Vol 24, No. 9;Sep 2017
PROTEOLYTIC ACTIVITY OF RECENT FORAMINIFERAL FUNGI ISOLATED FROM DIFFERENT COASTAL LAGOONS IN RED SEA COAST, EGYPT Mohamed Youssef1,2 and Abdel-Rahman Saleem3,4* 1
Department of Geology and Geophysics, Faculty of Science, King Saud University, 2455 Riyadh, 11451 Saudi Arabia. 2 Geology Department, Faculty of Science, South Valley University, 83523 Qena, Egypt 3 Biology Department, Faculty of Science, Taibah University, 344 Almadinah Almonawarah, Saudi Arabia 4 Botany Department, Faculty of Science, South Valley University, 83523 Qena, Egypt *Corresponding author: Tel: +966532806243 E-mail:
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Abstract Twenty nine fungal species were isolated from 30 samples of picked recent foraminferal tests belong to three different coastal lagoons (Abu-Shaar, Umm al-Huwaytāt and Marsa Shūni lagoons) in red sea coast, Egypt. Aspergillus, Emericella, Paecilomyces, Penicillium and Trichoderma were the most prevalent genera on both dextrose Czapeks and potato dextrose agar media at 28°C. From the above genera, the most dominant species were Aspergillus flavus, A. niger, A. terreus, Emericella nidulans, Paecilomyces variotii, Penicillium chrysogenum and Trichoderma harzianum. Screening of fungi for their abilities to produce protease enzyme indicated that, twelve species represented 41.38% of total fungi exhibited high and moderate protease activity each. However, five species (17.24%) were low producers. Aspergillus flavus, Fusarium oxysporum and Penicillium chrysogenum were the most active species for protease production. These fungi exhibited maximum protease activity after 6 days of incubation at 32°C with initial pH 6. The crude enzyme of A. flavus culture extract was concentrated by ammonium sulphate and dialysis. The partially purified enzyme produced 4.8 fold with recovery of 4 and the molecular weight estimated by SDSPAGE was about 48 kDa.
Key words: Foraminiferal fungi, protease activity, Red Sea coast Introduction Fungi is one of the most diverse groups of organisms on the earth, approximately 1.5 million species have been estimated, of which about 70.000 have been described (Hawksworth, 1991, 2001). They live as saprobes, decaying dead organic materials, or in symbiosis with other organisms (Alexopoulos et al. 1996).
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Study of foraminiferal fungi provides valuable information about diversity, structure and evolution of fungi (Hibbett et al., 1995, 1997). These studies have revealed the role of fungi in biological relationships with plants and animals (Taylor et al., 2009). Ascocarps of intertidal species or varieties of genera Arenariornyces, Corollospora and Lindra colonized foraminifera in the natural
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habitat and contributed to the breakup of the tests. Fungal fruiting structures, which are normally subglobose when developing in other substrates, adjust to the shape of the foraminifera chambers when growing within the tests. Pure cultures of L. thalassiae var. crassa yielded the same type of irregular fruiting bodies when grown at 30°C on foraminiferal tests as the sole source of nutrients (Kohlmeyer, 1985). The evolutionary history of fungi based on foraminifera is generally limited due to incomplete fungal foraminiferal record (Taylor et al., 2011). Only a few geologic deposits have yielded fungal preserved in sufficient detail to assignment the major evolution tree of fungal life. The most famous of these deposits is the Lower Devonian. Rhynie chert, which has been instrumental in our conception of fungi in early continental ecosystems (Taylor et al., 2004). Other well preserved fungi occur in Carboniferous chert and coal balls (Taylor et al. 2005; Krings et al. 2007, 2009, 2010, 2011; Dotzler et al. 2011), as well as in Triassic permineralized peat from Antarctica (Osborn and Taylor 1989; Taylor and White 1989; Phipps and Taylor 1996; Schwendemann et al. 2011). The record of fungi from Triassic peat deposits in Antarctica includes several types of sporocarps and sporangia/spores that have been belonging to the zygomycetes fungi. The presence of hyphae was located as suspensor cells or gametangia (Taylor and White 1989; White and Taylor 1989, 1991). These fungal structures are significant because they document the evidence of zygomycetes in such habitat (Taylor et al. 2009). These fungal remains were also similar to the zygosporangiumgametangia complex in certain modern Endogonales.
reproduced by an alternation of generations through haploid-diploid life cycles. Larger benthic foraminifera are highly specialized protests. Some species host photosynthetic algae as symbionts. This form of symbiosis is only profitable in warm, oligotrophic seas within the photic zone (Renema & Hart, 2012). In modern seas symbiont-bearing foraminifera are restricted to areas with a minimum sea surface temperature of 16° C in the coldest month (Langer & Hottinger, 2000). Fungi are utilized substrates producing various hydrolytic enzymes. Proteases are able to hydrolyze the peptide bond in a protein chain. They represent one of the largest groups of enzymes with increasing market demands due to their beneficial applications in industrial, biotechnological, medicinal and basic research fields (Souza et al. 2015). Protease biosynthesis by fungi belonging to the genera Aspergillus, Penicillium and Rhizopus was reported by several researchers (Hamzah et al. 2009; Saleem and El-Said 2009; Rodarte et al. 2011; Anitha and Palanivelu 2012; Choudhary and Jain 2012; Radha et al. 2012; Kutateladze et al. 2013; Bensmail et al. 2015). Recently, proteases from several Aspergillus species have been isolated and characterized (Muthulakshmi et al. 2011; Sharma and De 2011; Anitha and Palanivelu 2012; Palanivel et al. 2013; Niyonzima and More 2015). This study aimed to investigate the distribution and abundance of different fungal genera and species on large foraminiferal tests picked from three lagoons along the Red sea coast in Egypt as well as the proteolytic activity of foraminiferal fungi, its optimization conditions and partially purification and characterization of protease produced by Aspergillus flavus.
Larger foraminifera are unicellular protests housed within a hardened shell or test that is at least 3 mm3 in volume. The tests are made of calcium carbonate. It is
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Materials and Methods Environmental conditions and geology Abu Shaar lagoon is located to the north Hurghada (north of the Marine Biological Station, about 5 km) with maximum depth 8 m during the high tide level, its maximum length ca 1000m (Fig. 1). It is lying at latitude 27o 18\ 16\\ N and longitude 33o 45\ 04\\ E. This lagoon is often bordered by beach to the west, and fringing coral reefs to the east. It has two water entrances, the first entrance is narrow and situated in the southward, while the second one is relatively wide and situated at the north side. Its bottom floor is a mixture of sand, which consists of terrigenous materials, mud, and biogenic fragments. Umm al-Huwaytāt lagoon is situated 10km south Safaga city. It is located between latitude 26o 38\ 55. 6\\ N and longitude 33o 57\ 44\\ E. Generally, Safaga area is occupied by relatively low hill of sedimentary rocks surrounded by mountains of igneous and metamorphic rocks. It occupies about 1500 m 2 with gentle slope. its boundaries are fringing reefs to the east and beach to the west, (Fig. 1). It has one inlet from the north and the maximum recorded depth is about 11m with a sandy substrata. Marsa Shūni lagoon is located at 50 km north Marsa Alam city between latitude 25o 26\ 27\\ N and longitude 34o 41\ 37\\ E. The lagoon is parallel to the shore with 1 km long and more than 500m wide (Fig. 1). It is very shallow and can be divided into the following zones: 1) The shore is narrow and composed of fine sand, 2) The intertidal zone is formed of fine sand with no vegetation. Collection and pretreatment of samples Sediment samples were collected from the extension of three selected coastal lagoons along the Red sea coast at Abu-
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Shaar , Umm al-Huwaytāt and Marsa Shūni lagoons (Fig.1) with a small Van grab sampler and some samples were taken by scuba diving and preserved in ethyl alcohol (30 samples). Each sediment sample was washed on a 200 mish (aperture 74 μm) sieve, the residue stained with rose Bengal (Walton 1952), then sieved again to remove the surplus stain and finally dried in an oven at 70°C. The faunal investigations were carried out on four fractions of sediments (1.0, 0.5, 0.250, and 0.125 mm). Separate living and dead assemblage counts were made each of 250–300 individuals. The foraminiferal species in each sample were identified based on the classification of foraminifera by Loeblich & Tappan (1988). The most common foraminiferal genera which used in this study are: Borelis schlumbergeri, Peneroplis planatus, Sorites marginalis, Eliphidum crispum, Quinqueloculina agglutinans, Spiroloculina angulata (Madkour and Youssef, 2011) (Fig. 2). Determination of foraminiferal fungi The baiting technique was used for determination of foraminiferal fungi on two types of media, Dextrose-Czapek's and potato dextrose agar at 28ºC. A known weights of foraminifera were spread on the surface of the agar medium in each plate. Five replicates were prepared for each sample and each type of cultivation media. Cultures were incubated at 28ºC for 5-7 days. The developing fungi were counted, identified and calculated per g of foraminifera. Media used for isolation of fungi Dextrose-Czapek’s agar (sodium nitrate 3.0, potassium dihydrogen phosphate 1.0, Magnesium sulphate 0.5, potassium chloride 0.5, ferrous sulphate 0.01, dextrose 10.0, agar 15.0 g/L) and potato dextrose agar (200 potato, 20 dextrose and 15.0 agar g/L) were used. Amoxycillin (0.5 mg/ml) was used as antibacterial agent (Al-Doory 1980).
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Figure 1. Sampling locations along the Red Sea coast, Egypt. Screening of fungi for their abilities to produce protease enzymes Twenty nine fungal species belong to 15 genera, collected from foraminifera, were screened for their abilities to produce extracellular protease enzyme. Fifty ml of casein yeast extract broth with ingredients (g/L) Casein acid hydrolysate 30.0, Yeast
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extract 4.0, Dipotassium phosphate 0.5 and Dextrose 2.0 (pH at 25°C, 7.2±0.2) were dispensed into each 250 ml Erlenmeyer conical flasks. The flasks were autoclaved for 15 minutes at 1.5 atm. Each flask was inoculated with two agar mycelial discs (10 mm diameter) of tested fungi obtained from 7 days old cultures growing on the same agar medium.
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Cultures were incubated at 28ºC for 7 days. Then the mycelium was harvested by filtration. Filtrates were centrifuged at 5000 rpm for 10 minutes and the supernatant were assayed for proteolytic activity according to the method of casein agar cup plate clearing zone assay (ElGendy, 1966) as follow: Aliquots of 0.2 ml of the above filtrate was dropped into 10 mm cavities previously made in the above solid medium. After 48 hours of incubation at 28ºC, the plates were folded with 5% trichoroacetic acid solution. The average diameter of degradation zone (in mm) of the triplicates was measured. Assay for protease activity Protease activity was determined according to the method described by Sumantha et al. (2006). This method based on casein hydrolysis in which casein is used as a substrate. The release of soluble tyrosine was estimated by Folin reagent. The activity was calculated from the difference between the control and the experimental titration values. Enzyme activity was expressed by units of Ltyrosine liberated during the reaction. The units were determined by a standard curve early constructed using different concentrations of L-tyrosine. Partial purification and characterization of protease enzyme Crude extract of Aspergillus flavus was centrifuged and the supernatant was precipitated by 70% saturation with ammonium sulphate and then dialyzed against 50mM phosphate buffer (pH 7.0) for 24 hours at 4ºC. The filtrate was loaded onto a DEAE-Cellulose dialysis sack (Sigma, USA) equilibrated with phosphate buffer, 50mM, pH 7.0. Then the solution was centrifuged at 10000 rpm for 15 minutes at 4°C. The supernatant was discarded and the precipitate was dissolved in 1 ml phosphate buffer solution. SDS-PAGE electrophoresis was carried out and molecular weight was determined. The protein content was
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estimated by the method of Lowry et al. (1951). Statistical analysis Statistical analysis of data was carried out by one way analysis of variance. The means and standard deviations were separated by Tukey's honest significant difference test using Biostat 2008 statistical analysis program (Copyright © 2001-2009 Analystsoft). Results and Discussion Mycobiota of foraminifera Twenty nine fungal species were recovered from foraminiferal tests picked from 30 sediment samples on both dextrose-Czapek's (16 genera+28 species) and potato dextrose (16 genera+29 species) agar at 28ºC. Aspergillus (9), Emericella (1), Paecilomyces (1), Penicillium (3) and Trichoderma (2) were the most common genera on the two types of media. These genera represented 35.71, 2.5, 11.43, 16.43 and 6.07 % of total fungi (for Om Alhowaitat sector), 51.6, 7.83, 4.98, 1.68 and 8.18 % (for Marsa Alshoni sector) and 46.65, 9.45, 4.27, 11.28 and 7.93 % (for Abo shaar sector), respectively on dextrose Czapek’s agar at 28ºC. However on potato dextrose agar, these genera contributed 38.75, 6.92, 8.3, 15.92 and 6.92 % for Om Alhowaitat , 64.12, 9.05, 4.31,10.34 and 6.03 % for Marsa Alshoni and 44.52, 6.98, 2.99, 12.62 and 11.29 % of total fungi for Abo shaar sector, respectively. From the above genera, the most prevalent species were Aspergillus flavus, A. niger, A. terreus, Emericella nidulans, Paecilomyces variotii, Penicillium chrysogenum and Trichoderma harzianum. These fungi contributed 11.4316.16, 7.86-18.5, 3.93-7.32, 5.71-9.45, 4.27-11.43, 10.68-16.43 and 6.07-8.18% of total fungi, respectively in all sectors on dextrose-Czapek's agar. However, on potato dextrose agar they contributed 9.91-14.53, 3.32-15.52, 5.19-6.03, 6.92-
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9.05, 2.99-8.3, 4.74-7.97 and 3.02-9.3% of total fungi, respectively in all sectors (Tables 1,2). Kohlmeyer (1985) reported that ascocarps of five species or varieties of genera Arenariornyces, Corollospora and Lindra colonized foraminiferans in the natural habitat and contributed to the breakup of the tests. Pure cultures of L. thalassiae var. crassa yielded the same types of fruiting bodies when grown on foraminiferal tests as the sole source of nutrients. Recently, Krings et al. (2012) gave an evidence of zygomycetes fungi. Fungal fossil of Jimwhitea circumtecta occurs in permineralized peat from the Middle Triassic of Antarctica were interpreted as a mantled zygosporangium that initiate from a gametangium extend from sac-like suspensor. This structure is similar to the zygosporangium-gametangia complex appear in modern Endogonales. The sporangia are borne on ovoid or elongate gametangia. Screening of fungi for their abilities to produce protease enzymes Twenty nine fungal species representing 16 genera, collected from foraminiferal tests, were assayed for their abilities to produce protease enzymes. The result obtained revealed that, all fungi tested are able to produce protease enzymes, but with variable levels. Both high and moderate protease activity were twelve species represented 41.38% of total fungi. However, five species (17.24%) were low producers (Table 3). Aspergillus flavus, Fusarium oxysporum and Penicillium chrysogenum were the most active species for protease, so these fungi were used in further studies. Rodarte et al. (2011) isolated one hundred forty-four microorganisms from coffee fruit (Coffea arabica) on casein agar to evaluate their proteolytic activities. Fifty percent of filamentous fungi were able to secrete proteases. Aspergillus dimorphicus, A. ochraceus, Fusarium moniliforme, F. solani, Penicillium fellutanum and P. waksmanii showed the highest activities.
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Pangrikar et al. (2011) assayed proteolytic activity of rhizosphere fungi of cotton. Alternaria alternata, Fusarium oxysporum and Trichoderma viride showed maximum production of protease. Narendra et al. (2012) reported that among 25 isolates recovered from soil samples collected from different fields, five isolates showed good proteolytic activity, three isolates showed moderate activity, two isolates showed poor activity and eight isolates exhibited very poor activity. Aspergillus, Mucor and Penicillium were the most protease activity with deep cultivation conditions (Kutateladze et al., 2013). Effect of incubation periods Enzyme production by microorganisms is greatly influenced by environmental factors such as temperature, pH and incubation time. Protease activity was variable with different incubation periods. Maximum protease activity was achieved after 6 days of incubation by A. flavus, F. oxysporum and P. chrysogenum. The enzyme activity was decreased when the incubation periods were decreased or increased above the optimum one (Fig. 3). Muthulakshmi et al. (2011) reported that maximum protease activity of Aspergillus flavus was found to be at 7 days of incubation. Radha et al. (2012) studied the optimal factors influencing the production of protease from Aspergillus sp. under solid state fermentation. The optimal value of protease production was after 5 days of incubation. Effect of temperature Temperature is one of the most important physical factors that affects the growth and development of microorganisms. The optimum temperature for protease activity was recorded at 32°C. Decreasing of temperature below the optimum or increasing it above the optimum led to the decreasing of enzyme production by A. flavus, F. oxysporum and P. chrysogenum (Fig. 4). Muthulakshmi et al. (2011)
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reported that 30°C was the optimum temperature for protease production by Aspergillus flavus. Radha et al. (2012) studied the optimal factors influencing the production of protease from Aspergillus sp. under solid state fermentation. The optimal value of protease production was at temperature 32±2ºC. On the other hand, Kutateladze et al. (2013) reported
that the optimum temperature for protease was recorded at 35, 40 and 30ºC in Aspergillus, Mucor and Penicillium spp. respectively. Recently, Bensmail et al. (2015) reported that the maximum activity of milk protease by local isolate of Aspergillus niger was obtained at temperature 30°C.
Table 1. Total counts (TC, calculated per g in all samples), number of cases of isolation (NCI, out of 10 for each sector), and occurrence remarks (OR) of fungal genera and species recovered from foraminfera on dextrose-Czapek’s agar at 28°C. Om Alhowaitat sector Marsa Alshoni sector Abo shaar sector Fungal genera & species TC NCI & % TC NCI & % TC NCI & % OR OR OR Acremonium strictum 5 3L 1.78 6 4L 1.83 Alternaria alternata 9 5M 3.21 9 4L 3.20 8 5M 2.44 Aspergillus 100 10 H 35.71 145 10 H 51.60 153 10 H 46.65 A. candidus 4 2R 1.43 5 3L 1.78 4 3L 1.22 A. flavus 32 10 H 11.43 40 10 H 14.23 53 10 H 16.16 A. fumigatus 7 4L 2.50 12 5M 4.27 10 5M 3.05 A. niger 22 9H 7.86 52 10 H 18.50 46 10 H 14.02 A. sydowii 5 3L 1.78 7 3L 2.13 A. terreus 11 5M 3.93 19 6M 6.76 24 7M 7.32 A. ustus 9 4L 3.21 5 3L 1.78 A. versicolor 10 4L 3.57 12 5M 4.27 9 4L 2.74 Cladosporium cladosporioides 8 3L 2.86 5 3L 1.52 Cochliobolus 7 3L 2.50 7 3L 2.49 14 6M 4.27 C. lunatus 2 1R 0.71 6 3L 1.83 C. spicifer 7 3L 2.50 5 2R 1.78 8 4L 2.44 Emericella nidulans 16 8H 5.71 22 8H 7.83 31 9H 9.45 Fusarium oxysporum 4 2R 1.43 2 1R 0.71 3 2R 0.91 Gibberella fujikuroi 3 2R 1.07 2 2R 0.61 Mucor 10 4L 3.57 4 2R 1.42 6 3L 1.83 M. hiemalis 4 2R 1.43 M. racemosus 6 3L 2.14 4 2R 1.42 6 3L 1.83 Mycosphaerella tassiana 14 7M 5.00 8 4L 2.85 5 3L 1.52 Nectria haematococca 3 2R 1.07 Neurospora crassa 7 3L 2.49 10 5M 3.05 Paecilomyces variotii 32 10 H 11.43 14 6M 4.98 14 6M 4.27 Penicillium 46 10 H 16.43 30 10 H 10.68 37 10 H 11.28 P. chrysogenum 26 9H 9.28 16 7M 5.69 22 8H 6.71 P. funiculosum 12 6M 4.28 6 3L 2.13 5 3L 1.52 P. oxalicum 8 4L 2.86 8 4L 2.85 10 5M 3.05 Rhizopus stolonifer 6 3L 2.14 10 5M 3.56 8 4L 2.44 Trichoderma 17 8H 6.07 23 8H 8.18 26 9H 7.93 T. harzianum 11 6M 3.93 18 7M 6.40 22 8H 6.71 T. viride 6 3L 2.14 5 3L 1.78 4 2R 1.22 Total counts 280 100 281 100 328 100 Number of genera 15 12 15 Number of species 26 22 25 H= High occurrence, 8-10, M= Moderate occurrence, 5-7, L= Low occurrence, 3-4, R=Rare occurrence, 1-2.
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Table 2. Total counts (TC, calculated per g in all samples), number of cases of isolation (NCI, out of 10 for each sector), and occurrence remarks (OR) of fungal genera and species recovered from foraminfera on potato dextrose agar at 28°C.
Fungal genera & species Acremonium strictum Alternaria alternata Aspergillus A. candidus A. flavus A. fumigatus A. niger A. ochraceous A. sydowii A. terreus A. ustus A. versicolor Cladosporium cladosporioides Cochliobolus C. lunatus C. spicifer Emericella nidulans Fusarium oxysporum Gibberella fujikuroi Mucor M. hiemalis M. racemosus Mycosphaerella tassiana Nectria haematococca Neurospora crassa Paecilomyces variotii Penicillium P. chrysogenum P. funiculosum P. oxalicum Rhizopus stolonifer Trichoderma T. harzianum T. viride Total counts Number of genera Number of species
Om Alhowaitat sector TC NCI & % OR 4 3L 1.38 8 4L 2.77 112 10 H 38.75 7 2R 2.42 42 10 H 14.53 4 2R 1.38 29 10 H 10.03 15 7M 5.19 7 4L 2.44 8 4L 2.77 9 5M 3.11 6 3L 2.08 6 3L 2.08 20 8H 6.92 3 2R 1.04 3 2R 1.04 9 4L 3.11 2 2R 0.69 7 3L 2.44 6 2R 2.08 2 1 0.69 24 9H 8.30 46 10 H 15.92 23 9H 7.96 11 5M 3.81 12 6M 4.15 13 4L 4.50 20 9H 6.92 16 8H 5.54 4 2R 1.38 289 100 15 25
Marsa Alshoni sector TC NCI & % OR 6 3L 2.59 7 4L 3.02 107 10 H 64.12 4 3L 1.72 23 10 H 9.91 10 5M 4.31 36 10 H 15.52 7 3L 3.02 14 6M 6.03 3 2R 1.29 10 5M 4.31 6 3L 2.59 12 5M 5.17 5 3L 2.15 7 3L 3.02 21 8H 9.05 3 2R 1.29 3 2R 1.29 3 2R 1.29 6 3L 2.59 5 3L 2.15 10 5M 4.31 24 8H 10.34 11 6M 4.74 4 2R 1.72 9 4L 3.88 8 4L 3.45 14 8H 6.03 11 7M 3.02 3 2R 1.29 232 100 14 25
Abo shaar sector TC NCI & % OR 3 2R 1.00 11 5M 3.65 134 10 H 44.52 6 3L 1.99 38 10 H 12.62 9 4L 2.99 48 10 H 3.32 4 2R 1.33 4 3L 1.33 17 6M 5.65 8 4L 2.66 7 3L 2.32 10 4L 3.32 4 2R 1.33 6 3L 1.99 21 9H 6.98 2 1R 0.66 1 1R 0.33 6 3L 1.99 2 1R 0.66 4 2R 1.33 7 3L 2.32 6 3L 1.99 9 5M 2.99 38 10 H 12.62 24 9H 7.97 3 2R 1.00 11 5M 3.65 14 6M 4.65 34 10 H 11.29 28 9H 9.30 6 3L 1.99 301 100 15 27
H= High occurrence, 8-10, M= Moderate occurrence, 5-7, L= Low occurrence, 3-4, R=Rare occurrence, 1-2.
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Table 3. Screening of 29 fungal species recovered from foraminfera for their abilities to produce protease enzyme Protease activity Fungi clear zone (mm) Acremonium strictum 15 L Alternaria alternata 26 M Aspergillus candidus 20 L A. flavus 38 H A. fumigatus 31 H A. niger 33 H A. ochraceus 29 M A. sydowii 20 L A. terreus 26 M A. ustus 23 M A. versicolor 25 M Cladosporium cladosporioides 25 M Cochliobolus lunatus 19 L C. spicifer 32 H Emericella nidulans 32 H Fusarium oxysporum 36 H Gibberella fujikuroi 33 H Mucor hiemalis 30 H M. racemosus 27 M Mycosphaerella tassiana 24 M Nectria haematococca 28 M Neurospora crassa 20 L Paecilomyces variotii 32 H Penicillium chrysogenum 34 H P. funiculosum 25 M P. oxalicum 30 H Rhizopus stolonifer 31 H Trichoderma harzianum 29 M T. viride 27 M High =30-38 mm, Moderate = 22-29 mm, Low =15-21 mm
Effect of pH Protease activity was variable with different pH values. The optimum pH for protease activity was 6. The enzyme activity was decreased with shifting of pH to more acidic or alkaline medium by all fungi tested (Fig. 5). Muthulakshmi et al. (2011) reported that maximum protease production by Aspergillus flavus were found to be at pH 5. Similar results were obtained by Radha et al. (2012), they showed that among optimal factors influencing the production of protease from Aspergillus sp. under solid state
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fermentation, the optimal pH of protease production was 5. Recently, Bensmail et al. (2015) reported that the maximum protease activity of Aspergillus niger was obtained at initial pH 4. On the other hand, maximum alkaline protease activity was recorded in Aspergillus tamarii at pH 8-9 (Sharma and De, 2011). However, Kutateladze et al. (2013) reported that the optimum pH for protease was recorded at 6.5, 4.4 and 8.5 in Aspergillus, Mucor and Penicillium spp. respectively.
Figure 2: The most common foraminfera obtained during the study 1. Borelis schlumbergeri, Sh-4Peneroplis planatus, Sh-4Spiroloculina angulata, Sh8Quinqueloculina agglutinans, Sh3Sorites marginalis Sh-5Eliphidum crispum, Um-2. scale bar= 200 Purification of protease produced by Aspergillus flavus The purification of protease resulted in 1.3 fold purification with 69% of recovery by 70% ammonium sulphate precipitation. The purification of crude enzyme through DEAE-cellulose dialysis membrane gave 4.8 fold increases in purity with 4% recovery of protease from A. flavus (Table 4). SDS-PAGE analysis of A. flavus protease indicated that the molecular
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weight of the purified enzyme was about 48 kDa using standard molecular marker. Similar result was recorded for A. flavus protease under solid state fermentation (Muthulakshmi et al. 2011). These results are also more or less similar to those
A. flavus
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reported on Aspergillus proteases by several researchers (Sharma and De 2011; Anitha and Palanivelu 2012; Palanivel et al. 2013; Niyonzima and More 2015).
F. oxysporum
P. chrysogenum
Protease activity U/ml
40 35 30 25 20 15 10 5 0 2
4
6
8
10
12
Incubation periods (days)
Figure 3. Effect of incubation periods on protease activity of A. flavus, F. oxysporum and P. chrysogeum. A. flavus
Protease activity U/ml
60
F. oxysporum
P. chrysogenum
50 40 30 20 10 0 20
24
28
32
36
40
Temperature (°C)
Figure 4. Effect of temperature on protease activity of A. flavus, F. oxysporum and P. chrysogeum.
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A. flavus
F. oxysporum
P. chrysogenum
Protease activity U/ml
60 50 40 30 20 10 0 4
5
6
7
8
9
pH
Figure 5. Effect of pH on protease activity of A. flavus, F. oxysporum and P. chrysogeum.
Table 4. Purification of protease produced by Aspergillus flavus Purification step Culture supernatant 70 % Ammonium sulphate precipitation Dialysis
6580
Total protein (mg) 286
4540
152
29.9
1.3
69
264
2.4
110
4.8
4
Protease activity (U)
On the other hand, the molecular mass of the partially purified protease enzyme from Penicillium janthinellum and Neurospora crassa were found to be 33 kDa and 45 kDa respectively using SDS-PAGE (Abirami et al. 2011). Conclusion Although marine fungi have been described from calcareous tests of crustaceans and shells of mollusks, little information have been described on marine fungi associated with benthic foraminifera, the most abundant calcareous fossils frequently used in geochronology as well as the ability of
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Specific activity (U/mg)
Purification (fold)
Recovery (%)
23
1
100
these fungi to produce protease enzymes which play an important role in the biological degradation in marine habitat. Fungi have been adapted with physical changes in this habitat by forming carbonaceous fruiting bodies on calcareous and siliciferous substrates. This research gives an indication for the role of marine fungi in the decomposition of organic materials in such habitats due to their abilities to produce proteases. Acknowledgments This project was supported by King Saud University, Deanship of Scientific Research and College of Science Research Center.
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References Abirami, V., Meenakshi S.A., Kanthymathy K., Bharathidasan R., Mahalingam R., Panneerselvam A. 2011. Partial purification and characterization of an extracellular protease from Penicillium janthinellum and Neurospora crassa. European J. of Experimental Biology 1(3): 114-123. Al-Doory, Y. 1980. Laboratory medical mycology. Lea and Febiger, Philadelphia Kimpton publishers, London. Alexopoulos C.J., Mims C.W., Blackwell M. 1996. Introductory Mycology, 4th edn., Wiley, New York. Anitha T.S. and Palanivelu P. 2012. Production and characterization of keratinolytic protease (s) from the fungus, Aspergillus parasiticus. Inter. J. of Research in Biological Sciences 2 (2): 87-93. Bensmail S., Mechakra A., Fazouane-Naimi F. 2015. Optimization of milk-clotting protease production by a local isolate of Aspergillus niger ffb1 in solid-state fermentation. J. of Microbiol. Biotech. and Food Sci. 4 (5): 467472. Choudhary V. and Jain P.C. 2012. Screening of alkaline protease production by fungal isolates from different habitats of Sagar and Jabalpur district (MP). J. Acad. Indus. Res. 1 (4): 215–220. Dotzler N., Taylor T.N., Galtier J., Krings M. 2011. Sphenophyllum (Sphenophyllales) leaves colonized by fungi from the Upper Pennsylvanian Grand-Croix cherts of central France. Zitteliana A 51: 3–8. El-Gendy S.M. 1966. Microbial survey of blueveined cheese, Roqueforte-type. Bulletin of Science Technology 9: 367-384. Hamzah H.M., Ali A.H.L., HAMID G., Hassan H.G. 2009. Physiological regulation of protease and antibiotics in Penicillium sp. using submerged and solid state fermentation techniques. J. of Engineering Science and Technology 4 (1): 81-89. Hawksworth, D.L. 1991. The fungal dimension of biodiversity: magnitude, significance, and conservation. Mycol. Res. 95: 641-655. Hawksworth D.L. 2001. The magnitude of fungal diversity: the 1.5 million species estimate revisited. Mycol. Res. 105: 1422– 1432. Hibbett D.S., Grimaldi D. and Donoghue M.J. 1995. Cretaceous mushrooms in amber. Nature 377: 487. Hibbett D.S., Grimaldi D., Donoghue M.J. 1997. Fossil mushrooms from Miocene and
269
Cretaceous ambers and the evolution of Homobasidiomycetes. American Journal of Botany 84: 981–991. Kohlmeyer J. 1985. Marine fungi (Ascomycetes) within and on tests of Foraminifera. Marine Biology 90: 147-149. Krings M., Dotzler N., Galtier J., Taylor T.N. 2009. Microfungi from the upper Visean (Mississippian) of central France: Chytridiomycota and chytrid-like remains of uncertain affinity. Rev. Palaeobot. Palynol. 156: 319–328. Krings M., Dotzler N., Taylor T.N., Galtier J. 2007. A microfungal assemblage in Lepidodendron from the Upper Visean (Carboniferous) of central France. CR Palevol. 6: 431–436. Krings M., Dotzler N., Galtier J., Taylor T.N. 2010. A fungal community in plant tissue from the Lower Coal Measures (Langsettian, Lower Pennsylvanian) of Great Britain. Bull. Geosci. 85: 679–690. Krings M., Taylor T.N., Dotzler N., Galtier J. 2011. Fungal remains in cordaite (Cordaitales) leaves from the Upper Pennsylvanian of central France. Bull. Geosci. 86: 777–784. Krings M., Taylor T.N. Dotzler N., Persichini G. 2012. Fossil fungi with suggested affinities to the Endogonaceae from the Middle Triassic of Antarctica. Mycologia, 104 (4): 835–844. Kutateladze L., Zakariashvili N., Jobava M., Urushadze T., Khvedelidze R., Kvesitadze G. 2013. Selection of microscopic fungi-proteases producers. Bull. of the Georg. Nat. Acad. of Sci. 7 (3): 87-91. Langer, M.R. & Hottinger, L. 2000. Biogeography of selected “larger” foraminifera. Micropaleontology, 46 (1): 105-127. Loeblich, A.R. & Tappan, H. 1988. Foraminiferal evolution, diversification, and extinction. Journal of Paleontology, 62, 695714. Lowry, O.H., N.J. Rosebrough, A.L. Farr, and R.J. Randall 1951. Protein measurement with the folin phenol reagent. J. Biol. Chem. 193: 265-275. Madkour, H.A., Youssef, M. 2011. Distribution and diversity of benthic Foraminifera from the coastal lagoons along the Egyptian Red Sea. Egyptian J. of Aquatic Research, 37(1): 41-58. Muthulakshmi C., Gomathi D., Kumar D.G., Ravikumar G., Kalaiselvi M. and Uma C. 2011. Production, purification and characterization of protease by Aspergillus flavus under solid state fermentation. Jordan J. of Biological Sciences 4 (3): 137-148.
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Narasimha G., Radha S., Babu R.H., Sridevi A., Prasad N.B.L. 2012. Development of mutant fungal strains of Aspergillus niger for enhanced production of acid protease in submerged and solid state fermentation. European J. of Experimental Biology 2 (5): 1517-1528. Narendra D., Ramalakshmi N., Ramalakshmi M., Roja S., Archana B.K.N., Maanasaet G. 2012. Isolation and characterization of protease producing bacterial from soil and estimation of protease by spectrophotometer. The Experiment 1 (1): 1-7. Niyonzima F.N. and More S.S. 2015. Purification and characterization of detergentcompatible protease from Aspergillus terreus gr. Biotech. 5: 61-70. Osborn J.M. and Taylor T.N. 1989. Palaeofibulus gen. nov., a clamp-bearing fungus from the Triassic of Antarctica. Mycologia 81: 622–626. Palanivel P., Ashokkumar L., Balagurunathan R. 2013. Production and purification and fibrinolytic characterization of alkaline protease from extremophilic soil fungi. Inter. J. Pharm. Bio. Sci. 4 (2): 101-110. Pangrikar P.P., Wadikarl M.S., Borde V.U. and Chavan A.M. 2011. Producnon of protease enzyme from rhizospheric fungi of bt and non bt cotton varieties. Bioscience Discovery 2 (2): 249-250. Phipps C.J., Taylor T.N. 1996. Mixed arbuscular mycorrhizae from the Triassic of Antarctica. Mycologia 88: 707–714. Radha S., Sridevi A., HimakiranBabu R., Nithya V.J., Prasad N.B.L. and Radha G. 2012. Medium optimization for acid protease production from Aspergillus sps under solid state fermentation and mathematical modelling of protease activity. J. of Microbiol. and Biotech. Res. 2 (1): 6-16. Renema, W. and Hart, M.B. 2012. Larger benthic Foraminifera of the type Maastrichtian. Scripta Geol., Spec. Issue 8: 33-43. Rodarte M.P., Dias D.R., Vilela D.M. and Schwan R.F. 2011. Proteolytic activities of bacteria, yeasts and filamentous fungi isolated from coffee fruit (Coffea arabica L.). Maringa 33 (3): 457-464. Saleem A. and El-Said A.H.M. 2009. Proteolytic activity of beef luncheon fungi as affected by incorporation of some food
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preservatives. Acta Microbiol. et Immunol. Hung. 56 (4): 417-426. Schwendemann A.B., Decombeix A.L., Taylor T.N., Taylor E.L., Krings M. 2011. Morphological and functional stasis in mycorrhizal root nodules as exhibited by a Triassic conifer. Proc. Natl. Acad. Sci. USA 108:13630–13634. Sharma N. and De K. 2011. Production, purification and crystallization of an alkaline protease from Aspergillus tamarii [EF661565.1]. Agric. Biol. J. of North America 2 (7): 1135-1142. Souza P.M., Assis Bittencourt M.L., Caprara C.C., Freitas M., Almeida R.P.C., Silveira D., Fonseca Y.M., Filho E.X.F., Junior A.P., Magalhães P.O. 2015. A biotechnology perspective of fungal proteases. Brazilian J. of Microbiol. 46 (2): 337-346. Sumantha A., Larrochhe C. and Pandey A. 2006. Microbiology and industrial biotechnology of food-grade proteases: a perspective. Food Technol. Biotechnol. 44: 211-220. Taylor T.N., Klavins S.D., Krings M., Taylor E.L., Kerp H., Hass H. 2004. Fungi from the Rhynie chert: a view from the dark side. Trans. Roy. Soc. Edinburgh, Earth Sci. 94: 457–473. Taylor T.N., Krings M., Dotzler N., Galtier J. 2011. The advantage of thin sections over acetate peels in the study of late Paleozoic fungi and other microorganisms. Palaios 26: 239–244. Taylor T.N., Krings M., Klavins S.D., Taylor E.L. 2005. Protoascon missouriensis, a complex fossil microfungus revisited. Mycologia 97: 725–729. Taylor T.N., Taylor E.L., Krings M. 2009. Paleobotany. The biology and evolution of fossil plants. New York: Elsevier Academic Press Inc. xxi + 1230 p. Taylor T.N. and White J.F. 1989. Fossil fungi (Endogonaceae) from the Triassic of Antarctica. Am. J. Bot. 76: 389–396. White J.F. and Taylor T.N. 1989. Triassic fungi with suggested affinities to the Endogonales (Zygomycotina). Rev. Palaeobot. Palynol. 61: 53–61. White J.F. and Taylor T.N. 1991. Fungal sporocarps from Triassic peat deposits in Antarctica. Rev. Palaeobot. Palynol. 67: 229– 236.
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