Phosphorus physiological ecology and molecular mechanisms ... · Phosphorus (P) is an essential nutrient for marine phytoplankton and indeed all life forms. Current data show that

REVIEW

PHOSPHORUS PHYSIOLOGICAL ECOLOGY AND MOLECULAR MECHANISMS IN MARINEPHYTOPLANKTON1

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

Phosphorus (P) is an essential nutrient for marinephytoplankton and indeed all life forms. Currentdata show that P availability is growth-limiting incertain marine systems and can impact algal speciescomposition. Available P occurs in marine waters asdissolved inorganic phosphate (primarilyorthophosphate [Pi]) or as a myriad of dissolvedorganic phosphorus (DOP) compounds. Despitenumerous studies on P physiology and ecology andincreasing research on genomics in marinephytoplankton, there have been few attempts tosynthesize information from these differentdisciplines. This paper is aimed to integrate thephysiological and molecular information on theacquisition, utilization, and storage of P in marinephytoplankton and the strategies used by theseorganisms to acclimate and adapt to variations in Pavailability. Where applicable, we attempt to identifygaps in our current knowledge that warrant furtherresearch and examine possible metabolic pathwaysthat might occur in phytoplankton from well-studiedbacterial models. Physical and chemical limitationsgoverning cellular P uptake are explored along withphysiological and molecular mechanisms to adaptand acclimate to temporally and spatially varying Pnutrient regimes. Topics covered include cellular Piuptake and feedback regulation of uptake systems,enzymatic utilization of DOP, P acquisition byphagotrophy, P-limitation of phytoplankton growthin oceanic and coastal waters, and the role of P-limitation in regulating cell size and toxin levels inphytoplankton. Finally, we examine the role of Pand other nutrients in the transition ofphytoplankton communities from early successionspecies (diatoms) to late succession ones (e.g.,dinoflagellates and haptophytes).

Key index words: alkaline phosphatase; diatoms;dinoflagellates; dissolved organic phosphorus;

genetics; marine algae; nutrients; phosphate; phos-phorus uptake; transporter

Abbreviations: AMP, adenosine monophosphate; AP,alkaline phosphatase; ATP, adenosine triphosphate;DIN, dissolved inorganic nitrogen; DIP, Dissolvedinorganic phosphate; DOP, dissolved organic phos-phorus; HAB, harmful algal bloom; IP3, inositoltriphosphate; NADPH, reduced nicotinamide ade-nine dinucleotide phosphate; Pi, orthophosphate;RNA, ribonucleic acid; SRP, soluble reactive phos-phorus

Phosphorus (P) is an essential nutrient for allorganisms (Paytan and McLaughlin 2007). It is acentral component of nucleic acids (both DNA andRNA), and thus, plays a critical role in the storage,replication, and transcription of genetic informa-tion. It is present in phospholipids, a key compo-nent of cellular membranes. It also plays a centralrole in the production of chemical energy (adeno-sine triphosphate [ATP]) and of reducing equiva-lents (reduced nicotinamide adenine dinucleotide[NADH] and nicotinamide adenine dinucleotidephosphate [NADPH]) during photosynthesis andrespiration, which are required for carbon fixationand cell metabolism (Falkowski and Raven 2007).One of the highest P requirements is in the synthe-sis of proteins via ribosomal RNA (Geider and LaRoche 2002). Phosphorous also regulates the activityand function of many proteins and metabolic pro-cesses (via phosphorylation and dephosphorylation),and modulates signaling pathways in cells (e.g.,through adenosine monophosphate [AMP] or inosi-tol trisphosphate [IP3]) (Cooper 2000).Depending on the environment, the growth of

marine phytoplankton is typically limited by one ofthe major nutrients [phosphorous (P), nitrogen(N), and silicon (Si) (e.g., for diatoms)], and/orthe micronutrient iron [Fe] (Karl 2000, Paytan andMcLaughlin 2007, Moore et al. 2013). Growth

1Received 27 February 2015. Accepted 26 September 2015.2Author for correspondence: e-mail [email protected] Responsibility: M. Wood (Associate Editor)

J. Phycol. 52, 10–36 (2016)© 2015 Phycological Society of AmericaDOI: 10.1111/jpy.12365

10

limitation by P in the ocean occurs when bioavail-able P pools (orthophosphate ions [Pi = HPO4

2� +PO4

3�] and available dissolved organic P [DOP])drop below critical threshold concentrations relativeto levels of other required nutrients. Phytoplanktonpreferentially utilize Pi because it can be directlytaken up and assimilated to support algal metabo-lism and growth, while DOP generally requires con-version into Pi prior to its metabolic assimilation,which is more costly energetically (Falkowski andRaven 2007). However, when the external Pi pool isdepleted, phytoplankton growth often depends onthe ability to utilize the much more abundant DOPby its enzymatic hydrolysis to Pi.

The Pi concentration in the environment can bemeasured using standard colorimetric methods(Strickland and Parsons 1972) or the much moresensitive MAGIC coprecipitation method coupled tocolorimetry (Karl and Tien 1992). These methods,however, also typically measure some reactive DOPcompounds, and may overestimate the true dis-solved Pi concentration (Thomson-Bulldis and Karl1998, Laws et al. 2011a). Because of this, the Pimeasured by these methods is often referred to assoluble reactive phosphorus (SRP) with the knowl-edge that it likely includes some DOP and otherreactive P compounds, especially in low-Pi oceanicsurface waters with high DOP:Pi ratios (Fig. 1). Inthis review we will use what we believe is a moredescriptive term “measured Pi” for the SRP pool.The DOP concentration is determined operationallyby subtracting the initial measured Pi from total Pimeasured after oxidation of DOP to Pi using alka-line persulfate (Hosomi and Sudo 1986) or UVphoto-oxidation (Aminot and K�erouel 2001).

Dissolved organic P in the ocean consists of com-plex mixtures of compounds that have yet to be

fully characterized at the molecular level (Youngand Ingall 2010). It can largely be divided into twomajor groups of organic compounds: phosphoestersthat contain the C-O-P ester bond, and phospho-nates that contain the C-P bond. In analyses of highmolecular mass compounds (1–200 nm nominaldiameter), phosphoesters accounted for ~75% andphosphonates ~25% of DOP in ocean waters (Kolo-with et al. 2001). However, more recent analyses ofboth low and high molecular mass compoundsrevealed that 80%–85% of the measured DOP wascomprised of phosphate esters with the remainderconsisting of phosphonates (5%–10%) andpolyphosphates (8%–13%; Young and Ingall 2010).Because DOP is operationally defined as the differ-ence between total P and measured Pi, bothpolyphosphate esters and inorganic polyphosphateare included operationally in the measured DOP, aslikely are two other dissolved inorganic P (DIP) spe-cies: phosphite (PO3

3�) and phosphine (PH3).Although these latter two species have not yet beenchemically identified in seawater, their presence issuggested by the ability of the diatom Thalassiosirapseudonana to utilize dissolved PH3 (Fu et al. 2013)and presence of a phosphite transporter in thedinoflagellate 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 as0.2 nM in some surface waters of the Sargasso Seato 1–3 lM in upwelled water along the eastern mar-gins of the Atlantic and Pacific (Redfield et al.1963, Wu et al. 2000). Pi concentrations also canvary substantially over timescales ranging from hoursto seasons or even decades (Karl 2014). Further-more, large differences can occur in the growthdemand for P because of variations in specific

0 50 100 150 200 DOP (nmol L-1)

Atlantic Pacific

0

500

1000

1500

2000

2500

3000

3500

4000

0 1000 2000 3000

Dep

th (m

)

Measured Pi (nmol L-1)

Atlantic Pacific

FIG. 1. Vertical profiles ofmeasured Pi and DOP in theNorth Pacific and North AtlanticOceans (redrawn from Paytanand McLaughlin 2007 with thepermission of the AmericanChemical Society).

P GROWTH STRATEGIES AND MOLECULAR MECHANISMS 11

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growth rate linked to differences in temperature,growth-limitation by other nutrients (e.g., N andFe), and species-specific differences in maximumgrowth rates. Consequently, algal cells need thecapability to adjust their P uptake rates and storagecapacities to deal with these variations. To effectivelyexploit this variability in supply and demand cellsrequire coordinated cellular feedback systems thatcan simultaneously sense the Pi concentration atthe cell surface or within intracellular pools (e.g.,cytoplasmic Pi) and increase or decrease the num-ber and type of cellular Pi transporters and enzymesthat utilize DOP. Cells also require the capability tostore excess P when available and to draw uponinternal P stores as needed to support growth. Thisreview examines current knowledge concerning thephysiological and biochemical mechanisms andpathways by which phytoplankton sense, transport,assimilate, and store P.

The last decade has witnessed accelerated growthof information needed to understand how phyto-plankton respond to and cope with low-P stress andP-growth limitation (P-stress that reduces rates of cellgrowth and cell division) at the molecular, cellular,and ecological levels (Dyhrman et al. 2006a,Coleman and Chisholm 2010, Wurch et al. 2011,Dyhrman et al. 2012, Monier et al. 2012). This reviewintegrates new molecular discoveries with classicphysiological and biogeochemical studies to obtain abetter understanding of the mechanisms by which Pregulates cellular growth and species composition inmarine phytoplankton communities.

With the common coastal seasonal successionfrom a diatom-dominated spring bloom to a sum-mer dinoflagellate bloom in mind (Margalef 1978),this review focuses largely on dinoflagellates anddiatoms, with comparisons to other groups of phyto-plankton, including cyanobacteria. In addition,although this review deals mainly with P utilizationby marine phytoplankton, relevant insights were alsoderived from research on freshwater phytoplankton,other nutrients (e.g., N or Fe), and well-character-ized model bacterial molecular systems. Finally, it isbeyond the scope of this paper to exhaustively coverthe vast volume of literature on the physiologicalecology and biogeochemistry of P, for which readersare referred to other recent reviews (Falkowski et al.1998, Downing et al. 1999, Benitez-Nelson 2000,Elser et al. 2007, Paytan and McLaughlin 2007, Karl2014).

P AVAILABILITY AND GROWTH LIMITATION IN THE

OCEAN

In contrast to biologically available fixed N thathas a renewable source from N2 fixation by dia-zotrophic cyanobacteria, P in the global ocean ulti-mately comes from the slow process of rockweathering (Tyrrell 1999). However, its concentra-tion and chemical composition in marine waters

varies widely in time and space. As with all limitingnutrients, the abundance of P in the euphotic zone,where phytoplankton growth occurs, is determinedby the net balance between various local supply andremoval processes. Supply sources include terrige-nous and sedimentary inputs (coastal waters), localbiological regeneration, and resupply from deeperwaters containing high concentrations of Pi (Fig. 1)and other nutrients through upwelling or verticalmixing (Karl 2014). Dissolved Pi and DOP areremoved from euphotic waters by phytoplanktonand bacterial uptake, and the settling of planktonand biogenic detritus transfers this P to deeperwaters or to the sediments. This particulate organicP is then converted back into Pi by microbial degra-dation of the organic matter. The adsorption of Pionto the surfaces of iron oxides and other sinkingmineral particles also removes P from marine waters(Diaz et al. 2008).Due to its biological uptake in the euphotic zone

and regeneration in deeper waters, Pi is usuallydepleted near the surface and increases in concen-tration with depth (Fig. 1). In the subtropical NorthAtlantic, for instance, the measured Pi concentra-tion was 0.2–4 nM in surface waters and increasedby up to four orders of magnitude to 1.2 lM indeep waters (Wu et al. 2000). In contrast, DOP,which is much less readily utilized by phytoplank-ton, shows the opposite behavior and is highest nearthe surface owing to DOP inputs from the plank-tonic community (Paytan and McLaughlin 2007)(Fig. 1). Much of this DOP is composed of nucleicacids, free nucleotides, phospholipids, and phospho-rylated proteins and sugars released into the watercolumn by zooplankton grazing and excretion, virallysis of cells, plankton cell death, and bacterialdecomposition of organic matter (Young and Ingall2010). DOP concentrations in surface waters rangefrom 40 to 300 nM in the North and South Atlanticwith lowest values in the stratified subtropical watersof the North Atlantic during summer (Mather et al.2008). Because of the severe depletion of Pi in thesesubtropical waters, the large majority of total dis-solved P (90%–99%) exists as DOP (Wu et al. 2000,Lomas et al. 2010; Fig. 1).Marine primary productivity has traditionally been

characterized as being largely nitrogen- or iron-lim-ited (Falkowski et al. 1998, Downing et al. 1999,Tyrrell 1999, Moore et al. 2013). Historically, P-lim-itation was suspected in various areas in the globalocean (Rivkin and Swift 1982, Smith 1984), and thistopic has more recently seen renewed interest (Karl2000, Wu et al. 2000, Moore et al. 2008). Phospho-rus has been reported to occur at growth-limitinglevels for phytoplankton in the subtropical Atlantic(Rivkin and Swift 1979, Wu et al. 2000), Mediter-ranean Sea (Berland et al. 1972, Thingstad et al.2005), Red Sea (Fahmy 2003), and Gulf of Mexico(Hardison et al. 2013). High N:P ratios, elevatedalkaline phosphatase (AP) activity, molecular

12 SENJIE LIN ET AL.

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P-stress indicators, and nanomolar-measured Pi con-centrations all suggest that P limits phytoplanktongrowth in these regions (Fig. 2; Krom et al. 1991,Zohary and Roberts 1998, Wu et al. 2000, Kromet al. 2004, Thingstad et al. 2005, Bj€orkman et al.2012). However, recent nutrient addition experi-ments suggest that in the low-Pi waters of the sub-tropical Atlantic, phytoplankton growth is stillprimarily limited by N or colimited by N and P(Moore et al. 2008, 2013). But even when N is pri-marily limiting, P likely plays an important role incontrolling species composition in low-Pi oceanicregions (Moore et al. 2008).

Nitrogen is primarily limiting in the oceanbecause of iron limitation of cynaobactrial N2 fixa-tion, an iron-dependent metabolic process thatreplenishes oceanic inventories of fixed N by enzy-matically reducing N2 gas to ammonium (Sohmet al. 2011, Sunda 2012). However, in regionsreceiving high inputs of iron from aeolean dustdeposition (e.g., the subtropical North Atlantic),the elevated iron input rates fuel higher rates of N2-fixation and associated C-fixation, which drive thesesystems toward P-limitation (Benitez-Nelson 2000,Wu et al. 2000, Sa~nudo-Wilhelmy et al. 2001, Millset al. 2004, Dyhrman et al. 2006a, Meseck et al.2009, Paytan and McLaughlin 2007).

Concentrations of available P and other nutrientsare generally higher in coastal waters, but can varywidely in time and space with variations in nutrientinputs and algal growth. Algal growth in these sys-tems, where P concentration is typically higher and

more dynamics than the open ocean, can becomeP-limited from excess inputs of anthropogenic Nfrom N-rich fertilizers, municipal wastes, and NOx

from the burning of fossil fuels, which increases N:Pratios in the receiving waters (Cloern 2001, Huanget al. 2003, Scavia and Bricker 2006, Sylvan et al.2006, Zhang et al. 2007). Examples include manyeutrophic estuaries such as Chesapeake Bay (Fisheret al. 1992, 1999, Kemp et al. 2005) and Pearl RiverEstuaries (Huang et al. 2003, Xu et al. 2008),coastal regions of the Gulf of Mexico such as thosereceiving N-rich nutrient inputs from the MississippiRiver (Laurent et al. 2012, Turner and Rabalais2013), and various Chinese coastal regions (the EastChina and Yellow Seas; Harrison et al. 1990, Zhanget al. 2007, Fu et al. 2012, Fig. 2). These anthro-pogenic inputs are largely due to riverine or atmo-spheric sources, but in some coastal regions such asLong Island Sound, USA, N can be introduced byinputs of N-enriched ground water (Slomp and VanCappellen 2004).

CELLULAR PI UPTAKE AND ASSIMILATION

Fundamental constraints. The uptake of Pi by phy-toplankton cells is ultimately governed by three fun-damental constraints: the ambient Pi concentration;cell size and shape (which determines the cell’s sur-face to volume ratio and the thickness of its diffu-sive boundary layer); and the density, bindingaffinity, and turnover rate of the Pi transport pro-teins 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 phosphataseactivity (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).


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combination these factors impose an absolute limiton the rate at which Pi and other nutrients can besupplied to the cell surface and transported into thecell.

The external Pi concentration directly controlsthe net nutrient flux toward the cell surface (Hud-son and Morel 1993). That flux, however, must firstdiffuse through an aqueous boundary layer prior toreaching the cell surface (Pasciak and Gavis 1974).The rate of diffusion through the boundary layernormalized to cell volume varies with the inversesquare of the cell diameter, and consequently, nutri-ent uptake in larger cells is much more diffusionlimited than that in smaller cells (Pasciak and Gavis1974, Sunda and Hardison 2010). Thus, diffusionlimitation selects for smaller cells in nutrient-limitedenvironments such as open ocean surface waters(Chisholm 1992).

Once a nutrient such as Pi reaches the cell sur-face, it is transported into the cell by binding totransmembrane uptake proteins in the pla-malemma, the bilayer membrane surrounding thecytoplasm. P uptake is an energy consuming pro-cess that is often fueled directly or indirectly byATP, and like other protein-assisted nutrient uptakesystems, it is generally governed by the Michaelis–Menten equation (Gotham and Rhee 1981, Rhee1973):

VS ¼ Vmax½S �=ðKs þ ½S �Þ ð1Þwhere [S] is the extracellular substrate (Pi) concen-tration adjacent to the outer surface of the plasmamembrame, Ks is the half saturation constant (equalto the substrate concentration at which VS = 0.5Vmax), and Vmax is the maximum uptake rateachieved when [S] ≫ Ks. Vmax is equal to the num-ber of transport proteins on the cell membrane(normalized for example to cell volume or carbon)times the rate of intracellular transport and sub-strate release within the cell per transport protein(Hudson and Morel 1993). Ks provides a relativemeasure of the binding affinity of the uptake systemfor Pi or other nutrients (Dugdale 1967).

Transport proteins generally have a range of sub-strate binding affinities (i.e., Ks values) but are typi-cally divided into low- and high-affinity systems.Low-affinity transporters bind nutrients relativelyweakly. Consequently, they have high rates of disso-ciation back into the medium and release into thecell, and high Vmax values per individual transporter(Hudson and Morel 1993). These systems have highKs values and work most efficiently at high substrateconcentrations. Conversely, high-affinity transportsystems bind the substrate more tightly and releaseit into the cell more slowly. They exhibit lower Ks

and Vmax values and operate most efficiently at low-substrate concentrations. Thus, for a fully saturatedtransport system, a much higher number of high-affinity transporters are needed to yield the same

intracellular transport rate, compared to the num-ber required for a low-affinity transport system. As aresult, high-affinity transporters are inherently lessefficient at high substrate concentrations, but arenecessary for effective nutrient uptake at low con-centrations.Physiological studies of Pi uptake. Before the advent

of modern molecular methods, our understandingof P uptake in phytoplankton came primarily fromphysiological studies that used the Michaelis–Men-ten equation (eq. 1) to analyze Pi uptake kinetics(Gotham and Rhee 1981, Rhee 1973). In caseswhere Pi diffusion to the cell surface limits Piuptake rates, the Pi concentration at the cell surfaceis lower than that in the bulk solution and eq. 1needs to be modified to relate the uptake velocity Vsto the bulk solution Pi concentration (Laws et al.2013, Pasciak and Gavis 1974). Studies using bothkinetic models have consistently shown that cellsexhibit distinctly different Michaelis–Menten uptakecurves depending on the degree of physiologicalstress due to low Pi (referred to hereafter as low-Pistress) (Fig. 3). Most algal P uptake studies haveexamined only a narrow range of Pi concentrationsand fitted the data to a single Michaelis–Mentenequation. As the external Pi concentration and cellP-quota declined in these studies, there was nochange in the Ks value, but up to a 30-fold increasein the observed Vmax (Gotham and Rhee 1981, Jau-zein et al. 2010, Riegman and Mur 1984). In otherstudies with a wider range of Pi concentrations inthe medium, the cellular Pi uptake was consistentwith the presence of two or more transport systemswith differing Ks values, and thus, varying Pi affini-ties (Chisholm and Stross 1976, Rivkin and Swift1982, Spijkerman and Coesel 1996, Tomas 1979).The cells’ high-affinity systems appear to be up-regu-lated under low-Pi stress, which provides cells withthe ability to acquire Pi when external concentra-tions are low, but at the price of lower Vmax valuesper transporter as noted above. The exact nature ofthe negative feedback mechanism(s) by which the

[Pi]

Upt

ake

rate

Low affinity transporter used at high Pi

High affinity transporter upregulated at low external Pi and low

cellular Pi

Ks1 Ks2 Low

High

FIG. 3. Changes in uptake kinetics associated with differentialexpression of high and low affinity P-transporters in the cytoplas-mic membrane.


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internal P pool(s) or external Pi concentrations reg-ulate high- and low-affinity intracellular Pi transportsystems in eukaryotic phytoplankton, however, hasyet to be determined at the molecular level.

The diel light–dark cycle and associated dailygrowth cycle also affect the pattern of cellular Piuptake. For most species, Pi uptake rates increaseduring the day and decrease at night consistent withthe higher growth demand for P during daytimefrom light driven photosynthetic C-fixation(Chisholm and Stross 1976, Rivkin and Swift 1982,Ahn et al. 2002). Kinetic analysis indicates thatthese diel changes in uptake rates are often linkedto shifts in Vmax values rather than to variations inKs (Chisholm and Stross 1976, Rivkin and Swift1982). Thus, the total number of transporters onthe cell membrane likely varies over the L:D cycleto accommodate changes in P demand linked todiel changes in C-fixation and the cell division cycle.In the environment, diel changes in Pi uptake ratesmay also be caused by day/night differences in Piconcentrations, particularly in highly productive sys-tems where algal biomass and specific growth ratesare high and the cycling of Pi may be on the orderof hours (Nixon et al. 1976). Such diel changes inPi uptake rates, though experimentally inconve-nient, should not be overlooked, and may signifi-cantly impact ecosystem function.

An important unanswered question is how fastcells shift the relative abundance of low- and high-affinity membrane Pi transport systems in responseto variations in ambient nutrient concentrationsand growth demand for P. As Pi transport proteinsare identified in marine phytoplankton (see below),it may be possible to address this question if anti-bodies specific to high- and low-affinity Pi trans-porters can be developed. These antibodies couldbe used in conjunction with confocal light micro-scopy to document changes in the density of differ-ent transporters with changes in Pi concentration atthe cell surface and in intracellular P pools. Quanti-tative reverse transcription PCR an also be used tomeasure differential expression paterns of the dif-ferent P transporter genes, but only for species inwhich the transporter protein abundances are regu-lated at the transcriptional level.Molecular characterization of Pi transport sys-

tems. Current molecular work has begun to revealthe identity of some of the Pi transport proteins inmarine microorganisms that are responsible for theuptake kinetics noted above. Such uptake systemshave been best characterized in bacteria and areoften evolutionarily conserved (Pedersen et al.2013), so we will begin our discussion with adescription of bacterial Pi transporters. As appearsto occur in most microorganisms, heterotrophicbacteria contain both a low-affinity Pi transporter(PiT) that functions at high Pi concentrations and ahigh-affinity Pi transporter (PsT) which is up-regu-lated under low-Pi stress (van Veen 1997). Homo-

logs of these high- and low-affinity transporters havebeen found in marine cyanobacteria such asProchlorococcus (Martiny et al. 2006). Eukaryoticequivalents of PiT have been identified and includethe Pi transporter IPT and the sodium- or sulfate-dependent Pi transporter SPT (Fig. 4; Table 1). SPTis a symporter, which simultaneously transports Piand sodium or sulfate across the cell membrane.Recent research in the Lin laboratory revealed thepresence of putative homologs of IPT and SPT inthe genome of the dinoflagellate S. kawagutii (acces-sion number SRA148697 in NCBI SRA database)and in the transcriptome of another dinoflagellateProrocentrum donghaiense (GenBank no. KJ699385,KJ699384). IPT homologs were also found in thetranscriptomes of Karlodinium veneficum and Amphi-dinium carterae (KM881476, KM881477). Similarly,IPT homologs have been observed in the diatomsThalassiosira pseudonana and Phaeodactylum tricornu-tum (Bowler et al. 2008); the haptophyte Emilianiahuxleyi, the prasinophyte Ostreococcus spp., themamiellophytes Micromonas sp. and Batycoccus sp.(Monier et al. 2012, Worden et al. 2009); and thepelagophyte Aureococcus anophagefferens. It has alsobeen observed in unidentified eukaryotes detectedin the Global Ocean Sampling metagenomic dataset (Table 1). The IPT gene sequences from thesediverse eukaryotic algae are not strictly conservedsuggesting the homologs were derived from a com-mon ancestor that subsequently diverged as newgroups evolved. Eukaryotic phytoplankton virusescarrying IPT gene sequences have also been identi-fied (Lindell et al. 2004, Monier et al. 2012) andmay provide a mechanism for host phytoplanktonto acquire novel Pi transporter genes. Recombina-tion aided by viral transfer may therefore have con-tributed to the observed IPT gene diversification.Though less likely, another possibility is that conver-gent evolution of different genes encoding proteinswith IPT function produced the divergent IPThomologs.Comparable screening of cDNA libraries has

revealed only a few eukaryotic high-affinity PsTequivalents (Table 1). One of these is the high-affinity Pi transporter (PHO) identified in theprasinophyte Tetraselmis chui whose transcriptionalup-regulation under P-limitation was confirmedexperimentally (Chung et al. 2003). Using theamino acid sequence of PHO in T. chui as a queryin tBLASTn against the expressed sequence tag dataset in NCBI revealed a homologous gene in thedinoflagellate Alexandrium minutum (GenBank acces-sion number GW800973). S. Lin et al. (unpublisheddata) also identified a PsT homolog (KJ699386)from a transcriptome of Prorocentrum donghaiensegrown under Pi limitation. However, the function ofthe encoded protein as a high-affinity Pi transporterremains to be verified. In addition, a high-affinitytransport protein, phosphate-repressible phosphatepermease, was identified in P-limited cultures of the


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http://www.ncbi.nlm.nih.gov/nuccore/SRA148697

http://www.ncbi.nlm.nih.gov/nuccore/KJ699385


http://www.ncbi.nlm.nih.gov/nuccore/KM881476

http://www.ncbi.nlm.nih.gov/nuccore/KM881477

http://www.ncbi.nlm.nih.gov/nuccore/GW800973


haptophytes Emiliania huxleyi (Dyhrman et al.2006a) and Prymnesium parvum (Beszteri et al.2012). In gram-negative and gram-positive bacteria,archaea, and eukaryotes, phosphate permeases areH+ or Na+ symporters.

In yeast and land plants Pi uptake systems havefurther diversified and include two high-affinityPho84 and Pho89 Pi transporters, which are up-regulated under low-Pi stress, and two low-affinityPho87 and Pho90 transporters up-regulated at highPi concentrations (H€urlimann et al. 2009). Furtherinvestigation showed that the SPX domain in

Pho84, 87, 89, and 90 also plays a role in modulat-ing Pi uptake rates, suppressing Pi cellular efflux,and in regulating the phosphate signal transductionpathway (Secco et al. 2012). The extent to whichtransporters have diversified in different eukaryoticphytoplankton groups, and whether or not theycontain a similar SPX regulatory domain to main-tain P homeostasis remains largely unexplored. Asearch of the recently sequenced genome of thedinoflagellate S. kawagutii (Lin et al. 2015a; see Sup-plementary Table 33) reveals the presence of vari-ous proteins containing the SPX domain, including

R-OR-C

[Pi]

Outer cell membrane

PhnCDE

[Pi]

Promoter proteins PhoB/PhoP/etc.

(Up and down regulate Pi transporters)

PhosphoestersR-O-PO3

Low affinity Pi transporters

PhoR

Phosphonates(R-C-PO3)

R-O-PO3Transporter

Increase membrane sulfolipid to phospholipid

ratio under P stress

Up regulation of AP under low-P stress±Pi Internal Pi

Ene

rgy

R-O

R-C-PO3R-O-PO3

Cytoplasmic membrane

CP LyaseAP

(PhoA et al.)

PsTPiT/SPT

PiT/IPT

AP (PhoD, PhoX et al.)

High affinity Pi transporters

Metabolic P compounds (Nucleic acids,

phospholipids, etc.)

Phosphoester(R-O-PO3)

ADP

Phagotrophy (bacteria and

phytoplankton)

ATP

PPX PPK

PolyP

ADP

FIG. 4. Molecular mechanisms of phosphorus uptake, metabolism, and storage. This diagram represents a composite of known bacte-rial and eukaryotic phosphate acquisition and metabolic pathways which are likely to be present in some form in most phytoplankton spe-cies. Black arrows indicate pathways by which P is transported or metabolized. Red dashed arrows indicate regulatory systems that up- ordown-regulate specific proteins or protein complexes involved in the transport or metabolism of P. Cylinders indicate transporter proteincomplexes, ovals – specific phosphate pools, squares – phosphate-processing enzymes or regulatory proteins, and polygons – eitherphagotrophy or increases in the sulfolipid content in the membrane in response to low-P stress. Pi uptake involves low-affinity transporters(PiT) and high-affinity transporters (PsT). PiT includes subtypes of inorganic phosphate transporter (IPT) and ion-phosphate cotrans-porter or symporter (SPT). DOP uptake must involve transporters for phosphomonoesters (as yet unidentified) and phosphonates(PhnCDE in cyanobacteria). Alkaline phosphatase (AP) localized on the cell surface (or in the periplasm of cyanobacteria) hydrolyzesexternal phosphomonoesters to Pi followed by intracellular uptake of the released Pi. Inside the cell, phosphomonoesters are hydrolyzedby intracellular APs (or acid phosphatases) to simultaneously release Pi and R-O compounds while phosphonates are cleaved by phospho-nate lyase (in cyanobacteria) to release Pi and various R-C compounds. Intracellular Pi is assimilated via the photosynthetic and respiratoryformation of ATP, which is utilized as the major energy source for cell metabolism and as a phosphorus and energy source for the forma-tion of P-containing cellular moleculrs (e.g., phospholipids, DNA, RNA, and nucleotides). In cyanobacteria, ATP is converted by theenzyme polyphosphate kinase (PPK) into polyphosphate (polyP) with simultaneous release of ADP. PolyP serves as both a major cellularphosphate and an energy storage pool, and can be converted back into ATP (with reconsumption of ADP) by PPK. A PPK homologremains to be identified in eukaryotic phytoplankton, and here the formation of polyP is catalyped by other enzymes (e.g., Vtc 1-4). PolyPcan also be hydrolyzed to Pi by exophosphatase (PPX) in cyanobacteria and eukaryotic algae. Phagotrophy of bacteria and other phyto-plankton as well as potential endocytosis of surface adsorbed P represent further mechanisms for acquiring P in eukaryotic algae. UnderP-limitation of growth, many species can also increase the membrane sulfolipid to compensate for the reduction in phospholipid content.The present diagram is for a cyanobacterium, which in addition to the plasmamembrane, also contains a semipermeable outer membrane(shown here as a dotted line). By contrast, eukaryotic cells do not have an outer cell membrane, but instead usually possess an outer cellwall or cell plates (e.g., coccoliths in many prymnesiophytes).


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a sugar-phosphate exchanger (Skav202903). Thisgene belongs to the major facilitator superfamily,and is a sugar-phosphate antiporter that likely trans-ports sugar out of the cell and phosphate into thecell.Pi sensing and uptake regulatory systems. Up- and

down-regulation of Pi transport sytems is mediatedin microorganisms by two-component (sensor andresponse regulator) signal transduction systems(Wanner 1996, Stock et al. 2000). One componentof the system senses Pi at the cell surface or in thecell’s cytosol and the other regulates the expressionof various high and low-affinity Pi transport proteinsand other Pi acquisition proteins such as phos-phatases (Dick et al. 2011). One such regulatory sys-tem has been well-characterized at the molecularlevel in gram-negative bacteria. It consists of theprotein pair PhoB–PhoR, and is referred to as thePho regulon (Vershinina and Znamenskaya 2002).In this bacterial group, pores (porins) in the outermembrane allow Pi to diffuse into the periplasmand bind a periplasmic receptor site on PhoR,which is a transmembrane protein located in the

cytoplasmic membrane. As Pi in the environmentdeclines, concentrations in the periplasm drop,causing Pi to dissociate from PhoR. This dissocia-tion results in a conformational change in the pro-tein, which causes an intracellular kinase domain ofPhoR to transfer a phosphate group from ATP ontoanother cytoplasmic site on PhoR. This phosphate issubsequently used to phosphorylate the transcrip-tion regulator PhoB (response regulator) in thecytoplasm allowing it to bind regulatory regions ofDNA. This binding initiates transcription of thegenes involved in the synthesis of intracellular Pitransport proteins, and in many cases, thoseinvolved in the utilization of DOP (e.g., those cod-ing for APs; see section below). As Pi levels increase,the periplasmic receptor site on PhoR rebinds to Piand the process and resultant gene expression isdown-regulated.Similar regulatory systems are found in cyanobac-

teria (which are also gram-negative bacteria; Hiraniet al. 2001), archaea (Osorio and Jerez 1996), yeasts(Dick et al. 2011, Magbanua et al. 1997) and theroots of land plants (Dong et al. 2013, Ticconi and

TABLE 1. Protein or protein complexes involved in phosphorus acquisition in phytoplankton that have been recognized todate.

Species

Low-affinity DIP transporters (PiT-like)High-affinity DIP transporters

(PsT-like)

Inorganicphosphatetransporter

Pho4superfamily

Sodium-dependentphosphatetransporter

Phosphate/sulfate

permease

High-affinityphosphatetransporter

Phosphate-repressiblephosphatepermease

DinophytaAlexandrium catenella This studyAmphidinium caterae This studyKarlodinium veneficum This studyProrocentrum donghaiense This study This studyBacillariophytaThalassiosira pseudonana Bowler

et al. (2008)Phaeodactylum tricornutum Bowler

et al. (2008)HaptophytaEmiliania huxleyi Dyhrman

et al. (2006a)Dyhrmanet al. (2006a)

Prymnesium parvum Beszteriet al. (2012)

Beszteriet al. (2012)


PelagophytaAureococcus anophagefferens Wurch

et al. (2011)Aureoumbra lagnunaChlorophytaChlamydomonas reinhardtii Moseley

et al. (2006)Micromonas Worden

et al. (2009)Ostreococcus Worden

et al. (2009)Tetraselmis chui Chung

et al. (2003)CyanophytaProchlorococcus Martiny

et al. (2006)Synechodoccus Scanlan

et al. (2009)Trichodesmium


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Abel 2004). In addition, a second tier regulatory sys-tem dependent on PtrA (potential transcriptionalregulator) was identified in the cyanobacteriumSynechococcus (Ostrowski et al. 2010). In this system,low Pi concentration first triggers the PhoB-inducedresponse (a tier 1 response), leading to elevatedexpression of high-affinity Pi transporters. Whenthis response does not sufficiently alleviate low-Pistress, PtrA expression is increased. The higher PtrAprotein pool then binds to the promoters of phos-phatase (e.g., AP) genes, which up-regulates phos-phatase synthesis and increases cellular utilization ofDOP (a tier II response).

Recently, a response regulator receiver genecoding for a protein that is potentially part of a two-component P-response regulatory system was identi-fied in the proteome of the dinoflagellate Kareniamikimotoi (Lei and Lu 2011). Screening of genomicdatabases will likely reveal the presence of similarP-sensing/regulatory systems in many other eukary-otic phytoplankton. Determining the details of howthe Pi signal transduction pathways are regulated ineukaryotic algae, as well as how these pathways arecoregulated by internal P-pools, will likely prove animportant line of research in the future. However,these pathways may be substantially different indinoflagellates because much of the gene expres-sion is regulated post transcriptionally rather thanby direct regulation of RNA synthesis (for review seeLin 2011).Phosphate assimilation. Exactly how P is assimilated

into biomolecules needed for growth, metabolism,and cell division following cellular uptake of Pi orDOP or P acquisition by phagotropy has not beenextensively investigated in phytoplankton. MostDOP taken up into the cell must first be convertedto Pi and this may also be the case for acquisiton ofP by phagotrophy, which may involve a suite of lar-gely uncharacterized phosphatases. The majorbiosynthetic pathway for Pi assimilation in all cells,including phytoplankton, is the photosyntheticand/or respiratory production of ATP from Pi andadenosine diphosphate (ADP) via the enzyme ATPsynthase (Fig. 4). ATP is the major energy currencyof the cell and not only supplies energy for the syn-thesis of various organic biomolecules (e.g., in theCalvin–Benson cycle) but also supplies phosphatefor the synthesis of numerous phosphate-containingend product molecules such as phospholipids,nucleotides, polyphosphates, and phosphorolatedsugars and proteins. The assimilation of phosphateinto ATP takes place in three cellular compart-ments: the chloroplast, where ATP is a major pro-duct of photosynthesis; mitochondria, where it is amajor product of respiration; and in the outer cellmembrane, where it is synthesized by the light-acti-vated proton-pump proteorhodopsin. While thephotosynthetic and respiratory ATP synthesis sys-tems are universal in phytoplankton and indeed allphototrophs (Falkowski and Raven 2007), the puta-

tive energy-converting proteorhodopsin system isbest documented in certain marine bacteria (Fuhr-man et al. 2008 and references therein) and hasonly begun to be examined in eukaryotes, includingdinoflagellates (Lin et al. 2010, Guo et al. 2014, Shiet al. 2015), two diatoms, and a haptophyte (March-etti et al. 2012). The algal homologs of this proteinare similar to that in proteobacteria, where the pro-tein harvests solar energy and generates a protongradient across the plasma membrane for the pro-duction of ATP via the enzyme ATP synthase or tofuel the intracellular uptake of Pi or other smallnutrient molecules (e.g., via membrane symporters;B�ej�a et al. 2001, Fuhrman et al. 2008).P storage as polyphosphate. Phytoplankton are cap-

able of storing excess intracellular phosphate notneeded immediately to support cell metabolism andgrowth, such as that taken up at sustained high con-centrations or pulses of external Pi. The stored Pcan then support high population growth rates formultiple generations under subsequent low P condi-tions (Droop 1973, Ducobu et al. 1998, Morel1987). The major known mechanism for storing Pin phytoplankton (and indeed all organisms) is theformation of polyphosphate (polyP; Fig. 4), whichconsists of linear chains ranging from several tohundreds of phosphate residues linked by high-energy phosphoanhydride bonds (Kornberg et al.1999). Because of the high energy of the phospho-anhydride bonds, polyP is utilized by cells not onlyfor phosphate storage but also for energy storage,and can be used as a source of ATP by its enzymaticreaction with ADP (Kornberg et al. 1999, Achberg-erov�a and Nah�alka 2011):

PolyP(n)þ ADP $ PolyPðn� 1Þ þ ATP ð2Þ

where n is the number of phosphate residues in thepolyP chain. PolyP formation occurs in all organ-isms (Kornberg et al. 1999), including phytoplank-ton (Rhee 1973, Elgavish et al. 1982, Rivkin andSwift 1985).PolyP formation from ATP, and the reverse reac-

tion to reform ATP (eq. 2), is catalyzed by polyphos-phate kinase (PPK) in heterotrophic bacteria andcyanobacteria (Fig. 4), but this protein has not beenfound in eukaryotic cells (Kornberg et al. 1999,Rocap et al. 2003). In addition, two other enzymes,exophosphatase (PPX) and endophosphatase, cata-tyze the hydrolysis of polyP to Pi in bacteria andeukaryotes (Kornberg et al. 1999) (Fig. 4). In yeastand other eukaryotes polyP often occurs in vacuolesand its formation involves the vacuolar transporterchaperone (Vtc) 1–4 enzyme family (Ogawa et al.2000). Homologs of two Vtc family genes have beenidentified in the genome of the diatom Thalassiosirapseudonana (Dyhrman et al. 2012). Surprisingly,transcriptomic and proteomic analysis indicated thatthe Vtc 4 homolog is up-regulated under P limita-tion of growth rate, the opposite of what would be


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expected if polyP was merely a P-storage molecule(Dyhrman et al. 2012). Nuclear magnetic resonanceanalysis verified higher polyP levels in the P-limitedT. pseudonana cells and similar elevated polyP levelshave been observed in P-limited cells of the bac-terium Escherichia coli (Kornberg et al. 1999). Theseincreases appear to be a response to nutrientgrowth limitation in general as poly-P accumulationalso occurs when the growth rate of bacteria (Korn-berg et al. 1999) and diatoms (Perry 1976) are lim-ited by N. These findings appear to agree withrecent observations in low DIP and dissolved inor-ganic nitrogen (DIN) waters of the Sargasso Sea,where unexpectedly high polyP to particulate Pratios were observed, even though the systemappeared to be P-stressed based on high particulateAP activities and high levels of phospholipidreplacement by sulfolipids (Martin et al. 2014).These unexpected findings may be explained by themany other cellular functions of polyP other than Pstorage: particularly nutrient stress responses, energystorage, and storage of essential nutrient metalssuch as iron, which has been observed in yeast(Lesuisse and Labbe 1994).Surface phosphate adsorption. In addition to its

assimilation into cellular biomolecules, phosphatecan also adsorb directly onto the surface of phyto-plankton cells (Sa~nudo-Wilhelmy et al. 2004). Thisadsorbed phosphate can account for 14%–90% oftotal cell P and must be removed if true cellular Plevels are to be measured (Sa~nudo-Wilhelmy et al.2004, Fu et al. 2005). This adsorbed phosphate canbe removed with oxalate washes which dissolve andremove ferric (Fe[III]) oxyhydroxides and man-ganese (Mn[III and IV]) oxides via their reductionto soluble ferrous ions (Fe[II]) and manganous ions(Mn[II]) (Sa~nudo-Wilhelmy et al. 2004). Theadsorbed phosphate apparently is associated withthe oxides which precipitated on the cell surface(e.g., by oxidation of Mn (II) and Fe(II)), and areknown to strongly adsorb phosphate. A strong corre-lation between cell surface phosphate and Mn oxi-des (r2 = 0.81) supports the oxide adsorptionhypothesis (Sa~nudo-Wilhelmy et al. 2004). Theadsorbed phosphate may play an important, yetpoorly defined role in intracellular P uptake, partic-ularly in environments with fluctuating concentra-tions of Pi, Mn, and Fe (Fu et al. 2005). On the onehand, the abiotic adsorption of Pi on Mn and Feoxides removes Pi at the cell surface that wouldotherwise be available for intracellular transport.This could be particularly problematic for largercells where the diffusive flux of Pi to cell surface isalready severely limited. However, algal cells possessmechanisms to reductively dissolve Mn and Fe oxi-des (e.g., transmembrane reductases; Sunda 2012),which could release adsorbed Pi into solution forsubsequent uptake by the cell. Thus, if cells are ableto utilize this or other mechanisms (e.g., phagotro-phy) to take up the adsorbed phosphate, it could

function as an external phosphate storage pool thatcould be used to support cell growth during low-Pstress (Fu et al. 2005). The fraction of cellular Ppresent as surface-adsorbed phosphate increaseswith the external Pi concentration (Sa~nudo-Wil-helmy et al. 2004, Fu et al. 2005), which could makesuch an external storage mechanism particularlyeffective. Such possibilities clearly warrant furtherinvestigation, particularly from a biochemical andmolecular perspective.

ADAPTATION AND ACCLIMATION RESPONSES TO LOW-PSTRESS

In addition to the up-regulation of high-affinityPi uptake systems, phytoplankton show a variety ofother adaptation and acclimation responses to low-P stress. These include the substitution of sulfatefor phosphate in membrane lipids, the utilizationof DOP via hydrolytic enzymes, and the acquisitionof P via phagatrophic consumption of othermicroorganisms (Dyhrman et al. 2007, Hartmannet al. 2012). The specifics of these various low-P-coping mechanisms are discussed in the followingsections.Reducing cellular P demand. In many marine

cyanobacteria, particularly those residing in low-Poceanic waters, some phospholipids in cell mem-branes are replaced by sulfonated lipids to reducecellular 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 alsobeen found in the brown tide pelagophyteA. anophagefferens (Wurch et al. 2011). In addition,both A. anophagefferens and the dinoflagellateK. mikimotoi adjust their glycolytic pathway underlow-P stress to utilize alternate enzymes that requireless P, which enhances the ability of these species togrow in low-P environments (Lei and Lu 2011,Wurch et al. 2011). The molecular underpinning ofthe phospholipid-to-sulfolipid shift in phytoplanktonunder low-P stress remains to be elucidated.Utilization of phosphoesters via alkaline phos-

phatase. As Pi levels decline with algal growth, DOPconcentrations increase due to its release from thebiological community. So one of the most importantmechanisms for coping with low-Pi stress is the uti-lization of DOP. There is increasing evidence thatDOP is an important source of P to phytoplanktonin low-Pi regions, such as the Sargasso Sea whereDOP:Pi ratios in surface waters can exceed 100 (Wuet al. 2000, McLaughlin et al. 2013). For example,30% of primary production during the springbloom in the North Atlantic subtropical gyre wasestimated to be supported by DOP (Mather et al.2008). And 17%–82% of the P taken up by phyto-plankton in the Sargasso Sea is estimated to havebeen supplied from DOP (McLaughlin et al. 2013).However, the partitioning of P utilization betweenPi and DOP in stratified surface ocean waters with


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combined low-Pi and high-DOP concentrations isdifficult because true Pi values can be much lowerthan measured Pi levels (as discussed previously)and most of the DOP that is utilized must first beconverted to Pi by cell surface and extracellularAPs. Among the various forms of DOP, some studiessuggest that nucleotides are utilized preferentially(Wang et al. 2011) and are important in supportingphytoplankton growth in oceanic waters (Bj€orkmanand Karl 2005).

The most important DOP utilizing enzyme is AP,which hydrolyzes organic monophosphate esters toPi, often at the cell surface (Labry et al. 2005,Nicholson et al. 2006, Huang et al. 2007, Duhamelet al. 2010, 2011, Fig. 4). The released Pi is thentaken up intracellularly by Pi transport proteins. APis substantially up-regulated in low-P-stressed algalcells, allowing them to acquire Pi from extracellularDOP pools (Dyhrman et al. 2012). The activity ofAP 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 apH of 8.2 during preindustial times (Sunda and Cai2012). However, current and future ocean acidifica-tion from anthropogenic increases in atmosphericcarbon dioxide are decreasing surface ocean pH val-ues (Feely et al. 2009), which could potentiallydecrease the activity of AP in ocean waters, andthus, decrease the utilization of DOP by phytoplank-ton. Since DOP is thought to be a major source ofP to phytoplankton in ocean waters, this couldadversely affect P utilization and algal growth in thefuture ocean.

The enzymatic activity of AP has been widely uti-lized as an indicator of P stress (Dyhrman and Rutten-burg 2006, Lomas et al. 2010). To measure APactivity, phytoplankton and other microorganismsare incubated with a phosphoester substrate analogof AP to generate a product that can be measured flu-orometrically or colorimetrically, depending on thechemical nature of the added substrate (Gonz�alez-Gilet al. 1998). For the colorimetric assay, p-nitrophenylphosphate is used as a phosphatase substrate, whichturns yellow (kmax = 405 nm) when dephosphory-lated by AP. The substrates used for fluorescentassay include 2-(50-chloro-20-phosphoryloxyphenyl)-6-chloro-4-(3H)-quinazolinone (also known asenzyme-labeled fluorescence ELF-97� or ELF), 3-0-methylfluorescein phosphate, 3,6-fluorescein diphos-phate, and 4-methylumbelliferyl phosphate. Of theseELF-97 gives an insoluble fluorescent precipitate,allowing microscopic observation of the cellular andsubcellular localization of AP (Gonz�alez-Gil et al.1998). Bulk AP activity in a sample can be measuredusing a multiwell plate reader while the AP distribu-tion among cells can be measured with a flow cytome-ter. The quantitative AP activity is usually normalizedon a per cell basis, but a recent study showed thatnormalizing it to light absorbance at 450 nm, a proxyof algal cell biomass, increases the statistical power

and simplifies sample-handling (Peacock and Kudela2012).Compared to diatoms, dinoflagellates generally

exhibit higher AP activities on a per cell C or bio-volume basis. In a study in Monterey Bay, Califor-nia dinoflagellates accounted for the majority ofAP activity measured using the ELF substrate eventhough diatoms were dominant (Nicholson et al.2006). A similar trend was shown in a study con-ducted in the Taiwan Strait in August 2004 andMarch 2005 where the average ELF staining ratewas 75 � 16% for dinoflagellates and 29 � 19%for diatoms (Ou et al. 2006). The percentage ofELF labeling can also vary among dinoflagellatesranging from 17%–21% for Gonyaulax and Dinoph-ysis spp. to 82%–84% for Protoperidinium spp. andK. mikimotoi in the East China Sea (Huang et al.2007). These results indicate a wide variability inAP expression among species in response to low-Pstress.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 fordinoflagellates compared to 0.25–50 lM for othergroups of phytoplankton, with no clear lineage-based differences, although the highest thresholdsfor inducing AP activity tend to be in diatoms(Table 2). Some of these values exceed the maxi-mum Pi concentrations in ocean waters (2–3 lM).Studies show that AP activity is controlled more byintracellular P pools than by external Pi concentra-tions (Elgavish et al. 1982). This internal regulationcan complicate accurate determination of thresholdPi values and may account for the higher values forAP induction observed in Table 2. To complicatematters further, in some freshwater epiphytic algaeAP was expressed constitutively, even when Pi was1 mM (Young et al. 2010). Another potential issueassociated with the wide range of Pi thresholds isvariation associated with different methods fordetection of AP thresholds. For example, flowcytometer-based methods are often more sensitivethan those using a regular fluorometer, resulting indifferent threshold estimates (Jauzein et al. 2010).While AP activity is widely measured, relatively lit-

tle effort has been made to elucidate AP genesequences or regulation of gene expression in mar-ine eukaryotic phytoplankton (Lin et al. 2012a,b,2013, 2015b). Overall, AP gene sequences are highlyvariable among different microorganisms and thosefor heterotrophic bacteria, cyanobacteria, andeukaryotic algae can hardly be aligned, even at theamino acid level (Lin et al. 2012b). The high-sequence variability suggests rapid divergence ofgene homologs or converging evolution of differentAP genes, as in the case of the Pi transporter IPT.Three AP gene families, phoA, phoX, and phoD,

operate in heterotrophic bacteria and are oftenfound in different cell compartments (cytoplasm,periplasm, outer membrane, and extratracellular;


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TABLE2.

Values

(inlM

)ofPiuptake

,cellularPiquota,an

dgrowth

param

etersas

wellas

Pi-threshold

concentrationsreported

toinduce

alkalinephosphatase.

Species

Ks

(lM)

Vmax(p

mol�

cell�1�h

�1)

Referen

ceKl(lM)

Referen

ceQ0(p

mol�

lm

�3)

lmax

(d�1)

Referen

ce[P

i]threshold

Referen

ce

Dinophyta

“Alexandrium

catenella”

1.9

0.14

Nakam

ura

and

Watan

abe(198

3)2.28

Ouet

al.

(200

8)4.59

10�5

0.63

Ouet

al.(200

8)0.4–

1Jauzein

etal.

(201

0)“A

lexandrium

tamarense”

2.6

1.4

Yamam

oto

and

Tarutani(199

9)2.59

10�5

0.56

Yamaguch

ian

dItakura

(199

9)0.43

Ohet

al.(200

2)

Gym

nodinium

catenatum

3.4

1.42

Yamam

oto

etal.

(200

4)2.59

10�5

0.37

Yamam

oto

etal.

(200

4)3.3

Ohet

al.(200

2)

Heterocapsa

circularisquam

a3.39

10�5

Heterocapsa

triquetra

5.79

10�5

0.72

Tarutani(199

9)Karenia

mikimotoi

4.69

10�5

0.67

Yamaguch

ian

dItakura

(199

9)0.2

Yamaguch

iet

al.(200

4)Karenia

brevis

<0.5

Vargo

and

Shan

ley(198

5)Prorocentrum

donghaiense

1.73

Ouet

al.

2008

)6.79

10�5

0.72

Ouet

al.(200

8)

Prorocentrum

minimum

1.96

Cem

bella

etal.

(198

4)16

.42

Mesecket

al.

(200

9)Pyrocystisnoctiluca

1.9

Rivkinan

dSw

ift

(198

2)Bacillariophyta

Phaeodactylum

tricornutum

50Garc� ıaRuiz

etal.(199

7)Skeletonem

acostatum

0.68

0.03

8Tarutanian

dYamam

oto

(199

4)

0.58

Ouet

al.

2008

119

10�5

1.2

Ouet

al.(200

8)0.25

Yamaguch

iet

al.(200

4)

0.19

10�5

1.25

Tarutanian

dYamam

oto

(199

4)Chaetoceros

neogracile

12.1

Mesecket

al.

(200

9)Nitzschia

sp.

61.2

6.84

Yamam

oto

etal.(201

2)Yamam

oto

etal.(201

2)24

910

�5

0.48

Yamam

oto

etal.(201

2)Thalassiosira

pseudonan

a0.58

Fuhset

al.

(197

2)0.7

Perry

(197

6)Tha

lassiosira

weissflogii

1.72

Fuhset

al.

(197

2)Hap

tophyta

Emilianahu

xleyi

0.00

11Riegm

anet

al.(200

0)Riegm

anet

al.

(200

0)0.25

Dyh

rman

and

Palen

ik(200

3)Pha

eocystissp.

0.5

vanBoek

elan

dVeldhuis(199

0)Isochrysissp.

Mesecket

al.

(200

9)Pavlova

lutheri

0.00

26Law

set

al.

(201

1b)

(continued

)


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Luo et al. 2009, Sebastian and Ammerman 2009,White 2009). The greatest abundance of AP genesappear to code for cytoplasmic proteins, suggestingthat intracellular uptake of phosphoesters and sub-sequent hydrolysis within the cell may be moreprevalent in marine heterotrophic bacteria than pre-viously thought (Luo et al. 2009). For AP to func-tion intracellularly, however, would requirephosphoester transporters which have not yet beenidentified (Fig. 4). Alternatively, many of theseintracellular APs may be involved in the hydrolysisof phosphate esters produced within the cell thatare 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 arerequired for the hydrolytic activity of AP enzymes(Coleman 1992, Yong et al. 2014). Cyanobacteriacontain 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 amagnesium (Mg) ion in their active centers, whilePhoD utilizes calcium (Ca) instead of Mg ions andmay also contain Zn or other hydrolytically activemetal (although such a requirement has not yetbeen established; Roy et al. 1982, Coleman 1992,Kageyama et al. 2011). In contrast, recent structuralanalysis indicates that PhoX contains two ferric ionsand three Ca ions in its active center (Yong et al.2014). The metal ions that are utilized in the activesite of the enzyme is of interest, not only becausethis can affect the activity and specificity of theenzyme but also because regional variations in limit-ing metal concentrations may select for species con-taining APs with different metal requirements.Putative phoX genes have also been identified in

Prochlorococcus and Synechococcus (Kathuria and Mar-tiny 2011), and in two chlorophytes: Volvox carteri(Hallmann 1999) and Chlamydomonas reinhardtii (Qui-sel et al. 1996, Moseley et al. 2006). An AP has alsobeen identified by proteomics in the pelagophyteAureoumbra lagunensis, which was activated by Ca butinhibited by Zn, consistent with the behavior ofPhoX (Sun et al. 2012). However, an iron require-ment was not examined in this study; and even if ithad been, the standard chelator (EDTA) used toremove catalytically active metals from AP enzymes isnot strong enough to remove ferric ions from theactive site of PhoX (Yong et al. 2014). This fact hasled to much past confusion regarding the metalrequirements of this enzyme (Yong et al. 2014).A novel AP (EHAP1) has also been identified in

the widely distributed marine haptophyte Emililianiahuxleyi (Xu et al. 2006, 2010). Based on its aminoacid sequence, this enzyme is phylogeneticallyrelated to PhoD (Lin et al. 2013). It appears torequire either Zn or cobalt (Co) as a cofactor as theAP activity of E. huxleyi is greatly suppressed in theT

ABLE2.

(continued

)

Species

Ks

(lM)

Vmax(p

mol�

cell�1�h

�1)

Referen

ceKl(lM)

Referen

ceQ0(p

mol�

lm

�3)

lmax

(d�1)

Referen

ce[P

i]threshold

Referen

ce

Rap

hidophyta

Cha

ttonella

antiqua

1.76

0.14

Nakam

ura

and

Watan

abe(198

3),

Nakam

ura

(198

5)

1.69

10�5

0.86

Nakam

ura

and

Watan

abe(198

3)

Heterosigmaakashiwo

7.99

10�5

Watan

abe

etal.(198

2)Chlorophyta

Chlorella

autotrophica

13.6

Mesecket

al.

(200

9)Tetraselmischui

10.1

Mesecket

al.

(200

9)Tetraselmissuecica

0.00

345

Law

set

al.

(201

1a)

1.19

Law

set

al.

(201

1a)


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combined absence of Zn and Co and is enhancedby the addition of either metal (Shaked et al. 2006,Jakuba et al. 2008). The presence of Zn or Zn/Coin many AP enzyems (e.g., PhoA and EHAP1) sug-gests that P and Zn (and/or Co) may colimit DOPutilization and algal growth in some regions of theocean such as the Sargasso Sea where Pi, Zn, andCo occur at very low concentrations (Wu et al.2000, Shaked et al. 2006, Jakuba et al. 2008). How-ever, the presence of Fe in the active site of PhoXsuggests that colimitation by Fe and P may alsooccur in oceanic regions with low Fe and P concen-trations. Indeed, the relative abundance of Zn andFe in low-P ocean waters may influence the compo-

sition of phytoplankton communities since Fe- andZn-dependent APs often do not occur together inthe same species (Yong et al. 2014).Alkaline phosphatase genes have also been iden-

tified in diatoms and dinoflagellates, two dominantgroups of eukaryotic phytoplankton. AP genes havebeen identified in the genomes of the diatomsThalassiosira pseudonana and Phaeodactylum tricornu-tum (Armbrust et al. 2004, Bowler et al. 2008, Dyhr-man et al. 2012) and the dinoflagellatesAmphidinium carterae (Lin et al. 2011), Karenia brevis(Morey et al. 2011, Lin et al. 2012a), and Alexan-drium catenella (Lin et al. 2012b). A cell surfaceprotein showing AP activity was also identified in

TABLE 3. Alkaline phosphatase and other genes in phytoplankton that facilitate the utilization of dissolved organicphosphate identified to date.

Alkaline phosphatase (AP) Other enzymes

PhoA phoX phoD phoV Newtype

Acidphosphatase 50 nucleotidaseZn/Mg Fe/Ca Ca/? Zn/Mg

DinophytaAlexandrium“catenella”

Lin et al.(2012b)

Amphidiniumcarterae

Lin et al.(2011)

Karenia brevis Moreyet al. (2011)

Lin et al.(2012a)

BacillariophytaThalassiosirapseudonana

Armbrustet al. (2004)

Dhyrmanet al. (2012)

Phaeodactylumtricornutum

Bowler et al.(2008)

HaptophytaEmilianiahuxleyi

Dyhrmanet al. (2006a)

Xuet al.(2006)

Prymnesiumparvum


PelagophytaAureoumbralaguna

Sun et al.(2012)

Aureococcusanophagefferens

Wurchet al. (2011)

Wurchet al. (2011)

ChlorophytaChlamydomonasreinhardtii

Moseleyet al. 2006

Quiselet al. (1996);Moseleyet al. (2006)

Kruskopf andDu Plessis(2004)

Chlorella sp. Blanc et al.(2010)

Volvox carteri Volca_XP_002958226.1

Hallmann(1999)

CyanophytaAphanothecehalophytica

Kageyamaet al.(2011)

Prochlorococcus Mooreet al. 2005

Mooreet al. (2005)

Synechococcussp. PCC7942

Ray et al.1991

Mooreet al. (2005)

Wagneret al.(1995)

Trichodesmium Orchardet al. 2009

Orchardet al. (2009)


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the dinoflagellate Prorocentrum minimum (Dyhrmanand Palenik 1997). Algal species can contain multi-ple AP genes coding for different proteins. Forexample, at least four different putative APs havebeen identified in the diatom T. pseudonana, andall four were up-regulated under P-deficiency(Dyhrman et al. 2012).

Despite the high sequence divergence of alleukaryotic AP genes, those from dinoflagellates andsome from diatoms, pelagophytes, and haptophytesare slightly more similar to phoA genes than to theother bacterial AP types (Lin et al. 2012b). Further-more, these PhoA-like APs, from dinoflagellates, dia-toms, and haptophytes (the so-called red algallineage) are closer to one another than to those inchlorophytes, consistent with the known phyloge-netic and evolutionary relationships among thesealgal lineages (Lin et al. 2012b, Lin et al. 2015b).

Whether the high variability in AP sequences con-fers different substrate specificities or other func-tional differentiation is currently unknown. Inbacteria, the sequence variability among the differ-ent types of APs is related to the cellular localizationof the encoded enzymes (Luo et al. 2009). Similarly,in silico analysis of AP gene sequences in eukaryoticalgae predict that various AP enzyme types have dif-ferent localizations (extracellular, cell wall, plasmamembrane, or cytoplasm; Lin et al. 2012b). RecentELF staining of live cells of the dinoflagellatesAmphidinium carterae, Karenia brevis, and Alexandriumcatenella (=A. pacificum) shows AP localization pat-terns that largely agree with these predictions (Linet al. 2012b). The cellular localization of differentAP enzymes may enable species to utilize differentsources of DOP and to hydrolyze various DOP com-pounds in different cellular or extracellular loca-tions. It may also be related to variations inchemical environment, such as differences in pH orionic composition near the cell surface or withinthe cell. Much work remains to be done on the reg-ulation and localization of different AP proteins inphytoplankton.Utilization of phosphonates. Phosphonates con-

tribute 5%–25% to the total DOP in the ocean(Clark et al. 1998, Kolowith et al. 2001, Young andIngall 2010). They are likely produced in a widerange of organisms as constituents of phosphopro-teins and cell membrane phospholipids (Clark et al.1998, Villareal-Chiu et al. 2012). The commonlyoccurring phosphonate, 2-aminoethylphosphonicacid, for instance, is present in plant and animalcell membranes. Cyanobacteria, which can utilizephosphonates, can also produce them (Dyhrmanet al. 2009). Heterotrophic bacteria have long beenknown to take up and metabolize phosphonates(Shinabarger et al. 1984, Pipke et al. 1987, for areview see McGrath et al. 2013), but only recentlywas 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, whichis energetically more difficult than hydrolyzing aphosphoester bond. Utilization of phosphonates inheterotrophic bacteria is accomplished using eithera C-P hydrolase or a C-P lyase enzyme system. Incontrast, marine cyanobacteria, contain only the C-Plyase system (Dyhrman et al. 2006b, McGrath et al.2013). The proteins comprising the C-P hydrolaseenzyme system vary among bacteria, with PhnW andPhnX being the most common constituent proteins.By contrast, the C-P lyase system appears to be moreconserved, and consists of 14 proteins under Phoregulon control, which are capable of processing abroad range of phosphonate substrates (White andMetcalf 2004, Dyhrman et al. 2006b). The C-P lyasesystem proteins are encoded by a gene cluster,PhnCDEFGHIJKLMNOP. Within this clusterPhnCDE codes for a phosphonate ABC transporter(which includes an ATP binding subunit protein, aperiplasmic phosphonate binding subunit, andtransmembrane subunit), whereas PhnFGHIJKLM-NOP codes for the C-P bond cleaving enzymes(Fig. 4). Currently, there is no documented evi-dence that eukaryotic phytoplankton can utilizephosphonates, although some preliminary molecu-lar data indicate the presence of phosphonate-meta-bolizing enzyme genes in some dinoflagellates (Linet al.2015a; see Supplementary Table 33.P acquisition by phagotrophy. Phagatrophy occurs

widely in many groups of photosynthetic protistsand is now recognized as an important source of Pand 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 dinoflagel-lates, haptophytes, and pelagophytes, but does notoccur in diatoms. All dinoflagellates tested to dateare capable of phagotrophy, including species origi-nally considered obligate photoautotrophs; e.g., Pro-rocentum minimum (Stoecker et al. 1997), Akashiwosanguenium (Bockstahler and Coats 1993), Karlo-dinium veneficum (Li et al. 1996), Alexandrium osten-feldii (Jacobson and Anderson 1996), Gymnodiniumaureolum (Jeong et al. 2010a), and even the coralreef endosymbiont Symbiodinium (Jeong et al. 2012).Studies have shown that phagotrophy in dinoflagel-lates and other mixotrophic phytoplankton isinduced by low nutrient stress or nutrient limitationof growth rate (Stoecker et al. 1997, Litaker et al.2002, Carvalho and Gran�eli 2010, Jeong et al.2012). Potential phagotrophy-related genes havebeen identified in dinoflagellates and other phyto-plankton. Clathrin-mediated endocytosis proteinsand autophagy-related proteins were up-regulatedunder Pi limitation in the dinoflagellate K. mikimotoi(Lei and Lu 2011), the haptophyte Prymnesium par-vum (Beszteri et al. 2012), and the pelagophyteA. anophagefferens (Wurch et al. 2011). Phagotrophyrepresents an efficient means of acquiring P andother nutrients in situations where dissolved nutri-ent concentrations are low and a substantial propor-


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tion of the nutrients are contained in bacteria,microalgae, and other microorganisms (Smalleyet al. 2003, Flynn et al. 2013). The biochemicalpathways involved in phagotrophy and subsequentnutrient assimilation represent an understudiedaspect of nutrient acquisition worthy of furtherinvestigation. In particular, the potential involve-ment of acid phosphatase and nucleotidase(Table 3) in releasing phosphate during food diges-tion deserves attention.

GROWTH LIMITATION AND CELLULAR RESPONSES TO IT

Growth relation to cellular P. Decreases in P orother nutrients below critical threshold valuesresults in decreased rates of cellular growth andreproduction. Early models related nutrient limita-tion of growth rate to the external nutrient concen-tration (e.g., Monod 1949) based on chemostatstudies in which the external nutrient concentrationremained constant with time. However, concentra-tions of Pi and other nutrients in seawater are oftenvariable in time and space (Turpin and Harrison1979), and limitation of growth rate is now knownto be dependent on the cell nutrient quota(amount per cell) or concentration (e.g., amountper unit of cell carbon) rather than the externalconcentration (Droop 1968, Fuhs 1969, Grover1991). Depending on the growth status and previ-ous nutrient exposure, cell P quotas can vary widely(Jauzein et al. 2010, Pleissner and Eriksen 2012).

The model most often used to describe the rela-tionship between the cell nutrient quota (Q) andspecific growth rate (l) is the Droop equa-tion (Droop 1968, 1983, 2003):

l ¼ lmaxð1� Q0=Q Þ ð3Þ

where Q0 is the minimum cell quota at which thegrowth rate is reduced to zero, and lmax is the hypo-thetical maximum growth rate at an infinite cellquota. lmax is often unobtainable, and in reality isonly a fitting parameter for the equation (Droop1983, Laws et al. 2011a,b, 2013). Also Q should bethe average daily cell quota since quotas for P andother nutrients (e.g., N and Fe) typically vary withtime of day due to diel variations in rates of nutri-ent uptake, cell growth and cell division (Ahn et al.2002, Sunda and Huntsman 2004).

An advantage of this approach is that it allowsgrowth rate predictions based on cell quotas, whichare generally easier to measure than the low con-centrations of Pi and other nutrients that actuallylimit algal growth rates (Laws et al. 2011b, Sundaand Hardison 2010). In recent chemostat studiesunder continuous light conditions, the relationshipbetween growth rate and cell P quota was welldescribed by the Droop equation (Laws et al. 2013).

Although the classic Droop equation allows a pre-diction of growth versus cell quota (average amount

per cell) for a given species, it fails to provide ameasure of growth efficiency – the rate of cell car-bon production per unit of cell P. Relationshipsbetween growth rate and cell quota can vary widelyamong species because of large differences in cellsize, and resultant moles of P per cell. Normalizingcellular P on per unit carbon basis (i.e., the molarP:C ratio) eliminates this difficulty and allows car-bon growth per unit of cell P to be directly com-pared among species as was done in Jauzein et al.(2010). Only when more studies directly relategrowth rates to cell P:C ratios will it be possible todetermine if dinoflagellates, diatoms, or other phy-toplankton groups on average have different Pgrowth efficiencies. Such efficiencies are defined asthe net moles of cell C produced per mole of cell Pper unit time (i.e., per day) and equal the specificgrowth rate divided by average daily cellular P:Cratio.Changes in the cell P:C ratio (QC) with time is

governed both by the C-normalized cellular P-uptake 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 diellight cycle, all values in the above equations need tobe daily averages, which can usually be estimatedfrom measurements made in the middle of the lightperiod (Sunda and Hardison 2007). From the aboveequations it is evident that the cellular P:C ratio isdetermined by the balance between the cellular Puptake rate and the specific growth rate, and thatchanges in P:C ratios will be determined by the rela-tive 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 vari-ability in the uptake mechanisms and kinetics, cellsize and cellular P growth requirements, the Pilevels in any given ecosystem may be growth-suffi-cient for some species, but growth-limiting forothers (Sundareshwar et al. 2003, Nicholson et al.2006, Mackey et al. 2007). This is true even thoughthe phytoplankton community can adapt evolution-arily to a range of ambient concentrations of P, N,and other nutrients; for example, the consistentlylow nutrient levels in open ocean surface waters(Chisholm 1992, Sunda and Hardison 2007, 2010).Even when cellular P quotas are normalized to cellcarbon, these values still vary among species(Table 2). This variability likely results from differ-ences among algal species in their cellular P uptakerates and specific growth rates and to associated dif-ferences in their biochemical composition, particu-larly in the abundance of RNA, DNA,


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phospholipids, and polyphosphate (Falkowski 2000,Geider and La Roche 2002). As a result, N:P:C ratiosin phytoplankton species can be quite variable, withN:P ratios for nutrient replete algal species varyingfrom 5 to 19 (Geider and La Roche 2002). Molar N:P ratios >16:1 in seawater are typically regarded asP-limiting. However, in seawater environments withmeasured ratios of inorganic DIN:Pi as high as 25:1,the N:P ratios in plankton tend to be much less than16 (median 9) (Broecker and Henderson 1998,Geider and La Roche 2002). This discrepancy is mostlikely due to the accumulation of P storage pools(e.g., polyphosphate) and to variations in ribosomalRNA linked to changes in rates of growth and pro-tein synthesis (Geider and La Roche 2002). Theadsorption of phosphate on cell surfaces can also bea factor and can significantly skew N:P ratios in phy-toplankton toward lower values (Sa~nudo-Wilhelmyet al. 2004, Fu et al. 2005). The critical molar N:Pratio in phytoplankton that marks the transition fromN- to P-limitation is in the range of 20–50, signifi-cantly higher than the 16:1 Redfield ratio (Geiderand La Roche 2002). Variations in these critical N:Pratios among species may be an important factoraffecting algal species competition and the formationof species-specific algal blooms, including those ofdinoflagellates.Increased cellular toxins in P-limited cells. Growth

rate limitation by P and other nutrients (e.g., N) isoften accompanied by increases in toxin per cell orper mol of cell C in many toxic harmful algal bloomspecies. This has been observed for saxitoxins,which cause paralytic shellfish poisoning (Flynnet al. 1994, Maestrini et al. 2000, Anderson et al.2002), okadaic acid, which causes diarrhetic shell-fish poisoning (John and Flynn 2002), and domoicacid, which causes amnesic shellfish poisoning (Panet al. 1998). P limitation of growth rate alsoincreases cellular toxins in Chrysochromulina polylepis(Johansson and Graneli 1999a), Gambierdiscus polyne-siensis (Chinain et al. 2010), Karlodinium veneficum(Fu et al. 2010), Prymnesium parvum (Beszteri et al.2012, Johansson and Graneli 1999b), Protoceratiumreticulatum (Guerrini et al. 2007), and the N2-fixingcyanobacterium Nodularia spumigena (Sunda et al.2006). In a detailed study of the dinoflagellate Kare-nia brevis, Hardison et al. (2013) showed that theincrease in cellular toxins (bevetoxins):C ratios wasbest predicted by the degree of growth rate limita-tion by P and not by cell P:N or P:C ratios. Theyfound that the increased cellular toxin:C ratios werenot, however, due to an increase in the cellulartoxin production rate as might be expected intu-itively. Instead, as growth slowed under P limitation,cells down-regulated the rate of toxin synthesis, butto a lesser degree than the overall decrease in therate of cellular C production leading to higher cel-lular toxin:C ratios. Similar increases in cellularbrevetoxin:C ratios were also observed under bothN limitation (Hardison et al. 2013) and CO2 limia-

tion of growth rate (Hardison et al. 2014), suggest-ing that the increase in brevetoxins is inherentlylinked to the slower growth rates that occur duringnutrient limitation. Hardison et al. (2012, 2013)hypothesized that the increase in cellular toxins isevolutionarily advantageous, as elevated brevetoxinshave been shown to deter zooplankton grazing(Hong et al. 2012). The lower grazing rates wouldresult in higher net population growth rates thanwould occur otherwise as algal growth rates slowduring nutrient limitation.The actual biochemical pathways by which P limi-

tation of growth rate regulates toxin production arelargely unknown and researchers are just now begin-ning to isolate genes involved in the biosynthesis ofvarious toxins. Recently the polyketide synthasegene cluster responsible for the production ofbrevetoxins in Karenia brevis was identified (Monroeand Van Dolah 2008). Additionally, two genesbelieved to be involved in saxitoxin synthesis inAlexandrium spp. and other dinoflagellates were alsoidentified (Orr et al. 2013, St€uken et al. 2011).Determining how nutrient limitation of growth rateand other factors regulate these toxin pathwaygenes should prove a fruitful area for futureresearch.Encystment induced by P limitation. Dinoflagellates

can survive P deficiency by encystment (Andersonet al. 1985). However, laboratory experiments haveshown that motile cells need a minimum P contentto form cysts (Anderson et al. 1985), and that cystsmust contain sufficient ATP to germinate after dor-mancy (Lirdwitayaprasit et al. 1990). Furthermorein some dinoflagellates (e.g., Scrippsiella trochoidea),cysts can take up P intracellularly and their P con-tent increases with the external Pi concentration(Rengefors et al. 1996). This enables dinoflagellatecysts that settle to the bottom to accumulate P fromabundant sedimentary Pi pools for use during dor-mancy, germination, and subsequent growth. Themolecular mechanisms of P uptake, storage, andmetabolism during the encystment and germinationphases are currently unexplored.Cell enlargement and cell cycle arrest caused by P limita-

tion. Nutrient limitation of growth rate in phyto-plankton usually causes a decrease in cell size, asobserved for nitrogen (Sunda and Hardison 2007,2010, Hardison et al. 2012) and iron (Sunda andHuntsman 1995). Such decreases should be favoredby natural selection as they facilitate nutrient uptakeby increasing cell surface to volume ratios and therate of diffusive nutrient flux through the surfaceboundary layer normalized to cell volume (Sundaand Hardison 2007). In contrast, P limitation ofgrowth rate often leads to cell enlargement, asreported for dinoflagellates (Latasa and Berdalet1994, John and Flynn 2002, Lim et al. 2010, Varkitziet al. 2010, Hardison et al. 2013, Zhang et al. 2014),diatoms (Liu et al. 2011), and chlorophytes (Litch-man and Nguyen 2008). The increase in cell size


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under P limitation should decrease P uptake ratesper unit of cell volume, and thus, should be evolu-tionary disadvantageous. So, in contrast to othernutrients, there must be some factor peculiar tophosphorus that causes cell size to increase under Plimitation of growth rate.

To examine the effects of P limitation on cell sizewe must examine its effect on the cell’s growth anddivision cycle. This cycle consists of four discretephases: the G1 phase (gap 1 or growth stage 1)where a newly divided cell grows and increases insize prior to cell division, the S phase during whichDNA is replicated, the G2 phase (gap 2 or growthstage 2) where the cell continues to grow prior tomitosis, and the M phase during which nuclear divi-sion (mitosis) occurs leading to cell division (cytoki-nesis). Unlike other nutrients (e.g., Fe and N), Plimitation often results in a blockage of DNA repli-cation (the S phase; Vaulot 1995), which must pre-cede cell division into smaller daughter cells(Sclafani and Holzen 2007). P limitation of growthrate in the cyanobacteria Prochlorococcus and Syne-chococcus causes an arrest of the cell cycle progres-sion from G1 to S or G2 to M phases, and in thecase of P starvation (severe growth rate limitation),an arrest in the S phase (Parpais et al. 1996, Vaulotet al. 1996). Similarly, the few studies conducted sofar with dinoflagellates indicate an arrest of the cellcycle in G1 phase in response to P limitation ofgrowth rate (Lei and Lu 2011, Zhang et al. 2014;Li et al. 2015). This arrest is accompanied by theup-regulation of negative regulators (e.g., fizzy/celldivision cycle 20-related protein) and down-regula-tion of positive regulators of the cell cycle (e.g., cal-cium-dependent protein kinase) (Zhang et al.2014). With the arrest of the cell cycle, the cellscontinued to grow during an elongated G1 phase,resulting in an increase in average cell size. Theblockage of the cell cycle progression from G1 to S,or subsequent phase transitions, is likely linked to aneed for a sufficient supply of P for successful DNAreplication and for phosphorylation of key check-point enzymes that regulate DNA synthesis andnuclear and cell division in the S and M phases.Transitions in the cell cycle, including G2 to Mstages, are strictly regulated by a cascade of CDKphosphorylation and dephosphorylation events(Murray and Hunt 1993). The cell enlargement inP-limited cells further suggests that fulfilling CDKphosphorylation or other P-associated biochemicalrequirements (e.g., DNA replication) may supersedethat of a threshold in cell size in controlling theonset of cell division.

DIFFERENTIAL NUTRIENT ACQUISITION AND GROWTH

STRATEGIES AND SPECIES SUCCESSION

Differential P nutrient strategies may be one ofthe drivers of seasonal species succession that occursin many phytoplankton communities. The popula-

tion growth of diatoms with high nutrient-sufficientmaximum growth rates is often favored during earlysuccession (in late winter/early spring or in freshlyupwelled water) when the environment is character-ized by high concentrations of dissolved inorganicnutrients (Pi, DIN, and Fe), high turbulence, lowalgal biomass, and a low level of zooplanktongrazing (Margalef 1978, Sunda et al. 2006, Sundaand Hardison 2010, Fig. 5). As the season pro-gresses, the water column stabilizes with increasedsolar heating of surface water, and inorganic nutri-ent pools become depleted by the initial algalbloom which is often dominated by diatoms (Mar-galef 1978, Hood et al. 1990, Tiselius and Kuylen-stierna 1996, Yoshimura et al. 2014). As the DINand Pi pools decline during the bloom there is aprogressive buildup of organic nutrients (DOP andDON) linked to slopy grazing and excretion by zoo-plankton, viral and bacterial lysis of cells, andrelease by phytoplankton (van der Zee and Chou2005, Yoshimura et al. 2014). The combination of astable water column, higher phytoplankton biomass,low inorganic nutrients, increased DOP and DONlevels, and increased zooplankton grazing pressureno longer favors the population growth of diatomsor other early succession species and sets the stagefor a population shift to late succession species such

Time

P or

C

FIG. 5. Schematic for a typical diatom to dinoflagellate sea-sonal succession in a system whose biomass is limited by P. Theinitially high Pi levels, colder temperatures, and high turbulencelevels in the early spring favor diatoms, which are adapted forhigh growth rates under these conditions. The emerging diatombloom depletes the euphotic zone of Pi and fuels the growth ofzooplankton. Concomitantly, solar warming increases stratifica-tion of the water column, and decreases inputs of Pi and othernutrients from nutrient rich aphotic deeper waters. Phosphorusinputs during this time are mainly from recycling linked to zoo-plankton grazing and excretion, and much of that input is in theform of DOP. The combination of DOP inputs from grazing andPi uptake by phytoplankton increases the DOP concentration andgreatly increases the DOP:Pi ratio. These changes (decreased Piin surface waters, increased DOP, water column stratification, andincreased zooplankton grazing) sets the stage for a algal commu-nity shift from diatoms to dinoflagellates, whose growth and sur-vival are favored under these conditions due to their ability toobtain nutrients from alternate sources (diel vertical migration tonutrient rich deeper waters, utilization of DOP, and phagotro-phy) and their ability to minimize grazing losses (e.g., linked tolarge cell size and the production of toxins).


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as dinoflagellates, haptophytes, and pelagophytes,which are better adapted to this new set of environ-mental conditions (Margalef 1978, Sunda et al.2006).

In contrast to diatoms (which are non-motile),dinoflagellates and many other late sucession spe-cies possess flagella and are motile. This motilityallows them to vertically migrate out of nutrient-depleted surface waters into deeper nutrient-richwaters at the end of the light period and thenmigrate back into sunlit surface waters at the begin-ning of the light period (Sinclair and Kamykowski2008, Hall and Paerl 2011). In this way P and othernutrients that are depleted in surface waters duringblooms, but accumulate at depth or in bottom sedi-ments via POM settling and regeneration processes,can be utilized to sustain photosynthesis and growthduring the day (Hall and Paerl 2011, Sinclair andKamykowski 2008).

In addition, three other functional traits promoteincreased nutrient acquisition rates of dinoflagel-lates and other late succession species in low Pi andDIN surface waters. One is their higher capability toutilize DOP and DON than diatoms, which helpsfavor their growth in late succession waters withhigh ratios of DOP:Pi and DON:DIN (Sunda et al.2006, Burkholder et al. 2008). Indeed as noted ear-lier, dinoflagellates are observed to have higher APactivities during blooms (on either a per cell or cellvolume basis) than coexisting diatoms (Nicholsonet al. 2006, Ou et al. 2006). In addition, somedinoflagellates can grow as well on ATP as on DIP,suggesting an ability to directly utilize this DOPwithout the energy-costing hydrolysis (Li et al.2015). A second, perhaps more important func-tional trait, is the capability of dinoflagellates, hap-tophytes, and other late succession species toacquire P and other nutrients via phagotrophy(Stoecker et al. 1997, Hartmann et al. 2012, Flynnet al. 2013). This capability may have little survivalvalue prior to and during the early phase of the ini-tial diatom bloom, and indeed diatoms are one ofthe few groups of eukaryotic phytoplankton with noknown phagatrophic capability (Flynn et al. 2013).However, phagotrophy becomes an increasinglyimportant nutrient source as algal biomass increasesand Pi and DIN pools become depleted duringblooms. Dinoflagellates, especially athecategymnodinoid forms, are known to feed heavily ondiatoms (Sherr and Sherr 2007). Many mixotrophicdinoflagellates depend primarily on photosynthesisfor growth when nutrients are abundant, butbecome facultative heterotrophs once dissolved Piand DOP nutrients become growth limiting(Stoecker et al. 1997, Litaker et al. 2002, Smalleyet al. 2003). This capacity allows dinoflagellates toacquire P and other nutrients from the consump-tion of other algal species while at the same time toreduce competition for the remaining pool of dis-solved nutrients. The motility of dinoflagellates and

other flagellates enhances encounter rates withprey, and their diverse swimming speeds seem toenable them to effectively capture different types ofprey (Nielsen and Kiorboe 2015).The third potentially important late succession

trait in dinoflagellates is a greater cell P-storagecapacity. Although both diatoms and dinoflagellatescan store polyP, the large size of dinoflagellatespotentially allows for greater internal storage (Elgav-ish et al. 1980, Diaz et al. 2008). This greater stor-age capacity may allow some dinoflagellates tosustain relatively high growth rates during the latesuccession 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 growthand reproduction but also by grazing mortalitylosses (Sunda et al. 2006, Smayda 2008). Here too,dinoflagellates and other late succession speciesmay be better adapted to late succession environ-ments characterized by higher zooplankton popula-tions and higher grazing pressures. Manydinoflagellates and other late succession species(e.g., haptophytes such as Prymnesium parvum andpelagophytes such as A. lagunensis) appear to bewell defended from grazing due to high cellularconcentrations of grazing-deterrent compounds(e.g., toxins and mucilage layers), which increaseunder P or N limitation of growth rate (Sunda et al.2006, Hong et al. 2012, Waggett et al. 2012, Hardi-son et al. 2013). Thus, differences in P acquisition,P utilization, and grazing deterrence strategiesbetween diatoms and dinoflagellates contribute totheir respective dominance in early and late succes-sion environments (Fig. 5). However, many of themolecular mechanisms unlying these strategies arestill unclear and await more comparative genomicsstudies for key species from both phyla.

FUTURE DIRECTIONS

Considerable progress has been achieved in thelast two decades in our understanding of how Paffects the growth and ecology of phytoplankton.The molecular mechanisms regulating the uptakeand metabolism of P, and adaptation to low-P stresshave become clearer thanks to rapidly expandingdata sets on algal genomes, transcriptomes, and pro-teomes. However, this initial progress representsonly a small fraction of the information that can beobtained using careful chemical and physiologicalstudies coupled with modern molecular techniques.Some of the most pressing issues include: (i) obtain-ing a more detailed description of dissolved inor-ganic and organic P substrate pools in ocean waters,including actual measurements of the trueorthophosphate (Pi) concentrations in low-Pi ocea-nic waters; (ii) determining the extent to which Plimitation regulates the growth and species composi-tion of phytoplankton communities in various ocea-


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nic and coastal environments; (iii) determining theidentity, structure, binding affinities, and regulationof low- and high-affinity cellular transporters of Piand DOP; (iv) identifying specific molecular mark-ers (e.g., P sensitive proteins) that can be used toquantify P limitation in different phytoplanktongroups in the ocean, and developing proteomic ortranscriptomic methods to measure the distributionsof these markers in time and space; and (v) deter-mining the sequence variability among importantP transport and metabolic proteins (e.g., AP, trans-porters of Pi and DOP, and polyP metabolic pro-teins) and the extent to which this variability isrelated to different environmental and evolutionaryfactors. These questions are important in assessingthe role of phosphorus in regulating phytoplanktonproductivity, species diversity and carbon cycling inthe present day ocean. But perhaps, more impor-tantly, they may be critical to our understanding ofimpact of P on the biology of a warmer, more highlystratified and acidified future ocean. Tackling thesevarious questions will be greatly facilitated by thedevelopment of improved analytical methods; forexample, more sensitive and specific mass spectrome-try methods for individual DOP compounds andproteomic methods for measurement of specific pro-teins. These questions will also become more tract-able when more of the genomes and transcriptomesof dinoflagellates, diatoms, and other eukaryoticalgal groups become more readily available. Develop-ment of targeted gene transformation systems fordinoflagellates and other species, which has justbegun to gain momentum for diatoms, would alsofacilitate studies linking phosphorus physiology andecology to underlying molecular mechanisms.

We thank Dr. Xin Lin for assistance with making Figure 2.We are also indebted to Drs. Edward J. Carpenter andEdward Monahan and reviewers for their comments thathelped improve the manuscript. Mark Vandersea and StevenKibler provided thoughtful criticisms and edits. MichelleWood provided superb editorial guidance and insights forwhich we are grateful. This work was supported by theNational Science Foundation grant OCE-1212392 and NOAAprogram funds.

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Phosphorus physiological ecology and molecular mechanisms ... · Phosphorus (P) is an essential nutrient for marine phytoplankton and indeed all life forms. Current data show that