Characterization of Acid Phosphatase (AphC) from the Miso Koji Mold, Aspergillus oryzae KBN630: AphC is Mainly Responsible for Both Acid Phosphatase Activity and 5′-IMP Dephosphorylation Activity in Soybean-Koji Culture

Abstract

Single disruption of seven acid phosphatase genes (aphB-H) demonstrated that the aphC gene is mainly responsible for acid phosphatase production in the soybean-koji culture of the miso koji mold, A. oryzae KBN630. None of the single disruptions of aphB-H affected growth in the culture. Both the acid phosphatase activity and 5′-IMP dephosphorylation activity of the aphC disruptant decreased by 95% compared to the wild-type strain. Utilizing the promoter of the A. oryzae taaG2 gene, AphC was successfully overproduced in A. oryzae and secreted into liquid culture medium. The purified AphC had a molecular mass of 69.0 kDa, a pH optimum of 4.5, and a temperature optimum of 50°C. AphC exhibited high dephosphorylation activity towards the umami flavor enhancers, 5′-IMP and 5'-GMP, supporting the idea that AphC is involved in degradation of disodium 5′-ribonucleotides supplemented in miso products.

Introduction

Miso, fermented soybean paste, is a Japanese traditional seasoning which is indispensable for Japanese cuisine, as is soy sauce (shoyu). The taste, aroma and color are important parameters of miso quality. In order to improve the taste, miso products are often supplemented with disodium 5′-ribonucleotides such as disodium 5′-inosinate (5′-IMP) and disodium 5′-guanylate (5′-GMP). These umami flavor enhancers work synergistically with glutamic acid naturally contained in miso.

The filamentous fungus Aspergillus oryzae is used for miso brewing. During preparation of its koji culture (i.e., solid starter culture), A. oryzae secretes a large variety of enzymes such as amylases and proteases when grown on steamed rice (for rice-miso), soybean (for soybean-miso), or barley (for barley-miso). These enzymes are essential for efficient maceration and degradation of the ingredients. Among the enzymes secreted by A. oryzae, acid phosphatases appear to catalyze the hydrolysis of disodium 5′-ribonucleotides, yielding insipid ribonucleosides and phosphoric acid. Therefore, in order to prevent dephosphorylation of disodium 5′ribonucleotides, acid phosphatases must be inactivated by heating miso at 85°C for 15 min (Oike et al., 1984). This high-temperature heating process not only requires large equipment and a large amount of energy, but also reduces miso quality, causing browning and a burnt smell. Thus, an A. oryzae strain that produces acid phosphatases at a very low level is highly desirable as a miso koji mold for the production of seasoned miso.

A. oryzae has been reported to produce several acid phosphatases (Fujishima et al., 1964, Wang et al., 1980, Oike et al., 1984, Shimizu., 1993, Fujita et al., 2003a, 2003b). Eight aph genes encoding acid phosphatase with similarity to the well-characterized A. niger phyA gene were identified in the A. oryzae genome database by BLAST searches (Yoshino-Yasuda et al., 2012, Marui et al., 2012). In our previous report, we directly showed that acid phosphatase A encoded by the aphA gene (GenBank accession number AB042805 / AP007157) in A. oryzae possesses some level of 5′-IMP and 5′-GMP dephosphorylation (Yoshino-Yasuda et al., 2012). Disruption of the aphA gene in A. oryzae KBN630 resulted in only a 20% reduction in acid phosphatase activity in soybean-koji culture, a starter culture for soybean-miso brewing unique to the Chukyo area of Japan (Yoshino-Yasuda et al., 2012).

From these facts, we postulated that other Aphs are responsible for the remaining acid phosphatase activity. In order to clarify the major Aph with 5′-IMP dephosphorylation activity in A. oryzae, seven aph gene disruptants were constructed and their enzyme production in soybean-koji culture was examined. In addition, the aphC gene, demonstrated to be mainly responsible for both acid phosphatase activity and 5′-IMP dephosphorylation activity, was overexpressed under control of the A. oryzae taaG2 gene promoter (Tsukagoshi et al., 1989). Then AphC protein was purified and its enzymatic properties were characterized.

Materials and Methods

Fungal strains, culture media and transformation    A. oryzae KBN630, obtained from Bio'c (Toyohashi, Japan), was used for DNA isolation. Two strains were used for transformation. One was the pyrG, ku70 double deletion strain, A. oryzae KBN630-17K3 (Yoshino-Yasuda et al., 2011), derived from A. oryzae KBN630. The second was the alp, pyrG double deletion strain, A. oryzae PDE1, derived from the industrial shoyu koji mold A. oryzae KBN616. In A. oryzae PDE1, the alkaline protease gene (alp gene) (Murakami et al., 1991) was disrupted to produce recombinant proteins efficiently. The methods used to construct A. oryzae PDE1 will be published elsewhere. A. oryzae ΔaphA was used as an aphA gene disrupted strain (Yoshino-Yasuda et al., 2012). Rice starch (RS) medium containing 3% rice starch, 1% polypeptone, 1% NaNO₃, 0.2% KCl, 0.1% KH₂PO₄, and 0.05% MgSO₄7H₂O was used for liquid cultivation of A. oryzae. Soybean-koji culture was carried out at 30°C for 43 h on steamed soybean pellets as follows. Soybeans were soaked in water for 2 h, held overnight at 4°C, and then steamed for 30 min followed by autoclaving at 121°C for 5 min. Autoclaved soybeans were mashed, pelletized and coated with roasted barley flour before use. Protoplast transformation of A. oryzae was carried out as described previously (Kitamoto et al., 1995).

DNA techniques and PCR methods    Standard DNA techniques were used (Sambrook and Russell, 2001). Genomic DNA of A. oryzae was prepared using a previously described method (Kitamoto et al., 1993). PCR amplification was carried out with a GeneA-mp9700 thermal cycler (Applied Biosystems, Foster City, CA, USA). TaKaRa Ex Taq DNA polymerase (Takara Bio, Otsu, Shiga, Japan) and PfuUltra II Fusion HS DNA polymerase (Stratagene, La Jolla, CA, USA) were used in PCRs. Oligonucleotide primers used in this study are shown in Table 1. Essential cloning steps were confirmed by sequencing on a model 4000LS DNA sequencer (LICOR, Lincoln, NE, USA) and GenomeLab GeXP (Beckman Coulter, Brea, CA, USA).

Table 1. Oligonucleotide primers used in this study

Name	Sequence (5′ to 3')	Direction
Primers for gene disruption
The aphB gene
phyB1	CGGTACCCGGGGATCCCAACCGTACTCCTAGGAGTGG	Forward
phyB2	TTCCAGCAGGCCTTGAAAGCATGGCGCACTATCGCGG	Reverse
phyB3	ATTGATCAGGCCTTTCGACCAGCTAACCAAGACTAGT	Forward
phyB4	ATGCCTGCAGGTCGACATGATCAACGAATTCCTTCAACG	Reverse
The aphC gene
phyC1	CGGTACCCGGGGATCCTATGCCGGGATACTAACACAAT	Forward
phyC2	TTCCAGCAGGCCTTGCTATTAGTGGGTAATGCATAGTG	Reverse
phyC3	ATTGATCAGGCCTTTGAATGAGTGGGAGTATGTGCTC	Forward
phyC4	ATGCCTGCAGGTCGACAAGTCCAGATCTCGGAAATCAG	Reverse
The aphD gene
phyD1	CGGTACCCGGGGATCCATCCTTAACAGTAGACTACTTGG	Forward
phyD2	TTCCAGCAGGCCTTGCAGACTAACACTTCTAGACTAAG	Reverse
phyD3	ATTGATCAGGCCTTTAACTTCCGAACCCGCTATGCTAG	Forward
phyD4	ATGCCTGCAGGTCGACAACTTGACTCTCGTTCTTCGGTG	Reverse
The aphE gene
phyE1	CGGTACCCGGGGATCCCTGTGATCACTTTGAATAACACC	Forward
phyE2	TTCCAGCAGGCCTTGCAACATAACTTACTTGAAGACCAG	Reverse
phyE3	ATTGATCAGGCCTTTGGTCGTGATATTTTGCTTGCCAC	Forward
phyE4	ATGCCTGCAGGTCGACAGGTCCAGATTTCGGAGATTAG	Reverse
The aphF gene
phyF1	CGGTACCCGGGGATCCAGAATACAACATGCTGACTATGG	Forward
phyF2	TTCCAGCAGGCCTTGATGCGATACTAAGCAAAGCAATC	Reverse
phyF3	ATTGATCAGGCCTTTGCTCAAAGGGTCACTTGATCGCC	Forward
phyF4	ATGCCTGCAGGTCGACATACTAGATGGAACCGGAAAGG	Reverse
The aphG gene
phyG1	CGGTACCCGGGGATCCTTTAGACAATTTCAGTCCCGCTC	Forward
phyG2	TTCCAGCAGGCCTTGAAATGCAGCAGGTACGTTCTC	Reverse
phyG3	ATTGATCAGGCCTTTGGATCAAATACGGCAGGCTG	Forward
phyG4	ATGCCTGCAGGTCGACATAGAGGATCGGATCCAAAGAG	Reverse
The aphH gene
phyH1	CGGTACCCGGGGATCCACACAGAACAAACAAAAGATCTG	Forward
phyH2	TTCCAGCAGGCCTTGGTGAAGTTCTCTAGACTCTAGG	Reverse
phyH3	ATTGATCAGGCCTTTCCTTAATGACTGGAGCTATGGGC	Forward
phyH4	ATGCCTGCAGGTCGACCTTCATGAACGTGTCAAGCTCAC	Reverse
Primers for aphC expression vector
fupyrGN	CGGTACCCGGGGATCCAAGCCGCTGCTGGAATTGACA	Forward
pyrGC3	TCAGAAGAAAAGGATGATCAATACC	Reverse
pyrGtaa	ATCCTTTTCTTCTGAATTCATGGTGTTTTGATCATTTT	Forward
taaPrev	CATAAATGCCTTCTGTGGGGTTTATTGTT	Reverse
taaaphC	CAGAAGGCATTTATGCAGCAATTATTGCAATCAACGG	Forward
aphCSal	ATGCCTGCAGGTCGACGGGTTGATAGAGCTTGTTCTGGTGATC	Reverse

Disruption of aph genes    An aphB gene disruption vector, pDisAphB, was constructed as follows. A 1.0-kb 5′- and a 1.0-kb 3′-flanking region of the aphB gene were amplified from A. oryzae genomic DNA using the primer pairs phyB1/phyB2 and phyB3/phyB4. A 1.8-kb fragment of the pyrG gene was amplified from A. oryzae genomic DNA with the primer pair pyrGN2/pyrGC2 (Yoshino-Yasuda et al., 2011). The three PCR-products and the BamHI/SalI-digested pUC18 were joined in a four-piece In-Fusion reaction using an In-Fusion Advantage PCR Cloning Kit (Takara Bio). The resultant plasmid was designated as pDisAphB, in which the pyrG gene was located between the 5′- and 3′-flanking regions of the aphB gene. The gene disruption fragment for aphB was amplified with the primer pair phyB1/phyB4 and pDisAphB as the template, and introduced into A. oryzae KBN630-17K. Genomic DNAs from transformants were used as the template for PCR analysis using the primer pair phyB1/phyB4 for verification of the aphB disruption. Using the primers listed in Table 1, the other six aph gene disruption vectors, pDisAphC to pDisAphH, were constructed by the same procedure as for pDisAphB. The gene disruption fragments for the six aph genes were also amplified by the same procedure as for the aphB gene, and introduced into A. oryzae KBN630-17K. Genomic DNAs from the transformants were analyzed by PCR using specific primer pairs for aph genes (phyC1/phyC4 for aphC gene, phyD1/phyD4 for aphD gene, phyE1/phyE4 for aphE gene, phyF1/phyF4 for aphF gene, phyG1/phyG4 for aphG gene and phyH1/phyH4 for aphH gene, respectively) for verification of gene disruption.

Expression of aphC under control of the A. oryzae taaG2 promoter    To express the aphC gene under control of the A. oryzae taaG2 gene promoter, an expression plasmid, pTAAphC, was constructed as follows. A 1.8-kb fragment of the pyrG gene was amplified from A. oryzae genomic DNA using the primer pair fupyrGN/pyrGC3. A 0.6-kb fragment of the A. oryzae taaG2 gene promoter was amplified from A. oryzae genomic DNA using the primer pair pyrGtaa/taaPrev. A 2.0-kb fragment of the aphC gene was amplifed from A. oryzae genomic DNA using the primer pair taaaphC/aphCSal. The three PCR-products and the BamHI/SalI-digested pUC18 were joined in a four-piece In-Fusion reaction using an In-Fusion Advantage PCR Cloning Kit. The resultant plasmid was designated as pTAAphC, in which the A. oryzae taaG2 gene promoter was positioned precisely next to the aphC gene coding region. Then, plasmid pTAAphC was introduced into A. oryzae PDE1.

To confirm the introns of the aphC gene, RT-PCR was performed using a High Fidelity RNA PCR Kit (Takara Bio) with total RNA from the high-AphC-producing strain described below. Double stranded cDNA synthesis was performed using the primer pair Adaptor Primer FB and taaaphC. The amplified fragment was cloned into HincII-digested pUC118 for sequencing.

Purification, amino acid sequencing, and deglycosylation of AphC from the overproducing A. oryzae transformant    A. oryzae transformant APC15 was grown in RS medium for 5 days. After removing the mycelia by fltration, 50 mL of the culture fltrate was dialyzed against 10 mM Tris-HCl buffer (pH 7.0). The crude enzyme solution was loaded on an HR16/20 Fast Flow Q-Sepharose anion-exchange column (GE Healthcare, Buckinghamshire, UK) equilibrated in the same buffer and eluted with a linear gradient of 0 M to 0.25 M NaCl. The AphC-containing fractions were pooled, dialyzed against the same buffer, and rechromatographed on an HR16/20 Fast Flow Q-Sepharose anion-exchange column under the same conditions. The AphC-containing fractions were dialyzed against 10 mM Tris-HCl buffer (pH 7.0) and brought to 40% saturated (NH₄)₂SO₄, followed by loading on a HiLoad 26/10 Phenyl Sepharose HP (GE Healthcare) equilibrated in the same buffer containing 40% saturated (NH₄)₂SO₄. The column was washed with the equilibration buffer, and bound protein was eluted with a linear gradient of 40% to 0% saturated (NH₄)₂SO₄. The AphC-containing fractions were dialyzed against 10 mM Tris-HCl buffer (pH 7.0) and were determined to be pure by SDS-PAGE.

In order to analyze the N-terminal amino acid sequence, purified enzyme was applied to PVDF using a ProSorb device (Applied Biosystems) and sequenced on an Applied Biosystems Procise 491 sequencer (Applied Biosystems).

Purified enzyme was deglycosylated with endoglycosidase H (Glyko, Novato, CA, USA) according to the procedure provided by the manufacturer. The protein was denatured before addition of endoglycosidase H.

Enzyme assay    Acid phosphatase activity was measured using a slight modification of the procedure described by Oike et al. (Oike et al., 1984). An aliquot of the enzyme solution was incubated with 1 mM p-nitrophenylphosphate (PNPP) in 100 mM acetate buffer (pH 4.0) at 40°C for 10 min. The reaction was terminated by adding 10% trichloroacetic acid solution. After addition of 2 M Na₂CO₃ solution, the amount of liberated p-nitrophenol (PNP) was measured by absorbance at 405 nm. One enzyme unit was defined as the amount of enzyme that liberated 1 µmol of PNP per min under the assay conditions. In the case of soybean-koji culture, acid phosphatase activity was measured at 37°C for 20 min instead of 40°C for 10 min. The pH optimum of the enzyme was measured by incubating the enzyme for 10 min at 40°C in 100 mM sodium acetate buffers at various pHs (3.0 to 7.0). The temperature optimum was measured by incubation for 10 min at various temperatures (25°C to 65°C) in 100 mM sodium acetate buffer (pH 4.0). The thermal and pH stabilities were measured after incubation of the enzyme at various temperatures (25°C to 65°C) for 30 min and at various pHs (3.0 to 7.0) for 1 h at 30°C, respectively. Substrate specificity was investigated as follows. The enzyme solution was incubated with 2 mM of each substrate for 30 min at 40°C in 100 mM acetate buffer (pH 4.0). The amount of released inorganic phosphate was measured using a Phosphor C Test Kit (Wako Pure Chemical, Osaka, Japan).

Alpha-amylase activity was measured according the officially approved method (Nishiya, 1993). Neutral protease activity was measured according a slight modification of the officially approved method (Nishiya, 1993). McIlvaine buffer (pH 6.0) was used as a reaction buffer instead of McIlvaine buffer (pH 3.0). IMP dephosphorylation activity was measured as follows: 1 mL of 20 mM IMP was mixed with 0.2 mL of 200 mM acetate buffer (pH 4.0) and incubated with 0.1 mL of dialyzed culture extract at 37°C for 20 min. The reaction was terminated by adding 0.1 mL of 1 N NaOH, followed by neutralization with 0.1 mL of 1 N HCl. The liberated phosphate was measured using Pi ColorLock ALS (Innova Biosciences Ltd., Cambridge, UK). One unit of enzyme activity was defined as the amount of enzyme that liberated 1 µmol of phosphate per min under the assay conditions.

Computational sequence analysis    A homology search was performed using the BLAST search of DOGAN, the Database of the Genomes Analyzed at NITE (http://www.bio.nite.go.jp/dogan/top). The search for signal peptides was performed using SignalP 4.0 (http://www.cbs.dtu.dk/services/SignalP/). The search for glycosylation sites was performed using two online glycosylation site prediction servers, NetOGlyc 3.1 (http://www.cbs.dtu.dk/services/NetOGlyc/) for prediction of O-linked glycosylation sites and Net-NGlyc 1.0 (http://www.cbs.dtu.dk/services/NetNGlyc/) for prediction of N-linked glycosylation sites.

Results and Discussion

Construction and characterization of acid phosphatase gene disruptants    In our previous report, we identifed eight aph genes encoding acid phosphatase (Aph) with similarity to the well-characterized A. niger phyA by searching the A. oryzae genome database using BLAST (Table 2). Characterization results for the ahpA disruptant suggested that another Aph (or Aphs) is responsible for the majority of acid phosphatase activity. In order to identify the major Aph (or Aphs) possessing IMP dephosphorylation activity, we constructed seven aph disruptants in addition to the aphA disruptant. As an example, a construction scheme for the aphB gene disruption is shown in Fig. 1A. Genomic DNAs from eight randomly selected transformants were analyzed by PCR using the primer pair phyB1/phyB4. Only a 3.8-kb band was detected for fve of the transformants, indicating successful aphB gene disruption (data not shown). Disruption of the other genes was performed using the same procedure as for aphB. The efficiencies of the eight aph gene disruptions were: 62.5% for aphB, 100% for aphC, 87.5% for aphD, 100% for aphE, 62.5% for aphF, 87.5% for aphG and 100% for aphH. As shown in Fig. 1B, gene disruption of the targeted aph gene did not affect the other aph genes.

Table 2. Acid phosphatase genes in A. oryzae studied during this work

Gene	Gene ID	Genbank accession number
aphA	AO090023000692	AP007157 / AB042805
aphB	AO090120000167	AP007166
aphC	AO090010000202	AP007175 / AB775132
aphD	AO090011000174	AP007171
aphE	AO090023000448	AP007157
aphF	AO090023000481	AP007157
aphG	AO090124000063	AP007165
aphH	AO090005000912	AP007151

Fig. 1.

Construction scheme for aphB disruption (A) and confrmation of aphA-H single disruptants (B).

A. Scheme for aphB gene disruption is shown. PCR-amplifed DNA fragments from the aphB gene disruption vector, pDisAphB, were trans formed into A. oryzae KBN630-17K3. Gray and black boxes indicate the 5′-fanking region and a part of the coding region of the aphB gene, respectively, and closed and open arrows indicate the aphB gene and pyrG gene regions, respectively. The direction of the arrow indicates the orientation of the aphB and pyrG genes. Small arrows indicate the positions of the oligonucleotide primers used.

B. Agarose gel electrophoresis of amplifed DNA fragments in the aphA-H gene regions of the eight aph disruption strains using primer pairs phyB1/phyB4 to phyH1/phyH4 listed in Table 1, and phyA1/phyA6 from our previous report (Yoshino-Yasuda et al., 2012).

In order to investigate the effects of the single aphA-H gene disruption on acid phosphatase production, the enzyme productivity of the disruptants was compared to the wild-type strain, A. oryzae KBN630. The single disruptants of the aphA-H gene grew normally in soybean-koji culture, comparable to A. oryzae KBN630, indicating that aphA-H gene disruption did not affect growth when steamed soybeans were used as the source of nutrients. As shown in Table 3, aphA, aphC and aphE gene disruptants had lower acid phosphatase activity and 5′-IMP dephosphorylation activity than A. oryzae KBN630, whereas the enzyme activities of the other disruptants did not decrease. The acid phosphatase activity and 5′-IMP dephosphorylation activity of the aphC gene disruptant were severely decreased, to 3% and 6%, respectively, compared to A. oryzae KBN630. These results strongly indicate that AphC is mainly responsible for both acid phosphatase activity and 5′-IMP dephosphorylation activity in soybean-koji culture.

Table 3. Effects of aphA-H gene disruption on enzyme production in soybean-koji culture

Strain	Enzyme activity (units/g koji culture) (% of control)
	Acid phosphatase	5′-IMP dephosphorylation	α-Amylase	Neutral protease
ΔaphA	394 ± 39.4 (80)	547 ± 57.9 (89)	959 ± 50.0 (130)	20,800 ± 461 (106)
ΔaphB	482 ± 52.3 (98)	692 ± 18.7 (112)	894 ± 7.8 (121)	20,200 ± 385 (102)
ΔaphC	13 ± 0. 6 (3)	36 ± 10.3 (6)	899 ± 15.6 (122)	20,800 ± 307 (106)
ΔaphD	478 ± 27.2 (97)	632 ± 16.5 (103)	853 ± 4.2 (116)	20,400 ± 461 (104)
ΔaphE	378 ± 14.1 (76)	481 ± 1.0 (78)	869 ± 2.8 (118)	18,500 ± 461 (94)
ΔaphF	528 ± 1.4 (107)	667 ± 39.2 (108)	881 ± 7.1 (120)	22,000 ± 537 (112)
ΔaphG	496 ± 64.2 (100)	661 ± 75.7 (107)	924 ± 40.3 (125)	21,400 ± 615 (109)
ΔaphH	452 ± 20.7 (91)	590 ± 30.0 (96)	822 ± 24.0 (112)	21,000 ± 93(107)
KBN630	494 ± 28.3 (100)	615 ± 21.2 (100)	736 ± 42.4 (100)	19,700 ± 707 (100)

Activity of two independent experiments is presented as average ± standard deviation.

On the basis of transcriptional expression analysis of the aph genes in solid-state rice and soybean culture, the aph genes in A. oryzae were classifed into type R and type S due to their higher expression in solid-state Rice and Soybean culture, respectively (Marui et al. 2012). Disruption of the aphC gene, which was categorized as type S, resulted in a marked decrease in both acid phosphatase activity and 5′-IMP dephosphorylation activity in soybean-koji culture. However, disruption of the aphB and aphG genes, which were categorized as type S, reduced neither of the enzyme activities. In contrast, disruption of the aphA gene categorized as type R reduced both of the enzyme activities. These results indicate that transcriptional expression of the aph genes are regulated in a complicated manner by the PHO regulatory pathway, pH regulation, and other unknown mechanisms. Moreover, amylase activities in the aphA-H gene disruptants increased by 12 – 30% when compared to the wild-type strain, and the neutral protease activities increased by 2 – 9% except in the aphE disruptant. Although the reason is still unclear, these results suggest that disruption of the aphA-H genes have a positive effect on the production of other enzymes such as amylase. Therefore, further studies on the aphA-H gene disruptants will be valuable for elucidating the aph genes regulatory mechanisms in koji culture and be helpful for increasing the amylase and protease productivities of A. oryzae.

Overexpression and purification of AphC    In order to characterize the enzymatic properties of the aphC gene product, the aphC gene from A. oryzae KNB630 (GenBank accession number AB775132) was efficiently expressed under control of the A. oryzae taaG2 gene promoter, which is one of the strongest promoters in A. oryzae. During the construction of the aphC gene expression plasmid, pTAAphC, nucleotide sequence analysis revealed that the aphC gene from A. oryzae KBN630 consists of 1,678 bp, including two introns of 47 bp, and contains an open reading frame encoding 527 amino acids. The introns were subsequently confrmed by sequencing the cDNA fragment of the aphC gene. The aphC gene from A. oryzae KBN630 differs from that of A. oryzae RIB40 (GenBank accession number AP007157) by 25 bp in the nucleotide sequence, which led to two amino acid substitutions: Asn-279 to Tyr, and Ala-344 to Ser. After pTAAphC was introduced into A. oryzae PDE1, transformants grown in RS medium were assayed for extracellular acid phosphatase activity in order to select a high-AphC-producing strain for further study. The enzyme levels ranged from 203 to 490 U/mL (0.14 to 0.34 mg/L) for the seven transformants examined, whereas the control strain, A. oryzae PDE1 carrying the pyrG gene, showed no acid phosphatase activity.

AphC was purifed to homogeneity from the culture supernatant of the highest AphC producing strain, APC15, by two anion-exchange chromatography purifications followed by hydrophobic interaction chromatography, as described in the Materials and Methods. AphC was purifed 2.1-fold, with recovery of 49.4% of the initial activity. The specific activity of the purifed AphC was 1,459 U/mg of protein, which was approximately 13 times higher than that of AphA. The purifed AphC migrated as a band of approximately 69.0 kDa on SDS-PAGE (Fig. 2, lane 2).

Fig. 2.

SDS-polyacrylamide gel electrophoresis of AphC purifed from A. oryzae transformant APC15.

AphC was purifed as described in the Materials and Methods. The gel was stained with Coomassie Brilliant Blue. Lane 1, molecular-mass markers [E. coli β-galactosidase (116.0 kDa), rabbit muscle phosphor-ylase b (97.4 kDa), bovine serum albumin (69.0 kDa), glutamate dehydrogenase (55.0 kDa), porcine muscle lactate dehydrogenase (36.5 kDa), bovine liver carbonate anhydrase (29.0 kDa), soybean trypsin inhibitor (20.1 kDa), and egg white lysozyme (14.4 kDa)]; lane 2, purifed AphC; lane 3, endogylcosidase H treated AphC.

The N-terminal amino acid sequence of the purifed AphC protein was chemically determined as Ala-Pro-X-X-X-X-X-Ala-Ala-Ala (X denotes an unidentifed residue). This sequence corresponds to the deduced amino acid sequence from Ala-22 to Ala-31, except for the fve residues from Thr-24 to Ser-28. Thr-24 was one of the O-linked glycosylation sites predicted by NetOGlyc 3.1, and might be the reason why the residues from Thr-24 to Ser-28 could not be identified. Analysis using SignalP 4.0 predicted that the first 21 N-terminal amino acids of AphC function as a signal sequence, which is consistent with the results of N-terminal amino acid determination. Therefore, the mature AphC consists of 506 amino acids with a calculated molecular mass of 55,105 Da. Since the calculated molecular mass was smaller than that of the purifed AphC by approximately 14.0 kDa on SDS-PAGE (Fig. 2, lane 2), we assumed that the differences between the theoretical and apparent molecular mass are caused by protein glycosylation. This idea is partly supported by the presence of six putative N-linked glycosylation sites (Asn-106, Asn-113, Asn-178, Asn-213, Asn-257, and Asn-275) and three O-linked glycosylation sites (Thr-24, Ser-32, and Thr-35) in AphC predicted by the programs NetNGlyc 1.0 and at NetOGlyc 3.1, respectively. Treatment of the purified AphC with endoglycosidase H, which cleaves N-linked mannose-rich oligosaccharides, decreased the apparent molecular mass to approximately 58.0 kDa (Fig. 2, lane 3). However, this value is still larger than the theoretical mass and may be due to O-linked glycosylation, as described above.

Enzymatic properties of AphC    The enzymatic features of AphC, including pH and temperature optima and pH and thermal stabilities, were investigated with the purifed protein. AphC has a pH optimum of 4.5 and a temperature optimum of 50°C. AphC is stable at a wide range of pHs, from 3.5 to 6.5. AphC was moderately inactivated at temperatures up to 30°C, but sharply inactivated at temperatures above 30°C. No enzyme activity remained at temperatures above 50°C. This finding means that AphC has a low intrinsic resistance to heat inactivation.

The substrate specificity of AphC was examined with a series of phosphate compounds under the same conditions as AphA (Table 4, Yoshino-Yasuda et al., 2012). AphC displayed rather broad substrate specificity but had very little activity toward phytate. AphC showed considerably higher activity toward 5′-GMP (19.2%) and 5′-IMP (37.3%). As shown in Tables 4 and 5, the enzymatic properties and substrate specificity of AphC clearly differ from those described for other A. oryzae acid phosphatases, AphA, ACP-I, ACP-II, ACP-III, and AphKI. Based on the specific activities of AphC (1,459 U/mg) and AphA (108 U/mg) toward PNP, AphC had about 30-fold and 70-fold higher specific activity toward 5′-GMP and 5′-IMP than AphA, respectively. This result suggests that AphC is largely involved in disodium 5′-ribonucleotide dephosphorylation activity in A. oryzae in soybean-koji culture. This unique substrate specificity appeared to correlate with the significant decrease in 5′-ribonucleotide dephosphorylation activity observed in the soybean-koji culture of the aphC disruptant (Table 3). From a practical point of view, it will be extremely interesting to investigate the expression levels of aphC in A. oryzae KBN630 and other industrial strains used in soybean-koji. We anticipate that the degradation of 5′-ribonucleotides, such as 5′-IMP and 5′-GMP that are supplemented in miso products, can be prevented by use of the aphC disruptant. Further studies are contemplated to verify the effectiveness of the aphC disruptant in laboratory scale miso brewing using soybean-koji culture. It is also necessary to consider the possible effect of lactic acid bacteria and yeasts involved in miso brewing. Moreover, we will construct the triple gene disruptant of aphA, aphC, and aphE in order to further decrease acid phosphatase activity.

Table 4. Comparison of the properties of AphC with those of other acid phosphatases from A. oryzae

	AphC	AphAa	ACP-Ib	ACP-IIb	ACP-IIIb	AphK1c
Molecular mass (kDa)	69.0	58.0 to 65.0	110	58	56	70
pH optimum	4.5	4.0	4.5	5.0	5.5	5.5
pH stability	3.5 to 6.5	3.0 to 7.0	4.3 to 5.5	4.8 to 6.5	5.0 to 5.5	2.0 to 7.0
Optimum temperature (°C)	50	40	60	40	45	60
Thermal stability (°C)	< 25	< 35	< 45	< 35	< 40	NDd

^aThe results of AphA were taken from our previous report (Yoshino-Yasuda et al. 2012).

^bThe results of ACP-I, ACP-II and ACP-III were taken from the report by Fujita et al. (Fujita et al. 2003a).

^cThe results of AphK1 were taken from the report by Shimizu (Shimizu 1993).

^dND, not determined.

Table 5. Comparison of AphC substrate specificity with those of other acid phosphatases from A. oryzae

Substrate	Relative activitya (%)
	AphC	AphAb	ACP-Ic	ACP-IIc	ACP-IIIc	AphK1d
Sodium p-nitrophenylphosphate	100	100	100	100	100	100
Sodium phytate	0.04	54.0	14	255	23	3.3
Sodium glycerophosphate	12.1	49.2	10	0	81	93.3
Sodium pyrophosphate	16.2	33.3	NDe	ND	ND	183.3
D-Glucose-6-phosphate	6.5	43.4	52	0	45	ND
5′-GMP	19.2	8.1	ND	ND	ND	ND
5′-IMP	37.3	6.9	ND	ND	ND	ND

^aHydrolysis rate of p-nitrophenylphosphate was taken as 100%.

^bThe results of AphA were taken from our previous report (Yoshino-Yasuda et al. 2012).

^cThe results of ACP-I, ACP-II and ACP-III were taken from the report by Fujita et al. (Fujita et al. 2003a).

^dThe results of AphK1 were taken from the report by Shimizu (Shimizu 1993).

^eND, not determined.

Acknowledgements    We would like to thank Dr. Junichiro Marui in Japan International Research Center for Agricultural Sciences (JIRCAS) for valuable suggestions, and Asuka Ando for technical assistance. This study was partially supported by the Research and Development Projects for Application in Promoting New Policy of Agriculture Forestry and Fisheries, Japan.

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