 20 2.47 Caron et al (1985)
Paraphysomonas imperforata 20 2.55 Caron et al (1985)
Paraphysomonas imperforata 20 2.52 Caron et al (1985)
Paraphysomonas imperforata 20 2.10 Caron et al (1985)
Paraphysomonas imperforata 20 2.31 Caron et al (1985)
Paraphysomonas imperforata 24 3.48 Caron et al (1985)
Paraphysomonas imperforata 24 3.41 Caron et al (1985)
Paraphysomonas imperforata 24 3.50 Caron et al (1985)
Paraphysomonas imperforata 14 1.37 Caron et al (1986)
Paraphysomonas imperforata 18 1.92 Caron et al (1986)
Paraphysomonas imperforata 22 2.52 Caron et al (1986)
Paraphysomonas imperforata 26 4.04 Caron et al (1986)
Scuticociliate 18 6.70 Caron et al (1991)
Scuticociliate 18 6.20 Caron et al (1991)
Scuticociliate 18 0.50 Caron et al (1991)
Scuticociliate 18 1.90 Caron et al (1991)
Scuticociliate 18 1.40 Caron et al (1991)
Scuticociliate 18 6.50 Caron et al (1991)
Scuticociliate 18 6.50 Caron et al (1991)
Scuticociliate 18 7.00 Caron et al (1991)
Hymenostome ciliate 18 2.60 Caron et al (1991)
Hymenostome ciliate 18 3.10 Caron et al (1991)
Hymenostome ciliate 18 1.40 Caron et al (1991)
Hymenostome ciliate 18 1.90 Caron et al (1991)
Hymenostome ciliate 18 1.00 Caron et al (1991)
Hymenostome ciliate 18 3.60 Caron et al (1991)
Hymenostome ciliate 18 3.60 Caron et al (1991)
Hymenostome ciliate 18 2.40 Caron et al (1991)
Paraphysomonas sp. 18 5.50 Caron et al (1991)
Paraphysomonas sp. 18 5.30 Caron et al (1991)
Paraphysomonas sp. 18 3.40 Caron et al (1991)
Paraphysomonas sp. 18 4.10 Caron et al (1991)
Paraphysomonas sp. 18 4.30 Caron et al (1991)
Paraphysomonas sp. 18 4.80 Caron et al (1991)
Paraphysomonas sp. 18 5.30 Caron et al (1991)
Paraphysomonas sp. 18 5.50 Caron et al (1991)
Paraphysomonas imperforata -1.5 0.77 Choi and Peters (1992)
Paraphysomonas imperforata 6 1.53 Choi and Peters (1992)
Paraphysomonas imperforata 15 2.34 Choi and Peters (1992)
Paraphysomonas imperforata -1.5 0.53 Choi and Peters (1992)
Paraphysomonas imperforata 6 0.87 Choi and Peters (1992)
227
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Paraphysomonas imperforata 15 2.73 Choi and Peters (1992)
Paraphysomonas imperforata 15 1.29 Delaney (2003)
Paraphysomonas imperforata 15 2.24 Delaney (2003)
Paraphysomonas imperforata 10 0.60 Delaney (2003)
Paraphysomonas imperforata 10 0.91 Delaney (2003)
Paraphysomonas imperforata 5 0.36 Delaney (2003)
Paraphysomonas imperforata 5 0.46 Delaney (2003)
Paraphysomonas imperforata 0 0.22 Delaney (2003)
Paraphysomonas imperforata 0 0.36 Delaney (2003)
Euplotes vannus 23 1.66 Dini and Nyberg (1999)
Euplotes vannus 23 1.83 Dini and Nyberg (1999)
Euplotes vannus 23 1.87 Dini and Nyberg (1999)
Euplotes vannus 23 0.78 Dini and Nyberg (1999)
Euplotes vannus 23 0.97 Dini and Nyberg (1999)
Euplotes vannus 23 1.61 Dini and Nyberg (1999)
Euplotes vannus 23 1.90 Dini and Nyberg (1999)
Euplotes vannus 23 1.99 Dini and Nyberg (1999)
Euplotes vannus 23 0.74 Dini and Nyberg (1999)
Euplotes vannus 23 1.13 Dini and Nyberg (1999)
Euplotes vannus 23 1.56 Dini and Nyberg (1999)
Euplotes vannus 23 1.74 Dini and Nyberg (1999)
Euplotes vannus 23 1.66 Dini and Nyberg (1999)
Euplotes vannus 23 1.01 Dini and Nyberg (1999)
Euplotes vannus 23 1.09 Dini and Nyberg (1999)
Euplotes vannus 23 0.80 Dini and Nyberg (1999)
Euplotes vannus 23 1.43 Dini and Nyberg (1999)
Euplotes vannus 23 1.59 Dini and Nyberg (1999)
Euplotes vannus 23 1.53 Dini and Nyberg (1999)
Euplotes vannus 23 1.05 Dini and Nyberg (1999)
Euplotes vannus 23 1.15 Dini and Nyberg (1999)
Euplotes vannus 23 1.15 Dini and Nyberg (1999)
Euplotes vannus 23 1.41 Dini and Nyberg (1999)
Euplotes vannus 23 1.48 Dini and Nyberg (1999)
Euplotes vannus 23 1.12 Dini and Nyberg (1999)
Euplotes vannus 23 0.62 Dini and Nyberg (1999)
Euplotes vannus 23 0.82 Dini and Nyberg (1999)
Euplotes vannus 23 0.78 Dini and Nyberg (1999)
Euplotes vannus 23 1.38 Dini and Nyberg (1999)
Euplotes vannus 23 1.56 Dini and Nyberg (1999)
Euplotes vannus 23 1.39 Dini and Nyberg (1999)
Euplotes vannus 23 0.85 Dini and Nyberg (1999)
Euplotes vannus 23 0.72 Dini and Nyberg (1999)
Euplotes vannus 23 0.99 Dini and Nyberg (1999)
Euplotes crassus 23 1.28 Dini and Nyberg (1999)
Euplotes crassus 23 1.55 Dini and Nyberg (1999)
Euplotes crassus 23 1.76 Dini and Nyberg (1999)
228
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Euplotes crassus 23 0.91 Dini and Nyberg (1999)
Euplotes crassus 23 1.23 Dini and Nyberg (1999)
Euplotes crassus 23 2.05 Dini and Nyberg (1999)
Euplotes crassus 23 2.18 Dini and Nyberg (1999)
Euplotes crassus 23 2.48 Dini and Nyberg (1999)
Euplotes crassus 23 1.14 Dini and Nyberg (1999)
Euplotes crassus 23 1.63 Dini and Nyberg (1999)
Euplotes crassus 23 2.01 Dini and Nyberg (1999)
Euplotes crassus 23 2.05 Dini and Nyberg (1999)
Euplotes crassus 23 2.18 Dini and Nyberg (1999)
Euplotes crassus 23 1.48 Dini and Nyberg (1999)
Euplotes crassus 23 1.84 Dini and Nyberg (1999)
Euplotes crassus 23 1.82 Dini and Nyberg (1999)
Euplotes crassus 23 2.00 Dini and Nyberg (1999)
Euplotes crassus 23 1.98 Dini and Nyberg (1999)
Euplotes crassus 23 2.01 Dini and Nyberg (1999)
Euplotes crassus 23 1.16 Dini and Nyberg (1999)
Euplotes crassus 23 1.45 Dini and Nyberg (1999)
Euplotes crassus 23 1.64 Dini and Nyberg (1999)
Euplotes crassus 23 2.41 Dini and Nyberg (1999)
Euplotes crassus 23 2.22 Dini and Nyberg (1999)
Euplotes crassus 23 2.50 Dini and Nyberg (1999)
Euplotes crassus 23 1.47 Dini and Nyberg (1999)
Euplotes crassus 23 2.07 Dini and Nyberg (1999)
Euplotes crassus 23 2.13 Dini and Nyberg (1999)
Euplotes crassus 23 2.35 Dini and Nyberg (1999)
Euplotes crassus 23 2.25 Dini and Nyberg (1999)
Euplotes crassus 23 2.45 Dini and Nyberg (1999)
Euplotes crassus 23 1.16 Dini and Nyberg (1999)
Euplotes crassus 23 1.67 Dini and Nyberg (1999)
Euplotes crassus 23 1.77 Dini and Nyberg (1999)
Euplotes crassus 23 1.19 Dini and Nyberg (1999)
Euplotes crassus 23 1.32 Dini and Nyberg (1999)
Euplotes crassus 23 1.30 Dini and Nyberg (1999)
Euplotes crassus 23 0.95 Dini and Nyberg (1999)
Euplotes crassus 23 1.22 Dini and Nyberg (1999)
Euplotes crassus 23 1.22 Dini and Nyberg (1999)
Euplotes crassus 23 1.11 Dini and Nyberg (1999)
Euplotes crassus 23 1.56 Dini and Nyberg (1999)
Euplotes crassus 23 1.21 Dini and Nyberg (1999)
Euplotes crassus 23 1.11 Dini and Nyberg (1999)
Euplotes crassus 23 1.17 Dini and Nyberg (1999)
Euplotes crassus 23 2.30 Dini and Nyberg (1999)
Euplotes crassus 23 2.30 Dini and Nyberg (1999)
Euplotes crassus 23 2.28 Dini and Nyberg (1999)
Euplotes crassus 23 1.50 Dini and Nyberg (1999)
229
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Euplotes crassus 23 2.05 Dini and Nyberg (1999)
Euplotes crassus 23 2.24 Dini and Nyberg (1999)
Euplotes crassus 23 2.12 Dini and Nyberg (1999)
Euplotes crassus 23 2.08 Dini and Nyberg (1999)
Euplotes crassus 23 2.15 Dini and Nyberg (1999)
Euplotes crassus 23 1.06 Dini and Nyberg (1999)
Euplotes crassus 23 1.63 Dini and Nyberg (1999)
Euplotes crassus 23 1.72 Dini and Nyberg (1999)
Euplotes crassus 23 1.92 Dini and Nyberg (1999)
Euplotes crassus 23 2.07 Dini and Nyberg (1999)
Euplotes crassus 23 2.06 Dini and Nyberg (1999)
Euplotes crassus 23 1.53 Dini and Nyberg (1999)
Euplotes crassus 23 1.81 Dini and Nyberg (1999)
Euplotes crassus 23 1.92 Dini and Nyberg (1999)
Euplotes crassus 23 1.96 Dini and Nyberg (1999)
Euplotes crassus 23 1.94 Dini and Nyberg (1999)
Euplotes crassus 23 2.03 Dini and Nyberg (1999)
Euplotes crassus 23 1.23 Dini and Nyberg (1999)
Euplotes crassus 23 1.52 Dini and Nyberg (1999)
Euplotes crassus 23 1.72 Dini and Nyberg (1999)
Euplotes crassus 23 2.17 Dini and Nyberg (1999)
Euplotes crassus 23 2.21 Dini and Nyberg (1999)
Euplotes crassus 23 2.31 Dini and Nyberg (1999)
Euplotes crassus 23 1.39 Dini and Nyberg (1999)
Euplotes crassus 23 1.83 Dini and Nyberg (1999)
Euplotes crassus 23 2.15 Dini and Nyberg (1999)
Euplotes crassus 23 2.37 Dini and Nyberg (1999)
Euplotes crassus 23 2.28 Dini and Nyberg (1999)
Euplotes crassus 23 2.43 Dini and Nyberg (1999)
Euplotes crassus 23 1.70 Dini and Nyberg (1999)
Euplotes crassus 23 2.26 Dini and Nyberg (1999)
Euplotes crassus 23 2.23 Dini and Nyberg (1999)
Euplotes crassus 23 2.06 Dini and Nyberg (1999)
Euplotes crassus 23 2.06 Dini and Nyberg (1999)
Euplotes crassus 23 2.06 Dini and Nyberg (1999)
Euplotes crassus 23 0.71 Dini and Nyberg (1999)
Euplotes crassus 23 1.38 Dini and Nyberg (1999)
Euplotes crassus 23 1.46 Dini and Nyberg (1999)
Euplotes crassus 23 2.12 Dini and Nyberg (1999)
Euplotes crassus 23 2.07 Dini and Nyberg (1999)
Euplotes crassus 23 2.29 Dini and Nyberg (1999)
Euplotes crassus 23 1.98 Dini and Nyberg (1999)
Euplotes crassus 23 2.38 Dini and Nyberg (1999)
Euplotes crassus 23 2.38 Dini and Nyberg (1999)
Euplotes crassus 23 1.89 Dini and Nyberg (1999)
Euplotes crassus 23 1.73 Dini and Nyberg (1999)
230
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Euplotes crassus 23 1.84 Dini and Nyberg (1999)
Euplotes crassus 23 1.20 Dini and Nyberg (1999)
Euplotes crassus 23 1.83 Dini and Nyberg (1999)
Euplotes crassus 23 1.66 Dini and Nyberg (1999)
Euplotes minuta 23 1.83 Dini and Nyberg (1999)
Euplotes minuta 23 1.90 Dini and Nyberg (1999)
Euplotes minuta 23 2.10 Dini and Nyberg (1999)
Euplotes minuta 23 1.74 Dini and Nyberg (1999)
Euplotes minuta 23 1.63 Dini and Nyberg (1999)
Euplotes minuta 23 1.66 Dini and Nyberg (1999)
Euplotes minuta 23 1.86 Dini and Nyberg (1999)
Euplotes minuta 23 1.90 Dini and Nyberg (1999)
Euplotes minuta 23 2.13 Dini and Nyberg (1999)
Euplotes minuta 23 1.94 Dini and Nyberg (1999)
Euplotes minuta 23 1.75 Dini and Nyberg (1999)
Euplotes minuta 23 1.94 Dini and Nyberg (1999)
Euplotes minuta 23 1.62 Dini and Nyberg (1999)
Euplotes minuta 23 2.16 Dini and Nyberg (1999)
Euplotes minuta 23 1.88 Dini and Nyberg (1999)
Euplotes minuta 23 1.99 Dini and Nyberg (1999)
Euplotes minuta 23 2.57 Dini and Nyberg (1999)
Euplotes minuta 23 2.75 Dini and Nyberg (1999)
Euplotes minuta 23 1.50 Dini and Nyberg (1999)
Euplotes minuta 23 1.98 Dini and Nyberg (1999)
Euplotes minuta 23 1.92 Dini and Nyberg (1999)
Euplotes minuta 23 2.14 Dini and Nyberg (1999)
Euplotes minuta 23 2.73 Dini and Nyberg (1999)
Euplotes minuta 23 2.95 Dini and Nyberg (1999)
Euplotes minuta 23 1.63 Dini and Nyberg (1999)
Euplotes minuta 23 1.50 Dini and Nyberg (1999)
Euplotes minuta 23 1.73 Dini and Nyberg (1999)
Euplotes minuta 23 1.99 Dini and Nyberg (1999)
Euplotes minuta 23 2.14 Dini and Nyberg (1999)
Euplotes minuta 23 2.15 Dini and Nyberg (1999)
Euplotes minuta 23 1.10 Dini and Nyberg (1999)
Euplotes minuta 23 1.19 Dini and Nyberg (1999)
Euplotes minuta 23 1.21 Dini and Nyberg (1999)
Euplotes minuta 23 1.46 Dini and Nyberg (1999)
Euplotes minuta 23 1.66 Dini and Nyberg (1999)
Euplotes minuta 23 1.66 Dini and Nyberg (1999)
Euplotes minuta 23 1.73 Dini and Nyberg (1999)
Euplotes minuta 23 1.69 Dini and Nyberg (1999)
Euplotes minuta 23 1.73 Dini and Nyberg (1999)
Euplotes minuta 23 2.32 Dini and Nyberg (1999)
Euplotes minuta 23 2.38 Dini and Nyberg (1999)
Euplotes minuta 23 2.36 Dini and Nyberg (1999)
231
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Euplotes minuta 23 1.76 Dini and Nyberg (1999)
Euplotes minuta 23 1.66 Dini and Nyberg (1999)
Euplotes minuta 23 1.97 Dini and Nyberg (1999)
Euplotes minuta 23 2.32 Dini and Nyberg (1999)
Euplotes minuta 23 2.40 Dini and Nyberg (1999)
Euplotes minuta 23 2.32 Dini and Nyberg (1999)
Paraphysomonas imperforata 20 1.06 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 1.34 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 1.51 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 1.78 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 1.99 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 2.10 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 2.16 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 3.36 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 4.08 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 5.28 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 6.00 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 0.84 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 0.89 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 1.18 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 1.82 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 2.21 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 3.10 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 3.60 Eccleston-Parry and Leadbeater (1994)
Bodo designis 20 3.84 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 0.29 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 0.48 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 0.50 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 0.79 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 0.84 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 0.98 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 1.15 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 1.20 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 1.27 Eccleston-Parry and Leadbeater (1994)
Stephanoeca diplocostata 20 1.25 Eccleston-Parry and Leadbeater (1994)
Paraphysomonas imperforata 20 5.04 Eccleston-Parry and Leadbeater (1995)
Paraphysomonas imperforata 20 4.08 Eccleston-Parry and Leadbeater (1995)
Paraphysomonas imperforata 20 4.32 Eccleston-Parry and Leadbeater (1995)
Bodo designis 20 3.36 Eccleston-Parry and Leadbeater (1995)
Bodo designis 20 2.40 Eccleston-Parry and Leadbeater (1995)
Bodo designis 20 2.40 Eccleston-Parry and Leadbeater (1995)
Jakoba libera 20 0.62 Eccleston-Parry and Leadbeater (1995)
Jakoba libera 20 0.50 Eccleston-Parry and Leadbeater (1995)
Jakoba libera 20 0.53 Eccleston-Parry and Leadbeater (1995)
Stephanoeca diplocostata 20 0.70 Eccleston-Parry and Leadbeater (1995)
Stephanoeca diplocostata 20 0.67 Eccleston-Parry and Leadbeater (1995)
232
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Stephanoeca diplocostata 20 0.48 Eccleston-Parry and Leadbeater (1995)
Uronema marina 0 0.15 Fenchel (1968)
Uronema marina 4 0.50 Fenchel (1968)
Uronema marina 4 0.56 Fenchel (1968)
Uronema marina 9 1.90 Fenchel (1968)
Uronema marina 12 1.90 Fenchel (1968)
Uronema marina 17 4.50 Fenchel (1968)
Uronema marina 20 4.90 Fenchel (1968)
Uronema marina 20 6.80 Fenchel (1968)
Aspidisca angulata 4 0.42 Fenchel (1968)
Aspidisca angulata 7.6 0.75 Fenchel (1968)
Aspidisca angulata 9.3 0.91 Fenchel (1968)
Aspidisca angulata 12 1.30 Fenchel (1968)
Aspidisca angulata 17.2 2.00 Fenchel (1968)
Aspidisca angulata 20 3.70 Fenchel (1968)
Euplotes vannus 6 0.47 Fenchel (1968)
Euplotes vannus 14.5 1.10 Fenchel (1968)
Litonotus lamella 5 0.30 Fenchel (1968)
Litonotus lamella 10 0.60 Fenchel (1968)
Litonotus lamella 23 1.70 Fenchel (1968)
Diophrys scutum 15 0.61 Fenchel (1968)
Diophrys scutum 20 0.74 Fenchel (1968)
Diophrys scutum 27 1.60 Fenchel (1968)
Lacrymaria marina 15 0.43 Fenchel (1968)
Keronopsis rubra 15 0.29 Fenchel (1968)
Keronopsis rubra 20 0.51 Fenchel (1968)
Condylostoma patulum 15 0.18 Fenchel (1968)
Condylostoma patulum 20 0.37 Fenchel (1968)
Monosiga sp. 20 1.06 Fenchel (1982)
Monosiga sp. 20 1.75 Fenchel (1982)
Monosiga sp. 20 2.26 Fenchel (1982)
Monosiga sp. 20 2.88 Fenchel (1982)
Paraphysomonas vestita 20 1.66 Fenchel (1982)
Paraphysomonas vestita 20 3.12 Fenchel (1982)
Paraphysomonas vestita 20 4.08 Fenchel (1982)
Paraphysomonas vestita 20 5.04 Fenchel (1982)
Actinomonas mirabilis 20 3.60 Fenchel (1982)
Actinomonas mirabilis 20 4.56 Fenchel (1982)
Actinomonas mirabilis 20 5.04 Fenchel (1982)
Actinomonas mirabilis 20 5.28 Fenchel (1982)
Actinomonas mirabilis 20 5.52 Fenchel (1982)
Actinomonas mirabilis 20 5.76 Fenchel (1982)
Actinomonas mirabilis 20 5.76 Fenchel (1982)
Ochromonas sp. 20 0.72 Fenchel (1982)
Ochromonas sp. 20 0.86 Fenchel (1982)
Ochromonas sp. 20 1.30 Fenchel (1982)
233
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Ochromonas sp. 20 1.32 Fenchel (1982)
Ochromonas sp. 20 1.70 Fenchel (1982)
Ochromonas sp. 20 1.99 Fenchel (1982)
Ochromonas sp. 20 2.14 Fenchel (1982)
Ochromonas sp. 20 3.12 Fenchel (1982)
Ochromonas sp. 20 3.60 Fenchel (1982)
Ochromonas sp. 20 4.32 Fenchel (1982)
Ochromonas sp. 20 3.60 Fenchel (1982)
Psuedobodo tremulans 20 1.25 Fenchel (1982)
Psuedobodo tremulans 20 1.78 Fenchel (1982)
Psuedobodo tremulans 20 2.40 Fenchel (1982)
Psuedobodo tremulans 20 2.64 Fenchel (1982)
Psuedobodo tremulans 20 3.12 Fenchel (1982)
Strombidium sulcatum 20 0.81 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 0.98 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 1.15 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 1.42 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 1.85 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 2.21 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 2.14 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 2.40 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 1.22 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 2.02 Fenchel and Jonsson (1988)
Strombidium sulcatum 20 1.92 Fenchel and Jonsson (1988)
Vorticella microstoma 8.5 0.29 Finlay (1977)
Vorticella microstoma 15 1.45 Finlay (1977)
Vorticella microstoma 20 2.61 Finlay (1977)
Cyclidium glaucoma 8.5 0.22 Finlay (1977)
Cyclidium glaucoma 15 1.22 Finlay (1977)
Cyclidium glaucoma 20 2.37 Finlay (1977)
Tetrahymena pyriformis 8.5 0.53 Finlay (1977)
Tetrahymena pyriformis 15 1.30 Finlay (1977)
Tetrahymena pyriformis 20 2.05 Finlay (1977)
Chilodonella uncinata 8.5 0.72 Finlay (1977)
Chilodonella uncinata 15 1.19 Finlay (1977)
Chilodonella uncinata 20 1.56 Finlay (1977)
Colpidium campylum 8.5 0.34 Finlay (1977)
Colpidium campylum 15 0.87 Finlay (1977)
Colpidium campylum 20 1.32 Finlay (1977)
Loxocephalus plagius 8.5 0.23 Finlay (1977)
Loxocephalus plagius 15 0.54 Finlay (1977)
Loxocephalus plagius 20 1.19 Finlay (1977)
Paramecium aurelia 8.5 0.15 Finlay (1977)
Paramecium aurelia 15 0.55 Finlay (1977)
Paramecium aurelia 20 0.96 Finlay (1977)
Paramecium bursaria 8.5 0.19 Finlay (1977)
234
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Paramecium bursaria 15 0.46 Finlay (1977)
Paramecium bursaria 20 0.82 Finlay (1977)
Stentor polymorphus 8.5 0.12 Finlay (1977)
Stentor polymorphus 15 0.27 Finlay (1977)
Stentor polymorphus 20 0.39 Finlay (1977)
Spirostomum teres 8.5 0.02 Finlay (1977)
Spirostomum teres 15 0.18 Finlay (1977)
Spirostomum teres 20 0.20 Finlay (1977)
Paraphysomonas imperforata 21 3.77 Fu et al (2003)
Paraphysomonas imperforata 21 3.35 Fu et al (2003)
Paraphysomonas imperforata 21 2.76 Fu et al (2003)
Paraphysomonas imperforata 24 2.40 Goldman and Caron (1985)
Paraphysomonas imperforata 24 2.40 Goldman and Caron (1985)
Paraphysomonas imperforata 24 2.30 Goldman and Caron (1985)
Paraphysomonas imperforata 24 2.40 Goldman and Caron (1985)
Paraphysomonas imperforata 24 2.50 Goldman and Caron (1985)
Paraphysomonas imperforata 24 3.50 Goldman and Caron (1985)
Paraphysomonas imperforata 24 3.50 Goldman and Caron (1985)
Paraphysomonas imperforata 24 3.70 Goldman and Caron (1985)
Paraphysomonas imperforata 24 3.40 Goldman and Caron (1985)
Paraphysomonas imperforata 24 3.40 Goldman and Caron (1985)
Paraphysomonas imperforata 24 3.60 Goldman and Caron (1985)
Pfiesteria piscicida 23 0.78 Gransden and Lewitus (2003)
Cyptoperidiopsis sp. 23 0.22 Gransden and Lewitus (2003)
Euplotes vannus 23 0.31 Gransden and Lewitus (2003)
Euplotes vannus 23 0.30 Gransden and Lewitus (2003)
Euplotes vannus 23 0.67 Gransden and Lewitus (2003)
Euplotes woodruffii 23 0.86 Gransden and Lewitus (2003)
Euplotes woodruffii 23 0.59 Gransden and Lewitus (2003)
Euplotes woodruffii 23 0.66 Gransden and Lewitus (2003)
Strobilidium gyrans 20 0.69 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.77 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.77 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.12 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.76 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.61 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.30 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.38 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.41 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.39 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.45 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.84 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.87 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.71 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.68 Jack and Gilbert (1993)
Strobilidium gyrans 20 0.72 Jack and Gilbert (1993)
235
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Strobilidium gyrans 20 0.67 Jack and Gilbert (1993)
Bursaridium difficile 20 0.46 Jack and Gilbert (1993)
Bursaridium difficile 20 0.47 Jack and Gilbert (1993)
Bursaridium difficile 20 0.35 Jack and Gilbert (1993)
Bursaridium difficile 20 0.33 Jack and Gilbert (1993)
Bursaridium difficile 20 0.45 Jack and Gilbert (1993)
Bursaridium difficile 20 0.41 Jack and Gilbert (1993)
Bursaridium difficile 20 0.49 Jack and Gilbert (1993)
Bursaridium difficile 20 0.47 Jack and Gilbert (1993)
Bursaridium difficile 20 0.18 Jack and Gilbert (1993)
Bursaridium difficile 20 0.25 Jack and Gilbert (1993)
Bursaridium difficile 20 0.35 Jack and Gilbert (1993)
Bursaridium difficile 20 0.70 Jack and Gilbert (1993)
Bursaridium difficile 20 0.74 Jack and Gilbert (1993)
Bursaridium difficile 20 0.85 Jack and Gilbert (1993)
Bursaridium difficile 20 0.83 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.42 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.41 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.41 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.42 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.37 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.35 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.22 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.34 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.24 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.25 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.29 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.31 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.34 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.35 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.34 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.36 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.39 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.42 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.44 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.42 Jack and Gilbert (1993)
Euplotes eurystomus 20 0.45 Jack and Gilbert (1993)
Paramecium aurelia 20 0.39 Jack and Gilbert (1993)
Paramecium aurelia 20 0.38 Jack and Gilbert (1993)
Paramecium aurelia 20 0.52 Jack and Gilbert (1993)
Paramecium aurelia 20 0.45 Jack and Gilbert (1993)
Paramecium aurelia 20 0.43 Jack and Gilbert (1993)
Paramecium aurelia 20 0.34 Jack and Gilbert (1993)
Paramecium aurelia 20 0.37 Jack and Gilbert (1993)
Paramecium aurelia 20 0.23 Jack and Gilbert (1993)
Paramecium aurelia 20 0.11 Jack and Gilbert (1993)
236
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Paramecium aurelia 20 0.16 Jack and Gilbert (1993)
Paramecium aurelia 20 0.28 Jack and Gilbert (1993)
Paramecium aurelia 20 0.32 Jack and Gilbert (1993)
Paramecium aurelia 20 0.21 Jack and Gilbert (1993)
Paramecium aurelia 20 0.25 Jack and Gilbert (1993)
Paramecium aurelia 20 0.28 Jack and Gilbert (1993)
Paramecium aurelia 20 0.32 Jack and Gilbert (1993)
Paramecium aurelia 20 0.44 Jack and Gilbert (1993)
Paramecium aurelia 20 0.38 Jack and Gilbert (1993)
Paramecium aurelia 20 0.40 Jack and Gilbert (1993)
Balanion comatum 15 0.38 Jakobsen and Hansen (1997)
Balanion comatum 15 0.82 Jakobsen and Hansen (1997)
Balanion comatum 15 0.98 Jakobsen and Hansen (1997)
Balanion comatum 15 1.25 Jakobsen and Hansen (1997)
Balanion comatum 15 1.42 Jakobsen and Hansen (1997)
Balanion comatum 15 1.37 Jakobsen and Hansen (1997)
Balanion comatum 15 0.38 Jakobsen and Hansen (1997)
Balanion comatum 15 0.84 Jakobsen and Hansen (1997)
Balanion comatum 15 0.98 Jakobsen and Hansen (1997)
Balanion comatum 15 1.25 Jakobsen and Hansen (1997)
Balanion comatum 15 1.44 Jakobsen and Hansen (1997)
Balanion comatum 15 1.39 Jakobsen and Hansen (1997)
Balanion comatum 15 0.03 Jakobsen and Hansen (1997)
Balanion comatum 15 0.77 Jakobsen and Hansen (1997)
Balanion comatum 15 0.86 Jakobsen and Hansen (1997)
Balanion comatum 15 1.27 Jakobsen and Hansen (1997)
Balanion comatum 15 0.65 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.34 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.46 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.67 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.79 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.96 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.89 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 1.08 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 1.01 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.34 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.46 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.65 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.79 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.86 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.96 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.98 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 1.08 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.16 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.22 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.29 Jakobsen and Hansen (1997)
237
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Gymnodinium sp. 15 0.82 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.79 Jakobsen and Hansen (1997)
Gymnodinium sp. 15 0.38 Jakobsen and Hansen (1997)
Paraphysomonas vestita 18 1.20 John and Davidson (2001)
Paraphysomonas vestita 18 1.68 John and Davidson (2001)
Paraphysomonas vestita 18 1.20 John and Davidson (2001)
Paraphysomonas vestita 18 1.20 John and Davidson (2001)
Strombidium reticulatum 12 0.17 Jonsson (1986)
Strombidium reticulatum 12 0.34 Jonsson (1986)
Strombidium reticulatum 12 0.36 Jonsson (1986)
Strombidium reticulatum 12 0.48 Jonsson (1986)
Strombidium reticulatum 12 0.67 Jonsson (1986)
Strombidium reticulatum 12 0.77 Jonsson (1986)
Strombidium reticulatum 12 0.86 Jonsson (1986)
Strombidium reticulatum 12 0.74 Jonsson (1986)
Lohmanniella spiralis 12 0.23 Jonsson (1986)
Lohmanniella spiralis 12 0.43 Jonsson (1986)
Lohmanniella spiralis 12 0.43 Jonsson (1986)
Lohmanniella spiralis 12 0.62 Jonsson (1986)
Lohmanniella spiralis 12 0.89 Jonsson (1986)
Lohmanniella spiralis 12 1.00 Jonsson (1986)
Lohmanniella spiralis 12 1.06 Jonsson (1986)
Favella azorica 20 0.57 Kamiyama (1997)
Favella azorica 20 2.40 Kamiyama (1997)
Favella azorica 20 2.10 Kamiyama (1997)
Favella azorica 20 2.20 Kamiyama (1997)
Favella azorica 20 1.90 Kamiyama (1997)
Favella azorica 20 0.28 Kamiyama (1997)
Favella azorica 20 2.20 Kamiyama (1997)
Favella azorica 20 2.10 Kamiyama (1997)
Favella azorica 20 1.20 Kamiyama (1997)
Favella azorica 20 1.20 Kamiyama (1997)
Favella taraikaensis 20 0.37 Kamiyama (1997)
Favella taraikaensis 20 1.90 Kamiyama (1997)
Favella taraikaensis 20 1.60 Kamiyama (1997)
Favella taraikaensis 20 1.20 Kamiyama (1997)
Favella taraikaensis 20 0.52 Kamiyama (1997)
Favella taraikaensis 20 0.62 Kamiyama (1997)
Favella taraikaensis 20 1.30 Kamiyama (1997)
Favella taraikaensis 20 2.60 Kamiyama (1997)
Colpidium campylum 20 0.06 Laybourn and Stewart (1975)
Colpidium campylum 20 0.91 Laybourn and Stewart (1975)
Colpidium campylum 20 1.25 Laybourn and Stewart (1975)
Colpidium campylum 20 1.12 Laybourn and Stewart (1975)
Colpidium campylum 20 1.23 Laybourn and Stewart (1975)
Colpidium campylum 20 1.19 Laybourn and Stewart (1975)
238
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Colpidium campylum 15 0.60 Laybourn and Stewart (1975)
Colpidium campylum 15 0.62 Laybourn and Stewart (1975)
Colpidium campylum 15 0.70 Laybourn and Stewart (1975)
Colpidium campylum 15 0.75 Laybourn and Stewart (1975)
Colpidium campylum 15 0.74 Laybourn and Stewart (1975)
Colpidium campylum 15 0.69 Laybourn and Stewart (1975)
Colpidium campylum 15 0.61 Laybourn and Stewart (1975)
Colpidium campylum 10 0.25 Laybourn and Stewart (1975)
Colpidium campylum 10 0.51 Laybourn and Stewart (1975)
Colpidium campylum 10 0.27 Laybourn and Stewart (1975)
Colpidium campylum 10 0.49 Laybourn and Stewart (1975)
Colpidium campylum 10 0.37 Laybourn and Stewart (1975)
Colpidium campylum 10 0.46 Laybourn and Stewart (1975)
Arcella vulgaris 10 0.20 Laybourn and Whymant (1980)
Arcella vulgaris 15 0.22 Laybourn and Whymant (1980)
Arcella vulgaris 20 0.26 Laybourn and Whymant (1980)
Arcella vulgaris 25 0.26 Laybourn and Whymant (1980)
Euplotes vannus 8 0.23 Lee and Fenchel (1972)
Euplotes vannus 10 0.32 Lee and Fenchel (1972)
Euplotes vannus 15 0.68 Lee and Fenchel (1972)
Euplotes vannus 21 1.85 Lee and Fenchel (1972)
Euplotes vannus 26 2.31 Lee and Fenchel (1972)
Euplotes vannus 28 2.23 Lee and Fenchel (1972)
Euplotes balteatus 5 0.31 Lee and Fenchel (1972)
Euplotes balteatus 8 0.55 Lee and Fenchel (1972)
Euplotes balteatus 10 0.76 Lee and Fenchel (1972)
Euplotes balteatus 17 2.00 Lee and Fenchel (1972)
Euplotes balteatus 22 4.27 Lee and Fenchel (1972)
Euplotes balteatus 25 4.62 Lee and Fenchel (1972)
Euplotes balteatus 28 5.04 Lee and Fenchel (1972)
Euplotes balteatus 31 5.04 Lee and Fenchel (1972)
Euplotes balteatus 33 4.50 Lee and Fenchel (1972)
Euplotes balteatus 37 4.50 Lee and Fenchel (1972)
Euplotes balteatus 40 2.25 Lee and Fenchel (1972)
Holosticha sp -2 0.12 Lee and Fenchel (1972)
Euplotes antarcticus 0 0.16 Lee and Fenchel (1972)
Euplotes antarcticus 5 0.28 Lee and Fenchel (1972)
Euplotes antarcticus 10 0.28 Lee and Fenchel (1972)
Euplotes sp. 22 0.60 Liu and Buskey (2000)
Euplotes sp. 22 0.13 Liu and Buskey (2000)
Euplotes sp. 22 0.08 Liu and Buskey (2000)
Euplotes sp. 22 0.02 Liu and Buskey (2000)
Oxyrrhis marina 22 0.80 Liu and Buskey (2000)
Oxyrrhis marina 22 0.68 Liu and Buskey (2000)
Oxyrrhis marina 22 0.67 Liu and Buskey (2000)
Oxyrrhis marina 22 0.53 Liu and Buskey (2000)
239
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Aspidisca sp. 22 0.64 Liu and Buskey (2000)
Aspidisca sp. 22 0.42 Liu and Buskey (2000)
Aspidisca sp. 22 1.15 Liu and Buskey (2000)
Aspidisca sp. 22 0.77 Liu and Buskey (2000)
Uronema marinum 20 7.05 Martinez (1980)
Uronema marinum 25 8.32 Martinez (1980)
Uronema marinum 27 7.46 Martinez (1980)
Euplotes crassus 20 1.21 Martinez (1980)
Euplotes crassus 23 1.43 Martinez (1980)
Euplotes crassus 27 1.62 Martinez (1980)
Euplotes crassus 29 1.99 Martinez (1980)
Euplotes crassus 30 1.65 Martinez (1980)
Strombidium sulcatum 20 0.93 Martinez (1980)
Strombidium sulcatum 25 1.29 Martinez (1980)
Strombidium sulcatum 27 1.46 Martinez (1980)
Strombidium sulcatum 29 1.67 Martinez (1980)
Strombidium sulcatum 31 2.30 Martinez (1980)
Strombidium sulcatum 33 1.07 Martinez (1980)
Euplotes harpa 20 0.46 Martinez (1980)
Euplotes harpa 25 0.56 Martinez (1980)
Euplotes harpa 27 0.59 Martinez (1980)
Euplotes harpa 30 0.79 Martinez (1980)
Paratetrahymena wassi 20 0.44 Martinez (1980)
Paratetrahymena wassi 25 0.78 Martinez (1980)
Paratetrahymena wassi 27 1.07 Martinez (1980)
Paratetrahymena wassi 29 1.53 Martinez (1980)
Paratetrahymena wassi 30 1.90 Martinez (1980)
Paratetrahymena wassi 34 2.24 Martinez (1980)
Paratetrahymena wassi 35 1.74 Martinez (1980)
Condylostoma arenarium 20 0.39 Martinez (1980)
Condylostoma arenarium 25 0.52 Martinez (1980)
Condylostoma arenarium 27 0.64 Martinez (1980)
Condylostoma arenarium 29 0.78 Martinez (1980)
Condylostoma arenarium 31 0.69 Martinez (1980)
Euplotes trisulcatus 20 0.35 Martinez (1980)
Euplotes trisulcatus 25 0.64 Martinez (1980)
Euplotes trisulcatus 29 1.06 Martinez (1980)
Euplotes trisulcatus 31 1.28 Martinez (1980)
Euplotes trisulcatus 33 1.34 Martinez (1980)
Euplotes trisulcatus 35 0.82 Martinez (1980)
Platyamoeba australis 0 0.29 Mayes et. al (1997)
Platyamoeba australis -2 0.26 Mayes et. al (1997)
Platyamoeba australis 2 0.40 Mayes et. al (1997)
Platyamoeba australis 4 0.25 Mayes et. al (1997)
Mayorella sp. 0 0.03 Mayes et. al (1997)
Mayorella sp. -2 0.19 Mayes et. al (1997)
240
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Mayorella sp. 2 0.06 Mayes et. al (1997)
Mayorella sp. 4 0.04 Mayes et. al (1997)
Metachaos sp. 0 0.21 Mayes et. al (1997)
Metachaos sp. -2 0.07 Mayes et. al (1997)
Metachaos sp. 2 0.16 Mayes et. al (1997)
Metachaos sp. 4 0.10 Mayes et. al (1997)
Vannella sp. 0 0.11 Mayes et. al (1997)
Vannella sp. -2 0.10 Mayes et. al (1997)
Vannella sp. 2 0.23 Mayes et. al (1997)
Vannella sp. 4 0.13 Mayes et. al (1997)
Unidentified amoeba 0 0.14 Mayes et. al (1997)
Unidentified amoeba -2 0.19 Mayes et. al (1997)
Unidentified amoeba 2 0.70 Mayes et. al (1997)
Unidentified amoeba 4 0.25 Mayes et. al (1997)
Strobilidium neptuni 16 1.00 Montagnes (1996)
Strobilidium neptuni 16 1.10 Montagnes (1996)
Strobilidium neptuni 16 1.20 Montagnes (1996)
Strobilidium neptuni 16 1.30 Montagnes (1996)
Strobilidium neptuni 16 1.30 Montagnes (1996)
Strobilidium neptuni 16 1.40 Montagnes (1996)
Strobilidium neptuni 16 1.20 Montagnes (1996)
Strobilidium neptuni 16 1.50 Montagnes (1996)
Strobilidium neptuni 16 1.20 Montagnes (1996)
Strobilidium neptuni 16 1.40 Montagnes (1996)
Strobilidium veniliae 16 0.36 Montagnes (1996)
Strobilidium veniliae 16 0.38 Montagnes (1996)
Strobilidium veniliae 16 0.45 Montagnes (1996)
Strobilidium veniliae 16 0.53 Montagnes (1996)
Strobilidium veniliae 16 0.57 Montagnes (1996)
Strobilidium veniliae 16 0.63 Montagnes (1996)
Strobilidium veniliae 16 0.72 Montagnes (1996)
Strobilidium veniliae 16 0.63 Montagnes (1996)
Strobilidium veniliae 16 0.48 Montagnes (1996)
Strobilidium veniliae 16 0.60 Montagnes (1996)
Strobilidium veniliae 16 0.57 Montagnes (1996)
Strobilidium veniliae 16 0.70 Montagnes (1996)
Strobilidium veniliae 16 0.64 Montagnes (1996)
Strobilidium veniliae 16 0.55 Montagnes (1996)
Strobilidium veniliae 16 0.49 Montagnes (1996)
Strobilidium veniliae 16 0.45 Montagnes (1996)
Strobilidium veniliae 16 0.40 Montagnes (1996)
Strobilidium veniliae 16 0.38 Montagnes (1996)
Strobilidium veniliae 16 0.27 Montagnes (1996)
Strombidium capitatum 16 0.15 Montagnes (1996)
Strombidium capitatum 16 0.08 Montagnes (1996)
Strombidium capitatum 16 0.22 Montagnes (1996)
241
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Strombidium capitatum 16 0.32 Montagnes (1996)
Strombidium capitatum 16 0.64 Montagnes (1996)
Strombidium capitatum 16 0.20 Montagnes (1996)
Strombidium capitatum 16 0.25 Montagnes (1996)
Strombidium capitatum 16 0.35 Montagnes (1996)
Strombidium capitatum 16 0.60 Montagnes (1996)
Strombidium capitatum 16 0.64 Montagnes (1996)
Strombidium capitatum 16 0.74 Montagnes (1996)
Strombidium capitatum 16 0.54 Montagnes (1996)
Strombidium capitatum 16 0.75 Montagnes (1996)
Strombidium capitatum 16 0.78 Montagnes (1996)
Strombidium capitatum 16 0.42 Montagnes (1996)
Strombidium capitatum 16 0.48 Montagnes (1996)
Strombidium capitatum 16 0.60 Montagnes (1996)
Strombidium capitatum 16 0.67 Montagnes (1996)
Strombidium capitatum 16 0.83 Montagnes (1996)
Strombidium capitatum 16 0.59 Montagnes (1996)
Strombidium capitatum 16 0.83 Montagnes (1996)
Strombidium capitatum 16 0.31 Montagnes (1996)
Strombidium capitatum 16 0.59 Montagnes (1996)
Strombidium capitatum 16 0.63 Montagnes (1996)
Strombidium capitatum 16 0.11 Montagnes (1996)
Strombidium capitatum 16 0.28 Montagnes (1996)
Strombidium capitatum 16 0.41 Montagnes (1996)
Pseudobalanion planctonicum 5.5 0.46 Muller and Geller (1993)
Pseudobalanion planctonicum 9 0.83 Muller and Geller (1993)
Pseudobalanion planctonicum 12 0.94 Muller and Geller (1993)
Pseudobalanion planctonicum 15.5 1.24 Muller and Geller (1993)
Pseudobalanion planctonicum 18.5 1.52 Muller and Geller (1993)
Urotricha farcta 5.5 0.46 Muller and Geller (1993)
Urotricha farcta 9 0.52 Muller and Geller (1993)
Urotricha farcta 12 0.66 Muller and Geller (1993)
Urotricha farcta 15.5 0.75 Muller and Geller (1993)
Urotricha farcta 18.5 1.33 Muller and Geller (1993)
Urotricha farcta 21.5 1.72 Muller and Geller (1993)
Pelagostrombidium fallax 5.5 0.03 Muller and Geller (1993)
Pelagostrombidium fallax 9 0.21 Muller and Geller (1993)
Pelagostrombidium fallax 12 0.42 Muller and Geller (1993)
Pelagostrombidium fallax 15.5 0.57 Muller and Geller (1993)
Pelagostrombidium fallax 18.5 0.76 Muller and Geller (1993)
Pelagostrombidium fallax 21.5 0.90 Muller and Geller (1993)
Strobilidium lacustris 5.5 0.43 Muller and Geller (1993)
Strobilidium lacustris 9 0.60 Muller and Geller (1993)
Strobilidium lacustris 12 0.70 Muller and Geller (1993)
Strobilidium lacustris 15.5 0.99 Muller and Geller (1993)
Strobilidium lacustris 18.5 1.38 Muller and Geller (1993)
242
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Strobilidium lacustris 21.5 1.42 Muller and Geller (1993)
Noctiluca scintillans 24 0.03 Nakamura (1998)
Noctiluca scintillans 24 0.09 Nakamura (1998)
Noctiluca scintillans 24 0.13 Nakamura (1998)
Noctiluca scintillans 24 0.16 Nakamura (1998)
Noctiluca scintillans 24 0.20 Nakamura (1998)
Noctiluca scintillans 24 0.21 Nakamura (1998)
Noctiluca scintillans 24 0.27 Nakamura (1998)
Noctiluca scintillans 24 0.28 Nakamura (1998)
Noctiluca scintillans 24 0.01 Nakamura (1998)
Noctiluca scintillans 24 0.01 Nakamura (1998)
Noctiluca scintillans 24 0.01 Nakamura (1998)
Noctiluca scintillans 24 0.03 Nakamura (1998)
Noctiluca scintillans 24 0.03 Nakamura (1998)
Noctiluca scintillans 24 0.06 Nakamura (1998)
Noctiluca scintillans 24 0.07 Nakamura (1998)
Noctiluca scintillans 24 0.08 Nakamura (1998)
Noctiluca scintillans 24 0.09 Nakamura (1998)
Noctiluca scintillans 24 0.10 Nakamura (1998)
Noctiluca scintillans 24 0.17 Nakamura (1998)
Noctiluca scintillans 24 0.21 Nakamura (1998)
Noctiluca scintillans 24 0.27 Nakamura (1998)
Noctiluca scintillans 24 0.29 Nakamura (1998)
Gyrodinium dominans 11 0.08 Naustvoll (2000)
Gyrodinium dominans 11 0.19 Naustvoll (2000)
Gyrodinium dominans 11 0.16 Naustvoll (2000)
Gyrodinium dominans 11 0.02 Naustvoll (2000)
Gyrodinium dominans 11 0.02 Naustvoll (2000)
Gyrodinium dominans 11 0.05 Naustvoll (2000)
Gyrodinium dominans 11 0.10 Naustvoll (2000)
Gyrodinium fusiforme 11 0.10 Naustvoll (2000)
Gyrodinium fusiforme 11 0.09 Naustvoll (2000)
Gyrodinium fusiforme 11 0.08 Naustvoll (2000)
Gyrodinium fusiforme 11 0.12 Naustvoll (2000)
Gyrodinium fusiforme 11 0.39 Naustvoll (2000)
Katodinium glaucum 11 0.06 Naustvoll (2000)
Katodinium glaucum 11 0.13 Naustvoll (2000)
Katodinium glaucum 11 0.03 Naustvoll (2000)
Strombidium sp. 15 0.53 Ohman and Snyder (1991)
Strombidium sp. 15 0.45 Ohman and Snyder (1991)
Strombidium sp. 15 0.47 Ohman and Snyder (1991)
Strombidium sp. 15 0.33 Ohman and Snyder (1991)
Strombidium sp. 15 0.65 Ohman and Snyder (1991)
Strombidium sp. 15 1.00 Ohman and Snyder (1991)
Strombidium sp. 15 1.10 Ohman and Snyder (1991)
Strombidium sp. 15 1.30 Ohman and Snyder (1991)
243
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Strombidium sp. 20 0.71 Ohman and Snyder (1991)
Strombidium sp. 20 1.60 Ohman and Snyder (1991)
Strombidium sp. 20 1.80 Ohman and Snyder (1991)
Strombidium sp. 20 3.05 Ohman and Snyder (1991)
Strombidium sp. 20 2.30 Ohman and Snyder (1991)
Strombidium sp. 20 2.60 Ohman and Snyder (1991)
Rimostrombidium veniliae 15 1.05 Pedersen and Hansen (2003)
Rimostrombidium veniliae 15 1.06 Pedersen and Hansen (2003)
Rimostrombidium veniliae 15 0.92 Pedersen and Hansen (2003)
Rimostrombidium veniliae 15 0.75 Pedersen and Hansen (2003)
Rimostrombidium veniliae 15 0.19 Pedersen and Hansen (2003)
Rimostrombidium caudatum 15 0.43 Pedersen and Hansen (2003)
Rimostrombidium caudatum 15 0.61 Pedersen and Hansen (2003)
Rimostrombidium caudatum 15 0.41 Pedersen and Hansen (2003)
Rimostrombidium caudatum 15 0.51 Pedersen and Hansen (2003)
Balanion comatum 15 2.39 Pedersen and Hansen (2003)
Balanion comatum 15 2.48 Pedersen and Hansen (2003)
Balanion comatum 15 2.48 Pedersen and Hansen (2003)
Balanion comatum 15 2.54 Pedersen and Hansen (2003)
Balanion comatum 15 2.54 Pedersen and Hansen (2003)
Balanion comatum 15 2.63 Pedersen and Hansen (2003)
Balanion comatum 15 2.20 Pedersen and Hansen (2003)
Balanion comatum 15 2.11 Pedersen and Hansen (2003)
Balanion comatum 15 2.03 Pedersen and Hansen (2003)
Balanion comatum 15 1.83 Pedersen and Hansen (2003)
Balanion comatum 15 1.74 Pedersen and Hansen (2003)
Favella ehrenbergii 15 0.52 Pedersen and Hansen (2003)
Favella ehrenbergii 15 0.35 Pedersen and Hansen (2003)
Favella ehrenbergii 15 0.52 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.99 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.83 Pedersen and Hansen (2003)
Gyrodinium dominans 15 1.10 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.89 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.89 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.82 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.84 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.37 Pedersen and Hansen (2003)
Gyrodinium dominans 15 0.29 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.65 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.76 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.81 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.87 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.74 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.87 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.77 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.87 Pedersen and Hansen (2003)
244
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Oxyrrhis marina 15 0.87 Pedersen and Hansen (2003)
Oxyrrhis marina 15 0.87 Pedersen and Hansen (2003)
Uronema marinum 18 4.27 Perez-Uz (1995)
Uronema marinum 18 3.86 Perez-Uz (1995)
Uronema marinum 18 3.60 Perez-Uz (1995)
Uronema marinum 18 6.00 Perez-Uz (1995)
Uronema marinum 18 3.26 Perez-Uz (1995)
Uronema marinum 18 3.77 Perez-Uz (1995)
Uronema marinum 18 3.79 Perez-Uz (1995)
Uronema marinum 18 3.50 Perez-Uz (1995)
Uronema marinum 18 3.17 Perez-Uz (1995)
Uronema marinum 18 2.52 Perez-Uz (1995)
Uronema nigricans 18 2.74 Perez-Uz (1995)
Uronema nigricans 18 3.05 Perez-Uz (1995)
Uronema nigricans 18 2.11 Perez-Uz (1995)
Uronema nigricans 18 2.54 Perez-Uz (1995)
Parauronema acutum 18 2.62 Perez-Uz (1995)
Uronema sp. 18 3.22 Perez-Uz (1995)
Uronema elegans 18 2.26 Perez-Uz (1995)
Uronema marinum 18 4.49 Perez-Uz (1996)
Uronema marinum 18 4.27 Perez-Uz (1996)
Uronema marinum 18 3.26 Perez-Uz (1996)
Uronema marinum 18 4.13 Perez-Uz (1996)
Uronema marinum 18 3.24 Perez-Uz (1996)
Uronema marinum 18 1.85 Perez-Uz (1996)
Uronema marinum 18 4.32 Perez-Uz (1996)
Uronema marinum 18 3.84 Perez-Uz (1996)
Uronema marinum 18 2.11 Perez-Uz (1996)
Uronema nigricans 18 3.24 Perez-Uz (1996)
Uronema nigricans 18 3.07 Perez-Uz (1996)
Uronema nigricans 18 2.02 Perez-Uz (1996)
Uronema nigricans 18 2.52 Perez-Uz (1996)
Uronema nigricans 18 2.11 Perez-Uz (1996)
Uronema nigricans 18 1.68 Perez-Uz (1996)
Uronema nigricans 18 2.54 Perez-Uz (1996)
Uronema nigricans 18 2.28 Perez-Uz (1996)
Uronema nigricans 18 1.34 Perez-Uz (1996)
Ochromonas sp. 20 1.30 Sanders et al (2001)
Ochromonas sp. 20 1.10 Sanders et al (2001)
Ochromonas sp. 20 1.30 Sanders et al (2001)
Ochromonas sp. 20 1.60 Sanders et al (2001)
Ochromonas sp. 20 2.60 Sanders et al (2001)
Ochromonas sp. 20 1.70 Sanders et al (2001)
Paraphysomonas bandaiensis 18 0.19 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.67 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.72 Selph et al (2003)
245
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Paraphysomonas bandaiensis 18 0.03 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.75 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.85 Selph et al (2003)
Paraphysomonas bandaiensis 18 1.00 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.92 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.34 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.43 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.35 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.65 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.60 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.50 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.81 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.29 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.72 Selph et al (2003)
Paraphysomonas bandaiensis 18 1.20 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.12 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.75 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.61 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.54 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.27 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.42 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.50 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.82 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.79 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.70 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.68 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.34 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.20 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.67 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.45 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.90 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.45 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.07 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.52 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.43 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.68 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.13 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.82 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.35 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.89 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.33 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.76 Selph et al (2003)
Paraphysomonas bandaiensis 18 0.98 Selph et al (2003)
Monas sp. 3 0.78 Sherr et al (1983)
Monas sp. 18 3.10 Sherr et al (1983)
Monas sp. 23.5 3.30 Sherr et al (1983)
246
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Monas sp. 30 1.30 Sherr et al (1983)
Favella sp. 13 0.86 Strom and Morello (1998)
Strombidinopsis acuminatum 13 0.77 Strom and Morello (1998)
Uronema sp. 13 1.01 Strom and Morello (1998)
Gymnodinium sp. 13 0.48 Strom and Morello (1998)
Amphidinium sp. 13 0.47 Strom and Morello (1998)
Noctiluca scintillans 13 0.41 Strom and Morello (1998)
Euplotes focardii 4 0.23 Valbonesi and Luporini (1993)
Euplotes vannus 10 0.26 Walton et al (1995)
Euplotes vannus 15 0.62 Walton et al (1995)
Euplotes vannus 20 1.22 Walton et al (1995)
Euplotes vannus 25 1.01 Walton et al (1995)
Euplotes vannus 30 1.32 Walton et al (1995)
Euplotes vannus 10 0.31 Walton et al (1995)
Euplotes vannus 15 0.24 Walton et al (1995)
Euplotes vannus 20 1.13 Walton et al (1995)
Euplotes vannus 25 0.62 Walton et al (1995)
Euplotes vannus 30 0.86 Walton et al (1995)
Euplotes vannus 10 0.12 Walton et al (1995)
Euplotes vannus 15 0.62 Walton et al (1995)
Euplotes vannus 20 0.82 Walton et al (1995)
Euplotes vannus 25 0.82 Walton et al (1995)
Euplotes vannus 30 0.60 Walton et al (1995)
Euplotes vannus 10 0.58 Walton et al (1995)
Euplotes vannus 15 0.36 Walton et al (1995)
Euplotes vannus 20 0.82 Walton et al (1995)
Euplotes vannus 25 1.30 Walton et al (1995)
Euplotes vannus 30 0.82 Walton et al (1995)
Urotricha furcata 12.5 0.72 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.66 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.49 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.32 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.60 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.66 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.53 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.63 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.90 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.56 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.48 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.51 Weisse and Frahm (2002)
Urotricha furcata 12.5 0.85 Weisse and Frahm (2002)
Balanion planctonicum 12.5 0.96 Weisse and Frahm (2002)
Balanion planctonicum 12.5 0.60 Weisse and Frahm (2002)
Balanion planctonicum 12.5 1.03 Weisse and Frahm (2002)
Balanion planctonicum 12.5 0.66 Weisse and Frahm (2002)
Balanion planctonicum 12.5 1.00 Weisse and Frahm (2002)
247
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Urotricha furcata 10 0.19 Weisse and Montagnes (1998)
Urotricha furcata 10 0.25 Weisse and Montagnes (1998)
Urotricha furcata 15 0.58 Weisse and Montagnes (1998)
Urotricha furcata 15 0.77 Weisse and Montagnes (1998)
Urotricha furcata 15 0.80 Weisse and Montagnes (1998)
Urotricha furcata 20 1.00 Weisse and Montagnes (1998)
Urotricha furcata 25 0.90 Weisse and Montagnes (1998)
Urotricha furcata 25 0.97 Weisse and Montagnes (1998)
Urotricha furcata 25 1.00 Weisse and Montagnes (1998)
Urotricha furcata 10 0.35 Weisse and Montagnes (1998)
Urotricha furcata 10 0.43 Weisse and Montagnes (1998)
Urotricha furcata 15 0.68 Weisse and Montagnes (1998)
Urotricha furcata 15 0.84 Weisse and Montagnes (1998)
Urotricha furcata 25 0.85 Weisse and Montagnes (1998)
Urotricha furcata 25 0.95 Weisse and Montagnes (1998)
Urotricha furcata 10 0.40 Weisse and Montagnes (1998)
Urotricha furcata 10 0.49 Weisse and Montagnes (1998)
Urotricha furcata 15 0.42 Weisse and Montagnes (1998)
Urotricha furcata 15 0.51 Weisse and Montagnes (1998)
Urotricha furcata 20 0.80 Weisse and Montagnes (1998)
Urotricha furcata 25 0.91 Weisse and Montagnes (1998)
Urotricha furcata 5 0.14 Weisse and Montagnes (1998)
Urotricha furcata 10 0.25 Weisse and Montagnes (1998)
Urotricha furcata 10 0.45 Weisse and Montagnes (1998)
Urotricha furcata 10 0.54 Weisse and Montagnes (1998)
Urotricha furcata 15 0.67 Weisse and Montagnes (1998)
Urotricha furcata 15 0.77 Weisse and Montagnes (1998)
Urotricha furcata 20 0.85 Weisse and Montagnes (1998)
Urotricha furcata 20 1.00 Weisse and Montagnes (1998)
Urotricha furcata 20 1.10 Weisse and Montagnes (1998)
Urotricha furcata 25 1.30 Weisse and Montagnes (1998)
Urotricha furcata 25 1.20 Weisse and Montagnes (1998)
Urotricha farcta 5 0.07 Weisse and Montagnes (1998)
Urotricha farcta 5 0.15 Weisse and Montagnes (1998)
Urotricha farcta 10 0.67 Weisse and Montagnes (1998)
Urotricha farcta 10 0.72 Weisse and Montagnes (1998)
Urotricha farcta 15 1.40 Weisse and Montagnes (1998)
Urotricha farcta 20 1.60 Weisse and Montagnes (1998)
Urotricha farcta 25 1.80 Weisse and Montagnes (1998)
Urotricha farcta 25 1.90 Weisse and Montagnes (1998)
Urotricha farcta 30 1.70 Weisse and Montagnes (1998)
Urotricha farcta 30 1.80 Weisse and Montagnes (1998)
Urotricha farcta 10 0.39 Weisse and Montagnes (1998)
Urotricha farcta 10 0.45 Weisse and Montagnes (1998)
Urotricha farcta 15 0.89 Weisse and Montagnes (1998)
Urotricha farcta 20 1.00 Weisse and Montagnes (1998)
248
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Urotricha farcta 25 1.10 Weisse and Montagnes (1998)
Urotricha farcta 25 1.20 Weisse and Montagnes (1998)
Urotricha farcta 25 1.30 Weisse and Montagnes (1998)
Urotricha farcta 12 1.20 Weisse et al (2001)
Urotricha farcta 15 1.50 Weisse et al (2001)
Urotricha farcta 15 1.40 Weisse et al (2001)
Urotricha farcta 18 1.80 Weisse et al (2001)
Urotricha farcta 20 1.80 Weisse et al (2001)
Urotricha farcta 20 1.60 Weisse et al (2001)
Urotricha farcta 25 1.80 Weisse et al (2001)
Urotricha farcta 30 1.70 Weisse et al (2001)
Urotricha farcta 20 2.70 Weisse et al (2001)
Urotricha farcta 24 3.00 Weisse et al (2001)
Urotricha castalia 8 0.21 Weisse et al (2001)
Urotricha castalia 12 0.65 Weisse et al (2001)
Urotricha castalia 10 0.50 Weisse et al (2001)
Urotricha castalia 25 0.48 Weisse et al (2001)
Urotricha castalia 22 0.42 Weisse et al (2001)
Urotricha castalia 15 0.55 Weisse et al (2001)
Urotricha castalia 18 0.52 Weisse et al (2001)
Urotricha castalia 20 0.50 Weisse et al (2001)
Urotricha furcata 10 0.22 Weisse et al (2001)
Urotricha furcata 15 0.70 Weisse et al (2001)
Urotricha furcata 20 1.01 Weisse et al (2001)
Urotricha furcata 25 0.94 Weisse et al (2001)
Urotricha furcata 28 0.24 Weisse et al (2001)
Urotricha furcata 5 0.21 Weisse et al (2001)
Urotricha furcata 10 0.18 Weisse et al (2001)
Urotricha furcata 15 0.23 Weisse et al (2001)
Urotricha furcata 20 0.34 Weisse et al (2001)
Urotricha furcata 25 0.52 Weisse et al (2001)
Balanion planctonicum 6 0.30 Weisse et al (2001)
Balanion planctonicum 7.5 0.60 Weisse et al (2001)
Balanion planctonicum 9 0.90 Weisse et al (2001)
Balanion planctonicum 12 1.00 Weisse et al (2001)
Balanion planctonicum 15 1.40 Weisse et al (2001)
Balanion planctonicum 18 1.60 Weisse et al (2001)
Balanion planctonicum 19.5 1.40 Weisse et al (2001)
Balanion planctonicum 21 0.77 Weisse et al (2001)
Balanion planctonicum 5.5 0.50 Weisse et al (2001)
Balanion planctonicum 9 0.84 Weisse et al (2001)
Balanion planctonicum 12 0.95 Weisse et al (2001)
Balanion planctonicum 15.5 1.30 Weisse et al (2001)
Balanion planctonicum 18.5 1.50 Weisse et al (2001)
Balanion planctonicum 15 1.70 Weisse et al (2001)
Balanion planctonicum 15 0.70 Weisse et al (2001)
249
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Balanion planctonicum 15 0.92 Weisse et al (2001)
Balanion planctonicum 15 0.75 Weisse et al (2001)
Balanion planctonicum 15 0.75 Weisse et al (2001)
Balanion planctonicum 15 0.81 Weisse et al (2001)
Balanion planctonicum 15 0.86 Weisse et al (2001)
Balanion planctonicum 15 0.97 Weisse et al (2001)
Balanion planctonicum 15 1.10 Weisse et al (2001)
Balanion planctonicum 15 1.20 Weisse et al (2001)
Balanion planctonicum 15 1.40 Weisse et al (2001)
Balanion planctonicum 15 1.10 Weisse et al (2001)
Balanion planctonicum 15 0.70 Weisse et al (2001)
Balanion planctonicum 15 0.85 Weisse et al (2001)
Balanion planctonicum 15 0.89 Weisse et al (2001)
Balanion planctonicum 15 0.94 Weisse et al (2001)
Balanion planctonicum 15 1.50 Weisse et al (2001)
Balanion planctonicum 15 1.40 Weisse et al (2001)
Balanion planctonicum 15 1.40 Weisse et al (2001)
Urotricha furcata 15 0.42 Weisse et al (2001)
Urotricha furcata 15 0.48 Weisse et al (2001)
Urotricha furcata 15 0.50 Weisse et al (2001)
Urotricha furcata 15 0.67 Weisse et al (2001)
Urotricha furcata 15 0.54 Weisse et al (2001)
Urotricha furcata 15 0.59 Weisse et al (2001)
Urotricha furcata 15 0.60 Weisse et al (2001)
Urotricha furcata 15 0.68 Weisse et al (2001)
Urotricha furcata 15 0.65 Weisse et al (2001)
Urotricha furcata 15 0.64 Weisse et al (2001)
Urotricha furcata 15 0.67 Weisse et al (2001)
Urotricha furcata 15 0.68 Weisse et al (2001)
Urotricha furcata 15 0.79 Weisse et al (2001)
Urotricha farcta 15 0.78 Weisse et al (2001)
Urotricha farcta 15 0.80 Weisse et al (2001)
Urotricha farcta 15 0.81 Weisse et al (2001)
Urotricha farcta 15 0.85 Weisse et al (2001)
Urotricha farcta 15 0.95 Weisse et al (2001)
Urotricha farcta 15 0.90 Weisse et al (2001)
Urotricha farcta 15 0.91 Weisse et al (2001)
Urotricha farcta 15 0.91 Weisse et al (2001)
Urotricha farcta 15 1.00 Weisse et al (2001)
Urotricha farcta 15 1.10 Weisse et al (2001)
Urotricha farcta 15 1.10 Weisse et al (2001)
Urotricha farcta 15 1.10 Weisse et al (2001)
Urotricha farcta 15 1.20 Weisse et al (2001)
Urotricha farcta 15 1.20 Weisse et al (2001)
Urotricha farcta 15 1.40 Weisse et al (2001)
Urotricha farcta 15 1.50 Weisse et al (2001)
250
Appendix 3 (cont’d): Growth Rates of Heterotrophic Protists Within
Chapter 2 Data Set
Urotricha farcta 15 1.30 Weisse et al (2001)
Urotricha farcta 15 1.00 Weisse et al (2001)
Urotricha farcta 15 1.10 Weisse et al (2001)
Urotricha farcta 15 1.50 Weisse et al (2001)
Urotricha farcta 15 0.97 Weisse et al (2001)
Urotricha farcta 15 1.30 Weisse et al (2001)
Urotricha farcta 15 1.30 Weisse et al (2001)
Urotricha farcta 15 1.30 Weisse et al (2001)
Urotricha farcta 15 1.60 Weisse et al (2001)
Urotricha farcta 15 1.20 Weisse et al (2001)
Urotricha farcta 9 0.79 Weisse et al (2002)
Urotricha farcta 12 1.20 Weisse et al (2002)
Urotricha farcta 15 1.30 Weisse et al (2002)
Urotricha farcta 18 1.80 Weisse et al (2002)
Urotricha farcta 21 3.90 Weisse et al (2002)
Urotricha farcta 24 3.70 Weisse et al (2002)
Bodo saliens 10 0.34 Zubkov and Sleigh (2000)
Caecitellus parvulus 10 0.81 Zubkov and Sleigh (2000)
Cafeteria roenbergensis 10 0.54 Zubkov and Sleigh (2000)
Pteridomonas danica 10 1.29 Zubkov and Sleigh (2000)
Uronema marinum 10 2.36 Zubkov and Sleigh (2000)

251
-5
-4
-3
-2
-1
0
1
2
-5
-4
-3
-2
-1
0
1
2
-5
-4
-3
-2
-1
0
1
2
Slope = .06
r
2
= .076
Slope = .08
r
2
= .44
-5
-4
-3
-2
-1
0
1
2
Slope = .13
r
2
= .86
Temperature (
o
C)
A
B
C
D
-5
-4
-3
-2
-1
0
1
2
0 5 10 15 20 25 30
-5
-4
-3
-2
-1
0
1
2
0 5 10 15 20 25 30
Slope = 0.11
r
2
= 0.78
Slope = 0.12
r
2
= 0.80
E
F
Appendix 4. Sample generation of upper envelope of growth rates for herbivorous protists (A-D). Total
collected growth rates for herbivorous protists (A); maximal two growth rates collected at each temperature
over the entire temperature range (B); maximal growth rates after outlier removal (C); upper envelope
generated in (C) plotted against entire data set (D). Upper envelope generated in (C) with growth rates at
extreme low temperature removed (E). Upper envelope generated in (C) with growth rates at extreme high
temperature removed (F). Solid lines indicated least squares regression.
Appendix 4. Generation of upper envelope of growth rates
for herbivorous protists
252

Abstract (if available)

Abstract The Ross Sea, Antarctica experiences one of the world's largest annual phytoplankton blooms at exceptionally low environmental temperature (-1.5-0.5oC). Chlorophyll concentrations during the bloom can exceed 15 mg l-1. Heterotrophic protists seasonally dominate biomass within Antarctic marine ecosystems, and function as important players in nutrient remineralization and carbon flow. However, these potential algal grazers do not prevent the formation of phytoplankton blooms in this ecosystem. Low temperature represents a constant potential limit on maximal growth of all Antarctic species, yet the effect of temperature on growth of heterotrophic protists is not well characterized. Existing data for growth rates and growth efficiencies of polar protists are conflicting. I investigated top-down control of algal and bacterial standing stocks at ambient Ross Sea temperatures through a literature review of temperature effects on protistan growth rate, laboratory experiments with cultured Antarctic protists and field estimates of grazing rates by assemblages of Antarctic herbivorous protists. Growth rates of phototrophs and heterotrophs at ambient Antarctic temperatures were universally low, but growth rates of heterotrophs appeared to be more strongly affected at low temperature than phototrophs. Maximum growth rates of herbivores were equivalent to algae at 20oC, but a quarter of maximum algal growth rates at 5oC. Growth and nutrient remineralization rates of Antarctic bacterivorous protists were low at 0oC and increased sharply with increasing temperature. Growth efficiency and nutrient remineralization efficiency by Antarctic bacterivorous protists were comparable to temperate conspecifics at all temperatures examined. Specific ingestion rates of Antarctic herbivorous protists in culture and in field assemblages were extremely low when compared with temperate congeners.

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Asset Metadata

Creator Rose, Julie Marie (author)

Core Title Physiological and ecological consequences of environmental temperature on Antarctic protists

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Biology

Degree Conferral Date 2007-05

Publication Date 04/06/2007

Defense Date 03/31/2006

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag Antarctic protists,growth efficiencies,growth rates,heterotrophic protists,microzooplankton grazing,OAI-PMH Harvest,temperature

Language English

Advisor Caron, David A. (committee chair), Bakus, Gerald Joseph (committee member), Fuhrman, Jed Alan (committee member), Kiefer, Dale A. (committee member), N[illegible], K[illegible] (committee member), Sullivan, Cornelius W. (committee member)

Creator Email jrose@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m356

Unique identifier UC1186370

Identifier etd-Rose-20070406 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-402667 (legacy record id),usctheses-m356 (legacy record id)

Legacy Identifier etd-Rose-20070406.pdf

Dmrecord 402667

Document Type Dissertation

Rights Rose, Julie Marie

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu