extracellular acid phosphatases of lentinula edodes ... · assignment of an . in vivo . function to...

M U S H . J . T r o p i c s , 1989 9, 55-78

Extracellular acid phosphatases of Lentinula edodes: correlation of increased activity with fruit body development and enzyme localization, substrates, effectors and stability*

Gary F. Leatham†& Jane C. Hasselkus††

Forest Products Laboratory, Forest Service, U. S. Departrnent of Agriculture, Institute f o r Microbial and Biochemical Technology, O n e Gifford Pinchot Drive, Madison, Wisconsin 53705-2398, U . S . A .

Received 10 November 1988; accepted 15 February 1989

Summary

The dominant phosphatases in Lentinula edodes are acid phosphatases whose increased activity in cultures correlates with mushroom development. The majority of the activity is localized on the extracellular surfaces of primordia and fruit bodies. The purpose of this study was to determine the biochemical characteristics of the bulk of the phosphatase activity present in culture extracts. Rased on its response t o a range of substrates and inhibitors, the activity behaved as a single class o f Mn+++-dependent monophosphoesterase. Gel filtration chromatography showed the activity to be a single peak with an apparent molecular weight of 86,000. Anion exchange Chromatography showed one major and two minor peaks. The activity showed strong preference for phosphomonoesters a n d compounds containing aromatic groups. I t was capable of breaking terminal P -O , P-N, and P-S bonds, but showed little or no activity against either internal P - O or the external S - O bonds in sulfate. Its pH optimum varied between pH 4 and 5 depending on the buffer used. The activity was maximal at 50°C and markedly stable at up to 60°C. With p-nitrophenolmonophosphoester as substrate, the activity was insensitive to a large range of potential inhibitors. 'The few strong inhibitors found included

* This article was written and prepared by U.S. Government employees on official time. and i t i s therefore in the public domain [and n o t subject to copyright].

† To whom the correspondence should be addressed. †† Present address: 1910 Madison S t . , Madison, WI 53711, U.S.A.

56 Gary F. Leatham & Jane C. Hasselkus

p-nitrophenol (a reaction end-product), molybdate, o-vanadate, Fe3+, N-bromosuccinimide, and polyacrylic acid. In contrast, inorganic phosphate was a very weak inhibitor. Mn3+ and citric acid stimulated o r stabilized the activity. Activity taken either from cultures capable of fruiting or from actively growing fruit bodies was affected in a complex fashion by pH. Decreasing the pH from 5.0 to 4.0 caused a rapid. stepwise, and pH-irreversible loss in activity of 24 to 60% which was caused by a decreased Vmax. Proteins or phosphorylated substrates protected against the activity loss. The potential role of acid phosphatase activity in development is discussed.

Introduction

Lentinula (syn. Lentinus) edodes (Berk.) Pegler produces shiitake, the most commercially-important mushroom grown on wood (Ito 1978; San Antonio 1981; Leatham 1982). Previous studies indicated that its major phosphatases are highly active extracellular acid phosphatase(s) (orthophosphoric-monoester phosphohydrolyase; EC 3.1.3.2) whose total activity is highest after the cultures have ceased vegetative growth (Leatham 1985a). The bulk of the activity is located in the cell-wall region, hut is not secreted into the medium. It can be easily released by blending the cultures in water (“shockable fraction”; Leatham 1985a). Although these initial studies suggest i t may be of key importance to fruiting, the specific tissue localization, characteristics, and f unc t i o n ( s ) of the enzyme(s) are not known.

Assignment of an in vivo function to an enzyme requires knowledge of its specificity as well as the substrates to which i t has access. Function assignment is complicated for phosphatases by the large range of substrates possible. These include phosphorylated sugars, polyols, organic acids. nucleosides, amino acids, peptides. and proteins (Kuo & Blumenthal 1961; Ikawa et al. 1964). For instance, human prostatic acid phosphatase shows a preference for aromatic phosphates (Hollander 1971), shows decreasing activity with increasing phosphoester chain length (Staehelin 1964). and as typical for acid phosphatases, can cleave P - O , P-N, P-S, but not P-C bonds. Due to the frequent differences in the range of substrates tested, it is often difficult to compare a newly characterized enzyme with those reported in the literatue.

Sensitivity to inhibitors and other characteristics are helpful in enzyme classification. For example, prostatic acid phosphatase is a metalloenzyme whose essential manganic (Mn 3+) cofactors are proposed to he ligated in the enzyme by a tyrosyl and a sulfhydryl residue (Bobrzecka et al. 1968). It is inhibited by compounds which ligate Mn3+, such as L-(+)- (but not D-(-)- o r meso-) tartaric acid and F1- (Hollander 1971). In addition, it is inhibited by reagents which interfere with the binding of Mn3+ such as sulfhydryl reagents (Barron 1951; Bobrzecka et al. 1968), Ca2+ (Steen-Lievens & Tagnon 1962), heavy metals (e.g., Fe3+, C u 2 +, Hg2+ and Pb2+; Barron 1951; Hollander 1971), and the tyrosyl reagent iodine monochloride (especially at p H 8.1; Bobrzecka et al. 1968). Its sensitivity to N-bromosuccinimide also suggests that it has an essential tryptophyl residue (Bobrzecka et al. 1969). The enzyme shows reversible inhibition with diisopropylflurophosphate (Greenberg & Nachmansohn 1965). I t is very sensitive to surface inactivation caused by shaking. And it is markedly unstable to storage (Hollander 1971). Citrate, a mild chelator, stabilizes the enzyme during storage (Doe et a l . 1965). Based on sensitivity to inhibitors arid substrate preference, the less well characterized acid phosphatase of Neurospora crassa appears to be similar to the prostatic enzyme (Kuo & Blumenthal 1961).

Acid phosphatases of Lentinula edodes 57

Although differing in some respects, the Mn3+-dependent “purple” acid phosphatase o f sweet potatoes as well as the trivalent metal-dependent acid phosphatase ”uteroferrin” from mammals, appear to have active sites markedly similar to that of the prostatic enzyme (Sugiura et al. 1982; Antanaitis & Asien 1983). They show inhibition with many of the above compounds and also with the phosphate analogs molybdate and/or o-vanadate (Uehara et al. 1974a & 1974b: Davis et al. 1981; Antanaitis & Aisen 1985). The sweet potato enzyme differs in that it shows sensitivity to the chelators 1 ’ , 10’-(o)-phenanthroline and a , a ’-dipyridyl (Uehara et al. 1974).

In vivo acid phosphatases most commonly function in either o f two roles: 1) the liberation and recycling of phosphorous from phosphorylated compounds in the medium (e.g., phytase degradation of phytate in corn; Sloane-Stanley 1961), or 2) the regulation of metabolism o r growth (e.g., prostatic acid phosphatase dephosphorylation of phosphoproteins in animals; Lin et al. 1986). Developmentally-important roles are indicated for the extracellular cell wall-associated acid phosphatases of certain fungi (Pugh & Cawson 1977; Bojovic-cvetic & Vujicic 1982).

Rather than investigate a single enzyme, the purpose o f this initial study was t o partially characterize the range of extracellular acid phosphatase activities present in crude mycelial extracts of L . edodes. This overview information is needed to help determine their in v i vo function(s). Reported are the temporal correlation of increased activity with culture development, as well as the tissue. cellular localization and enzymatic characteristics of the activity in extracts made from different tissues, cellular localization and enzymatic characteristics of the activity in extracts made from different tissues. The enzymatic characteristics studied included temperature stability, p H optimum, substrate preferences, inhibitors, stimulators, and pH stability. Summarized are some of our initial attempts to purify the major enzyme(s) present.

Materials and methods

Chemicals All chemicals were reagent grade from the following sources: manganese(III)-oxidc and acetonate were from Morton Thiokol (Alfa Products) Inc. (Danvers. MA)†; psilocybin pure standard was from Supleco; all other enzyme substrates and inhibitors were from Sigma Chemical Co. (St. Louis, MO); manganese(IV)-dioxide (MnO2); and all other chemicals were from Aldrich Chemical Co. (Milwaukee, WI). Unless otherwise stated. all ionic compounds were either the sodium salts or chlorides.

Fungus, media, and culture conditions Lentinula edodes heterodikaryon strain ATCC #48085 was maintained on Sabouraud dextrose agar (Difco Laboratories, D e t r o i t , Mi.). As described previously (Leatham 1983), 25 ml of a chemically-defined liquid medium and mycelial homogenate inoculum were used for experimental cultures in cotton-stoppered 300 ml Erlenmeyer flasks. The cultures were grown in lighted (Leatham 1985a) chambers a t 2 2 ° C and 80% RH under an air atmosphere. Fall- harvested mushrooms of the same strain were obtained from a local commercial grower. They were grown on small-diameter Northern Ked Oak (Q u e r c u s r u b r a L.) logs as described

† The use of trade or firm names in this publication is for reader. i n f o r m a t i o n and does not imply endorsement by U.S. Department of Agriculture of any product or service.


previously (Leatham 1982). Fungal culture growth (culture mass formation) in liquid media was assessed by weight

after collecting the mycelia on preweighed filter disks, rinsing with distilled water, and drying overnight at 60°C. Data on the temporal observations for fungal development in a culture population were adapted from those reported previously (Leatham 1985a).

Preparation of enzyme extracts Extracellular enzyme extracts from the cell wall region of 35- to 50-day-old whole ‘cultures o r specific fruit body tissues were prepared by tissue suspension in distilled water using a Waring‘ blender followed by vacuum filtration through glass fiber filters as described previously (“shockable fraction”: Leatham 1985a). Protein content of the extracts was measured by Coomassie blue G-250 dye binding (Spector 1978). Unless otherwise stated, when used, enzyme extract dialysis was against water in a 25,000 MWT cut-off collodion bag (Schleicher & Schuell, Danvers, Ma.). When used, freeze-thawing was carried out by repetitive incubations in a dry ice/ethanol bath and 22°C water bath.

Column chromatography Anion exchange chromatography was on D E A E BioGel (BioRad Laboratories, Richmond, C A ) column - 14 ml of resin on a 1 X 16 cm column bed, eluting with a 100 ml linear gradient of KCl in 50 mM pH 5.25 sodium acetate (HCl) buffer. Gel filtration chromatography was on a SuperoseTM 12 HR-10/30 column in a FPLC system (Pharmacia Fine Chemicals AB, Uppsala, Sweden) eluting with 50 mM sodium citrate (HCl) p H 5.25 buffer. For either column, the source of activity was enzyme extract dialyzed through a 75,000 MWT cut-off collodion bag.

Enzyme assays Esterases capable of hydrolyzing p-nitrophenolphosphomonoester (PNP-P) o r other PNPanalogs were generally assayed with 3,33 mM substrate in triplicate or higher numbers of replicate samples at 22°C in 50 mM sodium acetate (HCl) pH 4.0 buffer as described previously (Leatham 1985b). Assays were carried out with or without the inclusion of enzyme inhibitors or stimulators. Unless stated otherwise, PNP-P was used as substrate in 15 min long assays. Assays were typically initiated by the addition of enzyme extract diluted four-fold with distilled water. Assays for the different p H incubations were initiated by adding substrate. Activity was also assayed by the Fiske-Subbarow method (Sigma Chemical Corp. 1974) using a dialyzed enzyme extract and 1 mM substrate in 50 mM sodium acetate (HCl) pH 4.0 buffer. Inorganic phosphate release was measured as increased absorbance at 660 nm. Assays for the determination of pH optima included 1 M NaCl (final concentration) to negate any significant differences in the concentration of buffer counter ions. Thermal stability was determined in the absente of substrate by incubation the enzyme extracts at different temperatures prior to assay at 22°C.

When used prior to the assay of extracts, single-step incubations at the different pH values specified were typically for 1 h at 4°C with or without the inclusion of potential protective compounds in 10 mM sodium citrate/TRIS (NaOH/HCl) buffer. This was followed by assay in 200 mM sodium acetate (HCl) p H 4.0 buffer. Two-step incubations were first in 10 mM sodium citrate/TRIS buffer at different pH values specified and 4°C for 1 h , followed


by a second incubation in 100 mM pH 4.0 sodium acetate buffer at 22°C for 4 h with assay as for the single-step incubation.

Results

Correlation with development Increased acid phosphatase activity in the extracellular extract (shockable fraction) of L . edodes was found to correlate with either culture development of physiological changes (Fig. 1). During the first six days of growth, the specific activity o f acid phosphatase in the cultures rose from essentially 0 to about 0.5 units/mg protein. It then slowly decreased during vegetative growth unitl net turnover of culture protein began. Next, correlating with onset of liquid droplet exudation from mushroom primordia and the exhaustion of the extracellular nitrogen source, it increased t o approximately 1.2 units/mg protein. During this period, only a limited number (less than 5%) of the cultures fruited. Then, as the intracellular nitrogen reserves were exhausted and net culture growth ceased, the largest numbers of cultures fruited (about 50%). Correlating with this period of increased development, the activity abruptly increased to its maximal level near 3.5 units/mg protein. Reflecting the lack of fruiting synchrony in the population, the largest variation among replicate cultures (largest relative error bars) occurred during this latter period of strong fruiting.

Fig. 1 Correlation of increased specific activity of the extracellular acid phosphatase(s) with the fruiting of L. edodes cultures. The developmentally-important events were those established previously (Leatham 1985a). (ML875453)


Tissue and cellular localization Explaining the increased activity during fruiting, the bulk of the acid phosphatase activity in flask-grown cultures was localized within expanding o r developing tissues. When separated from 40-day-old cultures, the primordia and mushrooms contained 80% o r more of the total extractable activity, with the residual being in the vegetative mycelium. In mushrooms grown out-of-doors on logs, the activity was found to be spread throughout all of the tissues tested at levels between 0.32 and 1.71 units/g tissue (Table 1). The highest activities were obtained from the tissues still expanding at the time of extraction (rim, gills & upper rind).

Table 1 Relative concentration of acid phosphatase activity extracted from different tissues taken from log-grown fruit bodies of Lentinula edodes.

Tissue Units/g fresh tissue Units/mg protein

lowerstipe upperstipe

gills innerpileustissue rim upperrind

0.64 1.32 0.32 0.47 1.50 0.54 0.96 0.95 1.71 0.81 1.42 1.32

In log-grown fruit bodies, the acid phosphatase activity was predominantly extracellular and non-particulate (soluble). Most of the activity (> 80% for log-grown and > 60% for flask-grown tissue) was extracted from the fruit bodies by simply cutting them into 3 to 4 mm3

pieces and shaking them in distilled water at 100 rpm on a gyratory shaker for 1 to 3 h. Once extracted, the activity was found to be soluble and did not bind to cell wall or membrane fragments. Essentially all of the activity remained in the supernant after centrifugation at 183,000 xG for 1 h.

Characteristics of the activity Thermal optimum and stability. Acid phosphatase activity in the crude aqueous extracts showed maximum activity when assayed at 50°C (Fig. 2) and little o r no activity above 80°C. Excellent thermal stability was found at up to 60°C (stability in the absence of substrate; Fig. 3). Activity was also stable at lower temperatures. Enzyme extracts could be stored a t 4°C for several days with < 5% activity loss. The activity was unaffected by freezing. Extracts freeze-thawed five times and subsequently assayed at 22°C underwent < 3% activity loss.


Fig. 2 Temperature optimum for acid phosphatase activity. (ML875452)

Fig. 3 Thermal stability for acid phosphatase activity. (ML875451)

pH optimum. The p H optimum for the activity varied between 4.0 and 5.0 depending on the buffer used. Teh p H optima with 50 mM citric, L-(+)-tartaric, succinic, or acetic acid (Na OH/HCl) buffers were 4.0. 4.2, 4.5, or 5.0, respectively. L-( +)-Tartaric acid, a Mn3+-binding ligand, gave a much narrower activity profile than did citric acid (Fig. 4) and the other two buffers tested (data not shown).

Fig. 4 pH optimum for the acid phosphatase activity in sodium citric or L-( +)-tartaric acid buffers. (ML875450)


Substrates. The use of different PNP-esters as substrates showed that the activity was due to a monophosphoesterase(s) which was markedly specific for terminal phosphates (Table 2). Less than 3% of the relative hydrolysis rate for PNP-P was seen with the five phosphodiesters tested. Similarly low hydrolysis rates occurred with naturally-occurring phosphodiesterase or other internal phosphate containing substrates - e.g., uridine-diphospho-sugars showed 7 to 9% and c-AMP only 0.6% the hydrolysis rate for PNP-P (Table 3). Failure to hydrolyze PNP-sulfate demonstrated that the major enzyme(s) present possessed no significant sulfatase activity (Table 2).

Table 2 Determination of the classes of esterase activities extracted

from flask-grown cultures of L. edodes.

PNP-ester Percent relative activity

PNP-phosphomonoester (PNP-P) a,b

PNP-phenylphosphate b

bis-BNP-phosphate b

thymidine-3'-phosphate-PNP b

thymidine-5'-phosphate-PNP b

PNP-phosphorylcholine c

PNP-sulfate d

100.0 ± 2.3

2.9 ± 0.6

0.7 ± 0.2

0.7 ± 0.5

0.3 ± 0.2

0.0 ± 0.0

0.0 ± 0.0

a Substrate for several classes of esterases including acid phosphatases,

alkaline phosphatases, and ATPases. b Phosphodiesterase substrate. c Phospholipase C substrate. d Sulfatase substrate.


Table 3 Comparison of the relative activity of acid phosphatase extracted from flask-grown cultures with different classes of phosphorylated substrates.

% Relative activity a % Relative activity a

Aromatic substrates (mean ± std dev) Non-aromatic substrates (mean ± std dev)

Substrates with P-O bonds

indoxyl-3-phosphateb,c 249.1 14.5 pyrophosphate (PP) b,c,d 84.8 4.0 psilocybin 1.4 1.6

O-phospho-L-threonineb 14.3 2.0 PNP-phosphomonoester (PNP-P)c 100.0 2.0 O-histidinol-L-phosphate 8.5 05 ß-naphthyl acid phosphateb 91.8 2.0 O-phospho-L-serine 4.5 0.4 phenylphosphate 74.7 1.2 a-naphthyl acidphosphate 64.9 2.9 D-glucosamine-6-phosphate b,c 37.9 0.8 phenolphthalein monophosphate 51.2 0.4 L-ribose-5-phosphateb 29.4 3.2 o-carboxyphenylphosphate 26.7 0.3 D-mannitol-1-phosphate b,f 26.0 0.8

L-fucose-l-phosphateb 20.3 2.5 adenosine-5'-monophosphate (5'-AMP) b,c 110.9 0.3 D-glucose-l-phosphateb 17.0 1.0 adenosine-5'diphosphate (ADP) b,d 75.5 3.4 D-mannosed-6-phosphateb 16.7 0.8 adenosine-5'-triphosphate (ATP) b,d 50.6 3.8 D-galactose-1-phosphate 14.1 1.3 adenosine-3'-monophosphate (3'-AMP)b 34.6 0.5 D-galactose-6-phosphate 10.7 0.6

D-mannose-1-phosphateb 5.9 0.8 uridine-5'-monophosphate (UMP)b 87.8 1.3 D-glucose-6-phosphate 4.9 1.0 uridine-5'-diphosphate (UDP) b,d 48.6 1.8 uridine-5'-triphosphate (UTP) b,d 30.8 0.3 ß-glycerophosphateb 72.3 5.3

L-a-glycerophosphate 43 6 1.9 O-phospho-L-tyrosine b,c 44.0 1.3 i-inositol-hexaphosphate (phytate) 7.5 1.3 riboflavine-5-phosphate (FMN) 32.7 0.6 pyridoxal-5-phosphate 30.1 8.2 phosphoenolpyruvate 46.7 4.1 ß-nicotinamide mononucleotide (NMN) 23.2 11.8 carbamylphosphate 18.6 2.9 pyridoxamine-5-phosphate 17.1 1.0

6-phospho-D-gluconateb 32.5 3.1 uridine-5'-diphospho-P-D-glucose (UDPG)d 8.7 0.4 3-phospho-D(-)-glycerate

phosphorylcholine b,c,g

21.8 19.0

1.1 0.4

0-phosphorylethanolamine 5.4 0.2 uridine-5'-diphospho-N-acetyl-ß-D-glucosamined 7.0 2.1 adenosine- c-3',5'-mnnophosphate (c:AMP) 0.6 0.2

Substrates with P-N and P-S bonds

adenylyl-5'-imidodiphosphate (AMP-PNP) c,e 45.6 1.1 imidodiphosphate (PNP) e 31.1 4.0 adenosine-5'-O-(2-thiodiphosphate)(ADP-ß-S)e 26.1 4.4 N-phosphocreatine 9.4 2.2

N5[phospho-L-arginine 2.1 0.3

a Inorganic phosphate release from 1 mM substrate relative to PNP-P assayed by the Fiske-Subbarow method.

b Less than 5% inhibition when using 1 mM of these substrates to compete with 3.33 mM PNP-P cleavage and under 50% inhibition when both were at 0.37 mM.

c Substrates used to test for the extent of activity loss after short-term incubation at pH 4.0 (single incubation; Fig. 6).

d The relative activity is over estimated by a factor of two or three because either two moles of inorganic phosphate are released per mole of substrate (pyrophosphate) cleaved or up to two or three phosphates can potentially be released.

e The activity is not overestimated because neither thiophosphate or aminophosphate should significantly interfere in the assay.

f The barium salt was used. g The calcium salt was used.


Among the substrates tested with P - O bonds, aromatic compounds - including aromatic phosphoesters and nucleotides were among the best substrates tested (Table 3). With the exception of ß-glycerophosphate, generally much less activity was noted for non-aromatic phospho-amino acid, -carbohydrate, -organic acid, and other compounds. For instance, the three aliphatic L-amino acid (threonine, histidine & serine) O-phosphornonoesters tested were hydrolyzed at three- to twenty-fold lower rates than was O-phospho-L-tyrosine. The highest activities were obtained with the following substrates (highest listed first): indoxyl-3-phosphate. 5’ -AMP, PNP-P, and ß-naphthyl acid phosphate. We did not detect any significant release of inorganic phosphate from phosphoproteins including phosvitin and casein (Fiske-Subbarow method and extended incubations used; data not shown).

Among the aromatic substrates, the activity showed specificity for both the location and number of phosphates in nucleotide phosphoester side chains as well as for the individual compound tested (Table 3). For instance. i n spite of the fact that they contain an identical phosphomonoester linkage. phosphate substitution of in the 5’ position AMP, gave a three-fold increase i n activity over substitution in the 3’ position. In fact, 3’-AMP was no better of a substrate than was ribose-5-phosphate. Activity decreased with increased phosphoester side chain length with both adenosine and uridine nucleotides. Finally, unlike indoxyl-3-phosphate and the many other aromatic phosphoesters tested, psilocybin (a naturally-occurring indole phosphate from mushrooms), was a very poor substrate. Control experiments showed that the psilocybin preparation used was indeed phosphorylated, that it contained no detectable level of inhibitors, and that it failed to compete with PNP-P cleavage (see below).

Competition with PNP-P cleavage was used t o determine the highest affinity substrate (the one with the lowest Km). Failure of a11 of the 21 substrates tested t o significantly compete ( < 5%) inhihition) indicated that PNP-P had a higher relative affinity (Table 3, footnote b). Using even more sensitive tests, where both PNP-P and the same potentially competitive substrates were tested at 0.37 mM, the only substrates that showed any significant ability to compete were nucleotides and aromatic phosphoesters including the following (10 to 40% inhibition; the most competitive listed first): 3’-AMP, A D P , ß-naphthylphosphate, 5’-AMP, and phosphotyrosine. Using an Eadie-Hofstee plot, the Km for PNP-P hydrolysis at 22°C was determined to be 0.37 mM with a relative Vmax of 3.7 µM (Fig. 5; at substrate concentrations greater than 3 mM

Fig. 5 Eadie-Hofstee plot of acid phosphatase activity with PNP-P as substrate at concentrations between 0.13 and 11 .24 mM. (ML875449)


The activity in crude extracts was capable of cleaving P-N bonds (in AMP-PNP, PNP and N-phosphoarginine) and P-S bonds (in ADP-ß-S) (Table 3). However, the cleavage rates were up to two-fold lower than that for P - O bonds in analogs of similar structure.

We could not conclusively demonstrate any ability of the extracts to cleave P-C bonds. Even with extended overnight incubations, the pyrophosphate analog methylenediphosphate (PCP) yielded little inorganic phosphate ( < 5%, the of the total possible; data not shown). The little release that did occur was all within the first 20 minutes of incubation. Thus, the release was apparently caused by the hydrolysis of a trace contaminant (e.g., pyrophosphate). Inhibitors and stimulators. The acid phosphatase activity was found to be remarkably insensitive to the majority of potential inhibitors or stirnulators that were tested at a concentration of generally 1 mM. A large range of enzyme inhibitors, enzyme active-site reagents, metal chelators, inorganic cations, inorganic anions, oxidants, reductants, detergents, organic acids, aromatic compounds, or nucleic acid base analogs failed either to significantly inhibit or t o stimulate the activity (Table 4).

Table 4 Failure of various classes of compounds at a concentration of 1 mM a to significantly inhibit or stimulateb acid phosphatase activity extracted from flask-grown cultures.

Enzyme inhibitors/active-site reagents N-acetylimidazole c

Chelators diethyldiothiocarbamate

Oxidants o-iodosobenzoic acid

2-bromoacetamido-4-nitrophenol c potassium ferricyanide butane-2.3-dione c EDTA c

p-chloromercuribenzoate (PCMB) c EGTA c Reductants 2’,4’dibromoacetophenone (p-bromophenacylbromide) c 8-hydroxyquinoline ascorbic acid dicyclohexylcarbodiimide (DCCD) c

diethylpyrocarbonate (DEPC) c,d " -5-sulfate c

iminodiacetic acid c dithionite dithiothreitol (DTT)

diisopropylfluorophosphate c nitrilotriacetic acid glutathione (reduced)

1,2-epoxy-3-p-nitrophenoxypropane (EPNP) c

N-ethylmaleimide (NEM) c Inorganic cations e Detergents

dimethylaminonaphthlene-5-sulfonyl chloride (Dansyl chloride) c l',10'-phenanthroline c p-hydroquinone

iodine c,e Li + b CHAPS c,g

iodine plus potassium iodide c,e Na + b polysorbate (Tween) 80 iodoacetamide c K + b

iodoacetic acid Rb + b Organic acids f

idomethane c Cs + b t-aconitic acid 4-iodophenylsulfonyl chloride (Pipsyl chloride) c m4

+ b,c citric acid 2-methoxy-5-nitrobenzyl bromide c Mg 2+ a 2,2'dimethylsuccinic acid phenylglyoxal c Ca 2+ a,c fumaric acid phenylisothiocyanate c Mn 2+ a D-glucuronic acid phenylmethyisulfonylfluoride (PMSF) c Zn 2+ c a-keto-glutaric acid tetranitromethane c Cu 2+ c

malic acid toluenesulfonyl chloride (Tosyl chloride) c Co 2+ pyruvic acid toluenesulfonyl fluoride (Tosyl fluoride) c Ni 2+ succinic acid

Hg 2+ c D-(-)-tartaricacid ATPase/phosphatase inhibitors Pb 2+ c meso-tartaric acid carbonylcyanide-m-chlorophenylhydrazine (CCCP) Ba 2+ benzoic acid diethylstilbestrol c Sr 2+ 3,4-dihydroxycinnamic acid oligomycin Sn 4+ D,L-dopa ouabain (strophanthin G) Ce 4+ h ferulic acid tetramisole indole-3-aceticacid

continued


trimethyltin c

valinomycin

Phosphodiesterase inhibitors 3'-AMP caffeine compounds theophylline

Nucleic base analogs adenine adenosine uracil uridine

Inorganic anions chlorided

bromide d

iodide d

arsenite c

azide c

cyanate aluminate borate iodate nitrate a

selenate (poly)15phosphate

phthalicacid salicylicacid phenylpyruvicacid sulfanilicacid a

Misc aromatic/indole

phenol quercetin a-amanitin bufoteine monooxalate 5-hydroxytryptamine 5-hydroxytryptophane psilocin

a If sparingly soluble, a saturated solution of the compound was used. Some common anions and cations

were tested at much higher concentrations (see footnotes f and g below). b Less than 20% inhibition or 10% stimulation. c These and the compounds listed in Table 7 failed to protect the acid phosphatase from activity loss during a

single-step incubation at pH 4.0. d Also tested was pre-incubation at pH 6.0 for 1 h before the assay at pH 4.0. e Also tested was pre-incubation at pH 8.0 in 5 mM TRIS buffer for 1 h before the assay at pH 4.0. f No significant effect when tested at concentrations of up to 1000 mM. g No significant effect when tested at concentrations of up to 100 mM. h Also an oxidant. i CHAPS = 3-((3-cholamidopropyl)-dimethyl-ammonio)-1-propanesulfonate. j Tested at 0.1%. k Added as the free acids. 1 An indole-containing mushroom peptide tested at 100 µg/ml.

However, a few compounds inhibited strongly in 10 to 100 µM concentration range. These included the following: the phosphate analogs molybdate and o-vanadate; poly-acrylic acid; Fe3+ ; and the enzyme active-site inhibitor N-bromosuccinimide (Table 5). In contrast. the following inhibited more weakly giving strong inhibition only at a concentration of 1 mM or higher: the phosphate analogs tungstate and arsentate; the tight-binding Mn3+ ligands Fland L-(+)- (but not either D-(-)- or meso-) tartaric acid; the metals Fe2+ (may simply be air

oxidized to Fe3+ ), Al3+, Cd3+ , Ag3+ , and Sn2+ (hut not Sn4+ ); the tyrosine reagent/oxidant iodine monochloride (especially at pH 8.1); the oxidant periodate; and the end-product of PNP-P cleavage, p-nitrophenol as well as its analog 2, 3-dinitrophenol (Table 5). Phosphate and common phosphate-like anions including pyrophosphate and sulfate were very weak inhibitors - e.g., inorganic phosphate only gave 76% inhibition at a concentration of 100 mM (Table 6).

-- -- --

-- -- --

-- -- -- --

-- -- --

--

-- --

- -

-- --

-- -- -- -- -- -- -- -- -- -- -- -- -- --

-- -- -- -- -- -- -- -- -- -- -- -- -- --

-- --

-- -- -- -- -- -- -- -- -- --

-- -- -- --

-- --


Table 5 Effects of selected inhibitors on acid phosphatase activity extracted from flask-grown cultures of L. edodes.

Percent inhibition

Inhibitor concentration (µM)

Inhibitor 0.1 1 .0 10.0 100.0 1000.0

molybdate a 0 23 88 100 100 o-vanadate a 7 32 73 95 100 tungstate 0 2 13 88 arsenate 35

polyacrylic acid 34 67 74 L-(+)-tartaric acid 31 oxalic acid 29 4-hydroxycinnamic acid 25 alginic acid 21

Fe 3+ 0 3 14 99 98 Fe 2+ 72 Al 3+ 50 Cd 2+ 50 Ag 3+ 36 Sn 2+ 35

N-bromosuccinimide (NBS) 0 66 100

iodine monochloride (8.1) c 9 85 iodine monochloride (5.5) c 29 periodate 29

p-nitrophenol d 7 99 2.4-dinitrophenol d 0 99

Fl- 0 6 71

denotes not tested. a Similarly strong inhibitions were obtained with phosphomolybdate and phosphotungstate. b Concentration based on the monomeric molecular weight of 72 per carboxyl group.

The actual average polymeric molecular weight was in excess of 250,000. Thus, 10 µM in carboxyl groups is 2.9 nM polyacrylic acid.

c Reacted for one hour at 22 °C in 5 mM sodium acetate (pH 5.5) or TRIS (pH 8.1) buffer

prior to enzyme assay.


Table 6 Inhibitory effect of phosphate and selected phosphate-like

anions on acid phosphatase activity extracted from flask-grown cultures

of L. edodes.

Percent inhibition

Concentration ( mM )

Anion 1 10 100

phosphate 7 33 76 pyrophosphate 6 44 73 sulfate 5 10 50

Only three types of coinpounds were able to stimulate the acid phosphatase activity when tested at 1 mM. These included (% stimulation) 1) sources of Mn3+ (but not Mn2+ ) including all those tested - MnO2 (manganese (IV)-dioxide; 58%), manganese (III)-oxide (16%), and manganese (III)-acetonate (25%); 2) the chelator citric acid (36%), and 3) the strong detergent methylbenzethonium chloride (37%) The MnO 2 , a relatively strong oxidant conveniently capable of supplying Mn3+ through nonspecific chemical reduction, was equally effective even when added at 10 µM

pH stability. Acid phosphatase activity in the extracts made from certain tissues was affected in a complex fashion by pH treatments given near the pH optimum for 1 h at 22°C (Fig. 6). The extracts themselves. which typically had an in vitro pH between 5.5 and 6.5, were stable. However, when the incubation pH was decreased from 5.0 to 4.0 (or lower) by extract dilution into buffer, 24 to 60% activity loss was found upon subsequent assay at pH 4.0 (single-step incubation; Fig. 6). The extent of loss was dependant on the individual extract. Essentially identical extents of loss were seen when seven other substrates were tested (Table 3; footnote c). Once the loss had occurred, changes in pH ( e.g, back to 5.0) did not restore the activity. Other than this loss. the activity was stable to incubation in the range of pH 2 to 4 and 5 to 8 (Fig. 6).


Fig. 6 Effects of pH treatments given prior to the assay of acid phosphatase activity at pH 4.0. Single-step incubations were at the pH value specified and 4°C for 1 hr. Two-step incubations were first at the pH value specified for 1 hr and 4°C followed by a second incubation at pH 4.0 and 22°C for 4 h. (ML875448)

Later we discovered why the effect of incubation at pH 4.0 could demonstrated in spite of the fact that we later assayed at pH 4.0. This was because the substrate (PNP-P) in the assay protected the phosphatase against activity loss. No loss occurred at the substrate concentration used in the assay (3.33 mM equals 10 times the substrate Km concentration for PNP-P).

The nature of this unexpected pH-dependent activity loss was investigated further. First we used the same single-step incubation procedure as above slowing the reaction rate by using a lower temperature of 4°C. Under these conditions, the activity loss was found to occur as a single stepwise event. Half of the loss occurred within 5 min (T1/2 = 4.5 min) after being exposed to pH 4.0 buffer (Fig. 7). The full extent of activity loss occurred within 1 h - i.e., no further loss occurred even with an additional 22 h of incubation at 4°C.

Fig. 7 Time course for the loss of acid phosphatase activity at 4°C caused by a single-step incubation at pH 4.0. (ML875447)

The kinetic basis for the activity loss in a typical enzyme extract was determined at 22°C using Eadie-Hofstee plots (Fig. 5). At substrate concentrations above the Km concentration (V/[S] 6 x the enzyme was protected and Km of 0.37 mM (as reported above) and relative Vmax of 3.7 µM were observed. However, at substrate concentrations below the Kin

4 xconcentration 10(V/[S] -3 ), the enzyme lost activity at the beginning of the assay. The overall observed Km decreased to 0.2.0 mM and the relative Vmax decreased to 2.9 µM. Thus, in spite of the tighter substrate binding constant (decreased Km), the step-wise activity loss occurred due to a decreased Vmax.

Next, we attempted to determine the biochemical basis for the activity loss by seeking a method to protect other than by adding excess substrate (PNP-P). A range of potential enzyme inhibitors were tested that had previously been determined to not inhibit the acid phosphatase. All failed to protect against the loss (Table 4, footnote c). Likewise, a range of potential

10 Gary F. Leatham & June C. Hasselkus

proteinase inhibitors all failed to protect against the loss (Table 7). However, Bovine serum albumin and all o f the five other proteins tested, successfully protected to various extents.

Table 7 Inhibitors that fail to significantly protect a from or potentiate the

loss of acid phosphatase activity during a single-step incubation at pH 4.0.

Compound Concentration tested

Proteinase inhibitors carbobenzoxy-(L-Ala-L-Phe-L-Leu-L-Ala) carbobenzoxy-(L-Phe-L-Leu-L-Ala-L-Ala) carbobenzoxy-(L-Ala-L-Phe-L-Leu-L-Ala) chloromethyl ketone-(L- Ala-L-Ala-L-Phe) chloromethyl ketone-(L-Leu) diazoacetyl-D,L-norleucine methyl ester phenylmethylsulfonylfluoride (PMSF) N-tosyl-L-Lys-chhomethylketone (TLCK) N-tosyl-L-Phe-chloromethylketone (TPCK)

phenylpyuvate antipain antitrypsin bestatin chymostatin leupeptin papain inhibitor (Gly-Gly-L-Tyr-L-Arg) pepstatin A trypsin inhibitors (four types) *

Phosphatase inhibitors arsenate periodate Fe3+ molybdate o-vanadate

a Less than 15% protection. b The types of individual trypsin inhibitors tested included the following:

Bovine pancreas, ovoinhibitor, ovomucoid, and soybean.


Both the activity loss (Table 8) and the tendency to show the bimodal kinetic curves (as seen in Fig. 5; data not shown) were either reduced o r no longer occurred. The protection was concentration dependent. Bovine serum albumin concentrations of 1 mg/ml or higher gave complete protection (Table 8).

Table 8 Protein protection against against acid phosphatase activity loss resulting from a single-step incubation at pH 4.0.

Protein Concentration Activityafter %Protection (mg/ml) incubation (%)

none

Bovine serum albumin a

lysozyme a-amylase ß-lactoglobulin glucose oxidase phosvitin

a Cohn Fraction V. b This particular extract underwent 24% activity loss during a single-step incubation

at pH 4.0.

We made three other discoveries while searching for compounds t o protect against the pH-dependent activity loss. First. preincubation o f the enzyme extract for 4 h at 22°C in pH 4.6 or higher buffers completely negated the ability of the extract t o undergo the step-wise loss during a second incubation in pH 4.0 buffer (see two-step incubation; Fig. 6). Secondly, when present in assays at levels only moderately inhibitory for the acid phosphatase. certain


inhibitors potentiated the loss. These included the following (concentration, inhibition factor): polyacrylic acid (100 µM, 2.9-fold), N-bromosuccinimide (100 µM, 2,4-fold), and L-(+)tartaric acid (1 mM, 1.9-fold). In contrast. at the concentrations tested, the loss was not potentiated by five other acid phosphatase inhibitors (Table 7). Thirdly, in many (but not all) trials, the addition of 10 µM MnO2 restored the activity lost due to incubation at pH 4.0.

Evidence was gained suggesting that the pH-dependent activity loss might be of significance to development. Phosphatase activity from certain tissues showed much less tendency to undergo the step-wise loss than that from others. Extracts prepared from either whole cultures with rapidly growing primordia or the vegetative mycelium from successfully fruiting cultures, showed the highest sensitivity to pH. They gave 24 to 60 (most typically 45 ± 5) % loss. In contrast, extracts prepared from older cultures that had failed to fruit or from fruit bodies that had completed expansion. showed markedly less sensitivity to pH. They typically gave only a 5 to 10% loss. Besides the extent of loss, other differences were noted with extracts made from different tissues. Even though they had similar protein contents, extracts made from actively growing primordia (which contained about 80% of the total culture activity) gave over 20-fold faster rates of activity loss during 4°C incubations at pH 4.0 than did extracts made form the underlying vegetative cells.

Initial attempts to purify Stability and other problems were encountered in our initial attempts to purify the individual acid phosphatase(s) present in the crude extracts. Activity in the extracts was stable to dilution and was retained after repeated dialysis through 75,000 MWT collodion bags. However. attempted precipitation with as little as 40% (w/v) ammonium sulfate irreversibly destroyed 85% of the activity. Neither 50 mM pH 5.25 citric acid buffer nor 10 µM MnO2 was able to protect against loss or restore activity. Precipitation with polyethylene glycol (8,000 mwt) was of little use. The unusually high affinity of the enzyme for the polyethylene glycol phase hindered glycol removal. Unless a citric acid buffer was used. chromatography on anion exchange columns in many cases resulted in nearly complete activity loss. However, in this case, the activity in the partially purified preparations could often be restored by adding 10 µM MnO2.

Column chromatography in the presence of citric acid buffer showed promise as a purification method as well as yielded useful information about the major phosphatases present. Gel filtration chromatography showed that all of the activity resided within a single peak with an apparent molecular weight of 86,000 (Fig. 8) Anion exchange chromatography revealed one major and two minor peaks (Fig. 9). After separation on anion exchange resin. all three of the partially-purified phosphatases still retained their sensitivity to acidic pH and MnO2. They showed similar extents of activity loss to that obtained with the crude extract prior to separation (dare not shown; tested fractions #9 and #25) and showed similar abilities to he stimulated by MnO2 (Fig. 9).

Fig. 8 Determination of the molecular weight of acid phosphatase activity by gel filtration chromatography ( ) The positions for different molecular weight markers are shown ( ) (ML875446)

Fraction number


Fig. 9 Separation of acid phosphatase activities by anion exchange chromatography. Assays were carried out both i n the absence ( -) and presence ( ) of 10 µM MnO2. (ML875445)

Discussion

In this investigation we demonstrated temporal correlation with development, and determined tissue localization, pH optimum, substrate preferences, inhibitors, arid stability of the bulk o f the acid phosphatase activity present in L . edodes. ‘The bulk o f the enzyme activity shows distinct similarities to human prostatic acid phosphatase and is apparently due t o one major enzyme. However, there are key differences between the two enzymes.

The similarities of the acid phosphatase activity of L. edodes with the prostatic enzyme include the following: 1) clear preference for aromatic substrates (suggesting that an aromatic binding site is present); 2) decreasing activity with increasing phosphomonoester chain length; 3) likely participation of Mn3+ as a co-factor; 4) inhibition by Mn3+ ligands including FI- and the L-(+)- isomer o f tartaric acid; 5) inhibition by phosphate analogs including molybdate and o-vanadate; 6) inhibition by Fe3+ ; 7) stabilization by citric acid; 8) inhihition by tyrosyl and tryptophanyl reagents including iodine monochloride and N-bromosuccinimide, respectively; and 9) instability.

The acid phosphatase activity of L. edodes differs from the prostatic enzyme in the following characteristics: 1 ) higher specificity for aromatic compounds (even more strongly suggesting that an aromatic binding site is present), 2) lack o f inhibition by sulfhydryl group reagents, 3) lack of inhibition by Ca2+ and certain heavy metals (e .g . , Pb2+ , Cu2+ & Hg2+ ), 4) lack of inhibition by diisopropylfluropliosphate, and 5) much greater stability.

The acid phosphatase activity o f L . edodes also shows distinct differences from the Mn3+ -dependent “purple” acid phosphatase o f sweet potato. I t i s not inhibited by the chelators 1’,10’-( o)-phenanthroline or ’ -dipyridyl.

The surprising lack of sensitivity of the acid phosphatase activity of L. edodes to the many diverse potentially inhibitory compounds tested indicates that the few strong inhibitors found were highly specific. Because, all of our inhibitor studies were done at ten times the Km concentration for substrate (PNP-P). we predict that the binding constants (Ki’s) for the strongest inhibitors (molybdate , o-vanadate, Fe 3+ , polyiterylic acid. and N-bromosuccinimide) are probably in the high nanomolar to low micromolar concentration range. However, purification of this enzyme is required before the magnitude of the inhibition constants or can be determined with accuracy. The lack of sensitivity to many inhibitors and the good thermal stability are noteworthy. They suggest that, when adequately stabilized, the purified enzyme may have practical commercial applications.

The inhibition of the acid phosphatase activity of L. edodes by Fe3+ is noteworthy. Native uteroferrin is a Fe3+ -dependent enzyme in which Mn3+ can function (Keough et al. 1980; Burman et al. 1986). Fe3+ is also the only other trivalent metal ion that can serve as a

--


cofactor in the Mn3+ - dependent purple acid phosphatase of sweet potatoes (Kawabe et al. 1984). In the case of the L. edodes enzyme, Fe3+ has strong affinity, but does not generate an active enzyme. It is possible that citric acid, a good Fe3+ chelator, stabilizes or stimulates prostatic-like phosphatases by protecting the Mn3+ -binding site form Fe3+ or other heavy metals.

As observed here, there are potential problems inherent in assaying for metalloenzymes such as acid phosphatases. The indiscriminate use of metal chelators as buffers in enzyme assays or growth media may cause assay artifacts or cause failure to detect certain enzymes. For Mn3+ -dependent enzymes, these buffers include the polyacrylic and L-(+)-tartaric acids reported here, and polyanions such as alginic and polyxenyl phosphate acids reported for other enzymes (Jeffree 1957; Kuo & Blumenthal 1961). The narrow pH profile found here with the L-(+)-tartaric acid buffer may simply he an artifact caused by the increased chelation of Mn3+

occurring with increased ionization of the buffer. The rapid, step-wise, acidic pH-induced loss in acid phosphatase activity described here

may be of significance in vivo. Collectively, our data indicates that the loss is due to regulation of the major enzyme(s) present rather than due to inactivation of one unstable enzyme in a mixture of multiple enzymes. Given the nature of the kinetic change, the regulatory effect is ideal for efficient degradation of substrate. At high substrate concentrations, the high relative Vmax gives the highest degradation rates possible. And as the substrate concentration is diminished, the change to it lower relative Km gives the ability to scavenge the residual substrate. This hypothesis should be tested once the in vivo substrate for the enzyme is identified.

The two most phausible mechanisms for the pH effect include the following: 1) an endogenous factor e.g., a change in enzyme conformation or dissociation of a multi-subunit complex or regulatory subunit. or 2) modification by an exogenous factor - e.g., an acid proteinase insensitive to the inhibitors tested or the release of a protective agent such as a substrate or protective protein. Whatever the mechanism, the obility of citric acid and MnO2

to restore activity as well as the potentiation of loss caused by Mn3+-binding ligands, suggest that the pH-dependent alteration leads to lower affinity for Mn3+ or perhaps the loss of one of multiple Mn3+ cofactors.

In support of an endogenous factor causing the pH effect are the following: 1) the retention of the transition mechanism after dialysis through 75,000 MWT membranes; 2) the retention of the transition mechanism after partial purification by anion exchange chromatography; and 3) the failure of the large range of potential enzyme inhibitors to protect against activity loss.

In support of an exogenous factor causion pH effect are the following: 1) the different rates of activity loss for extracts made from different tissues; 2) the loss of the pH effect when the extract is diluted into buffers with a pH above 4.6; 3) the fact that the enzyme remains sensitive to pH in spite of the pH of the crude natural extract being between 5.5 and 6.0; and 4) the protection by protein. The fact that phosphorylated substrates and protein specifically protect against the pH-dependent transition would be easily explained if a phosphoprotein was the in vivo substrate.

If caused by an exogenous factor, the data here do not yet rule out (and are consistent with) an acid proteinase-catalyzed mechanism. Extracellular acid proteinases in concentrations capable of extensive protein turnover are produced by L. edodes (Leatham 1985a). Having


affinity fo r protein, they may be difficult t o separate from the phosphatase. Consistent with the data here, the major acid proteinase of L. edodes is reported to have a p H optimum of 2.9, t o be unstable near p H 5 or higher (Terashita et al. 1981a & 1984), and t o be insensitive t o well-known acid proteinase inhibitors (Terashita et al. 1981b).

Regulatory proteolytic modifications known to occur for other phosphatases could account for the p H effects in L. edodes. For instance, in a Zn2+ -dependent alkaline phosphatase from Escherichia coli, proteolytic cleavage lowers the binding affinity for one of the three Z n2+ cofactors present in each enzyme subunit (Roberts & Chlebowski 1985). This reduces its specific activity. If the Zn2+ cofactor is later removed from a proteolytic-modified enzyme, unlike with the native enzyme, activity can no longer be restored by incubating with Z n 2+ . Purification of the acid phosphatase of L . edodes will help in the determination of the mechanism causing the p H effect.

Tyrosine phosphorylation/dephosphorylation of proteins is being recognized as having a major developmental role in the control of cell proliferation (Bishop 1983; Heldin & Westermark 1984). Some of the well-characterized acid phosphatases which show preference fo r aromatic substrates have recently been determined t o be phosphoprotein phosphatases in vivo. These include an enzyme from Saccharomyces cerevisiae (Pavlovic, et al. 1985) and the prostatic acid phosphatase from mammalian cancer cell-lines (Lin et al. 1986). Like our enzyme, the one from S. cerevisiae demonstrates no significant ability t o release inorganic phosphate from either phosvitin or casein. However, it shows high affinity for specifically 32 P-labeled peptides. T h e mammalian prostatic acid phosphatase shows high affinity for specific phosphoproteins and preferentially hydrolyzes phosphoproteins (Lin & Clinton 1986). In addition, the prostatic activity tissue cultures is inversely correlated with the tyrosyl-kinase activity responsible for protein phosphorylation (Lin et al. 1986).

The close temporal correlation, localization, and characteristics of the acid phosphatase activity of L. edodes are consistent with it having an important role in development. The activity is maximal at the same time and is secreted t o the same aerial extracellular primordial and fruit body surfaces as the developmentally-important laccase of L. edodes (Leatham & Stahmann 1981). Both enzymes have pH optima near 4.0 and both most likely function in the acidic liquid film (droplets) produced by these rapidly growing structures. Further research is warranted to determine the specific in vivo function(s) of the acid phosphatase(s) of L. edodes.

Acknowledgements

We thank Mark Higgs of Arena, Wisconsin, U.S.A. for the fresh log-grown mushrooms of L. edodes and Kris R . Norris, formerly of this laboratory, for technical assistance in carrying out some of the preliminary experiments.

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