J. Phycol. 52, 10–36 (2016)
© 2015 Phycological Society of America
DOI: 10.1111/jpy.12365
REVIEW
PHOSPHORUS PHYSIOLOGICAL ECOLOGY AND MOLECULAR MECHANISMS IN MARINE
PHYTOPLANKTON1
Senjie Lin2
Department of Marine Sciences, University of Connecticut, Groton, Connecticut 06340, USA
Richard Wayne Litaker and William G. Sunda
National Oceanic and Atmospheric Administration, National Ocean Service, Center for Coastal Fisheries and Habitat Research,
Beaufort, North Carolina 28516, USA
genetics; marine algae; nutrients; phosphate; phosphorus uptake; transporter
Phosphorus (P) is an essential nutrient for marine
phytoplankton and indeed all life forms. Current
data show that P availability is growth-limiting in
certain marine systems and can impact algal species
composition. Available P occurs in marine waters as
dissolved
inorganic
phosphate
(primarily
orthophosphate [Pi]) or as a myriad of dissolved
organic phosphorus (DOP) compounds. Despite
numerous studies on P physiology and ecology and
increasing research on genomics in marine
phytoplankton, there have been few attempts to
synthesize information from these different
disciplines. This paper is aimed to integrate the
physiological and molecular information on the
acquisition, utilization, and storage of P in marine
phytoplankton and the strategies used by these
organisms to acclimate and adapt to variations in P
availability. Where applicable, we attempt to identify
gaps in our current knowledge that warrant further
research and examine possible metabolic pathways
that might occur in phytoplankton from well-studied
bacterial models. Physical and chemical limitations
governing cellular P uptake are explored along with
physiological and molecular mechanisms to adapt
and acclimate to temporally and spatially varying P
nutrient regimes. Topics covered include cellular Pi
uptake and feedback regulation of uptake systems,
enzymatic utilization of DOP, P acquisition by
phagotrophy, P-limitation of phytoplankton growth
in oceanic and coastal waters, and the role of Plimitation in regulating cell size and toxin levels in
phytoplankton. Finally, we examine the role of P
and other nutrients in the transition of
phytoplankton communities from early succession
species (diatoms) to late succession ones (e.g.,
dinoflagellates and haptophytes).
Abbreviations: AMP, adenosine monophosphate; AP,
alkaline phosphatase; ATP, adenosine triphosphate;
DIN, dissolved inorganic nitrogen; DIP, Dissolved
inorganic phosphate; DOP, dissolved organic phosphorus; HAB, harmful algal bloom; IP3, inositol
triphosphate; NADPH, reduced nicotinamide adenine dinucleotide phosphate; Pi, orthophosphate;
RNA, ribonucleic acid; SRP, soluble reactive phosphorus
Phosphorus (P) is an essential nutrient for all
organisms (Paytan and McLaughlin 2007). It is a
central component of nucleic acids (both DNA and
RNA), and thus, plays a critical role in the storage,
replication, and transcription of genetic information. It is present in phospholipids, a key component of cellular membranes. It also plays a central
role in the production of chemical energy (adenosine triphosphate [ATP]) and of reducing equivalents (reduced nicotinamide adenine dinucleotide
[NADH] and nicotinamide adenine dinucleotide
phosphate [NADPH]) during photosynthesis and
respiration, which are required for carbon fixation
and cell metabolism (Falkowski and Raven 2007).
One of the highest P requirements is in the synthesis of proteins via ribosomal RNA (Geider and La
Roche 2002). Phosphorous also regulates the activity
and function of many proteins and metabolic processes (via phosphorylation and dephosphorylation),
and modulates signaling pathways in cells (e.g.,
through adenosine monophosphate [AMP] or inositol trisphosphate [IP3]) (Cooper 2000).
Depending on the environment, the growth of
marine phytoplankton is typically limited by one of
the major nutrients [phosphorous (P), nitrogen
(N), and silicon (Si) (e.g., for diatoms)], and/or
the micronutrient iron [Fe] (Karl 2000, Paytan and
McLaughlin 2007, Moore et al. 2013). Growth
Key index words: alkaline phosphatase; diatoms;
dinoflagellates; dissolved organic phosphorus;
1
Received 27 February 2015. Accepted 26 September 2015.
Author for correspondence: e-mail [email protected].
Editorial Responsibility: M. Wood (Associate Editor)
2
10
11
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
fully characterized at the molecular level (Young
and Ingall 2010). It can largely be divided into two
major groups of organic compounds: phosphoesters
that contain the C-O-P ester bond, and phosphonates that contain the C-P bond. In analyses of high
molecular mass compounds (1–200 nm nominal
diameter), phosphoesters accounted for ~75% and
phosphonates ~25% of DOP in ocean waters (Kolowith et al. 2001). However, more recent analyses of
both low and high molecular mass compounds
revealed that 80%–85% of the measured DOP was
comprised of phosphate esters with the remainder
consisting of phosphonates (5%–10%) and
polyphosphates (8%–13%; Young and Ingall 2010).
Because DOP is operationally defined as the difference between total P and measured Pi, both
polyphosphate esters and inorganic polyphosphate
are included operationally in the measured DOP, as
likely are two other dissolved inorganic P (DIP) species: phosphite (PO33) and phosphine (PH3).
Although these latter two species have not yet been
chemically identified in seawater, their presence is
suggested by the ability of the diatom Thalassiosira
pseudonana to utilize dissolved PH3 (Fu et al. 2013)
and presence of a phosphite transporter in the
dinoflagellate Symbiodinium kawagutii (Lin et al.
2015a; see Supplementary Table 33).
Measured Pi concentrations in surface ocean
waters vary by almost 10,000-fold, from as low as
0.2 nM in some surface waters of the Sargasso Sea
to 1–3 lM in upwelled water along the eastern margins of the Atlantic and Pacific (Redfield et al.
1963, Wu et al. 2000). Pi concentrations also can
vary substantially over timescales ranging from hours
to seasons or even decades (Karl 2014). Furthermore, large differences can occur in the growth
demand for P because of variations in specific
Measured Pi (nmol L-1)
0
1000
2000
3000
DOP (nmol L-1)
0
50
100
150
200
0
500
1000
1500
Depth (m)
FIG. 1. Vertical profiles of
measured Pi and DOP in the
North Pacific and North Atlantic
Oceans (redrawn from Paytan
and McLaughlin 2007 with the
permission of the American
Chemical Society).
2000
2500
3000
3500
Atlantic
Pacific
4000
Atlantic
Pacific
REVIEW
limitation by P in the ocean occurs when bioavailable P pools (orthophosphate ions [Pi = HPO42 +
PO43] and available dissolved organic P [DOP])
drop below critical threshold concentrations relative
to levels of other required nutrients. Phytoplankton
preferentially utilize Pi because it can be directly
taken up and assimilated to support algal metabolism and growth, while DOP generally requires conversion into Pi prior to its metabolic assimilation,
which is more costly energetically (Falkowski and
Raven 2007). However, when the external Pi pool is
depleted, phytoplankton growth often depends on
the ability to utilize the much more abundant DOP
by its enzymatic hydrolysis to Pi.
The Pi concentration in the environment can be
measured using standard colorimetric methods
(Strickland and Parsons 1972) or the much more
sensitive MAGIC coprecipitation method coupled to
colorimetry (Karl and Tien 1992). These methods,
however, also typically measure some reactive DOP
compounds, and may overestimate the true dissolved Pi concentration (Thomson-Bulldis and Karl
1998, Laws et al. 2011a). Because of this, the Pi
measured by these methods is often referred to as
soluble reactive phosphorus (SRP) with the knowledge that it likely includes some DOP and other
reactive P compounds, especially in low-Pi oceanic
surface waters with high DOP:Pi ratios (Fig. 1). In
this review we will use what we believe is a more
descriptive term “measured Pi” for the SRP pool.
The DOP concentration is determined operationally
by subtracting the initial measured Pi from total Pi
measured after oxidation of DOP to Pi using alkaline persulfate (Hosomi and Sudo 1986) or UV
photo-oxidation (Aminot and K
erouel 2001).
Dissolved organic P in the ocean consists of complex mixtures of compounds that have yet to be
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
logs of these high- and low-affinity transporters have
been found in marine cyanobacteria such as
Prochlorococcus (Martiny et al. 2006). Eukaryotic
equivalents of PiT have been identified and include
the Pi transporter IPT and the sodium- or sulfatedependent Pi transporter SPT (Fig. 4; Table 1). SPT
is a symporter, which simultaneously transports Pi
and sodium or sulfate across the cell membrane.
Recent research in the Lin laboratory revealed the
presence of putative homologs of IPT and SPT in
the genome of the dinoflagellate S. kawagutii (accession number SRA148697 in NCBI SRA database)
and in the transcriptome of another dinoflagellate
Prorocentrum donghaiense (GenBank no. KJ699385,
KJ699384). IPT homologs were also found in the
transcriptomes of Karlodinium veneficum and Amphidinium carterae (KM881476, KM881477). Similarly,
IPT homologs have been observed in the diatoms
Thalassiosira pseudonana and Phaeodactylum tricornutum (Bowler et al. 2008); the haptophyte Emiliania
huxleyi, the prasinophyte Ostreococcus spp., the
mamiellophytes Micromonas sp. and Batycoccus sp.
(Monier et al. 2012, Worden et al. 2009); and the
pelagophyte Aureococcus anophagefferens. It has also
been observed in unidentified eukaryotes detected
in the Global Ocean Sampling metagenomic data
set (Table 1). The IPT gene sequences from these
diverse eukaryotic algae are not strictly conserved
suggesting the homologs were derived from a common ancestor that subsequently diverged as new
groups evolved. Eukaryotic phytoplankton viruses
carrying IPT gene sequences have also been identified (Lindell et al. 2004, Monier et al. 2012) and
may provide a mechanism for host phytoplankton
to acquire novel Pi transporter genes. Recombination aided by viral transfer may therefore have contributed to the observed IPT gene diversification.
Though less likely, another possibility is that convergent evolution of different genes encoding proteins
with IPT function produced the divergent IPT
homologs.
Comparable screening of cDNA libraries has
revealed only a few eukaryotic high-affinity PsT
equivalents (Table 1). One of these is the highaffinity Pi transporter (PHO) identified in the
prasinophyte Tetraselmis chui whose transcriptional
up-regulation under P-limitation was confirmed
experimentally (Chung et al. 2003). Using the
amino acid sequence of PHO in T. chui as a query
in tBLASTn against the expressed sequence tag data
set in NCBI revealed a homologous gene in the
dinoflagellate Alexandrium minutum (GenBank accession number GW800973). S. Lin et al. (unpublished
data) also identified a PsT homolog (KJ699386)
from a transcriptome of Prorocentrum donghaiense
grown under Pi limitation. However, the function of
the encoded protein as a high-affinity Pi transporter
remains to be verified. In addition, a high-affinity
transport protein, phosphate-repressible phosphate
permease, was identified in P-limited cultures of the
REVIEW
internal P pool(s) or external Pi concentrations regulate high- and low-affinity intracellular Pi transport
systems in eukaryotic phytoplankton, however, has
yet to be determined at the molecular level.
The diel light–dark cycle and associated daily
growth cycle also affect the pattern of cellular Pi
uptake. For most species, Pi uptake rates increase
during the day and decrease at night consistent with
the higher growth demand for P during daytime
from light driven photosynthetic C-fixation
(Chisholm and Stross 1976, Rivkin and Swift 1982,
Ahn et al. 2002). Kinetic analysis indicates that
these diel changes in uptake rates are often linked
to shifts in Vmax values rather than to variations in
Ks (Chisholm and Stross 1976, Rivkin and Swift
1982). Thus, the total number of transporters on
the cell membrane likely varies over the L:D cycle
to accommodate changes in P demand linked to
diel changes in C-fixation and the cell division cycle.
In the environment, diel changes in Pi uptake rates
may also be caused by day/night differences in Pi
concentrations, particularly in highly productive systems where algal biomass and specific growth rates
are high and the cycling of Pi may be on the order
of hours (Nixon et al. 1976). Such diel changes in
Pi uptake rates, though experimentally inconvenient, should not be overlooked, and may significantly impact ecosystem function.
An important unanswered question is how fast
cells shift the relative abundance of low- and highaffinity membrane Pi transport systems in response
to variations in ambient nutrient concentrations
and growth demand for P. As Pi transport proteins
are identified in marine phytoplankton (see below),
it may be possible to address this question if antibodies specific to high- and low-affinity Pi transporters can be developed. These antibodies could
be used in conjunction with confocal light microscopy to document changes in the density of different transporters with changes in Pi concentration at
the cell surface and in intracellular P pools. Quantitative reverse transcription PCR an also be used to
measure differential expression paterns of the different P transporter genes, but only for species in
which the transporter protein abundances are regulated at the transcriptional level.
Molecular characterization of Pi transport systems. Current molecular work has begun to reveal
the identity of some of the Pi transport proteins in
marine microorganisms that are responsible for the
uptake kinetics noted above. Such uptake systems
have been best characterized in bacteria and are
often evolutionarily conserved (Pedersen et al.
2013), so we will begin our discussion with a
description of bacterial Pi transporters. As appears
to occur in most microorganisms, heterotrophic
bacteria contain both a low-affinity Pi transporter
(PiT) that functions at high Pi concentrations and a
high-affinity Pi transporter (PsT) which is up-regulated under low-Pi stress (van Veen 1997). Homo-
15
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
more dynamics than the open ocean, can become
P-limited from excess inputs of anthropogenic N
from N-rich fertilizers, municipal wastes, and NOx
from the burning of fossil fuels, which increases N:P
ratios in the receiving waters (Cloern 2001, Huang
et al. 2003, Scavia and Bricker 2006, Sylvan et al.
2006, Zhang et al. 2007). Examples include many
eutrophic estuaries such as Chesapeake Bay (Fisher
et al. 1992, 1999, Kemp et al. 2005) and Pearl River
Estuaries (Huang et al. 2003, Xu et al. 2008),
coastal regions of the Gulf of Mexico such as those
receiving N-rich nutrient inputs from the Mississippi
River (Laurent et al. 2012, Turner and Rabalais
2013), and various Chinese coastal regions (the East
China and Yellow Seas; Harrison et al. 1990, Zhang
et al. 2007, Fu et al. 2012, Fig. 2). These anthropogenic inputs are largely due to riverine or atmospheric sources, but in some coastal regions such as
Long Island Sound, USA, N can be introduced by
inputs of N-enriched ground water (Slomp and Van
Cappellen 2004).
CELLULAR PI UPTAKE AND ASSIMILATION
Fundamental constraints. The uptake of Pi by phytoplankton cells is ultimately governed by three fundamental constraints: the ambient Pi concentration;
cell size and shape (which determines the cell’s surface to volume ratio and the thickness of its diffusive boundary layer); and the density, binding
affinity, and turnover rate of the Pi transport proteins embedded in the cell’s plasma membrane. In
FIG. 2. Global ocean map indicating where low-Pi stress or P-growth limitation has been demonstrated from high alkaline phosphatase
activity (red stars), P-stress gene expression (green circles), nutrient (P, N, and N + P) addition incubations (bioassays, purple squares),
and elevated N:P ratios (>16; yellow triangles).
REVIEW
P-stress indicators, and nanomolar-measured Pi concentrations all suggest that P limits phytoplankton
growth in these regions (Fig. 2; Krom et al. 1991,
Zohary and Roberts 1998, Wu et al. 2000, Krom
et al. 2004, Thingstad et al. 2005, Bj€
orkman et al.
2012). However, recent nutrient addition experiments suggest that in the low-Pi waters of the subtropical Atlantic, phytoplankton growth is still
primarily limited by N or colimited by N and P
(Moore et al. 2008, 2013). But even when N is primarily limiting, P likely plays an important role in
controlling species composition in low-Pi oceanic
regions (Moore et al. 2008).
Nitrogen is primarily limiting in the ocean
because of iron limitation of cynaobactrial N2 fixation, an iron-dependent metabolic process that
replenishes oceanic inventories of fixed N by enzymatically reducing N2 gas to ammonium (Sohm
et al. 2011, Sunda 2012). However, in regions
receiving high inputs of iron from aeolean dust
deposition (e.g., the subtropical North Atlantic),
the elevated iron input rates fuel higher rates of N2fixation and associated C-fixation, which drive these
systems toward P-limitation (Benitez-Nelson 2000,
Wu et al. 2000, Sa~
nudo-Wilhelmy et al. 2001, Mills
et al. 2004, Dyhrman et al. 2006a, Meseck et al.
2009, Paytan and McLaughlin 2007).
Concentrations of available P and other nutrients
are generally higher in coastal waters, but can vary
widely in time and space with variations in nutrient
inputs and algal growth. Algal growth in these systems, where P concentration is typically higher and
13
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
function as an external phosphate storage pool that
could be used to support cell growth during low-P
stress (Fu et al. 2005). The fraction of cellular P
present as surface-adsorbed phosphate increases
with the external Pi concentration (Sa~
nudo-Wilhelmy et al. 2004, Fu et al. 2005), which could make
such an external storage mechanism particularly
effective. Such possibilities clearly warrant further
investigation, particularly from a biochemical and
molecular perspective.
ADAPTATION AND ACCLIMATION RESPONSES TO LOW - P
STRESS
In addition to the up-regulation of high-affinity
Pi uptake systems, phytoplankton show a variety of
other adaptation and acclimation responses to lowP stress. These include the substitution of sulfate
for phosphate in membrane lipids, the utilization
of DOP via hydrolytic enzymes, and the acquisition
of P via phagatrophic consumption of other
microorganisms (Dyhrman et al. 2007, Hartmann
et al. 2012). The specifics of these various low-Pcoping mechanisms are discussed in the following
sections.
Reducing cellular P demand. In many marine
cyanobacteria, particularly those residing in low-P
oceanic waters, some phospholipids in cell membranes are replaced by sulfonated lipids to reduce
cellular demand for P in response to low-P stress
(Van Mooy et al. 2006, 2009, Snyder et al. 2009).
Similar phospholipid-to-sulfolipid shifts have also
been found in the brown tide pelagophyte
A. anophagefferens (Wurch et al. 2011). In addition,
both A. anophagefferens and the dinoflagellate
K. mikimotoi adjust their glycolytic pathway under
low-P stress to utilize alternate enzymes that require
less P, which enhances the ability of these species to
grow in low-P environments (Lei and Lu 2011,
Wurch et al. 2011). The molecular underpinning of
the phospholipid-to-sulfolipid shift in phytoplankton
under low-P stress remains to be elucidated.
Utilization of phosphoesters via alkaline phosphatase. As Pi levels decline with algal growth, DOP
concentrations increase due to its release from the
biological community. So one of the most important
mechanisms for coping with low-Pi stress is the utilization of DOP. There is increasing evidence that
DOP is an important source of P to phytoplankton
in low-Pi regions, such as the Sargasso Sea where
DOP:Pi ratios in surface waters can exceed 100 (Wu
et al. 2000, McLaughlin et al. 2013). For example,
30% of primary production during the spring
bloom in the North Atlantic subtropical gyre was
estimated to be supported by DOP (Mather et al.
2008). And 17%–82% of the P taken up by phytoplankton in the Sargasso Sea is estimated to have
been supplied from DOP (McLaughlin et al. 2013).
However, the partitioning of P utilization between
Pi and DOP in stratified surface ocean waters with
REVIEW
expected if polyP was merely a P-storage molecule
(Dyhrman et al. 2012). Nuclear magnetic resonance
analysis verified higher polyP levels in the P-limited
T. pseudonana cells and similar elevated polyP levels
have been observed in P-limited cells of the bacterium Escherichia coli (Kornberg et al. 1999). These
increases appear to be a response to nutrient
growth limitation in general as poly-P accumulation
also occurs when the growth rate of bacteria (Kornberg et al. 1999) and diatoms (Perry 1976) are limited by N. These findings appear to agree with
recent observations in low DIP and dissolved inorganic nitrogen (DIN) waters of the Sargasso Sea,
where unexpectedly high polyP to particulate P
ratios were observed, even though the system
appeared to be P-stressed based on high particulate
AP activities and high levels of phospholipid
replacement by sulfolipids (Martin et al. 2014).
These unexpected findings may be explained by the
many other cellular functions of polyP other than P
storage: particularly nutrient stress responses, energy
storage, and storage of essential nutrient metals
such as iron, which has been observed in yeast
(Lesuisse and Labbe 1994).
Surface phosphate adsorption. In addition to its
assimilation into cellular biomolecules, phosphate
can also adsorb directly onto the surface of phytoplankton cells (Sa~
nudo-Wilhelmy et al. 2004). This
adsorbed phosphate can account for 14%–90% of
total cell P and must be removed if true cellular P
levels are to be measured (Sa~
nudo-Wilhelmy et al.
2004, Fu et al. 2005). This adsorbed phosphate can
be removed with oxalate washes which dissolve and
remove ferric (Fe[III]) oxyhydroxides and manganese (Mn[III and IV]) oxides via their reduction
to soluble ferrous ions (Fe[II]) and manganous ions
(Mn[II]) (Sa~
nudo-Wilhelmy et al. 2004). The
adsorbed phosphate apparently is associated with
the oxides which precipitated on the cell surface
(e.g., by oxidation of Mn (II) and Fe(II)), and are
known to strongly adsorb phosphate. A strong correlation between cell surface phosphate and Mn oxides (r2 = 0.81) supports the oxide adsorption
hypothesis (Sa~
nudo-Wilhelmy et al. 2004). The
adsorbed phosphate may play an important, yet
poorly defined role in intracellular P uptake, particularly in environments with fluctuating concentrations of Pi, Mn, and Fe (Fu et al. 2005). On the one
hand, the abiotic adsorption of Pi on Mn and Fe
oxides removes Pi at the cell surface that would
otherwise be available for intracellular transport.
This could be particularly problematic for larger
cells where the diffusive flux of Pi to cell surface is
already severely limited. However, algal cells possess
mechanisms to reductively dissolve Mn and Fe oxides (e.g., transmembrane reductases; Sunda 2012),
which could release adsorbed Pi into solution for
subsequent uptake by the cell. Thus, if cells are able
to utilize this or other mechanisms (e.g., phagotrophy) to take up the adsorbed phosphate, it could
19
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
logs of these high- and low-affinity transporters have
been found in marine cyanobacteria such as
Prochlorococcus (Martiny et al. 2006). Eukaryotic
equivalents of PiT have been identified and include
the Pi transporter IPT and the sodium- or sulfatedependent Pi transporter SPT (Fig. 4; Table 1). SPT
is a symporter, which simultaneously transports Pi
and sodium or sulfate across the cell membrane.
Recent research in the Lin laboratory revealed the
presence of putative homologs of IPT and SPT in
the genome of the dinoflagellate S. kawagutii (accession number SRA148697 in NCBI SRA database)
and in the transcriptome of another dinoflagellate
Prorocentrum donghaiense (GenBank no. KJ699385,
KJ699384). IPT homologs were also found in the
transcriptomes of Karlodinium veneficum and Amphidinium carterae (KM881476, KM881477). Similarly,
IPT homologs have been observed in the diatoms
Thalassiosira pseudonana and Phaeodactylum tricornutum (Bowler et al. 2008); the haptophyte Emiliania
huxleyi, the prasinophyte Ostreococcus spp., the
mamiellophytes Micromonas sp. and Batycoccus sp.
(Monier et al. 2012, Worden et al. 2009); and the
pelagophyte Aureococcus anophagefferens. It has also
been observed in unidentified eukaryotes detected
in the Global Ocean Sampling metagenomic data
set (Table 1). The IPT gene sequences from these
diverse eukaryotic algae are not strictly conserved
suggesting the homologs were derived from a common ancestor that subsequently diverged as new
groups evolved. Eukaryotic phytoplankton viruses
carrying IPT gene sequences have also been identified (Lindell et al. 2004, Monier et al. 2012) and
may provide a mechanism for host phytoplankton
to acquire novel Pi transporter genes. Recombination aided by viral transfer may therefore have contributed to the observed IPT gene diversification.
Though less likely, another possibility is that convergent evolution of different genes encoding proteins
with IPT function produced the divergent IPT
homologs.
Comparable screening of cDNA libraries has
revealed only a few eukaryotic high-affinity PsT
equivalents (Table 1). One of these is the highaffinity Pi transporter (PHO) identified in the
prasinophyte Tetraselmis chui whose transcriptional
up-regulation under P-limitation was confirmed
experimentally (Chung et al. 2003). Using the
amino acid sequence of PHO in T. chui as a query
in tBLASTn against the expressed sequence tag data
set in NCBI revealed a homologous gene in the
dinoflagellate Alexandrium minutum (GenBank accession number GW800973). S. Lin et al. (unpublished
data) also identified a PsT homolog (KJ699386)
from a transcriptome of Prorocentrum donghaiense
grown under Pi limitation. However, the function of
the encoded protein as a high-affinity Pi transporter
remains to be verified. In addition, a high-affinity
transport protein, phosphate-repressible phosphate
permease, was identified in P-limited cultures of the
REVIEW
internal P pool(s) or external Pi concentrations regulate high- and low-affinity intracellular Pi transport
systems in eukaryotic phytoplankton, however, has
yet to be determined at the molecular level.
The diel light–dark cycle and associated daily
growth cycle also affect the pattern of cellular Pi
uptake. For most species, Pi uptake rates increase
during the day and decrease at night consistent with
the higher growth demand for P during daytime
from light driven photosynthetic C-fixation
(Chisholm and Stross 1976, Rivkin and Swift 1982,
Ahn et al. 2002). Kinetic analysis indicates that
these diel changes in uptake rates are often linked
to shifts in Vmax values rather than to variations in
Ks (Chisholm and Stross 1976, Rivkin and Swift
1982). Thus, the total number of transporters on
the cell membrane likely varies over the L:D cycle
to accommodate changes in P demand linked to
diel changes in C-fixation and the cell division cycle.
In the environment, diel changes in Pi uptake rates
may also be caused by day/night differences in Pi
concentrations, particularly in highly productive systems where algal biomass and specific growth rates
are high and the cycling of Pi may be on the order
of hours (Nixon et al. 1976). Such diel changes in
Pi uptake rates, though experimentally inconvenient, should not be overlooked, and may significantly impact ecosystem function.
An important unanswered question is how fast
cells shift the relative abundance of low- and highaffinity membrane Pi transport systems in response
to variations in ambient nutrient concentrations
and growth demand for P. As Pi transport proteins
are identified in marine phytoplankton (see below),
it may be possible to address this question if antibodies specific to high- and low-affinity Pi transporters can be developed. These antibodies could
be used in conjunction with confocal light microscopy to document changes in the density of different transporters with changes in Pi concentration at
the cell surface and in intracellular P pools. Quantitative reverse transcription PCR an also be used to
measure differential expression paterns of the different P transporter genes, but only for species in
which the transporter protein abundances are regulated at the transcriptional level.
Molecular characterization of Pi transport systems. Current molecular work has begun to reveal
the identity of some of the Pi transport proteins in
marine microorganisms that are responsible for the
uptake kinetics noted above. Such uptake systems
have been best characterized in bacteria and are
often evolutionarily conserved (Pedersen et al.
2013), so we will begin our discussion with a
description of bacterial Pi transporters. As appears
to occur in most microorganisms, heterotrophic
bacteria contain both a low-affinity Pi transporter
(PiT) that functions at high Pi concentrations and a
high-affinity Pi transporter (PsT) which is up-regulated under low-Pi stress (van Veen 1997). Homo-
15
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
DIFFERENTIAL NUTRIENT ACQUISITION AND GROWTH
STRATEGIES AND SPECIES SUCCESSION
Differential P nutrient strategies may be one of
the drivers of seasonal species succession that occurs
in many phytoplankton communities. The popula-
P or C
tion growth of diatoms with high nutrient-sufficient
maximum growth rates is often favored during early
succession (in late winter/early spring or in freshly
upwelled water) when the environment is characterized by high concentrations of dissolved inorganic
nutrients (Pi, DIN, and Fe), high turbulence, low
algal biomass, and a low level of zooplankton
grazing (Margalef 1978, Sunda et al. 2006, Sunda
and Hardison 2010, Fig. 5). As the season progresses, the water column stabilizes with increased
solar heating of surface water, and inorganic nutrient pools become depleted by the initial algal
bloom which is often dominated by diatoms (Margalef 1978, Hood et al. 1990, Tiselius and Kuylenstierna 1996, Yoshimura et al. 2014). As the DIN
and Pi pools decline during the bloom there is a
progressive buildup of organic nutrients (DOP and
DON) linked to slopy grazing and excretion by zooplankton, viral and bacterial lysis of cells, and
release by phytoplankton (van der Zee and Chou
2005, Yoshimura et al. 2014). The combination of a
stable water column, higher phytoplankton biomass,
low inorganic nutrients, increased DOP and DON
levels, and increased zooplankton grazing pressure
no longer favors the population growth of diatoms
or other early succession species and sets the stage
for a population shift to late succession species such
Time
FIG. 5. Schematic for a typical diatom to dinoflagellate seasonal succession in a system whose biomass is limited by P. The
initially high Pi levels, colder temperatures, and high turbulence
levels in the early spring favor diatoms, which are adapted for
high growth rates under these conditions. The emerging diatom
bloom depletes the euphotic zone of Pi and fuels the growth of
zooplankton. Concomitantly, solar warming increases stratification of the water column, and decreases inputs of Pi and other
nutrients from nutrient rich aphotic deeper waters. Phosphorus
inputs during this time are mainly from recycling linked to zooplankton grazing and excretion, and much of that input is in the
form of DOP. The combination of DOP inputs from grazing and
Pi uptake by phytoplankton increases the DOP concentration and
greatly increases the DOP:Pi ratio. These changes (decreased Pi
in surface waters, increased DOP, water column stratification, and
increased zooplankton grazing) sets the stage for a algal community shift from diatoms to dinoflagellates, whose growth and survival are favored under these conditions due to their ability to
obtain nutrients from alternate sources (diel vertical migration to
nutrient rich deeper waters, utilization of DOP, and phagotrophy) and their ability to minimize grazing losses (e.g., linked to
large cell size and the production of toxins).
REVIEW
under P limitation should decrease P uptake rates
per unit of cell volume, and thus, should be evolutionary disadvantageous. So, in contrast to other
nutrients, there must be some factor peculiar to
phosphorus that causes cell size to increase under P
limitation of growth rate.
To examine the effects of P limitation on cell size
we must examine its effect on the cell’s growth and
division cycle. This cycle consists of four discrete
phases: the G1 phase (gap 1 or growth stage 1)
where a newly divided cell grows and increases in
size prior to cell division, the S phase during which
DNA is replicated, the G2 phase (gap 2 or growth
stage 2) where the cell continues to grow prior to
mitosis, and the M phase during which nuclear division (mitosis) occurs leading to cell division (cytokinesis). Unlike other nutrients (e.g., Fe and N), P
limitation often results in a blockage of DNA replication (the S phase; Vaulot 1995), which must precede cell division into smaller daughter cells
(Sclafani and Holzen 2007). P limitation of growth
rate in the cyanobacteria Prochlorococcus and Synechococcus causes an arrest of the cell cycle progression from G1 to S or G2 to M phases, and in the
case of P starvation (severe growth rate limitation),
an arrest in the S phase (Parpais et al. 1996, Vaulot
et al. 1996). Similarly, the few studies conducted so
far with dinoflagellates indicate an arrest of the cell
cycle in G1 phase in response to P limitation of
growth rate (Lei and Lu 2011, Zhang et al. 2014;
Li et al. 2015). This arrest is accompanied by the
up-regulation of negative regulators (e.g., fizzy/cell
division cycle 20-related protein) and down-regulation of positive regulators of the cell cycle (e.g., calcium-dependent protein kinase) (Zhang et al.
2014). With the arrest of the cell cycle, the cells
continued to grow during an elongated G1 phase,
resulting in an increase in average cell size. The
blockage of the cell cycle progression from G1 to S,
or subsequent phase transitions, is likely linked to a
need for a sufficient supply of P for successful DNA
replication and for phosphorylation of key checkpoint enzymes that regulate DNA synthesis and
nuclear and cell division in the S and M phases.
Transitions in the cell cycle, including G2 to M
stages, are strictly regulated by a cascade of CDK
phosphorylation and dephosphorylation events
(Murray and Hunt 1993). The cell enlargement in
P-limited cells further suggests that fulfilling CDK
phosphorylation or other P-associated biochemical
requirements (e.g., DNA replication) may supersede
that of a threshold in cell size in controlling the
onset of cell division.
27
17
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
TABLE 1. Protein or protein complexes involved in phosphorus acquisition in phytoplankton that have been recognized to
date.
High-affinity DIP transporters
(PsT-like)
Low-affinity DIP transporters (PiT-like)
Species
Pho4
superfamily
Sodiumdependent
phosphate
transporter
Phosphate/
sulfate
permease
This study
Bowler
et al. (2008)
Bowler
et al. (2008)
Haptophyta
Emiliania huxleyi
Pelagophyta
Aureococcus anophagefferens
Dyhrman
et al. (2006a)
Beszteri
et al. (2012)
Ostreococcus
Dyhrman
et al. (2006a)
Beszteri
et al. (2012)
Beszteri
et al. (2012)
Wurch
et al. (2011)
Aureoumbra lagnuna
Chlorophyta
Chlamydomonas reinhardtii
Micromonas
Phosphaterepressible
phosphate
permease
This study
This study
This study
This study
Phaeodactylum tricornutum
Prymnesium parvum
High-affinity
phosphate
transporter
REVIEW
Dinophyta
Alexandrium catenella
Amphidinium caterae
Karlodinium veneficum
Prorocentrum donghaiense
Bacillariophyta
Thalassiosira pseudonana
Inorganic
phosphate
transporter
Moseley
et al. (2006)
Worden
et al. (2009)
Worden
et al. (2009)
Tetraselmis chui
Cyanophyta
Prochlorococcus
Synechodoccus
Chung
et al. (2003)
Martiny
et al. (2006)
Scanlan
et al. (2009)
Trichodesmium
a sugar-phosphate exchanger (Skav202903). This
gene belongs to the major facilitator superfamily,
and is a sugar-phosphate antiporter that likely transports sugar out of the cell and phosphate into the
cell.
Pi sensing and uptake regulatory systems. Up- and
down-regulation of Pi transport sytems is mediated
in microorganisms by two-component (sensor and
response regulator) signal transduction systems
(Wanner 1996, Stock et al. 2000). One component
of the system senses Pi at the cell surface or in the
cell’s cytosol and the other regulates the expression
of various high and low-affinity Pi transport proteins
and other Pi acquisition proteins such as phosphatases (Dick et al. 2011). One such regulatory system has been well-characterized at the molecular
level in gram-negative bacteria. It consists of the
protein pair PhoB–PhoR, and is referred to as the
Pho regulon (Vershinina and Znamenskaya 2002).
In this bacterial group, pores (porins) in the outer
membrane allow Pi to diffuse into the periplasm
and bind a periplasmic receptor site on PhoR,
which is a transmembrane protein located in the
cytoplasmic membrane. As Pi in the environment
declines, concentrations in the periplasm drop,
causing Pi to dissociate from PhoR. This dissociation results in a conformational change in the protein, which causes an intracellular kinase domain of
PhoR to transfer a phosphate group from ATP onto
another cytoplasmic site on PhoR. This phosphate is
subsequently used to phosphorylate the transcription regulator PhoB (response regulator) in the
cytoplasm allowing it to bind regulatory regions of
DNA. This binding initiates transcription of the
genes involved in the synthesis of intracellular Pi
transport proteins, and in many cases, those
involved in the utilization of DOP (e.g., those coding for APs; see section below). As Pi levels increase,
the periplasmic receptor site on PhoR rebinds to Pi
and the process and resultant gene expression is
down-regulated.
Similar regulatory systems are found in cyanobacteria (which are also gram-negative bacteria; Hirani
et al. 2001), archaea (Osorio and Jerez 1996), yeasts
(Dick et al. 2011, Magbanua et al. 1997) and the
roots of land plants (Dong et al. 2013, Ticconi and
REVIEW
18
SENJIE LIN ET AL.
Abel 2004). In addition, a second tier regulatory system dependent on PtrA (potential transcriptional
regulator) was identified in the cyanobacterium
Synechococcus (Ostrowski et al. 2010). In this system,
low Pi concentration first triggers the PhoB-induced
response (a tier 1 response), leading to elevated
expression of high-affinity Pi transporters. When
this response does not sufficiently alleviate low-Pi
stress, PtrA expression is increased. The higher PtrA
protein pool then binds to the promoters of phosphatase (e.g., AP) genes, which up-regulates phosphatase synthesis and increases cellular utilization of
DOP (a tier II response).
Recently, a response regulator receiver gene
coding for a protein that is potentially part of a twocomponent P-response regulatory system was identified in the proteome of the dinoflagellate Karenia
mikimotoi (Lei and Lu 2011). Screening of genomic
databases will likely reveal the presence of similar
P-sensing/regulatory systems in many other eukaryotic phytoplankton. Determining the details of how
the Pi signal transduction pathways are regulated in
eukaryotic algae, as well as how these pathways are
coregulated by internal P-pools, will likely prove an
important line of research in the future. However,
these pathways may be substantially different in
dinoflagellates because much of the gene expression is regulated post transcriptionally rather than
by direct regulation of RNA synthesis (for review see
Lin 2011).
Phosphate assimilation. Exactly how P is assimilated
into biomolecules needed for growth, metabolism,
and cell division following cellular uptake of Pi or
DOP or P acquisition by phagotropy has not been
extensively investigated in phytoplankton. Most
DOP taken up into the cell must first be converted
to Pi and this may also be the case for acquisiton of
P by phagotrophy, which may involve a suite of largely uncharacterized phosphatases. The major
biosynthetic pathway for Pi assimilation in all cells,
including phytoplankton, is the photosynthetic
and/or respiratory production of ATP from Pi and
adenosine diphosphate (ADP) via the enzyme ATP
synthase (Fig. 4). ATP is the major energy currency
of the cell and not only supplies energy for the synthesis of various organic biomolecules (e.g., in the
Calvin–Benson cycle) but also supplies phosphate
for the synthesis of numerous phosphate-containing
end product molecules such as phospholipids,
nucleotides, polyphosphates, and phosphorolated
sugars and proteins. The assimilation of phosphate
into ATP takes place in three cellular compartments: the chloroplast, where ATP is a major product of photosynthesis; mitochondria, where it is a
major product of respiration; and in the outer cell
membrane, where it is synthesized by the light-activated proton-pump proteorhodopsin. While the
photosynthetic and respiratory ATP synthesis systems are universal in phytoplankton and indeed all
phototrophs (Falkowski and Raven 2007), the puta-
tive energy-converting proteorhodopsin system is
best documented in certain marine bacteria (Fuhrman et al. 2008 and references therein) and has
only begun to be examined in eukaryotes, including
dinoflagellates (Lin et al. 2010, Guo et al. 2014, Shi
et al. 2015), two diatoms, and a haptophyte (Marchetti et al. 2012). The algal homologs of this protein
are similar to that in proteobacteria, where the protein harvests solar energy and generates a proton
gradient across the plasma membrane for the production of ATP via the enzyme ATP synthase or to
fuel the intracellular uptake of Pi or other small
nutrient molecules (e.g., via membrane symporters;
Beja et al. 2001, Fuhrman et al. 2008).
P storage as polyphosphate. Phytoplankton are capable of storing excess intracellular phosphate not
needed immediately to support cell metabolism and
growth, such as that taken up at sustained high concentrations or pulses of external Pi. The stored P
can then support high population growth rates for
multiple generations under subsequent low P conditions (Droop 1973, Ducobu et al. 1998, Morel
1987). The major known mechanism for storing P
in phytoplankton (and indeed all organisms) is the
formation of polyphosphate (polyP; Fig. 4), which
consists of linear chains ranging from several to
hundreds of phosphate residues linked by highenergy phosphoanhydride bonds (Kornberg et al.
1999). Because of the high energy of the phosphoanhydride bonds, polyP is utilized by cells not only
for phosphate storage but also for energy storage,
and can be used as a source of ATP by its enzymatic
reaction with ADP (Kornberg et al. 1999, Achbergerova and Nahalka 2011):
PolyP(n) þ ADP $ PolyPðn 1Þ þ ATP
ð2Þ
where n is the number of phosphate residues in the
polyP chain. PolyP formation occurs in all organisms (Kornberg et al. 1999), including phytoplankton (Rhee 1973, Elgavish et al. 1982, Rivkin and
Swift 1985).
PolyP formation from ATP, and the reverse reaction to reform ATP (eq. 2), is catalyzed by polyphosphate kinase (PPK) in heterotrophic bacteria and
cyanobacteria (Fig. 4), but this protein has not been
found in eukaryotic cells (Kornberg et al. 1999,
Rocap et al. 2003). In addition, two other enzymes,
exophosphatase (PPX) and endophosphatase, catatyze the hydrolysis of polyP to Pi in bacteria and
eukaryotes (Kornberg et al. 1999) (Fig. 4). In yeast
and other eukaryotes polyP often occurs in vacuoles
and its formation involves the vacuolar transporter
chaperone (Vtc) 1–4 enzyme family (Ogawa et al.
2000). Homologs of two Vtc family genes have been
identified in the genome of the diatom Thalassiosira
pseudonana (Dyhrman et al. 2012). Surprisingly,
transcriptomic and proteomic analysis indicated that
the Vtc 4 homolog is up-regulated under P limitation of growth rate, the opposite of what would be
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
function as an external phosphate storage pool that
could be used to support cell growth during low-P
stress (Fu et al. 2005). The fraction of cellular P
present as surface-adsorbed phosphate increases
with the external Pi concentration (Sa~
nudo-Wilhelmy et al. 2004, Fu et al. 2005), which could make
such an external storage mechanism particularly
effective. Such possibilities clearly warrant further
investigation, particularly from a biochemical and
molecular perspective.
ADAPTATION AND ACCLIMATION RESPONSES TO LOW - P
STRESS
In addition to the up-regulation of high-affinity
Pi uptake systems, phytoplankton show a variety of
other adaptation and acclimation responses to lowP stress. These include the substitution of sulfate
for phosphate in membrane lipids, the utilization
of DOP via hydrolytic enzymes, and the acquisition
of P via phagatrophic consumption of other
microorganisms (Dyhrman et al. 2007, Hartmann
et al. 2012). The specifics of these various low-Pcoping mechanisms are discussed in the following
sections.
Reducing cellular P demand. In many marine
cyanobacteria, particularly those residing in low-P
oceanic waters, some phospholipids in cell membranes are replaced by sulfonated lipids to reduce
cellular demand for P in response to low-P stress
(Van Mooy et al. 2006, 2009, Snyder et al. 2009).
Similar phospholipid-to-sulfolipid shifts have also
been found in the brown tide pelagophyte
A. anophagefferens (Wurch et al. 2011). In addition,
both A. anophagefferens and the dinoflagellate
K. mikimotoi adjust their glycolytic pathway under
low-P stress to utilize alternate enzymes that require
less P, which enhances the ability of these species to
grow in low-P environments (Lei and Lu 2011,
Wurch et al. 2011). The molecular underpinning of
the phospholipid-to-sulfolipid shift in phytoplankton
under low-P stress remains to be elucidated.
Utilization of phosphoesters via alkaline phosphatase. As Pi levels decline with algal growth, DOP
concentrations increase due to its release from the
biological community. So one of the most important
mechanisms for coping with low-Pi stress is the utilization of DOP. There is increasing evidence that
DOP is an important source of P to phytoplankton
in low-Pi regions, such as the Sargasso Sea where
DOP:Pi ratios in surface waters can exceed 100 (Wu
et al. 2000, McLaughlin et al. 2013). For example,
30% of primary production during the spring
bloom in the North Atlantic subtropical gyre was
estimated to be supported by DOP (Mather et al.
2008). And 17%–82% of the P taken up by phytoplankton in the Sargasso Sea is estimated to have
been supplied from DOP (McLaughlin et al. 2013).
However, the partitioning of P utilization between
Pi and DOP in stratified surface ocean waters with
REVIEW
expected if polyP was merely a P-storage molecule
(Dyhrman et al. 2012). Nuclear magnetic resonance
analysis verified higher polyP levels in the P-limited
T. pseudonana cells and similar elevated polyP levels
have been observed in P-limited cells of the bacterium Escherichia coli (Kornberg et al. 1999). These
increases appear to be a response to nutrient
growth limitation in general as poly-P accumulation
also occurs when the growth rate of bacteria (Kornberg et al. 1999) and diatoms (Perry 1976) are limited by N. These findings appear to agree with
recent observations in low DIP and dissolved inorganic nitrogen (DIN) waters of the Sargasso Sea,
where unexpectedly high polyP to particulate P
ratios were observed, even though the system
appeared to be P-stressed based on high particulate
AP activities and high levels of phospholipid
replacement by sulfolipids (Martin et al. 2014).
These unexpected findings may be explained by the
many other cellular functions of polyP other than P
storage: particularly nutrient stress responses, energy
storage, and storage of essential nutrient metals
such as iron, which has been observed in yeast
(Lesuisse and Labbe 1994).
Surface phosphate adsorption. In addition to its
assimilation into cellular biomolecules, phosphate
can also adsorb directly onto the surface of phytoplankton cells (Sa~
nudo-Wilhelmy et al. 2004). This
adsorbed phosphate can account for 14%–90% of
total cell P and must be removed if true cellular P
levels are to be measured (Sa~
nudo-Wilhelmy et al.
2004, Fu et al. 2005). This adsorbed phosphate can
be removed with oxalate washes which dissolve and
remove ferric (Fe[III]) oxyhydroxides and manganese (Mn[III and IV]) oxides via their reduction
to soluble ferrous ions (Fe[II]) and manganous ions
(Mn[II]) (Sa~
nudo-Wilhelmy et al. 2004). The
adsorbed phosphate apparently is associated with
the oxides which precipitated on the cell surface
(e.g., by oxidation of Mn (II) and Fe(II)), and are
known to strongly adsorb phosphate. A strong correlation between cell surface phosphate and Mn oxides (r2 = 0.81) supports the oxide adsorption
hypothesis (Sa~
nudo-Wilhelmy et al. 2004). The
adsorbed phosphate may play an important, yet
poorly defined role in intracellular P uptake, particularly in environments with fluctuating concentrations of Pi, Mn, and Fe (Fu et al. 2005). On the one
hand, the abiotic adsorption of Pi on Mn and Fe
oxides removes Pi at the cell surface that would
otherwise be available for intracellular transport.
This could be particularly problematic for larger
cells where the diffusive flux of Pi to cell surface is
already severely limited. However, algal cells possess
mechanisms to reductively dissolve Mn and Fe oxides (e.g., transmembrane reductases; Sunda 2012),
which could release adsorbed Pi into solution for
subsequent uptake by the cell. Thus, if cells are able
to utilize this or other mechanisms (e.g., phagotrophy) to take up the adsorbed phosphate, it could
19
REVIEW
20
SENJIE LIN ET AL.
combined low-Pi and high-DOP concentrations is
difficult because true Pi values can be much lower
than measured Pi levels (as discussed previously)
and most of the DOP that is utilized must first be
converted to Pi by cell surface and extracellular
APs. Among the various forms of DOP, some studies
suggest that nucleotides are utilized preferentially
(Wang et al. 2011) and are important in supporting
phytoplankton growth in oceanic waters (Bj€
orkman
and Karl 2005).
The most important DOP utilizing enzyme is AP,
which hydrolyzes organic monophosphate esters to
Pi, often at the cell surface (Labry et al. 2005,
Nicholson et al. 2006, Huang et al. 2007, Duhamel
et al. 2010, 2011, Fig. 4). The released Pi is then
taken up intracellularly by Pi transport proteins. AP
is substantially up-regulated in low-P-stressed algal
cells, allowing them to acquire Pi from extracellular
DOP pools (Dyhrman et al. 2012). The activity of
AP is highest under alkaline conditions (pH ≥8),
and thus AP is well adapted to surface seawater,
which has a current average pH of 8.1 and had a
pH of 8.2 during preindustial times (Sunda and Cai
2012). However, current and future ocean acidification from anthropogenic increases in atmospheric
carbon dioxide are decreasing surface ocean pH values (Feely et al. 2009), which could potentially
decrease the activity of AP in ocean waters, and
thus, decrease the utilization of DOP by phytoplankton. Since DOP is thought to be a major source of
P to phytoplankton in ocean waters, this could
adversely affect P utilization and algal growth in the
future ocean.
The enzymatic activity of AP has been widely utilized as an indicator of P stress (Dyhrman and Ruttenburg 2006, Lomas et al. 2010). To measure AP
activity, phytoplankton and other microorganisms
are incubated with a phosphoester substrate analog
of AP to generate a product that can be measured fluorometrically or colorimetrically, depending on the
chemical nature of the added substrate (Gonzalez-Gil
et al. 1998). For the colorimetric assay, p-nitrophenyl
phosphate is used as a phosphatase substrate, which
turns yellow (kmax = 405 nm) when dephosphorylated by AP. The substrates used for fluorescent
assay include 2-(50 -chloro-20 -phosphoryloxyphenyl)6-chloro-4-(3H)-quinazolinone (also known as
enzyme-labeled fluorescence ELF-97â or ELF), 3-0methylfluorescein phosphate, 3,6-fluorescein diphosphate, and 4-methylumbelliferyl phosphate. Of these
ELF-97 gives an insoluble fluorescent precipitate,
allowing microscopic observation of the cellular and
subcellular localization of AP (Gonzalez-Gil et al.
1998). Bulk AP activity in a sample can be measured
using a multiwell plate reader while the AP distribution among cells can be measured with a flow cytometer. The quantitative AP activity is usually normalized
on a per cell basis, but a recent study showed that
normalizing it to light absorbance at 450 nm, a proxy
of algal cell biomass, increases the statistical power
and simplifies sample-handling (Peacock and Kudela
2012).
Compared to diatoms, dinoflagellates generally
exhibit higher AP activities on a per cell C or biovolume basis. In a study in Monterey Bay, California dinoflagellates accounted for the majority of
AP activity measured using the ELF substrate even
though diatoms were dominant (Nicholson et al.
2006). A similar trend was shown in a study conducted in the Taiwan Strait in August 2004 and
March 2005 where the average ELF staining rate
was 75 16% for dinoflagellates and 29 19%
for diatoms (Ou et al. 2006). The percentage of
ELF labeling can also vary among dinoflagellates
ranging from 17%–21% for Gonyaulax and Dinophysis spp. to 82%–84% for Protoperidinium spp. and
K. mikimotoi in the East China Sea (Huang et al.
2007). These results indicate a wide variability in
AP expression among species in response to low-P
stress.
The Pi threshold at which AP is induced has been
determined for only a limited number of species.
The results show a wide range, 0.4–16.4 lM for
dinoflagellates compared to 0.25–50 lM for other
groups of phytoplankton, with no clear lineagebased differences, although the highest thresholds
for inducing AP activity tend to be in diatoms
(Table 2). Some of these values exceed the maximum Pi concentrations in ocean waters (2–3 lM).
Studies show that AP activity is controlled more by
intracellular P pools than by external Pi concentrations (Elgavish et al. 1982). This internal regulation
can complicate accurate determination of threshold
Pi values and may account for the higher values for
AP induction observed in Table 2. To complicate
matters further, in some freshwater epiphytic algae
AP was expressed constitutively, even when Pi was
1 mM (Young et al. 2010). Another potential issue
associated with the wide range of Pi thresholds is
variation associated with different methods for
detection of AP thresholds. For example, flow
cytometer-based methods are often more sensitive
than those using a regular fluorometer, resulting in
different threshold estimates (Jauzein et al. 2010).
While AP activity is widely measured, relatively little effort has been made to elucidate AP gene
sequences or regulation of gene expression in marine eukaryotic phytoplankton (Lin et al. 2012a,b,
2013, 2015b). Overall, AP gene sequences are highly
variable among different microorganisms and those
for heterotrophic bacteria, cyanobacteria, and
eukaryotic algae can hardly be aligned, even at the
amino acid level (Lin et al. 2012b). The highsequence variability suggests rapid divergence of
gene homologs or converging evolution of different
AP genes, as in the case of the Pi transporter IPT.
Three AP gene families, phoA, phoX, and phoD,
operate in heterotrophic bacteria and are often
found in different cell compartments (cytoplasm,
periplasm, outer membrane, and extratracellular;
0.7
1.72
Thalassiosira weissflogii
Pavlova lutheri
Isochrysis sp.
Phaeocystis sp.
Haptophyta
Emiliana huxleyi
0.58
61.2
Thalassiosira pseudonana
Nitzschia sp.
Chaetoceros neogracile
Skeletonema costatum
0.68
1.9
Pyrocystis noctiluca
Bacillariophyta
Phaeodactylum tricornutum
1.96
Prorocentrum minimum
Prorocentrum donghaiense
Karenia brevis
6.84
0.038
1.42
3.4
Gymnodinium catenatum
Heterocapsa circularisquama
Heterocapsa triquetra
Karenia mikimotoi
1.4
2.6
“Alexandrium tamarense”
0.14
1.9
Vmax (pmol cell1 h1)
Dinophyta
“Alexandrium catenella”
Ks
(lM)
Yamamoto
et al. (2012)
Fuhs et al.
(1972)
Perry (1976)
Fuhs et al.
(1972)
Tarutani and
Yamamoto
(1994)
Cembella et al.
(1984)
Rivkin and Swift
(1982)
Nakamura and
Watanabe (1983)
Yamamoto and
Tarutani (1999)
Yamamoto et al.
(2004)
Reference
0.0026
0.0011
0.58
1.73
2.28
Kl (lM)
Laws et al.
(2011b)
Riegman
et al. (2000)
Yamamoto
et al. (2012)
Ou et al.
2008
Ou et al.
2008)
Ou et al.
(2008)
Reference
0.37
2.5 9 105
24 9 105
0.1 9 105
11 9 105
6.7 9 105
3.3 9 10
5.7 9 105
4.6 9 105
0.48
1.25
1.2
0.72
0.72
0.67
0.56
2.5 9 105
5
0.63
lmax
(d1)
4.5 9 105
Q0 (pmol lm3)
Riegman et al.
(2000)
Yamamoto
et al. (2012)
Tarutani and
Yamamoto (1994)
Ou et al. (2008)
Ou et al. (2008)
Tarutani (1999)
Yamaguchi and
Itakura (1999)
Yamaguchi and
Itakura (1999)
Yamamoto et al.
(2004)
Ou et al. (2008)
Reference
REVIEW
Species
Oh et al. (2002)
3.3
0.5
0.25
12.1
0.25
50
(continued)
Dyhrman and
Palenik (2003)
van Boekel and
Veldhuis (1990)
Meseck et al.
(2009)
Meseck et al.
(2009)
Garcıa Ruiz
et al. (1997)
Yamaguchi
et al. (2004)
Meseck et al.
(2009)
16.42
<0.5
Yamaguchi
et al. (2004)
Vargo and
Shanley (1985)
0.2
0.43
Jauzein et al.
(2010)
Oh et al. (2002)
Reference
0.4–1
[Pi]
threshold
TABLE 2. Values (in lM) of Pi uptake, cellular Pi quota, and growth parameters as well as Pi-threshold concentrations reported to induce alkaline phosphatase.
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
21
22
13.6
10.1
Chlorophyta
Chlorella autotrophica
Tetraselmis chui
1.19
Laws et al.
(2011a)
Watanabe
et al. (1982)
Nakamura and
Watanabe (1983)
Laws et al.
(2011a)
7.9 9 105
0.86
1.6 9 105
Nakamura and
Watanabe (1983),
Nakamura (1985)
0.14
Tetraselmis suecica
Heterosigma akashiwo
1.76
Raphidophyta
Chattonella antiqua
Species
TABLE 2. (continued)
Ks
(lM)
Vmax (pmol cell1 h1)
Reference
Kl (lM)
0.00345
Reference
Q0 (pmol lm3)
lmax
(d1)
Reference
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[Pi]
threshold
Reference
Meseck et al.
(2009)
Meseck et al.
(2009)
SENJIE LIN ET AL.
Luo et al. 2009, Sebastian and Ammerman 2009,
White 2009). The greatest abundance of AP genes
appear to code for cytoplasmic proteins, suggesting
that intracellular uptake of phosphoesters and subsequent hydrolysis within the cell may be more
prevalent in marine heterotrophic bacteria than previously thought (Luo et al. 2009). For AP to function intracellularly, however, would require
phosphoester transporters which have not yet been
identified (Fig. 4). Alternatively, many of these
intracellular APs may be involved in the hydrolysis
of phosphate esters produced within the cell that
are utilized in cell metabolism and cell signaling
(Dick et al. 2011).
The three bacterial AP proteins (PhoA, D, and X)
have active sites which contain different metal ions.
Fe, Zn, and sometimes Co occur in most (if not all)
AP-active centers as these reactive metal ions are
required for the hydrolytic activity of AP enzymes
(Coleman 1992, Yong et al. 2014). Cyanobacteria
contain genes for all three of these bacterial APs,
and in addition can contain genes of another, phoV
(Table 3). PhoA and phoV have two Zn ions and a
magnesium (Mg) ion in their active centers, while
PhoD utilizes calcium (Ca) instead of Mg ions and
may also contain Zn or other hydrolytically active
metal (although such a requirement has not yet
been established; Roy et al. 1982, Coleman 1992,
Kageyama et al. 2011). In contrast, recent structural
analysis indicates that PhoX contains two ferric ions
and three Ca ions in its active center (Yong et al.
2014). The metal ions that are utilized in the active
site of the enzyme is of interest, not only because
this can affect the activity and specificity of the
enzyme but also because regional variations in limiting metal concentrations may select for species containing APs with different metal requirements.
Putative phoX genes have also been identified in
Prochlorococcus and Synechococcus (Kathuria and Martiny 2011), and in two chlorophytes: Volvox carteri
(Hallmann 1999) and Chlamydomonas reinhardtii (Quisel et al. 1996, Moseley et al. 2006). An AP has also
been identified by proteomics in the pelagophyte
Aureoumbra lagunensis, which was activated by Ca but
inhibited by Zn, consistent with the behavior of
PhoX (Sun et al. 2012). However, an iron requirement was not examined in this study; and even if it
had been, the standard chelator (EDTA) used to
remove catalytically active metals from AP enzymes is
not strong enough to remove ferric ions from the
active site of PhoX (Yong et al. 2014). This fact has
led to much past confusion regarding the metal
requirements of this enzyme (Yong et al. 2014).
A novel AP (EHAP1) has also been identified in
the widely distributed marine haptophyte Emililiania
huxleyi (Xu et al. 2006, 2010). Based on its amino
acid sequence, this enzyme is phylogenetically
related to PhoD (Lin et al. 2013). It appears to
require either Zn or cobalt (Co) as a cofactor as the
AP activity of E. huxleyi is greatly suppressed in the
23
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
TABLE 3. Alkaline phosphatase and other genes in phytoplankton that facilitate the utilization of dissolved organic
phosphate identified to date.
Alkaline phosphatase (AP)
PhoA
Zn/Mg
Bacillariophyta
Thalassiosira
pseudonana
Phaeodactylum
tricornutum
Haptophyta
Emiliania
huxleyi
Prymnesium
parvum
Pelagophyta
Aureoumbra
laguna
Aureococcus
anophagefferens
Chlorophyta
Chlamydomonas
reinhardtii
phoD
Ca/?
Other enzymes
phoV
Zn/Mg
Synechococcus
sp. PCC7942
Trichodesmium
50 nucleotidase
Armbrust
et al. (2004)
Dhyrman
et al. (2012)
Bowler et al.
(2008)
Dyhrman
et al. (2006a)
Xu
et al.
(2006)
Beszteri
et al. (2012)
Sun et al.
(2012)
Wurch
et al. (2011)
Moseley
et al. 2006
Wurch
et al. (2011)
Quisel
et al. (1996);
Moseley
et al. (2006)
Kruskopf and
Du Plessis
(2004)
Blanc et al.
(2010)
Volca_XP_
002958226.1
Hallmann
(1999)
Cyanophyta
Aphanothece
halophytica
Prochlorococcus
Acid
phosphatase
Lin et al.
(2012b)
Lin et al.
(2011)
Morey
et al. (2011)
Lin et al.
(2012a)
Chlorella sp.
Volvox carteri
New
type
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Dinophyta
Alexandrium
“catenella”
Amphidinium
carterae
Karenia brevis
phoX
Fe/Ca
Kageyama
et al.
(2011)
Moore
et al. 2005
Ray et al.
1991
Moore
et al. (2005)
Moore
et al. (2005)
Orchard
et al. 2009
Orchard
et al. (2009)
combined absence of Zn and Co and is enhanced
by the addition of either metal (Shaked et al. 2006,
Jakuba et al. 2008). The presence of Zn or Zn/Co
in many AP enzyems (e.g., PhoA and EHAP1) suggests that P and Zn (and/or Co) may colimit DOP
utilization and algal growth in some regions of the
ocean such as the Sargasso Sea where Pi, Zn, and
Co occur at very low concentrations (Wu et al.
2000, Shaked et al. 2006, Jakuba et al. 2008). However, the presence of Fe in the active site of PhoX
suggests that colimitation by Fe and P may also
occur in oceanic regions with low Fe and P concentrations. Indeed, the relative abundance of Zn and
Fe in low-P ocean waters may influence the compo-
Wagner
et al.
(1995)
sition of phytoplankton communities since Fe- and
Zn-dependent APs often do not occur together in
the same species (Yong et al. 2014).
Alkaline phosphatase genes have also been identified in diatoms and dinoflagellates, two dominant
groups of eukaryotic phytoplankton. AP genes have
been identified in the genomes of the diatoms
Thalassiosira pseudonana and Phaeodactylum tricornutum (Armbrust et al. 2004, Bowler et al. 2008, Dyhrman et al. 2012) and the dinoflagellates
Amphidinium carterae (Lin et al. 2011), Karenia brevis
(Morey et al. 2011, Lin et al. 2012a), and Alexandrium catenella (Lin et al. 2012b). A cell surface
protein showing AP activity was also identified in
REVIEW
24
SENJIE LIN ET AL.
the dinoflagellate Prorocentrum minimum (Dyhrman
and Palenik 1997). Algal species can contain multiple AP genes coding for different proteins. For
example, at least four different putative APs have
been identified in the diatom T. pseudonana, and
all four were up-regulated under P-deficiency
(Dyhrman et al. 2012).
Despite the high sequence divergence of all
eukaryotic AP genes, those from dinoflagellates and
some from diatoms, pelagophytes, and haptophytes
are slightly more similar to phoA genes than to the
other bacterial AP types (Lin et al. 2012b). Furthermore, these PhoA-like APs, from dinoflagellates, diatoms, and haptophytes (the so-called red algal
lineage) are closer to one another than to those in
chlorophytes, consistent with the known phylogenetic and evolutionary relationships among these
algal lineages (Lin et al. 2012b, Lin et al. 2015b).
Whether the high variability in AP sequences confers different substrate specificities or other functional differentiation is currently unknown. In
bacteria, the sequence variability among the different types of APs is related to the cellular localization
of the encoded enzymes (Luo et al. 2009). Similarly,
in silico analysis of AP gene sequences in eukaryotic
algae predict that various AP enzyme types have different localizations (extracellular, cell wall, plasma
membrane, or cytoplasm; Lin et al. 2012b). Recent
ELF staining of live cells of the dinoflagellates
Amphidinium carterae, Karenia brevis, and Alexandrium
catenella (=A. pacificum) shows AP localization patterns that largely agree with these predictions (Lin
et al. 2012b). The cellular localization of different
AP enzymes may enable species to utilize different
sources of DOP and to hydrolyze various DOP compounds in different cellular or extracellular locations. It may also be related to variations in
chemical environment, such as differences in pH or
ionic composition near the cell surface or within
the cell. Much work remains to be done on the regulation and localization of different AP proteins in
phytoplankton.
Utilization of phosphonates. Phosphonates contribute 5%–25% to the total DOP in the ocean
(Clark et al. 1998, Kolowith et al. 2001, Young and
Ingall 2010). They are likely produced in a wide
range of organisms as constituents of phosphoproteins and cell membrane phospholipids (Clark et al.
1998, Villareal-Chiu et al. 2012). The commonly
occurring phosphonate, 2-aminoethylphosphonic
acid, for instance, is present in plant and animal
cell membranes. Cyanobacteria, which can utilize
phosphonates, can also produce them (Dyhrman
et al. 2009). Heterotrophic bacteria have long been
known to take up and metabolize phosphonates
(Shinabarger et al. 1984, Pipke et al. 1987, for a
review see McGrath et al. 2013), but only recently
was this capability found to occur in cyanobacteria
(Dyhrman et al. 2006b, Ilikchyan et al. 2009,
Gomez-Garcia et al. 2011). Utilization of phospho-
nates requires the cleavage of the C-P bond, which
is energetically more difficult than hydrolyzing a
phosphoester bond. Utilization of phosphonates in
heterotrophic bacteria is accomplished using either
a C-P hydrolase or a C-P lyase enzyme system. In
contrast, marine cyanobacteria, contain only the C-P
lyase system (Dyhrman et al. 2006b, McGrath et al.
2013). The proteins comprising the C-P hydrolase
enzyme system vary among bacteria, with PhnW and
PhnX being the most common constituent proteins.
By contrast, the C-P lyase system appears to be more
conserved, and consists of 14 proteins under Pho
regulon control, which are capable of processing a
broad range of phosphonate substrates (White and
Metcalf 2004, Dyhrman et al. 2006b). The C-P lyase
system proteins are encoded by a gene cluster,
PhnCDEFGHIJKLMNOP.
Within
this
cluster
PhnCDE codes for a phosphonate ABC transporter
(which includes an ATP binding subunit protein, a
periplasmic phosphonate binding subunit, and
transmembrane subunit), whereas PhnFGHIJKLMNOP codes for the C-P bond cleaving enzymes
(Fig. 4). Currently, there is no documented evidence that eukaryotic phytoplankton can utilize
phosphonates, although some preliminary molecular data indicate the presence of phosphonate-metabolizing enzyme genes in some dinoflagellates (Lin
et al.2015a; see Supplementary Table 33.
P acquisition by phagotrophy. Phagatrophy occurs
widely in many groups of photosynthetic protists
and is now recognized as an important source of P
and other nutrients (e.g., Fe) in low-nutrient waters
(Stoecker 1999, Jeong et al. 2010b, Hartmann et al.
2012, Flynn et al. 2013). It is common in dinoflagellates, haptophytes, and pelagophytes, but does not
occur in diatoms. All dinoflagellates tested to date
are capable of phagotrophy, including species originally considered obligate photoautotrophs; e.g., Prorocentum minimum (Stoecker et al. 1997), Akashiwo
sanguenium (Bockstahler and Coats 1993), Karlodinium veneficum (Li et al. 1996), Alexandrium ostenfeldii (Jacobson and Anderson 1996), Gymnodinium
aureolum (Jeong et al. 2010a), and even the coral
reef endosymbiont Symbiodinium (Jeong et al. 2012).
Studies have shown that phagotrophy in dinoflagellates and other mixotrophic phytoplankton is
induced by low nutrient stress or nutrient limitation
of growth rate (Stoecker et al. 1997, Litaker et al.
2002, Carvalho and Graneli 2010, Jeong et al.
2012). Potential phagotrophy-related genes have
been identified in dinoflagellates and other phytoplankton. Clathrin-mediated endocytosis proteins
and autophagy-related proteins were up-regulated
under Pi limitation in the dinoflagellate K. mikimotoi
(Lei and Lu 2011), the haptophyte Prymnesium parvum (Beszteri et al. 2012), and the pelagophyte
A. anophagefferens (Wurch et al. 2011). Phagotrophy
represents an efficient means of acquiring P and
other nutrients in situations where dissolved nutrient concentrations are low and a substantial propor-
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
tion of the nutrients are contained in bacteria,
microalgae, and other microorganisms (Smalley
et al. 2003, Flynn et al. 2013). The biochemical
pathways involved in phagotrophy and subsequent
nutrient assimilation represent an understudied
aspect of nutrient acquisition worthy of further
investigation. In particular, the potential involvement of acid phosphatase and nucleotidase
(Table 3) in releasing phosphate during food digestion deserves attention.
Growth relation to cellular P. Decreases in P or
other nutrients below critical threshold values
results in decreased rates of cellular growth and
reproduction. Early models related nutrient limitation of growth rate to the external nutrient concentration (e.g., Monod 1949) based on chemostat
studies in which the external nutrient concentration
remained constant with time. However, concentrations of Pi and other nutrients in seawater are often
variable in time and space (Turpin and Harrison
1979), and limitation of growth rate is now known
to be dependent on the cell nutrient quota
(amount per cell) or concentration (e.g., amount
per unit of cell carbon) rather than the external
concentration (Droop 1968, Fuhs 1969, Grover
1991). Depending on the growth status and previous nutrient exposure, cell P quotas can vary widely
(Jauzein et al. 2010, Pleissner and Eriksen 2012).
The model most often used to describe the relationship between the cell nutrient quota (Q) and
specific growth rate (l) is the Droop equation (Droop 1968, 1983, 2003):
l ¼ lmax ð1 Q0 =Q Þ
ð3Þ
where Q0 is the minimum cell quota at which the
growth rate is reduced to zero, and lmax is the hypothetical maximum growth rate at an infinite cell
quota. lmax is often unobtainable, and in reality is
only a fitting parameter for the equation (Droop
1983, Laws et al. 2011a,b, 2013). Also Q should be
the average daily cell quota since quotas for P and
other nutrients (e.g., N and Fe) typically vary with
time of day due to diel variations in rates of nutrient uptake, cell growth and cell division (Ahn et al.
2002, Sunda and Huntsman 2004).
An advantage of this approach is that it allows
growth rate predictions based on cell quotas, which
are generally easier to measure than the low concentrations of Pi and other nutrients that actually
limit algal growth rates (Laws et al. 2011b, Sunda
and Hardison 2010). In recent chemostat studies
under continuous light conditions, the relationship
between growth rate and cell P quota was well
described by the Droop equation (Laws et al. 2013).
Although the classic Droop equation allows a prediction of growth versus cell quota (average amount
per cell) for a given species, it fails to provide a
measure of growth efficiency – the rate of cell carbon production per unit of cell P. Relationships
between growth rate and cell quota can vary widely
among species because of large differences in cell
size, and resultant moles of P per cell. Normalizing
cellular P on per unit carbon basis (i.e., the molar
P:C ratio) eliminates this difficulty and allows carbon growth per unit of cell P to be directly compared among species as was done in Jauzein et al.
(2010). Only when more studies directly relate
growth rates to cell P:C ratios will it be possible to
determine if dinoflagellates, diatoms, or other phytoplankton groups on average have different P
growth efficiencies. Such efficiencies are defined as
the net moles of cell C produced per mole of cell P
per unit time (i.e., per day) and equal the specific
growth rate divided by average daily cellular P:C
ratio.
Changes in the cell P:C ratio (QC) with time is
governed both by the C-normalized cellular Puptake rate (VC) and the C-specific growth rate (lC)
according to the equation:
dQC =dt ¼ VC lC QC
ð4Þ
At steady state, this relationship collapses to:
QC ¼ VC =lC
ð5Þ
As noted previously, for cells growing under a diel
light cycle, all values in the above equations need to
be daily averages, which can usually be estimated
from measurements made in the middle of the light
period (Sunda and Hardison 2007). From the above
equations it is evident that the cellular P:C ratio is
determined by the balance between the cellular P
uptake rate and the specific growth rate, and that
changes in P:C ratios will be determined by the relative changes in the two factors.
Variations in N:P:C stoichiometry and its influence on
P versus N limitation of growth rate. Due to the variability in the uptake mechanisms and kinetics, cell
size and cellular P growth requirements, the Pi
levels in any given ecosystem may be growth-sufficient for some species, but growth-limiting for
others (Sundareshwar et al. 2003, Nicholson et al.
2006, Mackey et al. 2007). This is true even though
the phytoplankton community can adapt evolutionarily to a range of ambient concentrations of P, N,
and other nutrients; for example, the consistently
low nutrient levels in open ocean surface waters
(Chisholm 1992, Sunda and Hardison 2007, 2010).
Even when cellular P quotas are normalized to cell
carbon, these values still vary among species
(Table 2). This variability likely results from differences among algal species in their cellular P uptake
rates and specific growth rates and to associated differences in their biochemical composition, particularly in the abundance of RNA, DNA,
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GROWTH LIMITATION AND CELLULAR RESPONSES TO IT
25
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26
SENJIE LIN ET AL.
phospholipids, and polyphosphate (Falkowski 2000,
Geider and La Roche 2002). As a result, N:P:C ratios
in phytoplankton species can be quite variable, with
N:P ratios for nutrient replete algal species varying
from 5 to 19 (Geider and La Roche 2002). Molar N:
P ratios >16:1 in seawater are typically regarded as
P-limiting. However, in seawater environments with
measured ratios of inorganic DIN:Pi as high as 25:1,
the N:P ratios in plankton tend to be much less than
16 (median 9) (Broecker and Henderson 1998,
Geider and La Roche 2002). This discrepancy is most
likely due to the accumulation of P storage pools
(e.g., polyphosphate) and to variations in ribosomal
RNA linked to changes in rates of growth and protein synthesis (Geider and La Roche 2002). The
adsorption of phosphate on cell surfaces can also be
a factor and can significantly skew N:P ratios in phytoplankton toward lower values (Sa~
nudo-Wilhelmy
et al. 2004, Fu et al. 2005). The critical molar N:P
ratio in phytoplankton that marks the transition from
N- to P-limitation is in the range of 20–50, significantly higher than the 16:1 Redfield ratio (Geider
and La Roche 2002). Variations in these critical N:P
ratios among species may be an important factor
affecting algal species competition and the formation
of species-specific algal blooms, including those of
dinoflagellates.
Increased cellular toxins in P-limited cells. Growth
rate limitation by P and other nutrients (e.g., N) is
often accompanied by increases in toxin per cell or
per mol of cell C in many toxic harmful algal bloom
species. This has been observed for saxitoxins,
which cause paralytic shellfish poisoning (Flynn
et al. 1994, Maestrini et al. 2000, Anderson et al.
2002), okadaic acid, which causes diarrhetic shellfish poisoning (John and Flynn 2002), and domoic
acid, which causes amnesic shellfish poisoning (Pan
et al. 1998). P limitation of growth rate also
increases cellular toxins in Chrysochromulina polylepis
(Johansson and Graneli 1999a), Gambierdiscus polynesiensis (Chinain et al. 2010), Karlodinium veneficum
(Fu et al. 2010), Prymnesium parvum (Beszteri et al.
2012, Johansson and Graneli 1999b), Protoceratium
reticulatum (Guerrini et al. 2007), and the N2-fixing
cyanobacterium Nodularia spumigena (Sunda et al.
2006). In a detailed study of the dinoflagellate Karenia brevis, Hardison et al. (2013) showed that the
increase in cellular toxins (bevetoxins):C ratios was
best predicted by the degree of growth rate limitation by P and not by cell P:N or P:C ratios. They
found that the increased cellular toxin:C ratios were
not, however, due to an increase in the cellular
toxin production rate as might be expected intuitively. Instead, as growth slowed under P limitation,
cells down-regulated the rate of toxin synthesis, but
to a lesser degree than the overall decrease in the
rate of cellular C production leading to higher cellular toxin:C ratios. Similar increases in cellular
brevetoxin:C ratios were also observed under both
N limitation (Hardison et al. 2013) and CO2 limia-
tion of growth rate (Hardison et al. 2014), suggesting that the increase in brevetoxins is inherently
linked to the slower growth rates that occur during
nutrient limitation. Hardison et al. (2012, 2013)
hypothesized that the increase in cellular toxins is
evolutionarily advantageous, as elevated brevetoxins
have been shown to deter zooplankton grazing
(Hong et al. 2012). The lower grazing rates would
result in higher net population growth rates than
would occur otherwise as algal growth rates slow
during nutrient limitation.
The actual biochemical pathways by which P limitation of growth rate regulates toxin production are
largely unknown and researchers are just now beginning to isolate genes involved in the biosynthesis of
various toxins. Recently the polyketide synthase
gene cluster responsible for the production of
brevetoxins in Karenia brevis was identified (Monroe
and Van Dolah 2008). Additionally, two genes
believed to be involved in saxitoxin synthesis in
Alexandrium spp. and other dinoflagellates were also
identified (Orr et al. 2013, St€
uken et al. 2011).
Determining how nutrient limitation of growth rate
and other factors regulate these toxin pathway
genes should prove a fruitful area for future
research.
Encystment induced by P limitation. Dinoflagellates
can survive P deficiency by encystment (Anderson
et al. 1985). However, laboratory experiments have
shown that motile cells need a minimum P content
to form cysts (Anderson et al. 1985), and that cysts
must contain sufficient ATP to germinate after dormancy (Lirdwitayaprasit et al. 1990). Furthermore
in some dinoflagellates (e.g., Scrippsiella trochoidea),
cysts can take up P intracellularly and their P content increases with the external Pi concentration
(Rengefors et al. 1996). This enables dinoflagellate
cysts that settle to the bottom to accumulate P from
abundant sedimentary Pi pools for use during dormancy, germination, and subsequent growth. The
molecular mechanisms of P uptake, storage, and
metabolism during the encystment and germination
phases are currently unexplored.
Cell enlargement and cell cycle arrest caused by P limitation. Nutrient limitation of growth rate in phytoplankton usually causes a decrease in cell size, as
observed for nitrogen (Sunda and Hardison 2007,
2010, Hardison et al. 2012) and iron (Sunda and
Huntsman 1995). Such decreases should be favored
by natural selection as they facilitate nutrient uptake
by increasing cell surface to volume ratios and the
rate of diffusive nutrient flux through the surface
boundary layer normalized to cell volume (Sunda
and Hardison 2007). In contrast, P limitation of
growth rate often leads to cell enlargement, as
reported for dinoflagellates (Latasa and Berdalet
1994, John and Flynn 2002, Lim et al. 2010, Varkitzi
et al. 2010, Hardison et al. 2013, Zhang et al. 2014),
diatoms (Liu et al. 2011), and chlorophytes (Litchman and Nguyen 2008). The increase in cell size
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
DIFFERENTIAL NUTRIENT ACQUISITION AND GROWTH
STRATEGIES AND SPECIES SUCCESSION
Differential P nutrient strategies may be one of
the drivers of seasonal species succession that occurs
in many phytoplankton communities. The popula-
P or C
tion growth of diatoms with high nutrient-sufficient
maximum growth rates is often favored during early
succession (in late winter/early spring or in freshly
upwelled water) when the environment is characterized by high concentrations of dissolved inorganic
nutrients (Pi, DIN, and Fe), high turbulence, low
algal biomass, and a low level of zooplankton
grazing (Margalef 1978, Sunda et al. 2006, Sunda
and Hardison 2010, Fig. 5). As the season progresses, the water column stabilizes with increased
solar heating of surface water, and inorganic nutrient pools become depleted by the initial algal
bloom which is often dominated by diatoms (Margalef 1978, Hood et al. 1990, Tiselius and Kuylenstierna 1996, Yoshimura et al. 2014). As the DIN
and Pi pools decline during the bloom there is a
progressive buildup of organic nutrients (DOP and
DON) linked to slopy grazing and excretion by zooplankton, viral and bacterial lysis of cells, and
release by phytoplankton (van der Zee and Chou
2005, Yoshimura et al. 2014). The combination of a
stable water column, higher phytoplankton biomass,
low inorganic nutrients, increased DOP and DON
levels, and increased zooplankton grazing pressure
no longer favors the population growth of diatoms
or other early succession species and sets the stage
for a population shift to late succession species such
Time
FIG. 5. Schematic for a typical diatom to dinoflagellate seasonal succession in a system whose biomass is limited by P. The
initially high Pi levels, colder temperatures, and high turbulence
levels in the early spring favor diatoms, which are adapted for
high growth rates under these conditions. The emerging diatom
bloom depletes the euphotic zone of Pi and fuels the growth of
zooplankton. Concomitantly, solar warming increases stratification of the water column, and decreases inputs of Pi and other
nutrients from nutrient rich aphotic deeper waters. Phosphorus
inputs during this time are mainly from recycling linked to zooplankton grazing and excretion, and much of that input is in the
form of DOP. The combination of DOP inputs from grazing and
Pi uptake by phytoplankton increases the DOP concentration and
greatly increases the DOP:Pi ratio. These changes (decreased Pi
in surface waters, increased DOP, water column stratification, and
increased zooplankton grazing) sets the stage for a algal community shift from diatoms to dinoflagellates, whose growth and survival are favored under these conditions due to their ability to
obtain nutrients from alternate sources (diel vertical migration to
nutrient rich deeper waters, utilization of DOP, and phagotrophy) and their ability to minimize grazing losses (e.g., linked to
large cell size and the production of toxins).
REVIEW
under P limitation should decrease P uptake rates
per unit of cell volume, and thus, should be evolutionary disadvantageous. So, in contrast to other
nutrients, there must be some factor peculiar to
phosphorus that causes cell size to increase under P
limitation of growth rate.
To examine the effects of P limitation on cell size
we must examine its effect on the cell’s growth and
division cycle. This cycle consists of four discrete
phases: the G1 phase (gap 1 or growth stage 1)
where a newly divided cell grows and increases in
size prior to cell division, the S phase during which
DNA is replicated, the G2 phase (gap 2 or growth
stage 2) where the cell continues to grow prior to
mitosis, and the M phase during which nuclear division (mitosis) occurs leading to cell division (cytokinesis). Unlike other nutrients (e.g., Fe and N), P
limitation often results in a blockage of DNA replication (the S phase; Vaulot 1995), which must precede cell division into smaller daughter cells
(Sclafani and Holzen 2007). P limitation of growth
rate in the cyanobacteria Prochlorococcus and Synechococcus causes an arrest of the cell cycle progression from G1 to S or G2 to M phases, and in the
case of P starvation (severe growth rate limitation),
an arrest in the S phase (Parpais et al. 1996, Vaulot
et al. 1996). Similarly, the few studies conducted so
far with dinoflagellates indicate an arrest of the cell
cycle in G1 phase in response to P limitation of
growth rate (Lei and Lu 2011, Zhang et al. 2014;
Li et al. 2015). This arrest is accompanied by the
up-regulation of negative regulators (e.g., fizzy/cell
division cycle 20-related protein) and down-regulation of positive regulators of the cell cycle (e.g., calcium-dependent protein kinase) (Zhang et al.
2014). With the arrest of the cell cycle, the cells
continued to grow during an elongated G1 phase,
resulting in an increase in average cell size. The
blockage of the cell cycle progression from G1 to S,
or subsequent phase transitions, is likely linked to a
need for a sufficient supply of P for successful DNA
replication and for phosphorylation of key checkpoint enzymes that regulate DNA synthesis and
nuclear and cell division in the S and M phases.
Transitions in the cell cycle, including G2 to M
stages, are strictly regulated by a cascade of CDK
phosphorylation and dephosphorylation events
(Murray and Hunt 1993). The cell enlargement in
P-limited cells further suggests that fulfilling CDK
phosphorylation or other P-associated biochemical
requirements (e.g., DNA replication) may supersede
that of a threshold in cell size in controlling the
onset of cell division.
27
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28
SENJIE LIN ET AL.
as dinoflagellates, haptophytes, and pelagophytes,
which are better adapted to this new set of environmental conditions (Margalef 1978, Sunda et al.
2006).
In contrast to diatoms (which are non-motile),
dinoflagellates and many other late sucession species possess flagella and are motile. This motility
allows them to vertically migrate out of nutrientdepleted surface waters into deeper nutrient-rich
waters at the end of the light period and then
migrate back into sunlit surface waters at the beginning of the light period (Sinclair and Kamykowski
2008, Hall and Paerl 2011). In this way P and other
nutrients that are depleted in surface waters during
blooms, but accumulate at depth or in bottom sediments via POM settling and regeneration processes,
can be utilized to sustain photosynthesis and growth
during the day (Hall and Paerl 2011, Sinclair and
Kamykowski 2008).
In addition, three other functional traits promote
increased nutrient acquisition rates of dinoflagellates and other late succession species in low Pi and
DIN surface waters. One is their higher capability to
utilize DOP and DON than diatoms, which helps
favor their growth in late succession waters with
high ratios of DOP:Pi and DON:DIN (Sunda et al.
2006, Burkholder et al. 2008). Indeed as noted earlier, dinoflagellates are observed to have higher AP
activities during blooms (on either a per cell or cell
volume basis) than coexisting diatoms (Nicholson
et al. 2006, Ou et al. 2006). In addition, some
dinoflagellates can grow as well on ATP as on DIP,
suggesting an ability to directly utilize this DOP
without the energy-costing hydrolysis (Li et al.
2015). A second, perhaps more important functional trait, is the capability of dinoflagellates, haptophytes, and other late succession species to
acquire P and other nutrients via phagotrophy
(Stoecker et al. 1997, Hartmann et al. 2012, Flynn
et al. 2013). This capability may have little survival
value prior to and during the early phase of the initial diatom bloom, and indeed diatoms are one of
the few groups of eukaryotic phytoplankton with no
known phagatrophic capability (Flynn et al. 2013).
However, phagotrophy becomes an increasingly
important nutrient source as algal biomass increases
and Pi and DIN pools become depleted during
blooms.
Dinoflagellates,
especially
athecate
gymnodinoid forms, are known to feed heavily on
diatoms (Sherr and Sherr 2007). Many mixotrophic
dinoflagellates depend primarily on photosynthesis
for growth when nutrients are abundant, but
become facultative heterotrophs once dissolved Pi
and DOP nutrients become growth limiting
(Stoecker et al. 1997, Litaker et al. 2002, Smalley
et al. 2003). This capacity allows dinoflagellates to
acquire P and other nutrients from the consumption of other algal species while at the same time to
reduce competition for the remaining pool of dissolved nutrients. The motility of dinoflagellates and
other flagellates enhances encounter rates with
prey, and their diverse swimming speeds seem to
enable them to effectively capture different types of
prey (Nielsen and Kiorboe 2015).
The third potentially important late succession
trait in dinoflagellates is a greater cell P-storage
capacity. Although both diatoms and dinoflagellates
can store polyP, the large size of dinoflagellates
potentially allows for greater internal storage (Elgavish et al. 1980, Diaz et al. 2008). This greater storage capacity may allow some dinoflagellates to
sustain relatively high growth rates during the late
succession period of low Pi availability (Flynn et al.
1994, Hou et al. 2007).
The net growth of phytoplankton populations is
not only controlled by nutrient–dependent growth
and reproduction but also by grazing mortality
losses (Sunda et al. 2006, Smayda 2008). Here too,
dinoflagellates and other late succession species
may be better adapted to late succession environments characterized by higher zooplankton populations and higher grazing pressures. Many
dinoflagellates and other late succession species
(e.g., haptophytes such as Prymnesium parvum and
pelagophytes such as A. lagunensis) appear to be
well defended from grazing due to high cellular
concentrations of grazing-deterrent compounds
(e.g., toxins and mucilage layers), which increase
under P or N limitation of growth rate (Sunda et al.
2006, Hong et al. 2012, Waggett et al. 2012, Hardison et al. 2013). Thus, differences in P acquisition,
P utilization, and grazing deterrence strategies
between diatoms and dinoflagellates contribute to
their respective dominance in early and late succession environments (Fig. 5). However, many of the
molecular mechanisms unlying these strategies are
still unclear and await more comparative genomics
studies for key species from both phyla.
FUTURE DIRECTIONS
Considerable progress has been achieved in the
last two decades in our understanding of how P
affects the growth and ecology of phytoplankton.
The molecular mechanisms regulating the uptake
and metabolism of P, and adaptation to low-P stress
have become clearer thanks to rapidly expanding
data sets on algal genomes, transcriptomes, and proteomes. However, this initial progress represents
only a small fraction of the information that can be
obtained using careful chemical and physiological
studies coupled with modern molecular techniques.
Some of the most pressing issues include: (i) obtaining a more detailed description of dissolved inorganic and organic P substrate pools in ocean waters,
including actual measurements of the true
orthophosphate (Pi) concentrations in low-Pi oceanic waters; (ii) determining the extent to which P
limitation regulates the growth and species composition of phytoplankton communities in various ocea-
P G R O W T H S TR A T E G I E S A N D M O L E C U L A R ME C H A N I S M S
We thank Dr. Xin Lin for assistance with making Figure 2.
We are also indebted to Drs. Edward J. Carpenter and
Edward Monahan and reviewers for their comments that
helped improve the manuscript. Mark Vandersea and Steven
Kibler provided thoughtful criticisms and edits. Michelle
Wood provided superb editorial guidance and insights for
which we are grateful. This work was supported by the
National Science Foundation grant OCE-1212392 and NOAA
program funds.
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impact of P on the biology of a warmer, more highly
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29
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