J. Phycol. 38, 107–115 (2002)
PHOSPHORUS BIOAVAILABILITY MONITORING BY A BIOLUMINESCENT CYANOBACTERIAL SENSOR STRAIN 1 Osnat Gillor Environmental Sciences, the Fredy and Nadin Herrmann Graduate School of Applied Science, The Hebrew University, Jerusalem 91904, Israel
Ora Hadas Israel Oceanographic and Limnological Research, Yigal Allon Kinneret Limnological Laboratory, POB 345, Tiberias 14102, Israel
Anton F. Post H. Steinitz Marine Biology Laboratory, Interuniversity Institute for Marine Sciences, POB 469, Eilat 88103, Israel
and Shimshon Belkin2 Environmental Sciences, the Fredy and Nadin Herrmann Graduate School of Applied Science, The Hebrew University, Jerusalem 91904, Israel
trial habitats, phosphorus (P) is often considered to be the major limiting nutrient (Schindler 1977, Hecky and Kilham 1988). Typically, oligotrophic lakes have low to undetectable levels of both dissolved inorganic nitrogen and P compounds, and thus one of the major challenges facing phytoplankton ecologists is to identify the actual factor that limits their growth (Elser et al. 1990, Suttle et al. 1991, Istvanovics et al. 1992, Guildford et al. 1994, La Roche et al. 1999). In many cases, circumstantial evidence has been used to ascertain P limitation of phytoplankton in a given environment: low to undetectable concentrations (Hecky and Kilham 1988, Krom et al. 1991, Platt et al. 1992), enhanced uptake capacities (Reynold and Walsby 1975, Suttle et al. 1991), or enrichment bioassays (Elser and Kimmel 1986, Elser et al. 1990). Several methods were developed for a more direct assessment of P bioavailability to phytoplankton, based on physiological parameters of the photosynthetic apparatus (Platt et al. 1992, Greene et al. 1994), cell cycle properties (Parpais et al. 1996, Vaulot et al. 1996), enzymatic markers (Rose and Axler 1998, La Roche et al. 1999, Scanlan and Wilson 1999), or molecular and immunological probes (La Roche et al. 1999, Scanlan and Wilson 1999, Dyhrman and Palenik 2001). Cyanobacteria, as other prokaryotes, respond to P-limiting conditions by the synthesis of extracellular phosphatases, exuded to scavenge P from dissolved organic sources (Tandeau de Marsac and Houmard 1993, Grossman et al. 1994, Mann and Scanlan 1994). These enzymes hydrolyze a variety of organic P-containing molecules and release P available for uptake (Kuenzler and Perras 1965). In the freshwater unicellular cyanobacterium Synechococcus sp. PCC 7942, a two-component sensory system resembling the enterobacterial pho regulon (Tandeau de Marsac and Houmard 1993, Mann and Scanlan 1994) responds to P limitation by increasing the expression of alkaline
Phosphorus (P) is widely considered to be the main nutrient limiting the productivity of freshwater phytoplankton, but an assessment of its bioavailability in natural samples is highly complex. In an attempt to provide a novel tool for this purpose, the promoter of the alkaline phosphatase gene, phoA, from Synechococcus sp. PCC 7942 was fused to the luxAB luciferase genes of the bioluminescent bacterium Vibrio harveyi. The resulting construct was introduced into a neutral site on the Synechococcus sp. PCC 7942 genome to yield strain APL, which emitted light when inorganic P concentrations fell below 2.3 M. Light emission of P-deprived cells decreased rapidly upon inorganic P readdition. The reporter was demonstrated to be a sensitive tool for monitoring the bioavailability of both inorganic and organic P sources. In water samples taken from a natural freshwater environment (Lake Kinneret, Israel), the luminescence measured correlated with total dissolved phosphate concentrations. Key index words: alkaline phosphatase; bioluminescence; phoA; phosphorus bioavailability; phytoplankton; Synechococcus sp. PCC 7942 Abbreviations: AP, alkaline phosphatase; APA, alkaline phosphatase activity; P, phosphorus; RLU, relative light units; TDP, total dissolved phosphorus Cyanobacteria are a dominant component of marine and freshwater phytoplankton, the productivity of which is frequently limited by nutrient availability (Smith 1984, Hecky and Kilham 1988, Kilham and Hecky 1988). In freshwater systems, as in many terres-
1Received 2Author
9 April 2001. Accepted 27 October 2001. for correspondence: e-mail
[email protected].
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phosphatase (AP) and elevating P transport into the cell (Ray et al. 1991, Aiba et al. 1993, Aiba and Mizuno 1994, Nagaya et al. 1994). Two AP enzymes were identified and characterized: a P-irrepressible enzyme encoded by phoV (Wagner et al. 1995) and a P-repressible form, the product of phoA (Ray et al. 1991). The AP encoded by phoA was localized to the periplasmic space; its transcriptional and enzymatic activity vary with P availability (Ihlenfeldt and Gibson 1975, Block and Grossman 1988, Ray et al. 1991). In an attempt to construct a cyanobacterial P indicator, we chose to use the phoA promoter as the sensing element and fused it to bacterial luciferase (Vibrio harveyi luxAB) as its reporter. The fusion was integrated into the Synechococcus sp. PCC 7942 chromosome, yielding a strain that responds by dose-dependent light emission to a wide range of P concentrations. materials and methods Constructs. To integrate a fusion of the Synechococcus sp. PCC 7942 AP encoding gene (phoA) promoter with V. harveyi luxAB genes into the cyanobacterial chromosome, a plasmid (pAPL) was constructed. The APL plasmid was based on plasmid pAM1414 (Andersson et al. 2000), which contained the luciferase genes luxAB (V. harveyi) and a spectinomycin/streptomycin resistance cartridge. It also harbored intergenic regions of a neutral site (NSI, accession number U30252) of Synechococcus sp. PCC 7942 chromosomal DNA, which allowed the integration of the plasmid into the chromosome by a double homologous recombination (Li and Golden 1993, Liu et al. 1995). A 420-base pair (bp) fragment that included the phoA promoter (accession number D26162) was cloned by PCR using 32 base primers: 5 ataagtgcggccgctgagacagataatcaaca 3 at the 5 end of the promoter and 5 cgtaagggatcccgtcttgattactgaagaat 3 at its 3 end. The promoter was inserted into the NotI/ BamHI site of the plasmid, upstream of luxAB. The location and orientation of the fusion were confirmed by PCR and by restriction analysis with SalI. Wild-type Synechococcus sp. PCC 7942 was transformed by the recombinant and control plasmids pAPL and pAM1414 as previously described (Golden and Sherman 1984) to generate strains APL and M1414, respectively. Transformed cells were selected on agar plates that contained streptomycin and spectinomycin (25 gmL1 each) in BG-11 medium (Rippka et al. 1979). Escherichia coli DH5, used for plasmid propagation, was grown on Luria Bertani medium (Miller 1972) supplemented with spectinomycin and/or streptomycin (50 gmL1 each) when appropriate. Southern analysis. Southern blot hybridization was used to confirm the integration of pAPL into the target (NSI) site in the Synechococcus sp. PCC 7942 chromosome. Genomic DNA from the APL and wild-type strains was digested with Apa I and Mfe I and then run on a 1% agarose gel in TAE (Tris-acetate/ EDTA) buffer (Sambrook et al. 1989). DNA was transferred to an uncharged nylon membrane (NEN-New England Nuclear, Boston, MA) by capillary action in 20 SSC (sodium-chloride/ sodium-acetate) buffer (Sambrook et al. 1989). Prehybridization and hybridization were carried out in the presence of 2 SSC buffer, 0.5% blocking reagent (NEN), 5% dextran sulfate, 0.1% SDS, and 50 gmL1 denatured fragmented salmon testes sperm DNA (Sigma, Bornem, Belgium) at 60 C. Hybridization to probe was carried out overnight. The 382-bp NSI (accession number U30252) probe was cloned by PCR using a 19 base primer (5 cacatcgctatctcttagg 3) at the 5 end of the probe and an 18 base primer (5 tcgtcgaagatggaaaag 3) at its 3 end. The probe was labeled with a fluorescein nucleotide mix using a random labeling kit (NEN). Probe detection was carried out with the antifluorescein-AP antibody and a CDP- star nucleic
acid chemiluminescence reagent (NEN) followed by a 10-s exposure to x-ray film (Fuji). Growth and experimental conditions. Wild-type Synechococcus sp. PCC 7942 and its derived PphoA::luxAB engineered strain (APL) were grown in batch culture (50 mL in 250-mL flasks) at 30 C in BG-11 medium under constant fluorescent illumination (50 mol photonsm2s1) with continuous shaking (Innova 4340, New Brunswick Scientific Co., Edison, NJ; 125 rpm). For P deprivation experiments, the cells were first pregrown in a reduced-P BG-11 medium (containing 46 M K2HPO4, 20% of the original concentration) to an optical density at 750 nm (OD750) of approximately 0.6. Cells were then harvested by centrifugation (4000 rpm, Eppendorf centrifuge 5810 R [Eppendorf, Hamburg, Germany], 10 min, room temperature), washed twice in P-free BG-11 (K2HPO4 was replaced by equimolar KCl) without antibiotics, and resuspended in the same medium to a final OD750 of 0.1. For nitrogen deprivation experiments, cells were also pregrown to an OD750 of approximately 0.6, harvested by centrifugation, and washed twice with nitrogen-free BG-11 (NaNO 3 and ferric ammonium sulfate were replaced by equimolar concentration of NaCl and ferric chloride, respectively) without antibiotics, and resuspended in the same medium to a final OD 750 of 0.25. For experiments involving readdition of P to P-deprived cells, the cells were first incubated for 30 h in P-free BG-11 as described above and then supplemented either with different concentrations of K2HPO4 or with the following organic phosphates: -glycerol-phosphate (1 mM), p-nitrophenyl-phosphate (0.1 mM), D-glucose-6-phosphate (0.1 mM), D-fructose-6-phosphate (0.1 mM), ADP (0.05 mM), and K 2HPO4 (0.023 mM). These concentrations were selected in preliminary experiments as the lowest concentrations that inhibited phoA-driven bioluminescence. All organic P sources were dissolved in water and filter sterilized (0.22-m pore size filters, Schleicher & Schuell). Measurement of bioluminescence. At intervals, cell aliquots (3–5 mL) were removed from their growth flasks and brought to a uniform cell density (OD750 of 0.5). Subsamples (100 L) of the cell suspension were then transferred in duplicate to wells of an opaque, white, 96-well, microtiter plate (Nunc, Denmark). The reaction was started by addition of 100 L P-free BG-11 containing both the bacterial luciferase substrate nonanal (nonyl aldehyde, Sigma) and a detergent (Igepal CA-630, Sigma) to final concentrations of 0.002% and 0.005%, respectively. Nonyl aldehyde stock (1%) was prepared in dimethyl sulfoxide (Merck, Rahway, NJ)). The plate was immediately placed in a temperature-controlled (30 C) microtiter plate luminometer (Victor2, Wallac, Turku, Finland), and luminescence was measured every 10 s for 15 min. Light emission increased to a maximum during the first 5 min and then declined. Maximum luminescence at the peak, presented in the instrument’s arbitrary relative light units (RLU), is the value used in this communication. All experiments were conducted in duplicate and were repeated at least three times. Measurement of AP activity (APA). Whole cell APA of Synechococcus sp. PCC 7942 was measured as described previously (Ray et al 1991). Cells were collected and suspended in 0.5 mL of a buffer (0.2 M Tris-HCl, pH 8.5, 2 mM MgCl 2) containing 3.6 mM p-nitrophenyl-phosphate at 37 C for 20 min. The reaction was stopped by the addition of 50 L of 4 N NaOH and the samples were centrifuged (4 min at 7000 rpm, Eppendorf centrifuge 5417 C) to remove the cells. The OD 405 of the supernatant was measured, and concentrations were calculated using a p-nitrophenol (Sigma) standard curve. APA values are presented as nmol p-nitrophenol per h per OD750 or as a percentage of maximal APA determined with P-deprived cells at the same time point. APA of Lake Kinneret was measured as described previously (Hadas et al 1999). Water samples were filtered using 2- m polycarbonate filters (Poretics), which were then incubated in a reaction mixture similar to the one mentioned above but with 0.01 mM 4-methyl umbelliferyl phosphate (Sigma) as the sub-
MONITORING OF P BIOAVAILABILITY strate. After 0.5-h incubation, the enzymatic reaction was terminated by 1 N NaOH, and samples were filtered through 0.45- m cellulose nitrate filters. Fluorescence was determined in the filtrate using a Turner 430 spectrofluorometer (365 nm excitation, 460 nm emission), and activities were calculated against a 4-methyl umbelliferone standard curve. Field samples. Water samples were collected from Lake Kinneret, Israel, a warm monomictic freshwater lake with mean and maximum depths of 24 and 42 m, respectively. Water samples were collected in June 2000 and in February 2001 from station A (situated at the middle of the lake) to a depth of 38 m. Water temperature varied in June from 28.5 C at the surface to 15.9 C at the bottom, whereas in February it was nearly uniform throughout the water column: 17.2 C at the surface to 16.2 C at the bottom. Total dissolved P (TDP) in Lake Kinneret samples was determined according to standard methods (American Public Health Association 1995) and is defined as all organic and inorganic P that passed through a 0.45-m pore size filter. For assaying bioavailable P using the indicator strain developed in this study, 30-mL lake samples were filter sterilized (0.22-m pore size, Schleicher & Schuell) and kept in 125-mL sterile flasks. P-replete APL cells, pregrown and washed as mentioned above (see Growth and experimental conditions), were then added to the flasks to an OD750 of approximately 0.1, with three types of nutritional supplements: no addition, complete BG-11 ingredients, or P-free BG-11 ingredients. Cells suspended in complete or P-free BG-11 media served as negative and positive controls, respectively. The samples were incubated at 30 C with continuous shaking (125 rpm) under constant fluorescent illumination (50 mol photonsm2s1), and bioluminescence was assayed in 96-well, opaque, white, microtiter plates, as described above.
results Verification of APL strain chromosomal integration. To construct a P-stress reporter in Synechococcus sp. PCC 7942, we introduced a PphoA::luxAB fusion into pAM1414 to yield plasmid pAPL, which was then incorporated into the NSI site of the cyanobacterial chromosome. Homologous recombination at the target site was verified by Southern analysis using a fluorescein-labeled NSI PCR product as probe. ApaI and MfeI digests from wild-type and APL strains were hybridized with the probe (Fig. 1). In the APL strain the NSI probe hybridized to a larger fragment. The difference in size (approximately 5000 bp) corresponds to the size of the insert within the NSI in the pAPL plasmid (4894 bp), confirming that it was introduced into the genome at the NSI site by a double recombination event. Response of Synechococcus sp. strain APL to P availability. When untreated cells of Synechococcus sp. PCC 7942 and strain APL (harboring the PphoA::luxAB fusion) were transferred to a medium lacking a source of P, they continued to grow with a generation time of 7–9 h for approximately 24 h; growth then ceased and cells started to bleach, as was shown previously for this organism (Block and Grossman 1988, Collier and Grossman 1992). In strain APL, an initiation of bioluminescence was observed approximately 10–12 h after transfer of the cells to a P-deplete medium, and the intensity of light emission increased for about 45–50 h (Fig. 2). Longer incubation periods led to a decrease in light output. No luminescence whatsoever was observed in strain
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M1414 cells, which lacked the phoA promoter upstream of the luxAB genes. Neither did a limitation in nitrogen induce luminescence in APL. APA, which was monitored in parallel in the same cultures, also increased during that time period. Induction of APA, above its basic time zero level, was immediate; enzymatic activity appeared to reach a maximum after ca. 20–25 h and remained at that level for up to 50 h. When the same experiment was carried out in the presence of different PO43 concentrations (Fig. 3), the lag time preceding the onset of the luminescent response increased with increasing initial P concentration, possibly reflecting the time needed for exhaustion of P reserves: from approximately 10 h in the absence of P it increased to 20 h in the presence of 2.3 M K2HPO4. At initial P concentrations of 23 M or higher, no luminescence was detected even after 30 h. As a consequence of the different time courses observed in the presence of different initial PO43 concentrations, at any given time point the measured luminescence was dependent on the initial P concentration. In Figure 4, such a dose-response curve was plotted using the 30-h luminescence data from Figure 3. The system appeared to respond in a reproducible manner to the initial PO43 concentrations in the general range of 2 nM to 2 M. A 10-fold lower initial P concentration (0.2 nM) yielded a luminescent response similar to that of P-deprived cells (not shown). The enzymatic determination of APA, also presented in Figure 4 for the same time point, exhibits a similar response pattern but with a somewhat shifted linear detection range: around 20 nM to 20 M. To characterize the response of bioluminescent APL cultures to a renewed PO43 supply, they were prestarved for P for 30 h and then transferred to media with different PO43 concentrations (Fig. 5). In a P-free medium, luminescence continued to increase for approximately 20 more hours, whereas at P concentrations of 23 M and higher, light emission was completely inhibited within 10 h. For intermediate concentrations, a temporary decline in luminescence was followed by a renewed increase after 5 h (0.23 M) or 15 h (2.3 M). The decline, representing a temporary cessation of the phoA promoter activation and loss of active Lux protein, probably lasted for as long as PO43 was available. APA (not shown) maintained high levels throughout the transient period, probably as a result of the long lifetime of this enzyme. This difference in response sensitivity between the two activities is demonstrated in Figure 6, which records luminescence and APA 10 h after the beginning of the experiment. Quite clearly, changes in light emission were not only more rapid but were also more sensitive to low initial PO43 concentrations; the detection threshold appeared to be around 100-fold lower than that for APA. The reason for the differences may lie in the different half-lives of the two enzymes. From the kinetic data in Figure 5, a luciferase
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Fig. 2. Bioluminescence in Synechococcus sp. strain APL after transfer to a P-free (squares) or a nitrogen-free (diamonds) medium and in strain M1414 in a P-free medium (triangles). APA (circles) in strain APL, P-free medium.
Fig. 1. Southern analysis of Apa I and Mfe I digests of the genomic DNA of Synechococcus sp. PCC 7942 strain APL and wild-type. The blot was hybridized with a fluorescein-labeled probe based on a 382-bp NSI PCR product. Numbers on the right indicate molecular weights.
half-life of approximately 3 h under our experimental conditions can be calculated. AP, in contrast, is a much more stable enzyme (Li et al. 1998). From kinetic data (not shown), an approximate half-life of 10 h was derived; this value is supported by the data in Figures 6 and 7. Response to organic P. To examine the response of the reporter strain APL to P supplied in an organic form, the cells were prestarved for P for 30 h and then supplied with different organic P sources. Luminescence and APA were determined 10 h later, and the results are presented in Figure 7. All P sources used had some negative effect on bioluminescence, indicating that they served to supply the cells with P and thus repressed phoA induction at least to some extent. This inhibitory effect was almost complete for ADP and for D-fructose-6-phosphate and only partial for D-glucose-6-phosphate, p-nitrophenyl-phosphate, and -glycerol-phosphate. As seen in Figure 7, this pattern of expression was also apparent for APA, albeit at a much less pronounced manner. Availability of the organic P substrate, therefore, had a less apparent effect on long-term enzyme activity than on the induction of its promoter. Monitoring P bioavailability in natural water samples. To test the applicability of our reporter strain for as-
sessing bioavailability of P in a natural system, we exposed P-replete APL cells to water samples collected at different depths in Lake Kinneret (Sea of Galilee), Israel’s largest lake and main water reservoir, and luminescence was recorded after 25 h. Without mineral enrichment, the cells rapidly bleached and luminescence was very low; no luminescence at all was observed when excess P (230 M) was added. In contrast, after P-free mineral enrichment, a very clear luminescent signal was recorded in most Lake Kinneret samples. The example in Figure 8 presents data
Fig. 3. Bioluminescence in Synechococcus sp. strain APL after transfer to different K2HPO4 concentrations.
MONITORING OF P BIOAVAILABILITY
Fig. 4. Bioluminescence and APA of Synechococcus sp. strain APL after a 30-h exposure to various P concentrations. Data (averages standard deviation) are presented as percent of maximal luminescence (squares) or APA (circles) obtained at that time point for the P-free medium in each of three replicate experiments (1210 176 RLU and 101.3 7.4 nmol p-nitrophenolh1OD7501, respectively).
obtained in two sampling periods: during stratification in June 2000 (Fig. 8, A and B) and when the lake was mixed in February 2001 (Fig. 8, C and D). Figure 8 A and C depict the degree of luminescence induced by water samples from the different depths, along with the TDP concentrations in the same samples. Whereas in February neither TDP nor bioluminescence varied significantly throughout the depth profile, a very clear mirror image developed in June:
Fig. 5. Effects of adding different K2HPO4 concentrations to cells starved for P for 30 h.
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Fig. 6. Bioluminescence and APA in 30-h P-starved cells, 10 h after readdition of the indicated K 2HPO4 concentrations. Data (averages standard deviation) are presented as percent of maximal luminescence (squares) or APA (circles) obtained at that time point for the P-free medium in each of three replicate experiments (3070 654 RLU and 102.8 10.9 nmol p-nitrophenolh1OD750 1, respectively).
higher P concentrations near the bottom of the lake were reflected in a greatly reduced bioluminescence. Phytoplankton APA displayed a similar pattern (Fig. 8, B and D): relatively uniform water column activity in February and a much lower bottom activity during June stratification.
Fig. 7. Bioluminescence and APA in 30-h P-starved cells, 10 h after readdition of the following organic compounds at the indicated concentrations: -glycerol-phosphate (1 mM), pnitrophenyl-phosphate (0.1 mM), D-glucose-6-phosphate (0.1 mM), D-fructose-6-phosphate (0.1 mM), ADP (0.05 mM), and K2HPO4 (0.023 mM). Data ( standard deviation) are presented as percent of maximal luminescence (dark columns) or APA (open columns) obtained at that time point for the P-free medium in each of three replicate experiments (3015 519 RLU and 118.2 6.9 nmol p-nitrophenolh1O.D.7501, respectively).
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Fig. 8. Bioluminescence of Synechococcus sp. strain APL in water samples collected from Lake Kinneret in June 2000 (A and B) and February 2001 (C and D). Bioluminescence (squares) was measured 25 h after inoculation and compared with total dissolved phosphorous concentration (TDP, triangles) and to phytoplankton alkaline phosphatase activities (APA, circles) determined for the same samples. Temperature data (diamonds) are provided to demonstrate the stratification (June) and mixing (February) of the lake at the time of sampling.
MONITORING OF P BIOAVAILABILITY
discussion Analytical chemical methods can determine P concentrations with a high degree of accuracy and low detection thresholds. Nevertheless, these methodologies do not always report adequately on the availability of this element to aquatic microorganisms (Hecky and Kilham 1988, Scanlan and Wilson 1999). One complication is that P may appear in various organic and inorganic forms, dissolved and particulate, possibly in association with other molecules (Cotner and Wetzel 1992, Björkman and Karl 1994, Tezuka 1994). Another factor is that a chemical determination, as precise as it may be, presents a snapshot of ambient P concentrations without providing any information on its flux. It is not surprising, therefore, that there is a constant search for alternative methodologies to assay nutrient bioavailability as sensed by the cells (Platt et al. 1992, Parpais et al. 1996, Dyhrman and Palenik 1999, 2001). One such approach is the use of APA, known to be induced when P is scarce, as an indicator of P deficiency (Rose and Axler 1998). This widely used method suffers from several drawbacks: 1) in a natural community, it is difficult to distinguish among the different phosphatases produced by bacterioplankton, phytoplankton, and zooplankton (Boon 1994, Jamet et al. 1995); 2) there may be background levels of constitutive enzymes (Rose and Axler 1998); 3) substrate availability rather than enzyme concentration might limit the reaction rate in nature (Elser et al. 1990); and 4) AP is a highly stable enzyme that may retain its activity over extended periods (Li et al. 1998). In this communication we report a different approach to P bioavailability assessment: the fusion of the promoter of a P stress response gene (phoA) to reporter genes and monitoring its induction in real time. The use of luxAB reporter genes in bacterial constructs has previously been proposed and subsequently applied for the monitoring of environmental pollutants (Belkin et al. 1997, Erbe et al. 1996, Daunert et al. 2000, Kohler et al. 2000), and there has also been a report on assaying nutrient availability in this manner in an E. coli system (Prest et al. 1997). To our knowledge, the approach has not been implemented for assaying P limitation in photosynthetic organisms. A similar methodology was recently reported for determining iron bioavailability to Synechocystis sp. strain PCC 6803 (Kunert et al. 2000). We successfully integrated the PphoA::luxAB fusion into the chromosome of the cyanobacterium Synechococcus sp. PCC 7942, and light was emitted under P-limiting conditions. When P-replete cells were transferred to a P-free medium or when P-starved cells were exposed to fresh P, their luminescence was dependent on the initial P levels over a broad range of concentrations (Figs. 4 and 6). The detection threshold of P-replete cells was as low as 2.3 nM, whereas that of APA was 10-fold higher (Fig. 4).
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In contrast to the long lag exhibited before the development of bioluminescence upon P starvation, the increase in APA was much faster. Similarly, an 8- to 12-h lag period was noted in the same organism before AP mRNA transcripts accumulation (Ray et al. 1991). It is therefore possible that the rapid APA enhancement is due to activation of an existing enzyme and not to its de novo synthesis. Another possible explanation is that AP mRNA accumulates earlier but cannot be detected early enough by either northern blot (Ray et al. 1991) or bioluminescence (Fig. 2). In this case, the lag might be due to a slower maturation of the luciferase, perhaps because its mRNA may be less stable than the native AP mRNA. A likely explanation for the length of time required for lux and phoA mRNA expression to be apparent is the consumption of available internal P (Grillo and Gibson 1979); this is supported by the observation that in the presence of higher P concentrations the lag period was longer (Fig. 3). Similarly, the duration of the decline in luminescence after the readdition of P to induced cells was also dependent on P levels (Fig. 5). Assuming that phoA induction ceases immediately upon P replenishment to starved cells, the rate of decline of luminescence allows the calculation of an approximate 3-h half-life for luciferase activity. AP, in contrast, is a highly stable enzyme that can retain its activity over extended periods (Li et al. 1998). It is thus not surprising that the bioluminescent expression of phoA induction, besides permitting real-time biomonitoring of P availability, also exhibited a more sensitive response to changes in the concentrations of this nutrient (Fig. 6). When dissolved organic P sources were added to P-limited APL cells, luminescence declined but APA levels remained high (Fig. 7). Upon ADP addition, both luminescence and APA declined in a manner similar to that caused by inorganic phosphate. Our results correlate with the work of Cotner and Wetzel (1992) and of Björkman and Karl (1994), who investigated the bioavailability of organic P to microorganisms in freshwater and marine environments, respectively. Both reports showed that nucleotides are the most readily utilizable of the combined P sources investigated, whereas monoesters showed lower bioavailability. In a preliminary attempt to demonstrate the applicability of our reporter strain to P monitoring in natural environments, we tested its responses to samples collected along a depth profile of Lake Kinneret, Israel. Clearly, the luminescent response mirrored TDP concentrations (Fig. 8, A and C) and correlated with the phytoplanktonic APA measurements (Fig. 8, B and D), but a more detailed study is required before such data can be interpreted and translated into actual concentrations of “available P.” Nevertheless, data accumulated to date indeed suggest that reporter strain APL may serve as a useful tool for investigating and monitoring the relationships between P bioavailability and the fate of cyanobacterial popula-
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tions in Lake Kinneret or other freshwater bodies. Such data may be combined with a battery of additional assays, including APA (Rose and Axler 1998) and molecular probes (La Roche et al. 1999), to further a better understanding of the P status of phytoplankton in freshwater environments. Supported by Infrastructure grant number 1319-1-98 from the Israel Ministry of Science and the Arts and by project ENV4CT97-0493 of the European Community. We thank Dr. Susan Golden, Texas A&M University, for providing us with pAM1414. We also thank Assaf Sukenik, Riki Pinkas, Nehama MalinskyRushansky, and James Easton of the Yigal Allon Kinneret Limnological Laboratory for their help in supplying the water samples from Lake Kinneret. Support for equipment was provided in part by the Israel Science Foundation founded by the Israel Academy of Sciences and Humanities. A. F. Post received financial support from grant 525/98 from the Israel Science Foundation. Aiba, H. & Mizuno, T. 1994. A novel gene whose expression is regulated by the response-regulator, SphR, in response to phosphate limitation in Synechococcus species PCC7942. Mol. Microbiol. 13:25–34. Aiba, H., Nagaya, M. & Mizuno, T. 1993. Sensor and regulator proteins from the cyanobacterium Synechococcus species PCC7942 that belong to the bacterial signal-transduction protein families—implication in the adaptive response to phosphate limitation. Mol. Microbiol. 8:81–91. American Public Health Association. 1995. Standard Methods for Examination of Water and Waste Water, 19th ed. APHA, Washington, DC. Andersson, C. R., Tsinoremas, N. F., Shelton, J., Lebedeva, N. V., Yarrow, J., Min, H. T. & Golden, S. S. 2000. Application of bioluminescence to the study of circadian rhythms in cyanobacteria. Methods Enzymol. 305:527–42. Belkin, S., Smulski, D. R., Dadon, S., Vollmer, A. C., Van Dyk, T. K. & LaRossa, R. A. 1997. A panel of stress-responsive luminous bacteria for toxicity detection. Water Res. 31:3009–16. Björkman, K. & Karl, D. M. 1994. Bioavailability of inorganic and organic phosphorous-compounds to natural assemblages of microorganisms in Hawaiian coastal waters. Mar. Ecol. Prog. Ser. 111:265–73. Block, M. A. & Grossman, A. R. 1988. Identification and purification of a derepressible alkaline phosphatase from Anacystis nidulans R2. Plant Physiol. 86:1179–84. Boon, P. I. 1994. Discrimination of algal and bacterial alkaline phosphatases with a differential-inhibition technique. Aust. J. Mar. Freshwater Res. 45:83–107. Collier, J. L. & Grossman, A. R. 1992. Chlorosis induced by nutrient deprivation in Synechococcus sp. strain PCC 7942: not all bleaching is the same. J. Bacteriol. 174:4718–26. Cotner, J. B. J. & Wetzel, R. G. 1992. Uptake of dissolved inorganic and organic phosphorus compounds by phytoplankton and bacterioplankton. Limnol. Oceanogr. 37:232–43. Daunert, S., Barrett, G., Feliciano, J. S., Shetty, R. S., Shrestha, S. & Smith-Spencer, W. 2000. Genetically engineered whole-cell sensing systems: coupling biological recognition with reporter genes. Chem. Rev. 8:2705–38. Dyhrman, S. T. & Palenik, B. 1999. Phosphate stress in culture and field populations of the dinoflagellate Prorocentrum minimum detected by a single cell alkaline phosphatase assay. Appl. Environ. Microbiol. 65:3205–12. Dyhrman, S. T. & Palenik, B. 2001. A single-cell immunoassay for phosphate stress in the dinoflgellate Prorocentrumn minimum (Dinophyceae). J. Phycol. 37:400–10. Elser, J. J. & Kimmel, B. L. 1986. Alteration of phytoplankton phosphorus status during enrichment experiments: implications for interpreting nutrient enrichment bioassay results. Hydrobiologia 133:217–22.
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