Mar Biol
DOI 10.1007/s00227-011-1640-8
METHOD
Determining the viability of marine protists using a combination
of vital, fluorescent stains
Mia K. Steinberg • Edward J. Lemieux
Lisa A. Drake
•
Received: 14 September 2010 / Accepted: 4 February 2011
Ó Springer-Verlag (outside the USA) 2011
Abstract Determining the viability of protists and small
microzooplankton has long been a focus of studies in
marine biology and ecology. It is especially relevant in the
issue of shipborne invasive species, and impending international guidelines and various national regulations on the
allowable concentrations of organisms in discharged ballast water have spurred the growth of an industry that
develops and manufactures ballast water management
systems. The success of management systems and ability of
ships to meet ballast water discharge standards is determined by the number of viable organisms in treated water.
Here, we propose combining two vital, fluorescent stains
(fluorescein diacetate [FDA] and 5-chloromethylfluorescein diacetate [CMFDA]) with direct microscopic observation to enumerate viable organisms C10 and \50 lm in
minimum dimension (nominally protists). This approach
was validated in four locations in the United States to
determine the efficacy of the stains. Although the accuracy
of the stains varied by geographic location and the taxonomic composition of the planktonic assemblage, combining fluorescent stains is a robust, powerful tool that can
be optimized for the species present at each location. While
this method was developed for analyzing viable organisms
in treated ballast water, it may also be used or adapted for
Communicated by U.-G. Berninger.
M. K. Steinberg E. J. Lemieux
Naval Research Laboratory, Code 6130, Chemistry Division,
4555 Overlook Ave SW, Washington, DC 20375, USA
L. A. Drake (&)
Naval Research Laboratory, Code 6136, Chemistry Division,
Trumbo Point Annex, F-14, Key West, FL 33040, USA
e-mail: [email protected]
any field of research that examines a broad taxonomic
range of autotrophic and heterotrophic plankton.
Introduction
The need to accurately identify and enumerate microplankton has been addressed in numerous contexts: harmful
algal bloom investigations (Tester and Steidinger 1997;
Stumpf et al. 2003; Godhe et al. 2007), growth and toxicity
studies (Munawar and Munawar 1987; Lage et al. 2001;
Araújo et al. 2008), and diversity surveys (Bralewska and
Witek 1995; Anderson 1997; Savin et al. 2004). In a subset
of these studies, viability was also important (e.g., Jochem
1999; Kobiyama et al. 2010). Despite the effort in this
research area, no single method exists to determine reliably
the viability of microplankton in mixed assemblages.
A relatively new area where determining microplankton
viability is important is the transport of potential nuisance
species in ships’ ballast water. While ballast water-mediated introductions of aquatic nuisance species are not a new
subject (e.g., Carlton 1985; Williams et al. 1988; Carlton
and Geller 1993), recent national and international guidelines for controlling introductions have given rise to a new
industry of ballast water management systems that aim to
comply with these guidelines. On a national scale, the US
Coast Guard (USCG) is in the process of creating discharge
standards for vessels entering US waters (Federal Register
2009). In 2004, the international shipping community
(though the International Maritime Organization) adopted a
convention for managing ships’ ballast (IMO 2004). After
this convention is ratified and enters into force, up to 1
billion USD per year will be spent retrofitting existing
vessels with ballast water management systems (Tjallingii
2001). Once existing vessels are updated, the ballast water
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Mar Biol
industry could generate up to 327 million USD per year as
new constructions are outfitted (Tjallingii 2001).
The validation of a successful ballast water management
system hinges on the ability to detect organisms at the
specified discharge densities, which the IMO convention
and proposed US rule define as less than 10 viable
organisms C50 lm in minimum dimension per m3 of discharged water and less than 10 viable organisms C10 and
\50 lm in minimum dimension per ml of discharged
water (IMO 2004; Federal Register 2009; the discharge
standard for bacteria will not be considered here). The
Guidelines for Approval of Ballast Water Management
Systems define ‘‘viable’’ as ‘‘living,’’ which is the definition that will be used here (IMO 2005). For the purpose of
validating ballast water management systems and testing
water discharged from ships with low densities of viable
organisms, standard oceanographic methods must be
modified.
Organisms C50 lm are nominally zooplankton, whose
numbers and viability are often determined by direct counts
and mobility (Herwig et al. 2006). In contrast, the live/dead
status of organisms C10 and \50 lm (nominally protists),
which are often non-motile on scales in which they are
observed in the laboratory, is a question that has challenged
microbiologists for decades (e.g., Throndsen 1978; Gallagher 1984; Brussaard et al. 2001). Furthermore, coupling
the question of protist viability with ballast water technology testing adds additional constraints: analyses must
be relatively affordable because they should not inordinately inflate the cost of technology testing, they should be
uncomplicated and able to be performed at test facilities
around the world, they should produce results in a short
time frame (on a scale of hours, not days or weeks), and,
importantly, they must be applicable to the broad assemblage of organisms present in ballast water samples rather
than a monoculture grown in the laboratory.
One common method used for analyzing phytoplankton
abundance in marine and fresh waters is extracting and
measuring chlorophyll as an index of biomass (Yentsch and
Menzel 1963). Although used by a number of investigators
evaluating the efficacy of ballast water management systems, this technique has several disadvantages in this
application. First, chlorophyll molecules may remain intact
within a dead cell for up to 2 weeks (Veldhuis et al. 2001;
Garvey et al. 2007). Second, it is difficult—likely impossible—to estimate accurately the number of living cells per
volume of water from a biomass index, especially when the
species of cells and physiological states are unknown. Last,
this method does not address heterotrophic protists present
in this size class, which would lack chlorophyll.
Another method for determining the viability of protists
is the most probable number (MPN) technique based on
serial dilutions (Throndsen 1978); however, this approach
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requires incubations ranging from 24 h to months and will
only account for culturable species (Oemcke and van
Leeuwen 2005). Other techniques, such as ATP assays and
the uptake of radiolabeled substrates, rely on indices and
assumptions about the metabolism of organisms present in
the sample (Hunter and Laws 1981; Waite et al. 2003).
These techniques may be useful for preliminary tests to
indicate whether a management system grossly exceeds
IMO discharge standards, but they are unable to quantify
the number of viable organisms in this size class. Most
molecular techniques, such as quantitative PCR or RNA
hybridization, can analyze one or two target species
(Godhe et al. 2007) and are inadequate for assessing entire
plankton assemblages. Using direct cell counts, one can
accurately determine the density of organisms; however,
viability must be determined by another method, such as
using vital or mortal stains.
Regarding discharged ballast water that has undergone
treatment, it seems prudent to use a vital stain to enumerate
the small number of viable organisms. The alternative is to
use a mortal stain to enumerate the potentially large
number of dead organisms and a general nucleic acid stain
to enumerate all organisms, in which case the number of
live organisms is determined by subtraction. Membranepermeable vital stains, such as fluorescein diacetate
(FDA) and Calcein AM, have been used by researchers for
decades. Once inside a cell, non-specific esterases cleave
the molecules of stain to create a non-permeable, green
fluorescent product. The many studies employing FDA as
a measure of enzyme activity and viability span a wide
taxonomic range, including bacteria, diatoms, dinoflagellates, chlorophytes, prasinophytes, and prymnesiophytes; however, not all organisms stain with FDA, and
the fluorescent signal is often inconsistent (Dorsey et al.
1989; summarized in Garvey et al. 2007). To compensate
for these shortcomings, we investigated the addition of
other vital stains with the same fluorescence spectra as
FDA.
A newer vital stain, 5-chloromethylfluorescein diacetate
(CMFDA or CellTrackerTM Green [Invitrogen]), has been
used for multiple purposes including assessing viability and
labeling algae for feeding experiments (Li et al. 1996;
Murphy and Cowles 1997). CMFDA is similar to FDA in
that intracellular enzymes create a non-membrane permeable, fluorescent product; however, CMFDA is mildly thiol
reactive and has better cellular retention (Poot et al. 1991).
Since both FDA and CMFDA have similar emission
spectra, they can be combined and used simultaneously.
The goal of the research presented here was to develop
and validate a protocol to assess protist viability by combining two stains with the same fluorescence spectra. There
is need in the ballast water industry and in marine biology
and ecology as a whole for a robust viability analysis that is
Mar Biol
applicable to a broad range of organisms. To evaluate the
combination of FDA and CMFDA, samples from several
locations with different water characteristics and ambient
plankton communities were tested.
Materials and methods
Preliminary trials
In preliminary tests in our laboratory, we used FDA and
CMFDA to stain natural assemblages of protists collected
from Key West, FL as well as cultures of protists purchased
from commercial vendors. Cultures maintained in the
laboratory included the motile green flagellate Tetraselmis
sp. (Reed Mariculture; Campbell, California; strain PLY
429), the motile pennate diatoms Amphora coffeaeformis
(Algagen LLC; Vero Beach, FL) and Cylindrotheca closterium (Provasoli-Guillard National Center for Culture of
Marine Phytoplankton; West Boothbay Harbor, Maine;
strain CCMP340), the centric diatom Melosira octogona
(strain CCMP483), and the motile dinoflagellate Prorocentrum hoffmannianum (strain CCMP2804). Various
combinations of stain concentration (1–50 lM) and staining time (10–50 min) were used. Likewise, Calcein AM
and CFDA (other membrane-permeable vital stains) were
also tested.
Site locations and sample collection
Ten samples of ambient water were collected at the Naval
Research Laboratory in Key West, Florida (FL; 5–27 May
2009), and four samples were collected at the Maritime
Environmental Resource Center in Baltimore, Maryland
(MD; 2 July 2009–6 August 2009). Three samples were
collected at each the Pacific Northwest National Laboratory’s Marine Science Laboratory in Sequim, Washington
(WA; 14–16 September 2009) and the Bigelow Laboratory
for Ocean Sciences in West Boothbay Harbor, Maine (ME;
30 September–2 October 2009).
Water samples were taken from an unfiltered flowing
seawater system (FL) or from surface water collected using
buckets (MD, WA, ME). To collect a diverse and concentrated sample of protists, water was first filtered through
a 50-lm nylon mesh sieve to remove macrozooplankton
and then immediately through a 10-lm nylon mesh filter
bag. Mesh pore sizes were chosen to reflect the IMO size
class designations. The organisms concentrated in the
10-lm filter bag were analyzed at an onsite laboratory in
FL, WA, and ME, while samples from MD were transported in coolers to the Naval Research Laboratory’s main
campus in Washington, DC and analyzed within 4 h of
collection.
Sample staining and analysis
One milliliter subsamples from the concentrated sample
were placed in microfuge tubes and stained with 5 ll of 1
mM FDA and 10 ll of 250 lM CMFDA for final concentrations of 5 and 2.5 lM, respectively. Stained subsamples were incubated in the dark at room temperature for
10 min and then loaded onto 1-ml Sedgewick-Rafter
counting chambers etched with 1-mm2 grids. Chambers
were examined at 1009 magnification using compound
epifluorescent microscopes with standard blue light excitation—green bandpass emission filter cubes (e.g., exciter
BP 480/40; dichroic FT 505; barrier BP 530/30). Even
though fluorescein is polar and does not pass quickly
through cellular membranes, there is still some leakage
from cells that contributes to background fluorescence over
time (Jochem 1999). To maintain consistency among sites
and samples, chambers were only analyzed for 20 min after
incubation regardless of whether or not background fluorescence was seen at that time.
For each subsample, one row of the Sedgewick-Rafter
counting chamber was randomly selected, and every
organism in the row was scored as moving or non-moving
and fluorescing or non-fluorescing. Preliminary experiments with ambient plankton in FL showed that species
that fluoresced faintly when stained had absolutely no
fluorescence when heat-treated (described below); so for
these trials, any cell with fluorescence visible through the
oculars was considered living. Every field of view was
evaluated in brightfield illumination and under epifluorescence, and counting continued until either the row was
finished or the 20-minute time limit had been met.
For each sample, three un-manipulated subsamples
(considered ‘‘live’’) and three heat-treated subsamples
(considered ‘‘dead’’) were analyzed. For heat-treated
subsamples, a 1 ml sample was added to a microfuge tube,
placed in a 50°C water bath for 10 min, and then allowed
to cool to room temperature before staining (Table 1). In
this manner, the heat-treated subsamples served as negative
controls to the un-manipulated (‘‘live’’) samples. To compare methods of killing, subsamples from WA and ME
were cold-treated (also considered ‘‘dead’’) by placing
the subsample microfuge tubes in a -20°C freezer for
1 h and then allowing them to thaw to room temperature
before staining. Previous work shows the protocols used to
heat kill and freeze organisms is sufficient to kill all
organisms in the samples (Dressel et al. 1972; Seepersad
et al. 2004).
The organisms in the live and dead subsamples were
analyzed for the presence or absence of (1) movement and
(2) fluorescence, and the percentage of organisms in each
of the following categories were determined: moving/
fluorescing (M?/F?), moving/non-fluorescing (M?/F-),
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Table 1 Staining and treatments for plankton assemblages at four locations
Sampling locations
No treatment
‘‘live’’
Heat-treated
‘‘dead’’
Cold-treated
‘‘dead’’
Stains No stains* Stains No stains* Stains No stains*
FL
Naval Research Laboratory, Key West, Florida
X
MD
Maritime Environmental Resource Center, Baltimore, Maryland
X
WA Pacific Northwest National Laboratories, Sequim, Washington
X
ME Bigelow Laboratory for Ocean Sciences, West Boothbay Harbor, Maine X
X
X
X
X
X
X
X
X
X
X
* Any green fluorescence seen in unstained subsamples was attributed to natural green autofluorescence of the organisms
The preliminary investigations with natural communities
from Key West as well as cultures of protists showed
CMFDA did not stain as many species as FDA, but
CMFDA did stain some of the species that FDA stained
poorly or not at all, such as the pennate diatom Cylindrotheca closterium. In addition, FDA and CMFDA
products were often found in different regions of the cells
(Fig. 1). Calcein AM and CFDA showed no advantage
over FDA or CMFDA (data not shown).
The vital stains FDA and CMFDA were tested at four
locations using live and heat-treated samples. In the live,
stained samples, the percentage of organisms that fluoresced
green (F?) ranged from 74% in WA to 90% in ME (Fig. 2;
sum of M?/F? and M-/F? results). The FL and MD samples
had 2 and 1%, respectively, of the total organisms in the M?/
F- (false negative) category while WA and ME had none.
In the heat-treated, stained samples, none of the organisms at any location were moving; however, the number of
fluorescing organisms varied by site. Since all organisms in
the heat-treated samples were presumed dead, any organism that still fluoresced green was considered a false
positive. In FL and MD, the combination of stains worked
very well; the percentage of false positives was relatively
low at 5 and 3%, respectively (Fig. 3). The other two
locations had higher rates of false positives, where 36% of
the heat-treated samples in WA still fluoresced, as did 19%
in ME. In most cases, these false positives had very faint
green fluorescence; however, the signal was still identifiable from the background noise and was thus scored
as fluorescing to maintain consistency between sites.
Additionally, the false positives tended to be heterotrophic
or mixotrophic dinoflagellates such as Protoperidinium
bipes, Scrippsiella trochoidea, Prorocentrum micans, and
Dinophysis spp. Some spherical and elliptical cells that
Fig. 1 A laboratory culture of the diatom Amphora coffeaeformis
stained with FDA and CMFDA. a brightfield; b FDA only; c CMFDA
only. Each image is a different field of view of cells taken from the
same homogenous culture. Cells stained with FDA fluoresced mainly
in the center while cells stained with CMFDA fluoresced at either
ends of the cell
non-moving/fluorescing (M-/F?), and non-moving/nonfluorescing (M-/F-). Subsamples’ data were averaged to
create percentages for each day’s sample, and samples
were averaged for a total site percentage.
M?/F?, M?/F-, and M-/F? organisms were considered living by virtue of movement or staining or both,
while M-/F- organisms were considered dead. M?/Forganisms represented false negatives since they would not
be detected during an analysis using only the stains. All
fluorescing organisms in the dead subsamples represented
false positives.
To address the potentially confounding issue of naturally occurring green autofluorescence (GAF) in ambient
plankton assemblages (Elbrächter 1994; Tang and Dobbs
2007), one sample each from WA and ME was analyzed as
described above but without the addition of stains.
Results
Preliminary trials
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Fig. 2 Untreated ‘‘live,’’ stained ambient samples. Organisms at each
location were divided into four categories based on: M? Moving, MNot Moving, F? Fluorescing, F- Not Fluorescing. Error bars
represent one standard deviation. FL Florida, MD Maryland, WA
Washington, ME Maine
To test if the method of killing the sample affected green
fluorescence, three subsamples from WA were cold-treated
alongside the three heat-treated subsamples, and none of
the subsamples were stained. There was no significant
difference between the percentage of fluorescing organisms
in the heat-treated subsamples and the cold-treated subsamples (means of 6 and 11%, respectively; t test, t4 = 2.78,
P = 0.407), which suggests that the compound(s) responsible for GAF was not affected by temperature. A sample from
ME was cold-treated as well, and subsamples were stained
with FDA and CMFDA since it had been observed that there
was no GAF in the heat-treated subsamples. There was no
significant difference between the percentage of fluorescing
organisms in the heat-treated and cold-treated subsamples
(means of 7 and 9%, respectively; t test, t4 = 2.78,
P = 0.547).
Discussion
Fig. 3 Heat-treated ‘‘dead,’’ stained ambient samples. Organisms at
each location were divided into four categories based on: M?
Moving, M- Not Moving, F? Fluorescing, F- Not Fluorescing.
Error bars represent one standard deviation. FL Florida, MD
Maryland, WA Washington, ME Maine
fluoresced after heat-treatment may have been dinoflagellate cysts.
To evaluate green autofluorescence (GAF), one heattreated sample from WA was analyzed without stain, and
despite a lack of a fluorescent probe, 11% of the organisms
had green fluorescence, which was attributed to GAF (data
not shown). The stained, heat-treated subsamples from the
same day showed 45% of the organisms fluorescing, so
GAF could have accounted for up to one quarter of the
‘‘dead’’ organisms that were still fluorescing green. Similarly, one heat-treated sample from ME was analyzed
without stain, but no organisms had GAF. In this instance,
all of the green fluorescence in the ‘‘dead’’ cells could be
attributed to the stains. Samples from FL and MD were not
examined without stain since the percentages of fluorescing
organisms in the heat-treated samples were relatively low
(5 and 3%, respectively).
The combination of the fluorescent stains fluorescein
diacetate (FDA) and 5-chloromethylfluorescein diacetate
(CMFDA) was tested as a method for determining the
viability of organisms C10 and \50 lm (nominally protists) in minimum dimension, which is a size class specified
by the International Maritime Organization and also proposed for use by the USCG. Organisms were classified as
moving or not moving and fluorescing or not fluorescing.
Organisms that were moving but not fluorescing green
were considered false negatives since an assay only using
green fluorescence would incorrectly classify them as nonviable. ‘‘Dead’’ organisms that fluoresced green in heattreated or cold-treated subsamples were considered false
positives since an assay only using green fluorescence
would incorrectly classify them as viable.
At all sites, the false negative error rates were low,
0–2%, and considering the taxonomically diverse populations, we felt they were negligible. Regarding false positive
errors, trials in Baltimore, MD and Key West, FL yielded
low error rates (3 and 5%, respectively), but samples from
West Boothbay Harbor, ME and Sequim, WA had a high
incidence of false positives (19 and 36%, respectively).
The differences in false positives between sites are likely
because the WA and ME samples had a much higher
abundance and diversity of dinoflagellates than did the
samples from KW and MD. Most of the organisms in the
heat- and cold-treated samples from WA and ME had a
faint fluorescent signal; however, to remain consistent
among sites, any visible fluorescence was considered F?.
By choosing an appropriate threshold to discriminate
between a bright, live signal and a dim, dead signal, a
trained observer would likely be able to reduce the number
of false positives when using this method.
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One potential complication with using stains with green
wavelength emissions is overlap with green autofluorescence (GAF), which has been reported in many marine
taxa, including dinoflagellates, diatoms, and ciliates, and
could potentially be mistaken for stain fluorescence (LavalPeuto and Rassoulzadegan 1988; Carpenter et al. 1991;
Tang and Dobbs 2007). Researchers using flow cytometry
have been able to differentiate between GAF and SYTOX
Green or FDA signals in monocultured samples (Dorsey
et al. 1989; Lawrence et al. 2006), but ambient plankton
assemblages have much more variable GAF due to the
wide range of taxonomic groups, cell size, and shape (Tang
and Dobbs 2007). Of the four sites tested in this study, only
Washington had organisms with GAF after heat or cold
treatment. It is possible, however, that some organisms in
the WA samples had a broader thermal tolerance than
organisms found elsewhere, and that fluorescing cells in the
heat- and cold-treated samples were not well and truly
dead.
Low cellular metabolic activity in a cell (e.g., sequestered in a ballast tank) may prevent the vital stains from
becoming hydrolyzed, in which case, the fluorescent signal
would be weak or non-existent (Dorsey et al. 1989;
Brussaard et al. 2001). As a secondary measure of viability,
membrane-impermeable mortal stains such as SYTOXÒ or
cyanine nucleic acid stains (e.g., POPO-1 or BOBO-3)
could be used to test membrane integrity. Because some
studies showed differences in FDA staining within a single
species (Onji et al. 2000; Garvey et al. 2007), using a
mortal stain may ameliorate vital staining variation within
a species. If the mortal stain has different fluorescence
spectra from FDA and CMFDA, all three stains could be
used simultaneously.
Depending on the plankton assemblage, other green
fluorescent vital stains such as Calcein AM or CFDA may
be preferred over FDA or CMFDA, or only one stain may
be needed. FDA alone has been shown to be sufficient for
assessing viability of protists collected in Lake Superior
near Superior, WI (Reavie et al. 2010). Due to strong
chlorophyll autofluorescence in the orange and red wavelengths, only green and blue fluorochromes should be used.
In sum, the combination of FDA and CMFDA stains
proved a useful tool for enumerating viable protists in
Florida and Maryland but was less successful in Washington and Maine, owing largely to false positives from
heterotrophic and mixotrophic dinoflagellates. Using a
carefully selected threshold for separating the fluorescence
of live and dead organisms, the amount of error when using
this method would decrease. Despite these few limitations,
fluorescent vital stains are useful tools for evaluating the
efficacy of ballast water management systems against a
discharge standard and for any areas of research that
examine viability in diverse plankton assemblages. These
123
stains also help visualize and identify organisms in complex samples where debris and other flocculent may
obscure detection. To date, we know of no other viability
and enumeration method that is as robust as this procedure
for analyzing diverse plankton assemblages (including
heterotrophs) at low densities. Considering the variation in
false positive errors between sample sites, stains must be
validated at each location before use, and we suggest the
approach outlined here be used. This approach will also be
useful as a benchmark when validating new viability assays
and techniques in the future.
Acknowledgments This research was supported by the United
States Coast Guard (contract #HSCG23-09-X-MMS028) and does not
represent official USCG policy. Many thanks to Mr. Scott Riley, Ms.
Stephanie Robbins-Wamsley, and Dr. Matthew First at the Naval
Research Laboratory in Key West, Florida and to Dr. Richard Everett
for providing feedback that greatly improved this manuscript. Additional thanks to Dr. Mario Tamburri, Mr. Timothy Mullady, Mr.
George Smith, and Ms. Janet Barnes from the Maritime Environmental Resource Center; Dr. Andrea Copping, Dr. Dana Woodruff,
and Mr. William Pratt from the Pacific Northwest National Laboratory; and Dr. Michael Sieracki and Dr. Nicole Poulton from Bigelow
Laboratory for Ocean Sciences for their generous assistance during
this project.
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