Microbial Ecology

, Volume 63, Issue 1, pp 188–198

Molecular Response of the Bloom-Forming Cyanobacterium, Microcystis aeruginosa, to Phosphorus Limitation


  • Matthew J. Harke
    • School of Marine and Atmospheric SciencesStony Brook University
  • Dianna L. Berry
    • School of Marine and Atmospheric SciencesStony Brook University
  • James W. Ammerman
    • School of Marine and Atmospheric SciencesStony Brook University
    • School of Marine and Atmospheric SciencesStony Brook University
Microbiology of Aquatic Systems

DOI: 10.1007/s00248-011-9894-8

Cite this article as:
Harke, M.J., Berry, D.L., Ammerman, J.W. et al. Microb Ecol (2012) 63: 188. doi:10.1007/s00248-011-9894-8


Cyanobacteria blooms caused by species such as Microcystis have become commonplace in many freshwater ecosystems. Although phosphorus (P) typically limits the growth of freshwater phytoplankton populations, little is known regarding the molecular response of Microcystis to variation in P concentrations and sources. For this study, we examined genes involved in P acquisition in Microcystis including two high-affinity phosphate-binding proteins (pstS and sphX) and a putative alkaline phosphatase (phoX). Sequence analyses among ten clones of Microcystis aeruginosa and one clone of Microcystis wesenbergii indicates that these genes are present and conserved within the species, but perhaps not the genus, as phoX was not identified in M. wesenbergii. Experiments with clones of M. aeruginosa indicated that expression of these three genes was strongly upregulated (50- to 400-fold) under low inorganic P conditions and that the expression of phoX was correlated with alkaline phosphatase activity (p < 0.005). In contrast, cultures grown exclusively on high levels of organic phosphorus sources (adenosine 5′-monophosphate, β-glycerol phosphate, and d-glucose-6-phosphate) or under nitrogen-limited conditions displayed neither high levels of gene expression nor alkaline phosphatase activity. Since Microcystis dominates phytoplankton assemblages in summer when levels of inorganic P (Pi) are often low and/or dominate lakes with low Pi and high organic P, our findings suggest this cyanobacterium may rely on pstS, sphX, and phoX to efficiently transport Pi and exploit organic sources of P to form blooms.


Cyanobacteria blooms, such as Microcystis aeruginosa, have become increasingly common in freshwater ecosystems in recent decades [6, 21]. Since many cyanobacteria that bloom during summer months have the ability to fix nitrogen [35, 36] and since many freshwater ecosystems have been shown to be P-limited [18, 46, 50], P loading is hypothesized to play a key role in the occurrence of cyanobacteria blooms [35, 36]. As bodies of freshwater become enriched in total P, there is often a shift in the phytoplankton community towards dominance by cyanobacteria [51, 58, 65].

The types of P exploited by Microcystis during blooms have not been well studied. During summer months in temperate lakes, surface waters are often warm, stratified, and devoid of dissolved inorganic P [2, 19, 66] suggesting dissolved organic phosphorus (DOP) may be an important source of biologically available P. However, sources of DOP utilized and the mechanism by which DOP is obtained by Microcystis is unclear. Phosphomonoesters dominate the DOP pools in aquatic environments [3, 12, 26] and the degradation of these compounds requires the enzyme alkaline phosphatase [11]. While methodological limitations and the presence of dense bacterial populations within the polysaccharide sheath of Microcystis colonies [67, 69] have made investigating alkaline phosphatase in Microcystis challenging, recent advances in genomics [13, 24] provide a new means for examining this process.

The dominant molecular model used to describe bacterial P transformations and assimilation of P has been the Pho regulon of the gram-negative bacterium Escherichia coli; a suite of genes that encode for proteins that facilitate P assimilation under low P conditions [56, 62]. The Pho regulon includes genes coding P-binding proteins such as pstS and periplasmic enzymes like the alkaline phosphatase phoA [56, 62]. While the phoA alkaline phosphatase gene is an important component of the Pho regulon for many microbes, the gene phoX, a monomeric protein also within the Pho regulon (23) but requiring different metal co-factors than phoA (Ca2+ rather than Zn2+ and Mg2+), has recently been shown to be widely distributed among aquatic bacteria [25, 49].

Molecular aspects of phosphate acquisition in cyanobacteria have not been comprehensively assessed. While the general elements of the Pho regulon in cyanobacteria and its regulation may be similar to heterotrophic bacteria [57], its composition and arrangement maybe be more variable [55]. For instance, Prochlorococcus (MIT9313) has a pstS homologue, but apparently lacks phoA and may be deficient in the phosphate sensor response system [30]. Trichodesmium possesses both the phoA and phoX alkaline phosphatase genes, as well as the high-affinity phosphate-binding proteins, pstS and sphX [33]. The phoX gene also appears to be common among clones of Prochlorococcus and Synechococcus [25, 49]. Recently, the complete genome of M. aeruginosa has been sequenced [13, 24]. While this information has led to important new insights regarding the evolutionary diversity of prokaryotes in general and cyanobacteria in particular [13, 24], mechanisms of P acquisition in M. aeruginosa have not been robustly described or examined.

The goal of this study was to identify and characterize the expression of genes important for phosphate acquisition in the genus Microcystis. We specifically searched for the presence of two high-affinity phosphate-binding proteins (pstS and sphX) and a putative alkaline phosphatase (phoX) in 11 clones of M. aeruginosa and Microcystis wesenbergii. We additionally quantified the expression of these genes and alkaline phosphatase activity in cultures of M. aeruginosa grown under high and low inorganic P concentrations and supplemented with organic and inorganic P, providing insight into the molecular response of Microcystis to differing endogenous P sources and concentrations.


Microcystis Cultures

Eight strains of M. aeruginosa and one strain of M. wesenbergii were cultured for this study (Table 1). The identity of each culture was verified at the genus level by sequencing of the 16S rRNA gene [7, 32]. Cultures were grown in Blue Green (BG)-11 medium illuminated by a bank of fluorescent lights that provided a light intensity of ~100 μmol quanta m−2 s−1 on a 12:12 light/dark cycle and at 21°C. Bacterial densities in cultures were minimized following the methods of Carmichael et al. [5]. Briefly, a culture of M. aeruginosa LE3 was incubated in the dark for 48 h after which the antibiotic D-Cycloserine (Research Products International Corp., final concentration of 10 μM) was added. The culture was incubated for an additional 24 h in the dark and then gently filtered through a sterile 2 μm polycarbonate filter and collected cells were washed off the filter into sterile BG-11 media. Subsequent DAPI-staining of cultures [42] and an absence of alkaline phosphatase activity in the <2-μm size fraction of P-limited cultures demonstrated that this process reduced non-M. aeruginosa bacteria to levels undetectable by these methods.
Table 1

Microcystis cultured and nucleotide sequence comparison of P-scavenging genes explored in this study



Percent of identity to NIES-843





Lake Kasumigaura, Ibaraki, Japan





Lake Erie, USA





Lake Mendota, Madison, WI, USA





Little Rideau Lake, Ontario, Canada




NIVA-CYA 162/3a

Lake Arresø, Denmark




PCC 7806

Braakman Reservoir, The Netherlands




UTEX B2667a

Little Rideau Lake, Ontario, Canada




UTEX LB2061a

Lake Mendota, Madison, WI, USA




SAG 17.85a

Lake Neusiedl, Austria




EAWAG 198a

Lake Greifen, Switzerland




Microcystis wesenbergii SRa

Sassafras River, MD, USA



Nucleotide comparison for Microcystis cultures is the percent (%) identity top the gene fragment sequenced form the NIES-843 genome (24). Nucleotide comparisons for phoX with the Microcystis wesenbergii strain was not possible as this sequence was not amplified with our primers. All strains were identified at the genus level using a Microcystis-specific 16S rRNA gene (7, 27)

adesignates strains cultured during this study

Sequence Analysis

P-use and -scavenging genes (pstS, sphX, phoX) were identified from the National Center for Biotechnology Information GenBank database (http://www.ncbi.nlm.nih.gov/). PCR primers for target genes were designed using Primer-BLAST (www.ncbi.nlm.nih.gov/tools/primer-blast) against the most conserved regions based on the alignment of available nucleotide sequences with a minimum of 90% identity to M. aeruginosa NIES-843. Primers were also compared against other genomes using BLASTN to ensure they did not amplify other known freshwater prokaryotes. Primers were tested against eight cultured strains of M. aeruginosa and one strain of M. wesenbergii (Table 1). PCR was performed using 100 ng of genomic DNA in a final reaction mixture (25 μL) containing 0.4 μM of each primer and 12.5 μL of GoTaq® Green Master Mix (Promega Corporation, Madison, WI, USA). PCR reactions were carried out as follows: 1 cycle of 95°C for 5 min, followed by 30 cycles of 95°C for 1 min, 58°C for 2 min, 72°C for 3 min, and 1 cycle of 72°C for 30 min. PCR products were evaluated on a 2% agarose gel, stained with ethidium bromide and visualized on a UV transilluminator. Gel images were captured using the Doc-It®LS Image Analysis Software.

PCR products for each of the target genes from eight M. aeruginosa clones and one M. wesenbergii clone were directly sequenced using an ABI 3730 DNA Analyzer (Stony Brook University Sequencing Facility, Stony Brook, NY; Applied Biosystems, Carlsbad, California, USA). Resulting sequences were 264 bp for phoX, 192 bp for pstS, and 204 bp for sphX. Percent identity of each sequenced target gene to the NIES-843 sequence (Table 1) was determined using the Megablast algorithm (blast.ncbi.nlm.nih.gov). An amino-acid alignment of these sequences along with sequences for other marine and freshwater cyanobacteria homologs obtained from NCBI (BLASTX) was generated using ClustalW. Aligned sequences were trimmed to 72 bp for phoX, 195 bp for pstS, and 72 bp for sphX representing the most conserved regions in the sequences examined. Phylogenetic relationship among sequences was constructed using Bayesian inference carried out with MrBayes 3.1 using the following parameters: GTR substitution model, gamma rate variation, 1.1 million generations, and sampled every 200 generations [22]. Resulting trees were rooted to a non-cyanobacterium out member. Prediction of some protein encoding regions was performed with the Glimmer 3.0 program [9].

Phosphorus experiments

Experiments were conducted with M. aeruginosa [clone LE3, Lake Erie, USA; 4], to explore the expression of target genes under P-replete and P-stressed conditions and to explore the effect of organic monoesters on target gene expression. For all experiments, cultures were grown in triplicate 1-L flasks containing 500 mL of BG-11 media with differing levels and sources of P (see below) and were inoculated with M. aeruginosa LE3 (350,000 cells mL−1) which were harvested by centrifugation (Thermo Electron Corporation IEC CL31R Multispeed Centrifuge, 2,200×g for 10 min at 21°C) from a culture grown in P-replete or P-deficient media. Cultures were monitored daily at the same time through the entire experiment for in vivo chlorophyll a fluorescence, alkaline phosphatase activity, and dissolved phosphorus concentrations. At the same time, cell pellets were collected for subsequent RNA analysis (see below). In vivo chlorophyll a fluorescence was measured with 4 mL aliquots from each replicate flask using a Turner Designs TD-700 fluorometer (EM filter of 665 nm and EX filter of 340–500 nm). Bulk alkaline phosphatase activity was measured on a Turner Designs TD-700 fluorometer (EM filter of 410–600 nm and EX filter of 300–400 nm) as described by Hoppe [20] using 4-Methylumbelliferone phosphate (250-μM concentration) as the substrate. Dissolved organic and inorganic phosphorus concentrations in cultures were measured using standard wet chemical techniques [39, 59].

Samples for RNA analysis were harvested by centrifugation of 1.5 mL aliquots of culture into 2-mL microcentrifuge tubes. Each aliquot was centrifuged in an Eppendorf 5415D microcentrifuge at 16,110×g for 10 min and the supernatant poured off and replaced with an additional 1.5 mL of culture. This was repeated three times for a total volume of 4.5 mL. Collected supernatant of cultures was frozen for later analysis of dissolved inorganic and organic P as described above. Cell pellets were re-suspended in 1 mL of CTAB extraction buffer (100 mM Tris–HCl (pH 8), 1.4 M NaCl, 20 mM EDTA, 2% (w/v) cetyltrimethylammonium bromide (CTAB), 0.4% (v/v) β-mercaptoethanol, 1% (w/v) polyvinylpyrollidone) and incubated at 50°C for 20 min. Samples were stored at −80°C and later processed for DNA/RNA extraction [10].

Three different experiments were conducted to examine the expression of target genes under different P concentrations and sources. In the first experiment (P-starvation experiment), cultures were grown without P, harvested in late exponential phase growth, and seeded into media with P (100 μM K2HPO4, +P) or without P (−P). These cultures were monitored daily and when biomass in −P treatments was depressed relative to the +P treatment and alkaline phosphatase activity was elevated relative to the replete culture, 100 μM Pi (K2HPO4) was added back to both cultures. Daily monitoring continued for an additional two days to assess how replenishment of Pi affected target gene expression.

The second experiment examined how phosphomonoesters affected target gene expression in P-replete and P-deplete cultures (re-feed experiment). Experimental flasks were seeded from a P-replete culture and grown with 100 μM orthophosphate and without P as described above. When −P cultures displayed elevated alkaline phosphatase activity during late exponential phase growth, +P and −P cultures were assigned to one of four treatments: (1) 30 μM of the phosphomonoester d-glucose-6-phosphate, (2) 30 μM orthophosphate (K2HPO4), (3) a combination of 15 μM d-glucose-6-phosphate and 15 μM orthophosphate, and (4) a no addition control. Daily monitoring of cultures continued as described above for an additional 2 days to assess how the differing P sources affected target gene expression. In a third experiment (growth-on experiment), triplicate cultures were grown exclusively on orthophosphate or three different phosphomonoester substrates including d-glucose-6-phosphate (G6P; MP Biomedicals, LLC), adenosine 5′-monophosphate disodium salt (AMP; Sigma-Aldrich), and β-glycerol phosphate (BGP; Sigma-Aldrich) at similar concentrations (100 μM as P), and were monitored through the entire growth cycle as described above.

In addition to the three experiments conducted to examine the expression of target genes under different P concentrations and sources, a final experiment was conducted to determine if the target genes and APA were elicited by non-P cellular stress by specifically growing cultures into N-limitation. Triplicate cultures were grown under three conditions: Without N, without P, and with replete amounts of both nutrients. Cultures were monitored daily as described above and were harvested when the biomass in the cultures without P or N were depressed relative to replete cultures and the quantum efficiency of photosystem II (as measured from DCMU (3,4-dichlorophenyl-1,1-dimethylurea) enhanced in vivo fluorescence [15]) was reduced in the culture without N, a physiological indication of N-stress in phytoplankton [15, 38]

cDNA Synthesis and Examination of Gene Expression

Total nucleic acids were extracted using the CTAB technique [10] in which frozen nucleic acids suspended in CTAB were first subjected to an initial heating step at 65°C, followed by a double chloroform extraction, and an isopropanol precipitation. Extracted nucleic acids were re-suspended in 20 μL of LoTE (3 mmol L−1 Tris-HCl [pH 8.0], 0.2 mM EDTA [pH 8.0]). The quantity and quality of nucleic acids were assessed with a NanoDrop ND-1000 Spectrophotometer (NanoDrop Technologies, Inc.). For construction of cDNA, total nucleic acids were digested using DNase I (InvitrogenTM Cat# 18068–015) to remove genomic DNA according to the manufacturer's instructions. The remaining RNA was reverse transcribed into cDNA in 20 μL reactions using random hexamers with the SuperScriptTM III First-Strand Synthesis System for RT-PCR (InvitrogenTM Cat# 188–051) according to the manufacturer's instructions. Negative controls containing no reverse transcriptase (noRT) were run concurrently. The reverse transcribed cDNA (1 μL of a 1:10 dilution) was amplified in PCR reactions under the conditions described above. PCR products were evaluated on a 2% agarose gel, stained with ethidium bromide and visualized on a UV transilluminator.

Quantitative PCR

Triplicate reactions were run using Applied Biosystems Power SYBR® Green PCR Master Mix as follows: 10 μL SYBR Green Master Mix, 0.4 μM forward and reverse primers, 1 μL 1:10 dilution of cDNA, and nuclease free water for a final volume of 20 μL. The quantitative PCR (qPCR) program for most gene targets consisted of 50°C for 2 min, 95°C for 10 min, followed by 45 cycles of 95°C for 15 s, 62°C for 30 s, and 72°C for 30 s, followed by 1 cycle of 95°C for 15 s and 1 cycle of 60°C for 30 s in an Applied Biosystems 7300 Real Time PCR System (Applied Biosystems, Carlsbad, California, USA). The exception was the rpoC1 RNA polymerase gene which was used as the housekeeping gene, reference gene for these reactions for which the annealing temperature was lowered to 53°C [14]. For each sample and primer pair, no reverse transcriptase controls and no template controls were run to test for genomic DNA or PCR contamination. Dissociation curves for each reaction were performed as a quality control tool. Fold changes in gene expression were calculated using the Relative Expression Software Tool (REST 2009), which accounts for differences in reaction efficiency in its fold change calculation (http://www.gene-quantification.de/download.html) [41]. For each experiment, a single time point sample was used as a calibrator for calculating fold changes. Amplification efficiencies of primer pairs fell between 95% and 105% with r2 value of 0.98 or higher (Table 2).
Table 2

Amplification primers used in PCR, RT-qPCR, and sequencing reactions



Amplification primers 5′–3′

Product size [bp]

Efficiency (%)



Phosphate substrate binding protein




This study



Phosphate-binding periplasmic protein




This study



Alkaline phosphatase




This study



RNA polymerase




Ginn et al. (2010)

housekeeping gene



Ribosomal RNA




Neilan et al. (1997)


N.A. not applicable


Sequence Analysis

Sequence analyses among ten clones of M. aeruginosa and one cultured clone of M. wesenbergii indicates that the genes phoX, pstS, and sphX were present and conserved within the species, but not the genus. The putative phoX gene is present and highly conserved (96–98% identical at the nucleotide level) in ten M. aeruginosa clones examined but was not amplified by our primers in the single M. wesenbergii clone examined. The genes sphX and pstS were generally well conserved within Microcystis strains examined (ten M. aeruginosa, one M. wesenbergii) ranging from 91–99% identity to gene fragment sequences from the M. aeruginosa NIES-843 genome (Table 1). Phylogenetic analyses demonstrate that Microcystis formed distinct clusters within phylogenetic trees of phoX, pstS, and sphX sequences of selected cyanobacteria (Fig. S1, S2, and S3). For both the putative alkaline phosphatase phoX and the high affinity phosphate-binding protein pstS, the M. aeruginosa cluster appeared most closely related to clones of the unicellular, non-heterocystous diazotroph Cyanothece sp. (Fig. S1, S2).

Response to Varying P Sources and Concentrations

Cultures of M. aeruginosa grown without P displayed slower growth rates, higher alkaline phosphatase activity (APA), and greater expression of the target P-genes than P-replete cultures (Fig. 1). Cultures grown without P displayed rates of APA which increased from 0.87 ± 0.15 nM mL−1 h−1 on day 1 to 3.61 ± 0.15 nM mL−1 h−1 on day 5 (Fig. 1a) while DIP concentrations remained low throughout this period (0.39 ± 0.22 μM; Fig 1c). The expression of phoX in the −P culture correlated with APA activity (p < 0.005), increasing from low levels of expression at the start of the experiment to 123 ± 39-fold enrichment by day 5 (Fig. 1d). Both pstS and sphX in the −P cultures were strongly upregulated compared to the +P cultures throughout the experiment and reached their highest fold change on day 5 (419 ± 92- and 109 ± 22-fold, respectively; Fig. 1e, f). With the addition of Pi on day 7, APA in the −P culture decreased by 50% to 1.67 ± 0.91 nM mL−1 h−1 on day 8 and expression of phoX, pstS, and sphX, was downregulated on day 8 (Fig. 1). In contrast to the P-deficient culture, the P-replete culture averaged an APA of 0.04 ± 0.01 nM mL−1 h−1 (Fig. 1a) and did not display any significant changes in expression of phoX, sphX, and pstS throughout the experiment (data not shown).
Figure 1

Time series of alkaline phosphatase activity (APA) (a), in vivo Chl a fluorescence in relative fluorescence units (RFU) (b), dissolved inorganic phosphorus (c) for the P-starvation experiment. Error bars represent standard error among biological replicates. Plots d, e, and f represent the mean fold change in gene expression for each gene of the P-deplete (−P) treatment relative to the P-replete (+P) treatment. Error bars indicate the standard error of the fold change of biological replicates. Pi was added back to cultures on day 7

In a second experiment, M. aeruginosa cultures grown with and without orthophosphate (Pi) displayed patterns of growth and APA activity similar to the first experiment (Fig. 2). Upon supplementing late exponential phase, P-limited cultures with d-glucose-6-phosphate (DOP), orthophosphate (DIP), or a combination of DIP and DOP, alkaline phosphatase activity was reduced from 2.66 ± 0.23 nM mL−1 h−1 before the P additions to an average of 0.90 ± 0.16 nM mL−1 h−1 afterwards with little variation among the treatments (Fig. 2a). Furthermore, target P genes all displayed similar patterns of downregulation following the addition of the various P sources. For example, 24 h (day 8) after the addition of P sources, the gene phoX was downregulated 39 ± 12-fold in the DIP addition and 80 ± 29-fold in the DOP addition, and 22 ± 5-fold in the DIP/DOP addition (Fig. 2d). Similarly, pstS was downregulated in the DIP and DOP additions by 66 ± 38- and 52 ± 6-fold, respectively, and 23 ± 3-fold in the DIP/DOP treatment (Fig. 2e). Finally, sphX was downregulated in the DIP, DOP, and DIP/DOP additions by 31 ± 10-, 63 ± 10-, and 18 ± 2-fold, respectively (Fig. 2f). In contrast, the control (the P-limited culture which was not supplemented with P) continued to display elevated rates of APA (Fig. 2a) and expression of the target genes (data not shown). For the +P cultures, APA remained low throughout the experiment (0.03 ± 0.01 nM mL−1 h−1; Fig. 2a) and expression patterns among the three target genes did not change significantly (data not shown). In an experiment in which M. aeruginosa was grown exclusively on orthophosphate or three different phosphomonoester substrates (AMP, BGP, G6P), cultures did not display appreciable APA activity or expression of phoX, sphX, and pstS during exponential phase growth relative to those grown without P (Fig. 3). Similarly, N-limited cultures that displayed reduced photosynthetic efficiency, failed to display APA activity or expression of phoX, sphX, and pstS while concurrently P-limited treatments did (Fig. S4).
Figure 2

Time series of alkaline phosphatase activity (APA) (a), in vivo Chl a fluorescence in relative fluorescence units (RFU) (b), and dissolved inorganic phosphorus (c) for the re-feed experiment. Error bars represent standard error between biological replicates. Treatments were a negative control (+P; 100 μM Pi), a positive control (−P; no Pi added), an addition of 30 μM DIP (−P + DIP), an addition of 30 μM Glucose-6-phosphate (−P + DOP), and an addition of 15 μM DOP and 15 μM DIP (−P + DOP/DIP). Plots d, e, and f represent the mean fold change in expression of each target gene 24 h after the re-feed for the P-deplete cultures relative to the expression levels 24 h earlier. Error bars represent the standard error of biological replicates

Figure 3

a Alkaline phosphatase activity (APA) for cultures grown on dissolved organic phosphorus sources (AMP = Adenosine 5′-monophosphate, BGP = β-Glycerol phosphate, G6P = d-glucose-6-phosphate) as compared to both a positive (+P = 100 μM K2HPO4) and negative (−P = No Pi) control. Error bars represent the standard error among biological replicates. b The fold change in gene expression of cultures grown on organic phosphorus (BGP, AMP, G6P), orthophosphate (100 μM), or without orthophosphate. All data shown from mid-exponential phase growth with expression patterns being relative to +P for −P, BGP, AMP, and G6P treatments and early exponential phase growth for the +P treatment


Phosphorus is considered the primary limiting nutrient in most freshwater systems [52] and many freshwater ecosystems have experienced marked increases in the incident of cyanobacteria blooms during the past several decades [21]. Despite the sequencing of multiple cyanobacterial genomes during the past decades (e.g., [13, 23, 24]), the molecular response of cyanobacteria to specific environmental conditions are largely unknown. The cyanobacterium Microcystis commonly blooms in summer when temperatures are high and inorganic P levels can be low [2, 35, 47]. In the current study, the expression of the genes involved in the hydrolysis of phosphomonoesters (phoX) and high affinity P-transport (pstS and sphX) in M. aeruginosa were regulated by external orthophosphate concentrations. Expression of these genes increased when cells persisted for extended periods under low orthophosphate levels and displayed elevated alkaline phosphatase activity but decreased when cells were offered high levels of organic or inorganic P. Expression patterns did not differ significantly when cultures were grown on orthophosphate or organic monophosphate esters (AMP, BGP, G6P), suggesting these genes do not respond differentially to P sources. The conservation of these genes among ten strains of M. aeruginosa suggests these may be universal molecular responses to P-starvation in this species.

The alkaline phosphatase phoA is a metalloenzyme which catalyzes the non-specific hydrolysis of phosphomonoesters to an alcohol and Pi. It is regulated by external phosphate concentrations and upregulated during phosphate limitation. The enzyme is transported via the Sec pathway into the periplasm and becomes active after dimerization and binding of two zinc and one magnesium cations [57, 60, 63]. Analysis of sequenced M. aeruginosa genomes did not reveal the presence of phoA but rather a protein predicted to be phoX [24]. The enzyme phoX shares no homology with phoA and is thought to be secreted by the Tat pathway rather than the Sec system [29, 60, 68]. Sequence analysis of the two published M. aeruginosa genomes indicates that like all other phoX genes identified to date, the phoX within this species contains signature calcium-binding sites, further confirming their identity [25, 68]. During experiments, expression of phoX in this study was correlated with APA (p < 0.005), suggesting that phoX may be the primary APA gene in Microcystis as has been found for many marine bacteria and cyanobacteria [25, 49]. Moreover, its regulation by endogenous Pi concentrations, but not P source, suggests it is controlled by the Pho regulon, as has been found for heterotrophic bacteria (23). The ability of Microcystis cultures to grow for extended periods (8 days) at levels of DIP <1 μM but with >2 μM DOP coupled with the high rates of alkaline phosphatase activity and expression of phoX during experiments demonstrates that Microcystis grows robustly when DIP is low perhaps by relying on organic phosphomonesters as a source of P. The role of phosphonates and other less labile forms of DOP as a source of P for Microcystis has yet to be determined.

The gene phoX encodes for an alkaline phosphatase, an enzyme that hydrolyzes phosphomonoesters. This gene was not upregulated, however, when M. aeruginosa was grown exclusively on high levels (>100 μM) of three different types of phosphomonoesters. While this finding suggests that for M. aeruginosa, phoX lies within the Pho regulon that is controlled specifically by P stress [29, 56] and not P sources, the manner in which M. aeruginosa accesses phosphomonoesters under P-replete conditions is not currently known. As has been observed for Trichodesmium [33] and some marine bacterial genomes [49], M. aeruginosa may possess other alkaline phosphatase genes that may metabolize phosphomonoesters when P is not limiting. Alternatively, it is also plausible that substrate-specific transporters may target the low-molecular-weight P molecules used in experiments permitting direct transport through the cytoplasmic membrane [27, 64]. Finally, it is possible that the DOP substrates used in experiments were unstable and broke down to orthophosphate and their carbon moiety, providing enough free orthophosphate to suppress activation of the Pho regulon and upregulation of phoX.

The gene sphX was originally named due to its involvement in the Synechococcus phosphate regulatory circuit and has been shown to be transcriptionally regulated by the two component system sphS, a sensory kinase, and sphR, a response regulator which binds directly and specifically to the sequence upstream of the sphX promoter [1]. Within the Pst system, organic phosphates captured and processed by the Pst system travel from the outer membrane via the porin phoE where they are degraded by phosphatases in the periplasm. Once degraded, free Pi is captured by pstS proteins and transferred through the cytoplasmic membrane via a transmembrane protein channel consisting of two proteins (pstA and pstC) [44]. The ABC-type transporters examined in this study, sphX (MAE18390) and pstS (MAE18380) occur within the predicted sphX-pstS-C-A-B operon [24]. Additionally, further genomic analysis indicate that upstream of this operon, there is second putative phosphate transporter (MAE 18310) which may be the pst2 gene, similar to that found for Synechococcus elongatus (PCC 6301 and PCC 7942) and Thermosynechococcus elongatus (BP-1) [53]. Similar to Synechocystis PCC 6803 (freshwater strain), M. aeruginosa may have three ABC Pi transporters (sphX, pst1, and pst2) that, if they have different transport kinetic properties (as does PCC 6803), would allow Microcystis to extend the dynamic range over which it can incorporate Pi and provide it an ecological advantage over other microbial competitors with a less dynamic portfolio of transporters [40].

The expression of pstS paralleled sphX during this study and increased under P-starvation, suggesting that they are both regulated by external Pi concentration. The induction of gene expression of both genes under P-starvation is a pattern similar to those observed in P-stressed Prochlorococcus and Synechocystis [28, 54] but different than that reported for Trichodesmium which upregulated sphX, but not pstS, under P-limitation [33]. The induction of both pstS and sphX by P-limitation suggests that, like E. coli, both genes are within the Pho regulon [57]. While the Pst system is induced in E. coli, when external Pi is lower than 20 μM [44], we did not observe pstS expression until Pi levels were <5 μM.

Since cyanobacteria blooms have long been associated with P loading [35], many lake management plans seek to reduce total P concentrations to mitigate algal blooms [48]. While such plans have had some success, globally, Microcystis blooms continue to occur and in some cases expand [31, 34, 45]. Outbreaks of Microcystis blooms in some US lakes with low total phosphorus concentrations have also been associated with the arrival of recently established zebra mussel (Dreissena polymorpha) populations [43, 61] which are known to increase concentrations of dissolved organic P (DOP) as they filter feed [17]. Our findings suggest that this dominance of Microcystis in lakes with low P is enabled by this cyanobacterium's ability to efficiently access low levels of DIP via efficient P-transporters (pstS, sphX) and to hydrolyze phosphomonoesters via alkaline phosphatase (phoX).

Nitrogen loading is also an important promoter of algal blooms and it has been suggested that dual nutrient (N and P) reductions are necessary for controlling eutrophication [37]. Since, unlike several cyanobacteria, Microcystis is not diazotrophic, N may limit Microcystis growth in systems where there are adequate P supplies. For example, in some shallow systems, total Microcystis populations have been shown to be stimulated more frequently by N than P [8, 16] and abundances of toxic strains of Microcystis have been specifically enhanced by additions of inorganic N [8]. During the present study, cultures grown without N displayed neither increased APA nor expression of the three P genes (Fig S4). This suggests that these genes are not expressed by general cellular stress or N-stress but are upregulated under P-stress.

In summary, the expression of the genes involved in the hydrolysis of phosphomonoesters (phoX) and high affinity P-transport (pstS and sphX) in M. aeruginosa were regulated by external DIP concentration, but not by P sources (orthophosphate vs. BGP, AMP, G6P). The presence of these genes among ten clones of M. aeruginosa suggests this may be a universal response. Future studies may examine how these genes respond to other organic P compounds and how intracellular P-storage may regulate these genes and the growth of Microcystis. Finally, these genes represent useful molecular markers for assessing P-limitation of M. aeruginosa during bloom events.


We thank Dr. Jackie Collier for useful comments and feedback on this research. We thank Dr. Greg Boyer and Dr. Steve Wilhelm for supplying Microcystis cultures. This work was supported by the NOAA-ECOHAB program being funded by the National Oceanic and Atmospheric Administration Center for Sponsored Coastal Ocean Research under award no. NA10NOS4780140 to Stony Brook University.

Supplementary material

248_2011_9894_MOESM1_ESM.pdf (849 kb)
ESM 1(PDF 848 kb)

Copyright information

© Springer Science+Business Media, LLC 2011