biochemical characterization and subcellular localization ... · plant physiol. (1997) 114: 907-915...

Plant Physiol. (1997) 114: 907-915

Biochemical Characterization and Subcellular localization of the Red Kidney Bean Purple Acid Phosphatase’

Anil G . Cashikar, Rathinam Kumaresan, and N. Madhusudhana Rao*

Centre for Cellular and Molecular Biology, Uppal Road, Hyderabad 500 007, lndia

Phosphatases are known to play a crucial role in phosphate turnover in plants. However, the exact role of acid phosphatases in plants has been elusive because of insufficient knowledge of their in vivo substrate and subcellular localization. We investigated the biochemical properties of a purple acid phosphatase isolated from red kidney bean (Phaseolus vulgaris) (KBPAP) with respect to its substrate and inhibitor profiles. The kinetic parameters were estimated for five substrates. We used 31P nuclear magnetic resonance to investigate the in vivo substrate of KBPAP. Chemical and enzy- mological estimation of polyphosphates and ATP, respectively, indicated the absence of polyphosphates and the presence of ATP in trace amounts in the seed extracts. lmmunolocalization using antibodies raised against KBPAP was unsuccessful because of the non- specificity of the antiserum toward glycoproteins. Using histoenzymological methods with ATP as a substrate, we could localize KBPAP exclusively in the cell walls of the peripheral two to three rows of cells in the cotyledons. KBPAP activity was not detected in the embryo. In vitro experiments indicated that pectin, a major component of the cell wall, significantly altered the kinetic properties of KBPAP. The substrate profile and localization suggest that KBPAP may have a role in mobilizing organic phosphates in the soil during germination.

Phosphorus is an important macronutrient in plant growth and plays a vital functional role in energy transfer and metabolic regulation, constituting 0.3 to 0.5% of the dry weight of a plant. In ensuring sufficient Pi for the developing embryo, the seed has to rely on abundant organic phosphates in the soil rather than on soluble ri, which is only present at low concentrations (<I mM) (Bould and Hewitt, 1963). Extracellular phosphatases present in cell walls or in exudates from roots have been reported to be involved in mobilizing organic phosphates in the soil (Goldstein et al., 1989). Extracellular and secre- tory phosphatases have been demonstrated to be regulated by the Pi status of the cell, and may form a “salvage system,” which converts organic or leaked, esterified phosphate to Pi for reabsorption (Duff et al., 1991). On germination the amount of APase was shown to increase significantly in seeds, a process controlled by GA, (Bhargava and Sachar, 1987). Despite extensive reports of the occurrence of APases in a variety of plants and plant tissues, the exact

This work was supported by a senior research fellowship to A.G.C. from the Council for Scientific and Industrial Research, government of India.

* Corresponding author; e-mail [email protected]; fax 91-40-671195.

907

physiological role of these enzymes has yet to be established. Shortcomings of most of these reports are: (a) the inability to identify the in vivo substrate(s) and the lack of an attempt to investigate the kinetic differences between various substrates; (b) the inability to localize the enzyme at the subcellular level; and (c) the lack of thorough characterization of structural properties, unique features, if any, metal composition, and inhibitor / substrate profiles of the enzyme (Duff et al., 1994).

APases are enzymes that catalyze the hydrolysis of monoesters and anhydrides of phosphoric acid in the pH range of 4 to 7 (Vincent and Averill, 1990). The plant enzymes are mostly dimeric glycoproteins, and have subunit molecular masses of 50 to 60 kD. Differential glycosylation of the subunits, leading to a heterogeneity in the molecular mass of the subunits, has been reported for KBPAP (Stahl et al., 1994) and sunflower seed APases (Park and van Etten, 1986). Acid phosphatases are normally observed to be localized in the cytosol, vacuoles, or cell walls. Acid phosphatases from various sources have been shown to have specificities for various substrates such as 3-phosphoglycerate (Randall and Tolbert, 1971), PEP (Duff et al., 1989), phytate (Gibson and Ullah, 1988), and Tyr- phosphorylated proteins (Gellatly et al., 1994).

KBPAP was purified from red kidney bean (Phaseolus vulgaris) (Beck et al., 1986) and characterized with respect to its secondary structure and amino acid and metal composition (Cashikar and Rao, 1995). KBPAP is a dimeric glycoprotein (molecular mass 110 kD) with an intersubunit disulfide linkage. When antibody raised against KBPAP was used, immunological cross-reactivity was observed with potato tuber acid phosphatase but not with acid phosphatases from sweet potato or mammals (Cashikar and Rao, 1995). The primary structure of the protein and the structure of the oligosaccharide have been established (Klabunde et al., 1994; Stahl et al., 1994). Recently, the x-ray structure of this protein was solved at a 2.9-A resolution (Strater et al., 1995).

We have demonstrated the role of the intersubunit disulfide bond in the stability of the dimer (Cashikar and Rao, 1996a), as well as stabilization of the tertiary structure of the dimer upon binding phosphate, an inhibitor (Cash- ikar and Rao, 1996b). Despite extensive structural informa- tion on KBPAP and its abundance in the seed, its physio-

Abbreviations: APase, acid phosphatase; KBPAP, red kidney bean purple acid phosphatase; pNPP, p-nitrophenyl phosphate; polyP, polyphosphate.

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908 Cashikar et al. Plant Physiol. Vol. 11 4, 1997

logical role, localization, and substrate/ inhibitor profiles have not been clearly established. We present histoenzymological data on the subcellular localization of KBPAP, as well as a detailed kinetic characterization with severa1 substrates and inhibitors, with the objective of defining its physiological role.

MATERIALS A N D METHODS

KBPAP was purified in the laboratory as described by Cashikar and Rao (1995). A 200-fold purification was achieved, as confirmed by electrophoresis and spectroscopy. The substrates were from Sigma. pNPP was from Sisco Research Laboratory (Bombay, India). Pectin with about 6% methoxylation was obtained from S.D.Fine Chemicals, Ltd. (Hyderabad, India).

Enzyme Activity Assays

Enzyme assays were done in 50 mM acetate buffer (pH 5) and 500 mM NaCl containing the appropriate concentration of the required substrate. The reaction was started by the addition of 200 to 300 ng of enzyme per milliliter of the reaction mix. Phosphate released was estimated using the acid-acetone-molybdate reagent (Heinonen and Lahiti, 1981). At the end of the incubation time (5-10 min at room temperature) the reactions were stopped by the addition of the molybdate reagent, which brings the pH to <1. In assays for pNPP hydrolysis the reaction was stopped by the addition of 2 volumes of 0.5 M NaOH, and A410 was measured.

Enzyme assays were also performed in the presence of 0.1% pectin in the reaction mixture. In these assays the enzyme was preincubated with pectin for 5 min before it was added to the reaction mixture. The reactions were carried out at low ionic strength (in the absence of NaCl) in 10 mM Tris (pH 7) with ATP or pNPP as the substrate.

For determination of catalytic parameters such as K , and k,,, for various substrates, concentrations ranging from 0.01 to 10 mM were used. In the case of pNPP, concentrations ranging from 0.1 to 100 mM were used. K , and k,,, were calculated by the nonparametric method described by Cornish-Bowden and Wharton (1988).

For measuring the pH optimum, acetate buffer was used in the pH range of 3 to 6 and Tris-maleate-NaOH buffer was used in the pH range of 5 to 9. The activity measured in the two buffers in the overlapping pH range of 5 to 6 were identical for a11 substrates tested. The actual pH in the reaction mixture before addition of the enzyme was noted and used for representing the data.

The temperature optimum for activity was measured with pNPP as the substrate in the presence of 50 mM acetate (pH 5 ) and 500 mM NaCl. Enzyme a t a concentration of 6 ,ug/mL in 500 mM NaCl was incubated at the respective temperatures for 1 h. The reaction mixture was pre-equilibrated to the required temperature before the addition of the enzyme. The reactions were stopped by the addition of 2 volumes of ice-cold 0.5 M NaOH to stop the temperature-mediated hydrolysis of pNPP.

-

Extraction of Soluble Phosphorus Compounds from Seeds

Extraction of soluble phosphoesters was done by mild acid treatment of the seed meal at low temperature (Barany and Glonek, 1982). Perchloric acid (6 mL of 3.6% [VIVI acid) was added to the fine, dry seed meal(2 g) and mixed thoroughly. It was allowed to stand on ice for 10 min and was then centrifuged at 10,OOOg for 5 min. The supernatant was neutralized to pH 6 using 2 N KOH and kept on ice for another 10 min. The insoluble potassium salt of perchloric acid was removed by centrifugation at 10,OOOg for 5 min. Extracts of seeds soaked in water for 12 to 16 h were prepared using identical procedures. The weight of the dry seeds before soaking was considered for the preparation of the extracts. For NMR experiments EDTA was added to the extracts to a final concentration of 5 mM.

Measurement of the in Vivo ATP Concentration

The amount of ATP present in the extracts was estimated using a luciferase-luciferin-based ATP bioluminescence assay kit (CLS, Boehringer Mannheim). The light intensity was measured using a luminometer (LKB 1250). ATP was also estimated in extracts prepared from seeds soaked in water overnight.

3'P-NMR of the Seed Extract

31P-NMR spectra were recorded on a pulsed Fourier transform-NMR spectrometer (AM300, Bruker, Billerica, MA) operating at 121.5 MHz. The signals were collected with a spectral width of 8.9 Hz and a total of 2K time- domain data points were used. No decoupling of protons was employed. The sample was taken in a 10-mm tube and an interna1 lock with 4% D,O was used. The sample was spun at 15 Hz. Chemical shifts were measured with respect to methylene diphosphonic acid, the chemical shift of which was referenced to external 85% HJO,. About 400 signals were collected at 25°C. The signals were multiplied with an exponential filter and Fourier-transformed with a 4K memory size. In experiments involving the external addition of enzyme, 120 Fg of KBPAP in 10 pL was added to the sample. The spectra were recorded after 1 h of incubation at room temperature.

Activity Staining of Seed Sections

Light Microscopy

Red kidney bean (Phaseolus vulgaris) seeds were swollen in distilled water for about 12 h. The swollen seeds were flash-frozen in liquid nitrogen and thawed to -20°C in a cryomicrotome (Histostat, American Optical, Hong Kong). Ten-micrometer sections were attached to clean coverslips, air-dried for 30 min, and fixed in a fixative containing 3.5% formaldehyde, 5% acetic acid, and 50% ethanol. After fix- ing the sections were washed in water for 10 min. Staining of the sections for acid phosphatase activity was done using the lead sulfide method (Davenport, 1960). Reaction mixtures containing 50 mM acetate buffer (pH 5), 4 mM lead nitrate, and 1 mM ATP were preincubated at room temper-



Characterization of Red Kidney Bean Purple Acid Phosphatase 909

ature for about 2 h to allow precipitation of free phosphate that may have been present along with ATP. One-hundred microliters of this reaction mixture was placed on each coverslip and incubated at room temperature for 1 to 2 h. Another set of coverslips was incubated with reaction mixture not containing ATP, and these were used as substrate blanks. One set of sections was also treated with reaction mixture containing p-glycerophosphate as a substrate in- stead of ATP. At the end of 2 h of incubation the coverslips were washed thoroughly with water for about 10 to 15 min, and incubated in 0.5% ammonium sulfide solution for 3 to 5 min at room temperature. The sections were washed with water as before and mounted on glass slides. The stained sections were observed under a phase-contrast microscope (Dialux 2, Leitz, Wetzlar, Germany).

Transmission Electron Microscopy

Swollen red kidney bean seeds were cut into approximately l-mm3 blocks. Acid phosphatase ultrahistochemistry was performed using ATP as the substrate and lead nitrate as a capturing agent (Pearse, 1972). The cotyledon blocks were fixed for 10 min at 4°C in 10% neutralized formaldehyde or 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7), washed with deionized water, and then incubated for the required duration (typically 1-2 h) at 37°C in a mixture containing 50 mM acetate buffer (pH 5), 4 mM lead nitrate, and 1 mM ATP. Blocks incubated in reaction mixtures not containing the substrate were used as negative controls. After washing the samples in water, they were fixed in 3% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.3) for 12 h and postfixed in 1% osmium tetroxide (Pearse, 1972). The blocks were further embedded in Araldite (CIBA-CIGY, Bombay, India) and 0.5-pm sections were cut using a microtome (Ultracut E, Riechert-Jung, Vienna, Austria). The sections were attached to copper grids, stained for 2 h with 2% aqueous uranyl acetate, and viewed under an electron microscope (JEM 100 CX, Jeol) using 120 kV of accelerating voltage.

RESULTS AND DlSCUSSlON

Substrate Specificity

A common difficulty in defining the physiological role of phosphatases has been the identification of their in vivo substrates. In only a few studies have attempts been made to identify kinetic differences among the substrates, which help in defining sharply the functional role of these enzymes (Duff et al., 1994). KBPAP purified in the present study was tested for its activity on several substrates (Table I). The activity against pNPP was taken to be 100%. We observed that activity was highest against polyP (average chain length, approximately 16 residues), followed by activity against ATP. Acid and alkaline phosphatases are thought to be nonspecific toward the leaving group of the substrate (Vincent et al., 1992). We found that KBPAP could not hydrolyze a number of substrates, such as P-glycerophosphate, that are regularly used in the detection of acid phosphatases. The enzyme also could not hydrolyze the plant phosphate-storage compound phytate.

Table 1. Substrate specificity for the KBPAP

are represented relative to the activity for pNPP (taken as 100%). Each substrate was used a t a concentration of 1 mM. All activities

Substrate Activity

pNPP ATP ADP dATP Pyrophosphate polyP (approximately 16 residues) PEP Fru-l,6-diP Phosphocreatine 2,3-Diphosphoglycerate Phosphoglycolate AMP Phytate P-Gl ycerophosphate 3-Phosphogl ycerate 2-Phosphogl ycerate CIC-6-P CIC-1-P Fru-6-P 5-Phosphoribosyl 1 -pyrophosphate Phosphotyrosine Phosphothreonine cAMP

%

1 O0 596 134 547 372 597 266

26 16

7 2

N.D." N.D. N.D. N.D. N.D. N.D. N.D. N.D. N.D. N.D. N.D. N.D.

a N.D., Not detected.

When the substrates were categorized based on their phosphate-transfer potential (Jencks, 1976), ATP, ADP, pyrophosphate, polyP, and PEP, with a phosphorylating potential greater than 5 kcallmol, were hydrolyzed by KBPAP efficiently. Creatine phosphate, with a phosphorylating potential of 10.3 kcal/mol, was hydrolyzed very poorly, probably because of the P-N bond. The substrates that KBPAP hydrolyzes are, in general, more anionic, containing a second negative charge in the leaving group. An anion-binding pocket has been observed approximately 9 A away from the zinc atom of the active site of the enzyme (Strater et al., 1995). Therefore, it is plausible that the KBPAP active site may be more suited to hydrolyzing phosphates esterified to anionic leaving groups.

Since the enzyme was maximally active against polyp, we looked for the presence of polyP in kidney bean seeds. polyPs, found in a number of microorganisms (Harold, 1966) and very few plants (DeMason and Stillman, 1986), were thought to be reserves of phosphate and energy. The presence of polyPs in higher plants has not been clearly established. The soybean vegetative storage proteins VSPa and VSPP were shown to possess polyphosphatase activity (DeWald et al., 1992). We have extracted the seed powder by several procedures using both weak acids and alkalies, and estimated polyP in these extracts (Wood and Clark, 1988). polyP-binding dyes such as DAPI, which bind to polyanions, could detect only the DNA present in the nuclei but not polyP. 31P-NMR of seed extracts to detect polyp, which has a characteristic chemical shift, could not demonstrate the presence of polyP in the seeds.



91 O Cashikar et al.

The absence of polyP in the seed suggests that KBPAP may not function in vivo as a polyphosphatase.

The K,, k,,, and specificity constants were determined for many of the substrates that were hydrolyzed by KBPAP (Table 11). The K , for ATP, the second-best substrate in vitro, was estimated to be 0.53 mM at pH 5, and the specificity constant (kcat/Km) was about 10-fold lower than that for polyP. The highest turnover number (kcat) observed so far for any phosphatase is about 1 O O O / s for the bovine spleen purple acid phosphatase (Vincent et al., 1992). The k,,, for ATP hydrolysis by KBPAP at 25°C and pH 7 (the pH optimum for ATPase activity) was found to be 1294/s. These studies indicate that ATP is probably the in vivo substrate for KBPAP. Presentation of ATP as Mg-ATP had no effect on the hydrolysis of ATP at a range of ATE’ concentrations. The mung bean cotyledon acid phosphatase, which has a molecular mass similar to that of KBPAP, was also shown to be an ATPase involved in the recycling of phosphate (Haraguchi et al., 1990). We estimated the amount of ATP present in the seed extracts to be 22.28 t 1.68 nmollg (or 3.72 -C 0.28 PM) in dry seeds and 36.95 t- 2.87 nmol/g (6.17 t 0.48 PM) in soaked seeds. The observed values were much below the K , of KBPAP for ATP.

Protein Tyr phosphatase activity has been observed in the potato tuber acid phosphatase (Gellatly et al., 1994). Earlier studies in our laboratory have shown that antibodies to KBPAP cross-react with potato tuber acid phosphatase (Cashikar and Rao, 1995). The inability of KBPAP to hydrolyze both phosphotyrosine and phosphothreonine rules out the possibility of its being a protein phosphatase. An acid phosphatase from Penicillium funiculosum was shown to be capable of hydrolyzing synthetic phosphodiesters (Yoshida et al., 1989). We investigated if KBPAP could hydrolyze a nonsynthetic phosphodiester such as cAMP and found that it was inactive against this compound.

pH Optima

Severa1 studies carried out on the functions of various acid phosphatases do not report the pH optimum for enzymatic activity. It is important to know the pH optimum of activity to understand the in vivo function and also the regulation of enzyme activity of these enzymes. We tested

Table II. Kinetic parameters for the KBPAP Assays were performed in 50 mM acetate buffer (pH 5)/0.5 M NaCl

at 25°C. K, and k,,, values were determined by the method described by Cornish-Bowden and Wharton (1988), with at least 10 substrate concentrations. Enzyme concentration was normally between 2 and 5 nM.

Substrate Knl k C , l kca/Krr s- I M- 1 IllM s- 7

ATP 0.53 2 0.10 390 2 20 7.3 x 105

pNPP 7.1 2 1.5 145 +- 17 2.0 x 104

PPi (pH 6.2) 1.8 ir 0.2 315 +- 32 1.8 x 105 PEP 0.85 2 0.1 1 84 +- 8 9.8 x 104

ATP (pH 7) 1.68 2 0.22 1294 2 324 7.7 X I O5

POlYP 0.060 ? 0.021 500 +- 40 8.3 X 106

1

O

1

h

> a .3

‘.5 o

1 2

O

1

O

Plant Physiol. Vol. 114, 1997

I ATP

I pNPP

I Polyphosphate

I Pyrophosphate

3 4 5 6 7 8 9

PH Figure 1. p H optima for KBPAP activity with various substrates. Enzymatic activity was measured in 50 mM acetate buffer (between p H 3 and 6) or 50 mM Tris-maleate-NaOH buffer (between p H 5 and 9) containing 0.5 M NaCI. Taking the highest activity as 1, the activity was normalized with each substrate.

the pH optimum of KBPAP activity on four substrates and found it to be dependent on the substrate used (Fig. 1). To ensure proper measurement of pH, it was measured in the reaction mixture containing a11 of the components. Acetate buffer was used in the pH range 3 to 6 and Tris-maleate- NaOH buffer was used in the pH range 5 to 9 at a concentration of 50 mM. An overlapping pH range between 5 and 6 ensured that there was no effect of the two buffer systems on the enzymatic activity.

We observed a pH optimum of about 5.2 for pNPP, whereas the pH optima were 6 and 6.2 for polyP and PPi as substrates, respectively. For ATP the optimum pH of activity was 7. The pK, values for the secondary phosphate ionization in the case of ATP, PPi, and tripolypho- sphate have been reported to be 7.68, 6.61, and 6.54, respectively (Phillips et al., 1965). The pH optima of various substrates may reflect the ionic state of the substrate and enzyme complex. Smooth pH versus activity profiles suggest that one enzyme-substrate complex was present in the range of pH values tested, unlike the soybean cell wall phosphatase, in which more than one complex was postulated to be present (Ferte et al., 1993). It is of partic- ular interest that KBPAP shows a pH optimum of 7 for its ATPase activity, because this means that it probably functions as a neutra1 rather than as an acid ATPase in vivo.



Characterization of Red Kidney Bean Purple Acid Phosphatase 91 1

Phosphorus NMR Studies on Seed Extracts

From the data reported above it would appear that the natural substrate for KBPAP may be ATP, PPi, polyP, or PEP. In an attempt to identify the in vivo substrate, we extracted the kidney bean seed powder from both dry and soaked seeds with mild acid. The acid extracts were sub- jected to 31P-NMR spectroscopy after the addition of EDTA to a final concentration of 5 mM, so as to chelate the metal ions that might cause interference in the spectra. Several phosphate signals were well resolved in the region between 3 and 5 ppm, which indicates the presence of severa1 phosphomonoesters and Pi (Fig. 2) . The spectral region below the 2.5 ppm was silent. To this extract an excess of purified KBPAP was added with the supposition that externally added enzyme may selectively hydrolyze one or a11 of the esters that could be detected online as a decrease in the peak area of phosphate ester. Even after a 1-h incubation with excess purified KBPAP, no difference could be detected in the peak areas upon integration of the spectra before and after addition of the enzyme. The ability to hydrolyze externally added pNPP indicates that the enzyme was fully functional under NMR experimental conditions and that the free phosphate present in the extract was not inhibitory. Extraction of the seed meal using perchloric, trichloroacetic, or hydrochloric acid resulted in similar NMR profiles. The observed phosphoesters could be sugar phosphates, which could not be hydrolyzed by KBPAP (see Table I).

R

I " " " " ' " I 8 , ' , I I , , , I , , ,

5 : g a 1 5 4 1 8 3.- PPM

Figure 2. 31 P-NMR spectrum of the acid-soluble phosphoesters from the red kidney bean seed meal. Under the experimental conditions the compounds with the chemical shift between 3 and 5 ppm were mainly phosphomonoesters and Pi. Several sugar monophosphates and P-glycerophosphate showed chemical shifts between 3.5 and 5 ppm. Individual peaks were not conclusively identified. The NMR spectrum after the externa1 addition of KBPAP was identical to the spectrum shown.

Table 111. Effect o f va r iou anions and metal ions on the activity of KBPAP

Each inhibitor was used at a concentration of 10 mM. Enzyme activity against pNPP was measured at 25°C in 50 mM acetate buffer (pH 5)/0.5 M NaCI. Only sodium salts of the anions and chloride salts of cations were used to avoid nonspecific counter-ion effects. Activ- ity in the absence of any inhibitor was taken as 100%.

lnhibitor Activity

%

Anions Control Borate Carbonate Nitrate Sulfate Sulfite Tartarate Citrate Arsenate Fluoride Chloride Bromide lodide Phosphate POW Mo1 ybdate Pol ymolybdate

Metal ions C02+ CU*+ Fe2+ Ca2+ Mg2+ Mn2+ zn2+

1 O0 134 134 102 81

3 83

106 79 20

1 O0 93

102 14

1 O O

80 58 69

1 O 0 1 O0 1 O0 23

lnhibitors of KBPAP

Several anions and cations were analyzed for inhibition of pNPP hydrolysis at pH 5 by KBPAP (Table 111). We found that sulfite, fluoride, phosphate, and molybdate were the strongest inhibitors of enzymatic activity of KBPAP. As expected of purple acid phosphatases, the enzyme was not inhibited by tartarate (Vincent and Averill, 1990). Polyan- ionic inhibitors such as polyphosphate and polymolybdate inhibited the enzyme activity completely. This behavior was similar to our observations that substrates with anionic leaving groups were preferred at the active site. Among anions only borate and carbonate were observed to enhance the activity by about 30%.

Among the cations tested Zn2+ had an inhibitory effect and the inhibition was of mixed type. Zn2+ has been shown to be essential for the enzymatic activity of KBPAP. When KBPAP was dialyzed against EDTA, the activity was reduced to 50% of the initial value, which could be restored by adding Zn2+. A complete loss of activity could be observed upon prolonged dialysis against EDTA at higher temperatures (Suerbaum et al., 1993). However, at 10 miv Zn2+ about 80% of the activity was inhibited. The inhibitory effect of other divalent metal ions such as Co2+, Cu2+, and Fez+ was weaker compared with Zn2+. Other cations



91 2 Cashikar et al. Plant Physiol. Vol. 114, 1997

such as Ca2+, Mg2+, and Mn2+ did not affect enzyme activity (Table 111).

Thermal Stability of KBPAP

The acid phosphatase from soybean seeds (Ullah and Gibson, 1988) and the cell wall-localized acid phosphatase from Brassica nigra suspension cells (Duff et al., 1991) were shown to be heat-stable. To study the thermal stability of KBPAP, the protein was incubated at various temperatures for 1 h and activity against pNPP was measured at the incubation temperature (Fig. 3). It was found that the rate of enzyme activity increases linearly up to 60"C, after which the activity decreases very sharply. The activity at 60°C was about 3.6 times that observed at 30°C. Activity measurements done at 25°C after incubating the enzyme at various temperatures for 1 h indicated that the Tm of KBPAP was 65°C (data not shown). The sharp loss in enzymatic activity after 6OoC indicates that the protein undergoes an irreversible denaturation beyond 60°C. Stud- ies in our laboratory have shown that KBPAP, when bound to phosphate, is stabilized against heat denaturation (Cash- ikar, 1996) or guanidinium chloride (Cashikar and Rao, 199613). From the temperature-versus-activity data, activa- tion energy of pNPP hydrolysis was calculated to be 10.86 kcal / mol.

Subcellular Localization of KBPAP

Subcellular localization helps in defining the function of an enzyme in vivo. Based on fractionation procedures (Ferte et al., 1993) and on the requirement of high salt for extraction (Cashikar and Rao, 1995), KBPAP was thought to be associated with the cell wall. Glycosylation patterns suggested that KBPAP may be localized in the protein bodies of the seed (Stahl et al., 1994). We investigated the localization of KBPAP in the seed by activity-based histochemical methods using light and electron microscopy. Immunofluorescence and western analysis (data not shown) using polyclonal antiserum raised against KBPAP in rabbits were unsuccessful due to the nonspecific nature of the antiserum (an observation made by others as well when plant glycoproteins were used as antigens [Ramirez-

0- 20 30 40 50 60 70 80

Temperature ( O C ) Figure 3. Effect of temperature on KBPAP activity. The enzyme was incubated at the indicated temperatures for 1 h, followed by measurement of the enzymatic activity at the corresponding temperature. Enzyme blanks at each temperature were done separately and used for correction of nonenzymatic hydrolysis of pNPP.

Soto and Poretz, 19911). Generally, acid phosphatases are localized using p-glycerophosphate as a substrate. The use of ATP in the lead-precipitate method of localization of phosphatases has also been reported in eukaryotic cells (Novikoff, 1970). Since KBPAP could not hydrolyze P-glycerophosphate, we used ATP as the substrate (see Table I). Although the pH optimum for ATPase activity of KBPAP was found to be 7, we performed the localization studies at pH 5 to prevent the detection of various other ATPases, which would normally be expected to be active at pH 7 and not at pH 5. KBPAP is active over a broad pH range (Fig. 1). Ten-micrometer sections of the frozen samples were treated with the substrate for light microscopy as detailed in "Materials and Methods." Free phosphate released on KBPAP action was precipitated as insoluble lead phosphate using lead nitrate (Davenport, 1960).

Upon microscopic examination of the sections, the cotyledon cells were found to be packed without intercellular spaces. Each cell contained between 5 and 10 highly re- fractile starch granules, the identity of which was confirmed by staining the sections with iodine (data not shown). These studies clearly demonstrated that the activity was predominantly present in two to three rows of cells in the periphery of the cotyledon (Fig. 4, A and B). The epidermal cells did not show the presence of KBPAP activity. Very little or no activity was observed in cells toward the center of the cotyledons. Activity was also not observed in the embryo (Fig. 4B). The specific activity (units min-' mg-I protein) of KBPAP in crude extracts of embryos was less than 5% compared with cotyledon extracts under identical conditions. At higher magnifica- tions the ATPase activity of KBPAP was found to be localized mainly in the periphery of the cotyledon cells (data not shown). The sections that were treated with reaction mixture not containing the substrate did not show staining in any region (Fig. 4C).

Protein bodies do not show good contrast in light microscopy; so we used electron microscopy to accurately identify KBPAP activity at the subcellular level. Blocks of approximately 1 mm3 were cut from the peripheral regions of the cotyledons and were processed as described in "Ma- terials and Methods." The electron-microscopic studies indicated that the lead precipitate was present only in the cell wall of the cotyledon cells (Fig. 5). The protein bodies were densely packed and appeared in the cytosol as electron- dense organelles that were free of any lead precipitates.

lnfluence of Pectin on Substrate Binding

Since microscopy established the exclusive presence of KBPAP in the cell walls, we studied the influence of pectin, the major component of the cell wall, on KBPAP function. Pectin, a polymer of a-galacturonic acids, imparts a strong negative charge to the cell wall. It constitutes nearly 35% of the cell wall material in dicotyledons. KBPAP is a cationic protein that strongly binds to the cell wall and requires high salt (0.5 M NaC1) for desorption. The strong negatively charged surface of the cell wall has a pronounced effect on the distribution of anions such as phosphatase substrates,



Characterization of Red Kidney Bean Purple Acid Phosphatase 913

Figure 4. Light microscopy of activity staining of seed sections usingATP as a substrate. Activity staining was carried out as given in"Materials and Methods" using ATP as substrate at pH 5. A schematiccross-section of the seed shown at the left indicates the position ofthe sections in A and B. A, Staining pattern in a transverse sectiontaken from the peripheral region showing the seed coat and the outerlayer of the cotyledon. B, Staining pattern in a transverse sectiontaken from the peripheral region of the cotyledon adjacent to theembryo. No activity was observed in the embryo. Staining was absentin cells toward the center of the cotyledon. Sections that were treatedidentically in the absence of the substrate were negative for staining(C). Staining with /3-glycerophosphate was negative. SC, Seed coat; E,embryo.

and may affect the binding constants of these substrates.We estimated the kinetic parameters of ATP and pNPP forKBPAP in the presence of pectin and in the absence ofsodium chloride at pH 7. The Km for ATP in the absence ofpectin was 1.15 mM. It increased 2-fold, to 3.1 HIM, in thepresence of 0.1% pectin. Excess pectin did not further affectthe Km of ATP. The effect of pectin on pNPP binding wassimilar. The Xm for pNPP at pH 7 changed from 14 mM inthe absence of pectin to 31.4 HIM in its presence. Theobserved effects of pectin would only be apparent becausethe high pi (>9) value of KBPAP suggests a strong bindingof KBPAP to the polyanionic pectin, which reduces theconcentration of anions, i.e. substrates, in the vicinity andthereby increases the substrate concentration required forhalf-maximal velocity.

Possible in Vivo Functions of KBPAP

Based on the sequence of oligosaccharide glycans linkedto KBPAP, Stahl et al. (1994) suggested that the enzymewas present in the protein bodies. Our studies using phos-phatase activity staining could not detect any activity in theprotein bodies. Furthermore, in the acidic environment ofprotein bodies and vacuoles, KBPAP would function inef-ficiently as an ATPase. Our histoenzymological studieshave indicated that KBPAP was localized in the cell wallsof the peripheral two to three rows of cotyledon cells.Using the staining patterns in sections taken at variousregions of the seed, we tried to reconstruct the KBPAPlocation in the mature seed with the help of computergraphics programs. From the reconstruction it appears thatbecause of its peripheral localization KBPAP may play arole in the supply of phosphate to the developing embryo.Exclusive abundance and avid binding with the cell wallpectin strongly argues for a role in supplying phosphate todeveloping embryos from soil phosphate esters. Based onthe following observations, it is clear that natural substratefor KBPAP is not present in the seed: (a) ATP and polypho-sphate may not be the in vivo substrates, given their verylow concentration in the seed; (b) none of the monoestersand diesters of phosphates present (31P-NMR study) in theseed could be hydrolyzed; and (c) organic phosphoestersare absent in the cell wall.

We tested for the transphosphorylating ability ofKBPAP with a few phosphate acceptors such as Tris andmannitol. No transphosphorylating ability was observedfor KBPAP, ruling out the possibility that KBPAP mightgenerate phosphoesters. KBPAP also demonstrates nega-tive cooperative behavior with both ATP and pNPP assubstrates (preliminary observations). Negative cooperat-ivity imparts on KBPAP an ability to respond linearlyto wide fluctuations in the concentration of substrates.KBPAP is approximately 1% of the seed protein and hasthe highest turnover number among all of the reported

Figure 5. Electron microscopy of the KBPAP activity staining in theperipheral cotyledon cells. One-cubic-millimeter blocks of the cot-yledon were used for activity staining. Activity was restricted to thecell walls only and was absent in the protein bodies. CW, Cell wall;PB, protein bodies. www.plantphysiol.orgon January 3, 2020 - Published by Downloaded from

Copyright © 1997 American Society of Plant Biologists. All rights reserved.


914 Cashikar e t al. Plant Physiol. Vol. 114, 1997

phosphatases (Vincent e t al., 1992). Quantitative a n d catalytic abundance of KBPAP combined with (a) an absence of hydrolyzable phosphoesters native t o the seed i n the cell wall, (b) localization exclusively in the peripheral cotyledon cells, (c) a requirement for high salt for extraction, which indicates strong binding to the cell wall (therefore, KBPAP may not be present in the exudates), a n d (d) an inability to transphosphorylate argue strongly that the role of KBPAP is well suited for generating Pi for the developing seed. Given the ability t o hydrolyze nu- cleotide diphosphates and triphosphates, KBPAP may help reabsorb the residual phosphates, a role similar t o that proposed for soybean cell wall phosphatase (Ferte, e t al., 1993).

A n antioxidant role has previously been suggested for KBPAP, based on the findings that the Fe(II1) of KBPAP could be reduced to Fe(I1) i n the presence of ascorbic acid, which could reduce oxygen to water a n d thereby decrease the damaging free radical concentration in the seed (Kla- bunde et al., 1995). However, we could not demonstrate a peroxidative role for KBPAP using o-phenyldiamine (preliminary observations).

Cell wall turnover accompanies storage tissue break- down during seed germination (Labavitch, 1981). An inter- esting possibility is the involvement of KBPAP in the degradation of the cell wall of the peripheral cotyledon cells itself a t the start of germination. This process may be ATP-dependent, a n d KBPAP m a y provide the essential function. A possible role for the cell wall phosphatases i n plant-microbe interactions has also been proposed (Boller, 1987). Apart from substrate-specific phosphatases, e.g. Tyr phosphatases, defining the role for nonspecific phosphatases h a s been a confounding issue, given their abundance and wide occurrence. Systematic investigations of the tem- poral expression and activity patterns of KBPAP may help in further defining its role i n vivo.

ACKNOWLEDCMENTS

We wish to acknowledge the help received from Dr. C.S. Sundaram and Mr. V.M. Shanmugam with our NMR experiments.

Received January 14, 1997; accepted March 24, 1997. Copyright Clearance Center: 0032-0889/97/ 114/0907/09.

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