P Metabolism in the Bean-Rhizobium tropici Symbiosis' · mineral nitrogen, but little is known about the basis for the higher P requirement. Experiments were conducted to determine

Plant Physiol. (1 997) 1 13: 1233-1 242

P Metabolism in the Bean-Rhizobium tropici Symbiosis'

Thamir S. AI-Niemi, Michael 1. Kahn, and Timothy R. McDermott*

Department of Plant, Soil, and Environmental Sciences, Montana State University, Bozeman, Montana 5971 7- 031 2 (T.S.A.-N., T.R.M.); and lnstitute of Biological Chemistry and Department of Microbiology, Washington

State University, Pullman, Washington 991 64-6340 (M.L.K.)

Nodulated legumes require more P than legumes growing on mineral nitrogen, but little is known about the basis for the higher P requirement. Experiments were conducted to determine how Rhizobium tropici responds to P limitation and to understand how P is partitioned between the symbionts under conditions of adequate or limiting P. Free-living R. tropici responds to P stress by increasing P transport capacity and inducing both an acid and an alkaline phosphatase. This P-stress response occurs when the medium P concentration decreases below 1 p ~ . Both P-stress- inducible phosphatases are found in bacteroids taken from plants growing with adequate P, suggesting that P levels in the symbiosome space is low enough to induce the expression of these enzymes. Bacteroid alkaline phosphatase-specific activity was highest during vegetative growth of the bean plant, but decreased approximately 75% during the host reproductive stages. In hydroponic experiments 32P-tracer studies showed that in vivo rates of P accumulation were significantly higher in bacteroids from P-limited plants com- pared with those from plants that had been supplied with adequate P. In contrast, label accumulation in leaves was greatest in plants grown with adequate P.

On a worldwide basis, most cultivated soils have insuf- ficient P for maximum crop yields. Legumes are particu- larly affected because they are typically cultured symbiotically, and it has been shown that legumes dependent on symbiotic nitrogen fixation have higher P requirements than legumes grown with a nitrogen fertilizer. Soybean (Cassman et al., 1981~; Israel, 1987), clover (Powell, 1977), common bean (Pereira and Bliss, 1987, 1989), pigeon pea (Itoh, 1987), and cowpea (Cassman, et al., 1981~) a11 respond positively to P fertilization. Growth and symbiotic parameters increased by P fertilization include whole-plant N concentration, plant dry matter, nodule number, nodule mass, nitrogenase activity, and specific nodule nitrogenase activity.

The reasons for this P response are poorly understood. In soybean symbiotic N, fixation places a P tax on the host plant (Israel, 1987). Field studies with peanuts have found that P concentrations in the stems and seeds of nodulating

' This work was initiated while T.R.M. was a National Science Foundation postdoctoral fellow in plant biology (BIR-9203796) in the laboratory of M.L.K. This work was also supported by U.S. Department of Agriculture Competitive Grants Office grants 90- 37262-5166 (M.L.K.) and 94-37305-0574 (T.R.M.). This is Montana State Agricultura1 Experiment Station Contribution no. 1-5114.

* Corresponding author; e-mail [email protected]. edu; fax 1-406-994-3933.

lines are lower than those in nonnodulating isolines (Sahra- wat, et al., 1988), suggesting that P distribution within the plant is different when the legume plant grows symbiotically. The higher P requirement of symbiotically grown legumes is apparently not due to different abilities of roots in the two N environments to absorb P (Israel, 1987), which implies that an optimum symbiotic interaction between the host plant and rhizobia depends on an efficient allocation and use of available P.

Most P metabolism studies with rhizobia have used free- living cells. P requirements for growth (Keyser and Munns, 1979; Cassman et al., 1981a, 1981b), P transport rates, and P storage (Cassman et al., 1981b; Beck and Munns, 1984) vary between strains. Many rhizobia are similar to other gram- negative bacteria (Torriani, 1990; Wanner, 1993) in that P stress induces phosphatases and leads to significantly increased P transport rates (Smart et al., 1984a). In addition, activities important in plant-microbe interactions, such as synthesis of nodulation metabolites in Rhizobium leguminosarum bv trifolii (McKay and Djordjevic, 1993), polysaccha- ride production in Rhizobium meliloti (Zhan et al., 1991), and regulation of virulence genes in Agrobacterium tuinefaciens (Winans, 1990; Mantis and Winans, 1993), are sensitive to P availability. Since even in fertile soils the soil solution P concentration may be 510 FM (Barber, 1984), it is important to understand how both symbionts respond to low P during nodule formation and during symbiosis.

In conjunction with studies addressing the genetics of P metabolism, we conducted experiments to more thor- oughly understand the P requirements of the rhizobia during symbiosis. The objectives of this study were to compare P metabolism in Rhizobium tropici bacteroids and free-living cells, investigate how bacteroids respond when the host plant is P-stressed, and, to a limited extent, assess how P is partitioned between the symbionts.

MATERIALS A N D METHODS

Bacterial Strains and Plant Material

Rhizobium tropici strain CIAT899 was used throughout these experiments. It was cultured in yeast extract mannitol broth (Somerville and Kahn, 1983) when used to inoculate

Abbreviations: MMOP, minimal mannitol-zero phosphate; +P, 750 FM P; -P, without P; PEB, periplasm extraction buffer; PEB-C, PEB supplemented with 50 pg/mL chloramphenicol; TRB, transport rinse buffer; TRB-arsenate, TRB amended with 0.5 m M arsenate.

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1234 AI-Niemi et al. Plant Physiol. Vol. 11 3, 1997

common bean (Phaseolus uulgaris cv Viva Pink) for symbiosis studies. For P-starvation and P-induction experiments, the cells were grown in a minimal-mannitol broth (Somer- ville and Kahn, 1983) that contained no P (MMOP) but was buffered with 5 mM Mes and 10 mM Mops, pH 6.7. Potas- sium phosphate was added to this medium as required.

Plant Crowth Conditions

Bean seeds were surface-sterilized by washing with 95% ethanol for 30 s and 3% sodium hypochlorite for 5 min, and then rinsed six times with sterile, distilled water. Seeds were planted in washed quartz sand in 8-inch-diameter pots and inoculated with R. fropici strain CIAT899 (approximately 5.0 X 109 cells per pot). The plants were grown in a greenhouse under natural light supplemented with artificial light (80 p~ s-l m-'; 16-h day, 8-h night) and were irrigated with a nutrient solution as described by McDermott and Kahn (1992) that contained 750 p~ P, but modified such that FeC12.3H,0 was replaced by an equivalent amount of che- lated iron (Sprint 330, Ciba-Geigy, Hummert International, St. Louis, MO). The daytime temperature ranged from 22 to 27"C, and at night the temperature was 19°C.

lnduction Studies

Phosphatase Activity

R. tropici was grown to mid-log phase in MMOP containing 5 mM K,HP04, washed twice in MMOP, and then resuspended in MMOP. At various times, culture aliquots were processed to determine acid phosphatase, alkaline phosphatase, or P transport activities. To extract phosphatases and other periplasmic proteins, cells were pelleted, washed once in PEB consisting of 30 mM Tris and 20% (w/v) SUC, pH 8.0, then resuspended in PEB, and extracted on ice in the same buffer with lysozyme (0.5 mg/mL) for 5 min. EDTA was then added to the cell suspension to a final concentration of 1 mM and the cells were incubated on ice for an additional 15 min. The preparation was then centrifuged for 5 min at 12,OOOg. The supernatant containing periplasmic proteins was used for the enzyme assays or for nondenaturing polyacrylamide gel analysis (described be- l o ~ ) of phosphatase electrophoretic patterns. The cyto- plasm marker enzyme malate dehydrogenase was not released by this procedure (data not shown). Alkaline phosphatase activity was measured by recording the hy- drolysis of p-nitrophenyl phosphate (Smart et al., 1984b). Units for these assays are reported as nanomoles or micro- moles per minute per milligram of protein. Protein concentration was determined using a protein assay kit (Bio-Rad).

In P-depletion experiments alkaline phosphatase induction was monitored in parallel with the P concentration in the medium. R. tropici cells were washed twice in MMOP-50 p~ K,HPO, at room temperature to avoid cold shock, and then the cells were resuspended in MMOP-50 p~ K,HPO,. At 1-h intervals samples were assayed for alkaline phosphatase activity, and the P concentration in the medium was measured (Chen et al., 1956). Culture pH was monitored to verify that the Mes I Mops buffers in MMOP were maintaining pH. In these experiments alkaline phosphatase

was measured with intact cells; units were based on culture A595 and are reported as nanomoles substrate hydrolyzed per minute per A595.

Nondenaturing polyacrylamide gels (McDermott and Kahn, 1992) were used to separate phosphatase activities in periplasmic extracts of free-living cells and bacteroids. Alkaline phosphatase activity was visualized by incubat- ing the gels in 30 mM Tris, pH 8.5, 50 mM MgCl,, 0.5% (w / w) Fast Blue RR (4-benzoylamino-2,5-dimethoxy- benzenediazonium chloride hemi[zinc chloride] salt), and 0.5% (w / w) a-naphthylphosphate, monosodium salt. To detect acid phosphatases, the gels were presoaked in 30 mM acetate buffer (pH 5.5) until the equilibrium pH of the presoak buffer was 6.0. Fresh acetate buffer containing the staining reagents was then added. The gels were dried between cellulose films (gel drying kit, Promega), the image was scanned using a ScanJet scanner (IIcx, Hewlett- Packard), and then the NIH Image program (version 1.52) for the Macintosh computer was used to analyze the scan to determine phosphatase activity band intensities (see Fig. 2A).

P Transport

P transport rates in R. tropici were determined using [32P]KH,P0, which was used at an assay concentration of 50 p~ and a specific activity of 22.5 pCi pmol-'. The 32P-labeled P solution was filtered through a 0.45-pm pore size nitrocellulose filter (Schleicher & Schuell) prior to use. Cell concentration was 50 pg (dry weight) mL-'. During the 40-s transport assay, 0.5-mL samples were withdrawn every 10 s, rapidly filtered through 0.45-pm pore size nitrocellulose filters, and immediately washed with 10 mL of the TRB, which was composed of 20 mM Mes and 5 mM KH,PO,, pH 6.5. Filters were presoaked in TRB to minimize the binding of label that had not been taken up by the cells. Accumulation of label in the cells was measured by a direct counting of the filters using Cerenkov radiation (Hariland and Bieber, 1970).

Bacteroid Studies

Phosphatase Activity

Bacteroids were harvested at the V3, R1, and R4 growth stages (Fehr et al., 1971), which corresponded to approximately 32 to 36, 45 to 49, and 65 to 69 d after planting, respectively. At each harvest, crown nodules were rapidly stripped from the roots, briefly washed in cold, distilled water, and then homogenized in ice-cold PEB-C. The nodule homogenate was filtered through four layers of Mira- cloth (Calbiochem) and then centrifuged for 5 min at 3508 at 4°C to pellet the remaining nodule debris. The supernatant was transferred to a different tube and centrifuged for 8 min at 3,OOOg. This low-speed centrifugation is not quantitative for bacteroid recovery, but avoids plant organelles (Day et al., 1986; see "Bacteroid 32P Accumulation in Vivo," below). The supernatant was discarded and the surface of the bacteroid pellet was gently washed twice with PEB-C, and then resuspended in PEB-C. The bacteroid suspension was treated with lysozyme and incubated on ice for 5 min,

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Rhizobium tropici P Metabolism 1 2 3 5

and then EDTA was added and the incubation continued for another 5 min. The bacteroid suspension was then centrifuged at 12,OOOg for 5 min to separate the bacteroids from the periplasmic proteins released during the lysozyme-EDTA treatment. The supernatant contained the periplasmic proteins. The volume of PEB-C used to resus- pend the bacteroid pellet and the volumes of the EDTA and lysozyme stock solutions used to extract bacteroid periplasmic proteins were determined in preliminary experiments. The optimum volume of PEB-C, EDTA (100 mM), and lysozyme (10 mg mL-' in PEB) per gram of bacteroid pellet wet weight was 2.3, 0.12, 0.05 mL, respectively. Incubation of the bacteroids in EDTA was optimal at 5 min, since this releases periplasmic proteins without cell lysis. Longer exposures (e.g. 10 min) led to bacteroid lysis, as determined by the release of malate dehydrogenase, an intracellular enzyme, into the periplasmic extracts. Malate dehydrogenase is an extremely active enzyme in bacteroids and thus is a sensitive measure of bacteroid lysis. The periplasmic extracts were then assayed for alkaline phosphatase activity or electrophoresed in nondenaturing polyacrylamide gels to detect alkaline phosphatase and acid phosphatases, as described above for free-living cells.

Bacteroid P Transport

Bacteroids were harvested from nodules using Perco11 gradients and methods described by Day et al. (1989). Washed bacteroids were then assayed for P transport using the methods described above for free-living cells.

Whole-Plant Labeling Experiments

Hydroponically grown plants were required for the 32P- labeling studies. Plants were first grown in a sand culture with the +P nutrient solution, and then at the V2 growth stage (approximately 15-20 d after planting; visually in- spected to verify that nodulation was well established). Seedlings were gently removed from the sand by submerg- ing the pot in tap water and slowly pouring out the water- saturated sand. The plants were transferred to 25-L con- tainers outfitted with perforated air lines that were secured to the bottom of each container. Twelve plants per container were supported in holes drilled in sheets of 1.5-inch- thick rigid foam. The plants were separated into +P and -P treatments, receiving the same nutrient solution as described above for plants grown in pots, except that Mes buffer (1 mM) was included to provide some pH buffering capacity in -P treatments, and was also added to the +P nutrient solution as a control. Plants were cultured hydroponically for 30 d under the greenhouse conditions described above. The nutrient solutions were completely replaced on alternate days.

For 32P labeling, individual plants were transferred to 500 mL of the +P nutrient solution containing [32P]P0, (750 p~ I'; specific activity = 0.45 pCi pmol-I P) in 900-mL beakers. In each case, a soft foam-rubber collar was at- tached to the stem to suspend the plant at a height at which the entire root system was submerged. The beakers and plants were gently swirled at intervals during the labeling

period to maintain an even distribution of the nutrients per label.

Bacteroid 32P Accumulation in Vivo

At each time point, two to three nodules were carefully removed to avoid rupturing the nodule cortex and washed three times in TRB-arsenate. For each wash, nodules were vortexed in a 1.8-mL microcentrifuge tube, and the wash solution was removed by vacuum. The nodules were crushed in the microcentrifuge tube in 0.5 mL of fresh TRB-arsenate and then briefly centrifuged in the same microcentrifuge tube to pellet the nodule debris (5 s at 12,OOOg). To avoid disturbing the loose pellet, 250 pL was carefully removed from the top of the supernatant, transferred to a fresh microcentrifuge tube, and centrifuged at 3,OOOg for 4 min. The supernatant was completely removed by vacuum, and the pellet was resuspended in 1.0 mL of TRB-arsenate and centrifuged again at 3,0008 for 4 min. The supernatant was removed and the bacteroid pellet was resuspended in 1.0 mL of TRB-arsenate. This protocol did not give quantitative bacteroid recovery, but because we were concerned about the possible loss of label by P exchange from bacteroids that might result from prolonged bacteroid purification protocols, our objective was to avoid plant organelles and minimize the time interval following nodule detachment. The total time from nodule crushing to bacteroid sampling was less than 10 min. In addition to arsenate, which is an inhibitor of l' transport in bacteria, the 5 mM P in TRB lowered the specific activity of the label during bacteroid isolation, effectively stopping label incorporation into the bacteroids.

In separate experiments, the relative enrichment of the bacteroids in the final sample that was used to assess bacteroid label accumulation was estimated by comparing the levels of the mitochondria marker enzyme Cyt c oxidase with that of the bacteroid marker enzymes Ala dehydrogenase and p-hydroxybutyrate dehydrogenase. A11 fractions (including supernatant fractions that were collected by pi- pet) in the above bacteroid isolation protocol were sonicated three times for 30 s on ice and centrifuged to remove the debris. Cyt c oxidase was measured using the method described by Smith (1955). P-Hydroxybutyrate dehydrogenase was measured by following the increase in A,,, in a reaction that contained 25 mM P-hydroxybutyrate, 0.5 mM NAD', 150 mM Tris buffer (pH &O), and cell extract (Reibach et al., 1981). Ala dehydrogenase was determined from the decrease in A,,, in a reaction that contained 0.1 mM NADH, 5 mM sodium pyruvate, 10 mM NH,C1, 50 mM Tris buffer (pH 8.5), and cell extract (Reibach et al., 1981). The final bacteroid fraction contained 54% of the 0-hydroxybutyrate dehydrogenase activity, 44% of the Ala dehydrogenase activity, and less than 5% of the Cyt c oxidase activity. No Ala dehydrogenase or 0-hydroxybutyrate dehydrogenase activity was detectable in any of the samples unless the samples were first sonicated. This was evidence that the cytoplasmic membrane of the bacteroids was intact and that the distribution of the enzymes was a reliable indication of bacteroid distribution in the samples and of the purity of the bacteroid fraction. The specific activities of the bacteroids P-



1236 AI-Niemi et al.

hydroxybutyrate dehydrogenase and Ala dehydrogenase were 2.1 and 0.31 pmol min-' mg-' protein, respectively. This leve1 of p-hydroxybutyrate dehydrogenase is considerably higher than that reported for soybean bacteroids (Wong and Evans, 1971; Karr et al., 1984), but is similar to that previously reported for R. fropici bacteroids (Romanov et al., 1994).

Based on the relative distribution of enzymes among the fractions, it was concluded that the protocol selected for bacteroids and only limited amounts of plant organelles were present. Using the Percoll gradient technique, we obtained a similar segregation of bacteroids and organelles, except bacteroid yields were always nearly quantitative. Percoll gradients are not the method of choice in these particular experiments because they require more time to run and the bacteroids must be washed to remove the Percoll, which would otherwise interfere with bacteroid mass estimates based on absorbance or protein content (Reibach et al., 1981). Such manipulations must be mini- mized when trying to estimate in vivo uptake of the P ion, which is subject to exchange. Even though our method did not quantitatively recover bacteroids, it was rapid and efficient for obtaining bacteroids that were relatively free of plant organelles.

Aliquots of the resuspended final bacteroid pellet were used to determine A,,,. Optical density was converted to dry weight using the empirical equation y = 0.803~ - 0.0006, r2 = 0.992; where y = milligrams of bacteroid cell dry weight and x = As,,. To determine radioactivity in bacteroids, 0.5-mL aliquots of the bacteroid suspension were collected by suction on 0.45-pm nitrocellulose filters that had been presoaked in 10 mM KH,PO, to limit the binding of any label released by bacteroids or carried through the wash protocol. The filters were washed with 10 mL of TRB-arsenate, and 32P was quantified by measuring Cerenkov radiation.

Nodule 32P Accumulation

Two to three nodules were washed three times in TRB- arsenate, briefly dried with tissue paper, weighed, and then extracted in Soluene-350 (Packard Instrument Co., Downers Grove, IL) for 24 h. For nodule and bacteroid samples (above), a single plant represented one replicate, with a11 of the sampled nodules removed from that plant during the labeling period. A11 nodules were removed from the same part of the root system of each plant, using younger nodules from the outer region of the nodule clus- ter that was located at the root crown. Nodule dry weight was not determined, since labeled nodules were placed directly into the digestion solution immediately after washing and weighing. However, using unlabeled nodules, bean nodule moisture content was determined to be 87.9 2 0.3% (mean 2 SE of 11 replicate samples), and there was no significant difference in moisture content between +P nodules and -P nodules. This value was used to convert nodule fresh weight to nodule dry weight so that direct comparisons to bacteroid values could be made.

Plant Physiol. Vol. 11 3 , 1997

Leaf 32P Accumulation

Preliminary experiments showed that 32P accumulation differed significantly between leaves at different positions on the stem and between leaflets in a trifoliate leaf. There- fore, a different sampling scheme was used for leaves. Instead of using a single plant per replicate throughout the entire labeling period, which was done with the bacteroid and nodule samples, a single leaflet was removed from one plant and then the plant was discarded. At each time point for each treatment, the middle leaflet of the uppermost (youngest), fully opened trifoliate leaf was taken from three different plants. Leaf samples were weighed and extracted in Soluene-350.

After the leaf and nodule samples were extracted, Hionic-Fluor scintillation cocktail (Packard Instrument Co.) was added and the amount of 32P was quantified in a TRI-CARB liquid scintillation analyzer (model 2200CA, Packard Instrument Co.). Sample debris was allowed to settle prior to counting. Leaf extracts contained chlorophyll, so a quench curve was constructed by varying the amount of unlabeled leaf extract in the scintillation fluid and estimating the chlorophyll content by its A,,, using the relationship (Arnon, 1949):

chlorophyll ( p g mL-') = 29 x A,,,.

Each sample was then spiked with an equal amount of [32P]P0,. Throughout the range of the chlorophyll stan- dards measured (A,,, = 0.1-1.5), A,,, followed the equation y = 1.415x1.004 (y = micrograms of chlorophyll, x = A,,, r2 = 1.0). A quench curve was then determined by comparing apparent radioactivity with sample chlorophyll content. This relationship followed the equation y = -461.44~ + 45,699 (r2 =' 0.97), with y units in cpm and x units in micrograms of chlorophyll per milliliter. A11 radio- label data were corrected for background counts.

Statistical Analysis

Where statistical treatment was appropriate, the data were analyzed using analysis of variance, with means com- pared by protected LSD (Steel and Torrie, 1980).

RESULTS

Free-Living R. tropici

Initial studies were designed to determine what environmental conditions were required to induce a P-stress response in R. tropici and whether the induced response was like that in other gram-negative bacteria. After resus- pending free-living cells in zero-P medium, alkaline phosphatase-specific activity and the rate of P transport quickly increased (Fig. 1). After approximately 4 h, alkaline phosphatase activity appeared to reach a maximum of 1800 nmol min-' mg-' protein. Alkaline phosphatase activity in P-starved cultures varied between 1500 and 1800 nmol min-l mg-' protein, with the extent of induction always at least 35-fold greater than in +P cells, where the alkaline phosphatase-specific activity ranged from 20 to 40 nmol min-l mg-' protein. At time O, I' transport rates of both +P



Rhizobium tropici P Metabolism 1237

o Phosphorus Uptake 0 Phos• Phosphorus Uptake + Phos

n Alkaline Phosphatase Activity 0 Phos• Alkaline Phosphatase Activity + Phos

r2000

Incubation Period (h)

Figure 1. Induction of alkaline phosphatase activity and enhanced Ptransport in free-living R. tropici in response to P starvation. Alkalinephosphatase activity was monitored using p-nitrophenyl-P as a sub-strate, and P uptake was measured using 32P. Log-phase cells grownin a minimal-mannitol ammonium broth (starting P concentration =10 mM, also buffered by 10 mM Mops and 5 mM Mes) were washedin the same medium that lacked P and then resuspended in thatmedium containing 10 mM P ( + P) or no added P ( — P). Results arefrom a representative experiment showing a typical induction of theP-stress response. Details of cell harvest and assays are described in"Materials and Methods." Phos, P; dry wt., dry weight.

Neither of the acid phosphatase bands were visible in gelsstained at pH 8.5.

The P concentration required to induce the P-stress re-sponse was determined in P-depletion experiments by si-multaneously monitoring alkaline phosphatase activityand the P concentration in the medium. Mid-log-phasecells were washed and resuspended in MMOP amended to50 JU.M KH2PO4, and then P concentration in the mediumand alkaline phosphatase activity were measured at vari-ous times (Fig. 3). Preliminary experiments had shown thatalkaline phosphatase assays with whole cells and withextracted periplasmic proteins were superimposable underthese conditions. Therefore, the whole-cell assay was usedto follow the induction of the enzyme because it was more

Time of Induction (h)

and — P cells were approximately 2 nmol min"1 mg"1 celldry weight. Transport rates in — P cells increased rapidly toapproximately 95 nmol min"1 mg"1 cell dry weight, whichrepresents an almost 50-fold increase in response to Plimitation. Such induction responses are consistent withthose found for other rhizobia (Smart et alv 1984b) and inEscherichia coli (Torriani, 1990; Wanner, 1993).

There were two bands of acid phosphatase activity in R.tropici periplasmic extracts that were well separated fromthe position of the alkaline phosphatase bands (Fig. 2). Wehave not yet determined the conditions under which theacid phosphatase activities can be measured individuallyin extracts without interference by each other or by thehighly expressed alkaline phosphatase, so the activities ofthe acid phosphatases were qualitatively measured in non-denaturing polyacrylamide gels. The slower-migratingacid phosphatase (Fig. 2, band I) appeared to be inducibleby P starvation, since it was seen only in P-stressed cells,typically appearing after about 1 h of induction. The stain-ing intensity of the faster-migrating acid phosphatase band(Fig. 2, band R) decreased after incubation under P-stressconditions, suggesting that it was repressed by P-stressconditions. Relative to that necessary for alkaline phospha-tase, large amounts of periplasmic extract and long incu-bations were necessary to observe the acid phosphatasebands. This resulted in the alkaline phosphatase formingbands in gels stained for acid phosphatase, but alsoshowed that the acid phosphatase bands were electro-phoretically distinct from the alkaline phosphatase bands.

0 2 4 6 8

Time of Induction (h)

Figure 2. Acid phosphatase activity stains in nondenaturing poly-acrylamide gels. Periplasmic extracts of free-living R. tropici wereobtained from cells incubated in a minimal-mannitol broth contain-ing no added P but buffered with 5 mM Mes and 10 mM Mops. Detailsof induction, protein extraction, electrophoresis, and gel staining aredescribed in "Materials and Methods." Incubation times are shown inhours, and lanes were loaded with 20 .̂g of protein. A, The band ofthe slower-migrating, P-starvation-inducible phosphatase is desig-nated I, and the band of the faster-migrating, P-starvation-repressedphosphatase is designated R. Gel polarity is indicated by + and -.The pH of the staining buffer was not optimum for alkaline phospha-tase staining activity, but the alkaline phosphatase enzyme bandswere still evident and are identified as AP. B, Relative stainingintensity of the I and R acid phosphatases was determined usingscanning densitometry as described in "Materials and Methods."Intensities are plotted relative to 100%, defined as the R band attime 0. www.plantphysiol.orgon June 1, 2020 - Published by Downloaded from

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1238

--o- Culture Viable Counts

+ Alkaline Phosphatase Activity

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Plant Physiol. Vol. 113, 1997

--D- Culture Optical Density

---C Medium P concentration

I 50 0.25 7

40

30

20

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Figure 3. lnduction of alkaline phosphatase during medium P depletion. Representative results from one of severa1 experiments that document this response. Log-phase cells were washed and resuspended in a minimal-mannitol broth containing 50 p~ K,HPO, and incubated at 30°C with shaking. The P-stress response was monitored by assaying for alkaline phosphatase activity in whole-cell suspensions. At the indicated times, cell samples were withdrawn, centrifuged (supernatant completely removed by suction to avoid any inhibitory effects of remaining P on alkaline phosphatase activity), resuspended in 30 mM Tris (pH 8.5) and 50 mM MgCI,, and assayed for phosphatase activity as described in “Materials and Methods.” Samples of the cleared supernatant were saved and later measured for remaining P. Results are plotted as a function of culture-viable counts (A) and culture optical density and medium P concentration (B).

convenient and because the experiment only required measuring relative amounts of alkaline phosphatase during induction.

Alkaline phosphatase activity was not detected until the P concentration in the spent medium was less than 1.0 WM

(Fig. 3). Even though the washed cells had not been grown in low-P medium, they rapidly removed P from the new medium, suggesting that they could take up P at reasonably high rates using constitutively expressed P permeases. Culture-viable counts increased throughout the course of this experiment. Cells were dividing even after the culture P concentration had been reduced to micromolar levels, typically completing approximately two rounds of division.

Bacteroid and Plant Studies

Bacteroid Phosphatase Activity in Sand-Cultured Plants Grown with the + P Nutrient Solution

At different stages of the symbiosis, alkaline phosphatase activity was determined in bacteroids taken from sand-cultured plants irrigated with the 750 PM P nutrient solution. Three separate greenhouse experiments were conducted, with the results of a representative experiment shown in Figure 4. Bacteroids harvested at the host V3 growth stage had extremely high alkaline phosphatase specific activities and were consistently higher than those observed with P-stressed free-living cells. However, at later stages of the symbiosis, bacteroid alkaline phosphatase activities were much lower. At the R4 stage, alkaline phosphatase activity was approximately 25% of that measured in the host vegetative stage. The alkaline phosphatase activity stain patterns obtained with periplasmic extracts of P-stressed free-living cells and of bacteroids taken from V3 plants grown in sand with the +P nutrient solution are shown in Figure 48. In other samples of I‘-starved free- living cells, alkaline phosphatase banding patterns were

indistinguishable from those of the bacteroid preparations (results not shown). Even though bacteroid alkaline phosphatase specific activity decreased with time, the bacteroid alkaline phosphatase activity stain profile did not differ between the bacteroid samples that were taken at the different harvest dates.

Acid phosphatase activity was also detected in bacteroids throughout the growth cycle. The pattern of bacteroid acid phosphatase bands in native gels was similar to that of the P-starved free-living cells; the low-mobility, P-stress- inducible acid phosphatase was present (Fig. 5, band I), and the high-mobility, P-stress-repressible acid phosphatase (Fig. 5, band R) was absent.

P Transport in Bacteroids Harvested from + P and - P Hydroponic Plants

P transport rates were determined with isolated bacteroids that were harvested from hydroponically grown plants that had been cultured for 30 d with either the +P or - P nutrient solution. In these experiments the Perco11 gradient technique (Day et al., 1989) was used for quantitative recovery of purified bacteroids, and chloramphenicol was not included in the isolation buffers. Regardless of the host-plant P nutritional treatment, bacteroid P-transport rates were considerably lower than those of P-stressed free-living cells (Table I). These rates could be increased by starving the bacteroids for I’, but even after 6 h the rates were still much lower than in similarly treated free-living cells. Alkaline phosphatase activity in these bacteroids was approximately 1.0 Fmol min-l mg-l protein and did not differ between P treatments. This leve1 of alkaline phosphatase activity in bacteroids of hydroponic plants was similar to bacteroids of same-age plants that were grown in sand and irrigated with the high-P nutrient solution (45 d after planting; Fig. 4A).




>.*J'>t)

0)c

£ 4o

I ^

I 2

Hs ^=20 30 40 50 60 70

Days After Planting

B 0);oSv

Figure 4. Alkaline phosphatase activity in R. tropici bacteroids andfree-living cells. A, Alkaline phosphatase-specific activity in R.tropici bacteroids at the V3, R1, and R4 growth stages. The results arerepresentative of one of the three experiments conducted, in whichbacteroids were purified from nodules of bean plants grown in sandand irrigated with the +P nutrient solution. Each data point is themean of three replicate samples, and error bars (where visible)indicate 1 SE. Differences between data points are statistically signif-icant as determined using analysis of variance and protectedLSDa=005. B, Alkaline phosphatase activity stains obtained from R.tropici periplasmic proteins separated by electrophoresis in nonde-naturing polyacrylamide gels. Periplasmic proteins were extractedfrom P-stressed free-living cells and from bacteroids purified fromnodules harvested from bean plants (V3 growth stage) grown in sandand irrigated with the +P nutrient solution. Each lane was loadedwith 10 fAg of protein. Polarity of the gel is indicated by + and -.

Free-Living Bacteroid-P +P

Figure 5. Acid phosphatase activity stains obtained from electro-phoresis of R. tropici periplasmic proteins in nondenaturing poly-acrylamide gels. Extracts were taken from R. tropici free-living cellsand bacteroids. For free-living cells, periplasmic proteins were ex-tracted from cultures grown with 10 mM P (+P) or washed andincubated for 4 h in a minimal-mannitol broth containing no addedP (-P), but buffered with 5 mM Mes and 10 mM Mops. Bacteroidperiplasmic proteins were extracted from bacteroids harvested fromnodules at the V3 growth stage. Details of induction, periplasmicprotein extraction, and gel staining are described in "Materials andMethods." Lanes were loaded with 20 ^g of protein. The inducible,slower-mobility acid phosphatase and the P-stress-repressible, faster-mobility acid phosphatase are indicated by I and R, respectively, asin Figure 2. Gel polarity is indicated by + and -.

+P plants had taken up only 32 nmol P g"1 bacteroid dryweight. By contrast, 32P accumulation in leaves of the +Pplants was significantly greater than in leaves of the -Pplants (Fig. 7). After the 2-h labeling period, P accumulatedin the +P leaves was approximately 24.2 nmol g~a freshweight and that in the — P leaves was approximately 3.2nmol g"1 fresh weight.

DISCUSSION

In R. tropici the coordinated increase in P transport andphosphatase activity in response to P stress resembled thatfound in other bacteria (Torriani, 1990; Wanner, 1993) andoccurred when the medium P concentration was ^1 /U.M(Figs. 1-3). Alkaline phosphatase was clearly the dominantp-nitrophenyl-P phosphatase present in the P-starved cells.Alkaline phosphatase activity stains in nondenaturing

Labeling Studies with Hydroponic Plants Cultured in the+ P and -P Nutrient Solutions

To further assess bacteroid P acquisition capability andinvestigate how bacteroids respond to host-plant P nutri-tion, bacteroid P accumulation was measured in plantausing 32Pi. Radioactivity was rapidly taken up by the nod-ules of both P-stressed and +P plants (Fig. 6A). Uptake wasslightly higher in nodules of P-stressed plants, although thedifference was not statistically significant until 60 min.Transfer of phosphate to the bacteroids was significantlygreater in P-stressed plants than in +P plants (Fig. 6B),began immediately, and was nearly linear with time. At theend of the 1-h labeling period, bacteroids of the -P plantswere estimated to have accumulated approximately 440nmol P g"1 bacteroid dry weight, whereas bacteroids of the

Table I. Phosphate transport by R. tropici bacteroids isolated fromnodules of hydroponic bean plants cultured with +P nutrient solu-tion or -P nutrient solution for 30 d

Data are from a single representative experiment. Bacteroids wereisolated from nodules as described by Day et al. (1989), washed, andresuspended in MMOP, except that mannitol was replaced with 5mM succinate. After measuring P transport at time 0, the bacteroidsuspension was split into equal portions, with P (10 mM) added to oneportion and no P (0 mM) added to the other.

BacteroidOrigin

+ P plants

-P plants

IncubationTime

h

0606

P Transport Rate of

10 mM P

nmol P min~ 'mg~ '

1.7

2.2

Bacteroids

0 mM P

cell dry wt

3.628.0

1.618.5



1240 AI-Niemi et al. Plant Physiol. Vol. 11 3, 1997

600 -

u - P

o 20 40 60 80

6

T

- - I I I

20 40 60 80

Time (min)

Figure 6 . A, P accumulation in nodules. Plants were grown hydroponically for 30 d in the + P nutrient solution or in the -P nutrient solution, then labeled with 32P as described in "Materials and Meth- ods." Each point and error bar is the average and range of two mean values (three replicate samples per mean) obtained from different sets of plants from two independent greenhouse experiments. Differences are not statistically significant except at 60 min (determined by analysis of variance; protected LSD,=,,~~). B, P accumulation by R. tropici bacteroids in planta. Plants were grown hydroponically for 30 d in the +P nutrient solution or in the -P nutrient solution, and then labeled with 32P as described in "Materials and Methods." Results are from a typical experiment. Each point represents the mean of three replicates (+P) or two replicates (-P). Error bars, where visible, represent the SE. Differences at each time point are statistically significant as determined with analysis of variance (protected LSD,,,,,,). dry wt, Dry weight.

polyacrylamide gels revealed severa1 bands in the periplasmic extracts of bacteroids and P-starved free-living cells. The bacteroid phosphatase staining pattern gave a constant number of bands at a11 of the harvest dates. In free-living cells the pattern and intensity of the bands did not appear to vary during induction, but would vary between cultures and sometimes differed from bacteroids, as shown in Fig- ure 4B. In E. coli alkaline phosphatase staining can vary with growth conditions (Signer et al., 1961; Piggot et al., 1972; Nakata et al., 1977) and occurs because of two possible signal peptide processing sites. This results in the active dimer being composed of monomer combinations that vary in net charge and therefore have different electrophoretic mobilities in native gels (Nakata et al., 1978; Karamyshev et al., 1994).

Acid phosphatase activity in nondenaturing polyacrylamide gels revealed one acid phosphatase staining band that was seen only after shifting the cells to a zero-P medium. The appearance of this acid phosphatase band was associated with the reduction in staining of a constitutively expressed acid phosphatase. It is possible that this represents induction and repression of two proteins. Alterna- tively, the P-stress-inducible acid phosphatase band may result from a posttranslational modification of the consti- tutive phosphatase that yields a native protein differing in net charge. Neither of the acid phosphatase bands was observed in gels stained at an alkaline pH.

Bacteroids harvested from plants grown with the non- limiting P nutrient solution closely resemble the P-stressed free-living cells by having very high levels of alkaline phosphatase activity and only the slow-mobility form of acid phosphatase. This observation suggests that the inor- ganic P concentration in the symbiosome space is low. The high alkaline phosphatase activity in bacteroids was prob- ably not due to protein synthesis during the bacteroid preparation, since bacteroids were kept well chilled throughout the short isolation procedure (approximately 15 min) and chloramphenicol was included in the extraction buffer. In the control experiments, the induction of alkaline phosphatase in free-living R. tropici incubated in MMOP was inhibited by the addition of 50 p g mL-' chloramphenicol (L.M. Botero and T.R. McDermott, unpub- lished data). The high alkaline phosphatase activity in bacteroids was also not due to contamination by host enzymes. Nodule cytosol fractions electrophoresed in nondenaturing polyacrylamide gels and stained for phosphatase activity show three bands of acid phosphatase activity, but no bands are seen w h e n duplicate gels are stained at pH 8.5 (McDermott and Kahn, 1994).

The decrease in bacteroid alkaline phosphatase activity during the plant reproductive stages may have been due to

30' T 25- - +p - -P

20 -

O 1 2 1 3

Time (h) Figure 7. P accumulation in leaves. Plants were grown hydroponically for 30 d in the + P nutrient solution or in the -P nutrient solution. Details of plant culture and 32P labeling are in "Materials and Methods." Each point and error bar (where visible) i s the average and range of two mean values (three replicate samples per mean) obtained from different sets of plants from two independent greenhouse experiments. Differences are statistically significant at each time point (determined by analysis of variance; protected LSD,= , ,~~) .




more P being made available to the bacteroids, resulting in repression of alkaline phosphatase. However, developing seeds are a major P sink and would be expected to compete for P, perhaps limiting P flow to the nodule. It is possible that phoA was actively expressed at this stage, but that a large fraction of alkaline phosphatase was lost from the older bacteroids. The potential loss of periplasmic proteins through an unstable bacteroid outer membrane (Bal et al., 1980; Bal and Wong, 1982; Katinakis et al., 1988; Kinnback and Werner, 1991) was a concern, so a rapid, differential centrifugation technique was employed to reduce both the processing steps and the time. Percoll gradients were not used to isolate the bacteroids because with this bean culti- var, published Percoll procedures offer no advantage in maintaining intact symbiosomes during isolation.

P transport rates of the isolated R. tropici bacteroids (Table I) were similar to those reported by Smart et al. (1984a), who found that the P transport rate in isolated snake bean bacteroids was lower than that in free-living cells and was not substantially influenced by P nutrition of the host plant. Whereas the phosphatase activities in R. fvopici bacteroids suggested low-P conditions, high-P transport activity was not observed with Percoll gradient- purified bacteroids. The P transport capacity of Percoll- purified R. tropici bacteroids incubated in the absence of P increased with time of incubation, but the final rate was substantially lower than that in comparably induced free- living cells. Low-P transport activity may be an artifact, perhaps resulting from the loss of a low-molecular-weight periplasmic phosphate-binding protein during bacteroid purification. Such proteins are required for P-specific transport systems in many gram-negative bacteria (Willsky and Malamy, 1976; Gerdes et al., 1977; Poole and Hancock, 1984), and synthesis of this protein is inhibited by high-P growth conditions. Snake bean rhizobia apparently do not require such a protein for P transport (Smart et al., 1984a), but a binding protein may be involved in the P transport in R. melilofi (Bardin et al., 1996; T. Finan, personal communication).

Because of the potential difficulties in quantifying P transport in isolated bacteroids, 32P uptake and partition- ing between the symbionts was measured in vivo, using a rapid protocol for isolating bacteroids. In contrast to the lack of difference in in vitro P transport rates measured with isolated bacteroids from +P and -P hydroponic plants, the supply of P to the host significantly influenced bacteroid P acquisition in planta. Comparing the rates of bacteroid P accumulation between P treatments with nodule P uptake rates suggested that increased bacteroid P uptake in response to P stress in the symbiosis was independent of possible differences in nodule permeability to P. Additional experimentation will be required to determine if increased in vivo bacteroid P accumulation rates are due to a nove1 mechanism that increases the P supply to the bacteroids or if a trivial explanation could account for this observation. An example of the latter might be larger P pools in +P nodules leading to isotope dilution, which in turn would lead to an underestimation of P accumulation in +P bacteroids. It should be noted, however, that the

changes in apparent P accumulation rates in leaves were opposite those observed in bacteroids, even though this same isotope dilution effect might be expected in these tissues as well.

Only limited amounts of plant shoot material were analyzed for label incorporation; consequently, any assessment of label distribution throughout the -P and +P hydroponic plants must likewise be limited. However, it is evident that the label accumulation profiles of the leaves were very different from those observed in the nodules and bacteroids. This argues that the significant differences in P accumulation between -P and +P bacteroids did not sim- ply reflect the proportional differences in P accumulation in plants that may have been taking up P at substantially different rates.

A C K N O W L E D G M E N T

The authors thank Andrew Glenn for stimulating discussions.

Received August 16, 1996; accepted December 18, 1996. Copyright Clearance Center: 0032-0889/97/ 113 / 1233/ 10.

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Documents

P Metabolism in the Bean-Rhizobium tropici Symbiosis' · mineral nitrogen, but little is known about the basis for the higher P requirement. Experiments were conducted to determine