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- Materials and Methods
- Results and Discussion
Phytic acid (PA) frequently occurs in nature and constitutes the principal storage form of phosphorus (60–90%) and inositol in plants, legumes and oil seeds (Reddy et al. 1982; Konietzny and Greiner 2002). It is primarily present as a salt of monovalent and divalent cations (Fe2+, Mn2+, K+, Mg2+ and Ca2+) and accumulates in seeds during the ripening period. PA is therefore a common constituent of plant-derived foods.
PA exists as a highly negatively charged ion over a broad pH range and therefore has a tremendous affinity for food components with positive charge(s), such as minerals, trace elements and proteins (Greiner and Konietzny 2006). It is considered an anti-nutrient because it acts as a strong chelator of divalent and trivalent minerals such as Mn2+, Ca2+, Mg2+, Zn2+ and Fe2+/Fe3+ (Harland and Oberleas 1999). The formation of insoluble mineral–PA complexes at physiological pH is regarded as the main reason for the poor mineral bioavailability, essentially because these complexes are difficult to assimilate in the animal and human gastrointestinal tract (Greiner and Konietzny 2006). Minerals are involved in activation of intracellular and extracellular enzymes, in regulation of critical pH levels in body fluids necessary for the control of metabolic reactions and in osmotic balance between the cell and its environment. Deficiency of any one of the essential minerals can result in severe metabolic disorders and compromise the health of the organism (Lopez et al. 2002).
Legumes and cereals are good sources of proteins as well as dietary minerals such as Fe2+, Zn2+, Mg2+ and Ca2+. The content of Fe2+, Zn2+ and other minerals is generally high in legumes (Jambunathan and Singh 1981; Attia et al. 1994; Nestares et al. 1997). However, their applications as food are limited by PA, which acts as an anti-nutrient, resulting in reduction of nutritive value of these legumes. To increase the bioavailability of minerals from legume-based foods, enzymatic degradation of PA and its dephosphorylated isomer inositol pentaphosphate is desirable (Sandberg et al. 1999).
Phytases (myoinositol hexaphosphate phosphohydrolase EC 22.214.171.124 and 126.96.36.199) are PA degrading enzymes present in plants, animal tissues and also produced by a large number of bacteria and fungi (Vohra and Satyanarayana 2003; Cao et al. 2007). However, monogastric animals like poultry, pigs, fishes and humans cannot utilize dietary PA, due to the lack of adequate levels of phytases (Vats and Banerjee 2004; Rao et al. 2009). Therefore, the reduction of PA content in foods and feed using phytase is used to improve their nutritional value. Several reports have shown that supplementation of phytase in animal feed increases the availability of phosphorus for animal digestion through degradation of the PA (Pandey et al. 2001; Vohra et al. 2006). The great potential for use of phytase in processing and manufacturing of food for human consumption has been proposed (Haros et al. 2001; Hurrell et al. 2003), but it has not been applied to date. Research in this field has focused on obtaining the appropriate phytase producing microbial strain, improvement of nutritional value of plant-based foods as well as improving the techniques of food processing (Greiner and Konietzny 2006).
Multicellular fungal phytases have been studied extensively for their application in animal feed. However, there are very few reports on yeast phytases due to lack of cost-effective production and low expression levels. Hence, there is a need to identify yeasts with higher production and explore other applications of phytases. The present work was therefore directed toward screening and selection of phytase producing yeast cultures followed by exploitation of their role in mineral mobilization and dephytinization of chickpea flour (CPF).
Results and Discussion
- Top of page
- Materials and Methods
- Results and Discussion
About 600 yeast strains were inoculated on PSM plates to assess their growth on PA as the sole phosphorus source. All yeasts were able to grow on PSM, but only 40 yeasts showed zone of clearance around the colony after 4 to 7 days of incubation at 28C. It was observed that the five yeasts, Z. bisporus NCIM 3265 and NCIM 3296, W. saturnus NCIM 3298, Z. priorionus NCIM 3299 and S. octosporus NCIM 3297, which showed zone of clearance on PSM, have not been previously reported for phytase production. Along with them, 26 strains of Saccharomyces cerevisiae, 4 strains of Pachysolen tannophilus, 2 strains of Candida and a strain of Torulaspora delbrueckii, Zygosaccharomyces rouxi and Metschnikwoia pulcherrima also showed zones of clearance on PSM, which have already been reported to produce phytase.
Hydrolysis of Ca–phytate from PSM did not provide a clear evaluation of phytase activity, and an alternative testing method in LMM was therefore used according to Olstorpe et al. (2009). Growth at 28C was monitored by measuring optical density (OD) at 600 nm after 48 h. The relative capabilities of strains to utilize PA as sole phosphorus source were determined by comparing OD600 in medium with negative control (PA-depleted minimal medium). Forty strains showing zones of clearance on PSM, which were selected for further evaluation, grew in LMM but to varying extents. Of the tested yeast strains, six strains of S. cerevisiae, a strain of Z. biosporus NCIM 3265 and NCIM 3296, W. saturnus NCIM 3298, Z. priorionus NCIM 3299 and S. octosporus NCIM3297 grew very well. Four strains of P. tannophilus, two strains of Candida, a strain of T. delbrueckii, Z. rouxii and 19 strains of S. cerevisiae were unable to enter the log phase; and two strains were unable to grow at all, although they exhibited zones of clearance on PSM. Thus, a total of 11 yeast strains were found to grow in LMM. Five yeasts (viz. Z. bisporus NCIM 3265, Z. bisporus NCIM 3296, S. octosporus NCIM 3297, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299) showed extensive growth in LMM after plate screening method were selected for further study because of lack of significant study for cell associated phytase production in these strains.
Cell-associated and extracellular activities obtained for Z. bisporus NCIM 3265, Z. bisporus NCIM 3296, W. saturnus NCIM 3298, S. octosporus NCIM 3297 and Z. priorionus NCIM 3299, respectively, are shown in Table 1. Highest extracellular phytase activities were detected after 48 h, while cell-associated phytase activities were detected at 24 h of fermentation. Cell-associated activities were higher than extracellular; therefore, further studies were conducted with cell-associated phytase. Cell associated activities showed by Z. bisporus NCIM 3265, Z. bisporus NCIM 3296, S. octosporus NCIM 3297, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299 were higher or comparable with Candida krusei (201 U/mg cells), Pichia anomala (6 U/g cells) and Cryptococcus laurentii ABO 510 (4.55 U/g cells) as reported by Quan et al. (2001), Vohra and Satyanarayana (2001) and Van Staden et al. (2007), respectively; although it should be noted that these assays were not all performed under similar conditions. Z. bisporus NCIM 3265, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299 showed higher phytase activities than Z. bisporus NCIM 3296 and S. octosporus NCIM 3297. Though Z. bisporus NCIM 3265 and Z. bisporus NCIM 3296 belong to the same species, a large difference in phytase production was observed between them, indicating that phytase activity may be strain specific.
Table 1. Extracellular (secreted) and cell-associated phytase activity
|Yeast strain||Phytase activity||Cell biomass (g/L)|
|Extratracellular (U/mL)||Cell bound (U/g dry cell biomass)|
|Z. bisporus NCIM 3265||0.147||13||5.8|
|Z. bisporus NCIM 3296||0.076||6||5.8|
|S. octoporus NCIM 3297||0.134||5||6.0|
|W. saturnus NCIM 3298||0.065||7||5.6|
|Z. priorionus NCIM 3299||0.134||10||5.4|
The production of phytase was assessed in response to medium components other than Na–phytate (Table 2). The highest phytase production were observed in cane juice and cane molasses medium, which were 11, 15, 13, 25 and 18 times more than phytase production in LMM for Z. bisporus NCIM 3265, Z. bisporus NCIM 3296, S. octosporus NCIM 3297, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299, respectively. Yeast strains, Z. bisporus NCIM 3265 (145 U/g DCB), W. saturnus NCIM 3298 (198 U/g DCB) and Z. priorionus NCIM 3299 (190 U/g DCB) exhibited higher phytase production as compared with Z. bisporus NCIM 3296 (92 U/g DCB) and S. octosporus NCIM 3297(69 U/g DCB). Cane molasses and cane juice media (70–80 μM) contained more free inorganic phosphate compared with LMM and YG (20 μM) media and still showed higher phytase production. This implies that inorganic phosphorus in these media does not suppress the phytase production. Fredrikson et al. (2002) suggested that, either inorganic phosphorus in complex medium was not as effective in repressing phytase production, or promotion of phytase synthesis in complex medium was governed by a component absent in the minimal medium. According to Teixeira et al. (2011), the high production of sugarcane and its processing technology make its use as a carbon source for microbial enzyme production very promising. The cane juice and cane molasses also contains other elements like iron, potassium, sodium, etc., which support cell growth. The phytase activity of Z. bisporus NCIM 3265, Z. bisporus NCIM 3296, S. octosporus NCIM 3297, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299 was associated with the whole cells, whereas no phytase activity was found in the cell-free extract after permeabilization or treatment with ionic and detergent solutions. Correspondingly, treatment with the lysing enzyme or sonication increased the phytase activity in the cell-free extract, suggesting that the phytases were cell wall associated. Permeabilization with 0.2% Triton X 100 increased the cell-associated phytase activity in all strains; 0.2% SDS decreased the cell-associated activities by 40–50% in Z. bisporus NCIM 3265, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299 and by about 70–80% in Z. bisporus NCIM 3296 and S. octosporus NCIM 3297. Presence of EDTA had no effect on cell-associated phytase activity (data not shown). Cell-wall-associated phytases were also reported in Rhodotorula gracilis, C. krusei and P. anomala (Bindu et al. 1998; Quan et al. 2001; Vohra and Satyanarayana 2001) and were considered to have potential application in food processing because they remained stable, even at high temperature and in acidic environment (Kumar et al. 2010).
Table 2. Cell-associated phytase production by yeasts in different media after 24 h incubation at 28C (pH 5.5)
|Medium*||Enzyme production (U/g of dry biomass)|
|Z. bisporus||Z. bisporus||S. octoporus||W. saturnus||Z. priorionus|
|NCIM 3265||NCIM 3296||NCIM 3297||NCIM 3298||NCIM 3299|
|5% Cane molasses||142||86||69||153||185|
|50% Cane juice||145||92||48||198||190|
As depicted in Fig. 1a,b, the pH and temperature optima for all analyzed yeast phytases were pH 4.0 and 50C, respectively. The phytases were active over pH range 3.0 to 7.0 and temperature range of 30 to 80C. More than 60% of the residual activity was observed at acidic pH values between 4.0 and 5.0. Yeasts retained about 75–80% phytase activity after exposure to a temperature of 70C for 5 min, but no residual activity was observed after exposure to a temperature of 80C for 5 min (data not shown). Z. bisporus NCIM 3265 phytase retained more activity at alkaline pH and at higher temperature (60–70C). Phytase from Z. bisporus NCIM 3296 and S. octoporus NCIM 3297 was very sensitive to changes in pH and temperature and retained only 10% activity at pH 6.0 and 70C. All investigated yeast phytases had pH optima at 4 and were comparable with the phytases from Saccharomyces castellii (Segueilha et al. 1992), Arxula adeninivorans (Sano et al. 1999) and P. anomala (Vohra and Satyanarayana 2001), which were optimally active at pH 4.4, 4.0 and 4.0, respectively. In this study, all yeast phytases were optimally active at 50C. Similarly in C. krusei WZ-001, phytase had the temperature optima at 40C (Quan et al. 2002). This is in contrast to the phytases from S. castellii (Segueilha et al. 1992), A. adeninivorans (Sano et al. 1999) and P. anomala (Vohra and Satyanarayana 2001), which had temperature optima at 77, 75 and 60C, respectively. Most of the fungal phytases reported showed temperature optima of 50C. About 40–50% activity was retained at 37C, which is also the temperature generally used for food fermentations (In et al. 2009), suggesting its potential application for the same.
Figure 1. Effect of pH and Temperature on Phytase Activity
(a) pH optima measured as relative activity (%) of the enzyme after being incubated at 50C in buffer with various pH values from 2.0 to 9.0. (b) Temperature optima measured as relative activity (%) of enzyme at different temperatures in the range of 30–80C in 0.2 M acetate buffer (pH 5.0). Results are expressed as the mean of three replicated measurements (n = 3).
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In vitro system was used to show liberation of inorganic phosphate from CPF after phytase treatment. Given the same enzyme activity, the amount of inorganic phosphorus released by the five phytases were determined after incubation with CPF (Fig. 2). Supplementation of the CPF with yeast phytases resulted in similar initial rates of phosphate liberation. However, at later stages of incubation, significantly more inorganic phosphate was liberated by Z. bisporus NCIM 3265 and S. octoporus NCIM 3297 phytases in contrast to others. The increase in inorganic phosphate liberation was correlated with increase in incubation time of phytase. About 0.9–1 mM of inorganic phosphate were released after phytase treatment, which was significantly higher than control (0.4 mM). That is, about 2.5–3 times more inorganic phosphate was liberated in the presence of phytase. The release of inorganic phosphate as a result of phytase treatment was also observed in rice, wheat and corn containing poultry feed (Wu et al. 2004; Liang et al. 2009).
Figure 2. Release of Phosphate (mM) from CPF after the Treatment of Crude Cell-Free Extracts at 37C in 10 mL Acetate Buffer (pH 5.0) for 5 h
Results are expressed as the mean ± SD of three replicated measurements (n = 3).
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PA and mineral content (Zn2+, Fe2+ and Ca2+) in phytase untreated and treated CPF are shown in Figs. 3 and 4, respectively. The average concentration of Zn2+, Fe2+ and Ca2+ in untreated CPF were 2.1, 2.4 and 136 mg per 100 g of CPF, respectively, on a dry weight basis by ICP-AES analysis. The estimated concentration of minerals in untreated CPF was comparable with that of Srinivasan et al. (2007). Mineral accessibility in phytase untreated CPF were 40%, 32.1% and 42% for Fe2+, Zn2+ and Ca2+, respectively. In phytase-treated CPF, the mobilization of Fe2+, Zn2+ and Ca2+ increased to 68–79%, 52–61% and 66–82%, respectively, which was significantly higher than untreated CPF. Phytase from Z. bisporus NCIM 3265 and S. octosporus NCIM 3297 was more efficient in increasing the release of Zn2+, Mn2+ and Fe2+ than other yeast phytases. Z. bisporus NCIM 3265 and Z. bisporus NCIM 3296 belong to the same species but showed different efficacies in improving mineral mobilization. Phytase from Z. bisporus NCIM 3296 was least efficient, while Z. bisporus NCIM 3265 was most efficient in releasing minerals. The correlation between release of minerals and liberation of phosphate from PA showed that about 30–40% increase in mineral release was observed per 2.5–3 times more phosphate released from phytase treated CPF. In vitro solubility of Ca2+, Fe2+ and Zn2+ in CP was improved after the application of phytase. The minerals were released in the order Ca2+ > Fe2+ > Zn2+. It was earlier reported that cation capacity to bind PA was in the order Cu2+ > Zn2+ > Fe2+ > Ca2+, and the stability of the mineral–phytate complex was in the order Zn2+ > Cu2+ > Ca2+ (Jin and Ma 2005). It was observed that Zn2+ was less accessible as compared with Ca2+, which may be due to weaker binding power of Ca–phytate complex than Zn–phytate complex. The mineral content of legumes is generally high, but the presence of PA limits its mobilization since it is the main inhibitor of Fe2+ and Zn2+ absorption (Sandberg 2002). Sandberg and Svanberg (1991) have shown that the digestion of cereal samples with phytase led to increase in accessibility of Fe2+ significantly. Similar observation was also recorded in the present study. The increase in mobilization of Zn2+, Fe2+ and Ca2+ was 20–28%, 26–37% and 24–42%, respectively, suggesting that improved accessibility was achievable.
Figure 3. Phytic Acid Content (mg/g) and the Phytase Activities in Untreated and Treated CPF at 37C in 10 mL Acetate Buffer (pH 5.0) over 3 h
Results are expressed as the mean ± SD of triplicate samples.
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Figure 4. % Mineral Mobilization for CPF after Treatment of Crude Phytase at 37C in 10 mL Acetate Buffer (pH 5.0) over 2 h
Results are expressed as the mean ± SD of duplicate samples.
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PA content in untreated CPF, estimated using HPLC was 9.6 mg/g, and only 12–25% residual PA was observed in CPF after 2 h treatment with phytases. PA content of untreated CPF was found to be comparable with previously reported values (Chitra et al. 1996). Sandberg and Svanberg (1991) have reported that phytase was able to digest PA completely within 2 h. It was reported that cooking of soaked chick pea seeds lowered PA content by 20–26% (Duhan et al. 1989), whereas fermentation reduced the PA content by 26–39% (Chitra et al. 1996). Our results showed that enzymatic treatment in the processing of CPF was very efficient in hydrolyzing PA as compared to the previously reported treatments (Duhan et al. 1989; Chitra et al. 1996). This suggests the potential of yeast phytase in present study for its use in releasing PA-bound phosphorus from various commercial food and feed.
In conclusion, this study presents the first report on yeasts (i.e., Z. bisporus NCIM 3265, Z. bisporus NCIM 3296, S. octosporus NCIM 3297, W. saturnus NCIM 3298 and Z. priorionus NCIM 3299) producing cell-wall-associated phytase. Moreover, these yeasts exhibited higher cell-wall-associated phytase activity than previously reported yeast phytases. The fact that these yeast phytases possess the ability to degrade PA and enhance mineral mobilization from CPF suggests that yeast phytases may be utilized for processing and manufacturing of food for human consumption. Furthermore, genetic improvement of these natural yeast strains will broaden its future industrial applications as dietary yeast supplement and whole cell bio-catalyst.