Evaluating the Biodeterioration Enzymatic Activities of Fungal Contamination Isolated from Some Ancient Yemeni Mummies Preserved in the National Museum

Naji, Khalid Mohammed; Abdullah, Qais Yusuf M.; AL-Zaqri, Aida Qaseem M.; Alghalibi, Saeed M.

doi:https://doi.org/10.1155/2014/481508

Biochemistry Research International

On this page

Abstract Introduction Materials and Methods Results and Discussion Conclusion References Copyright Related Articles

Research Article | Open Access

Volume 2014 | Article ID 481508 | https://doi.org/10.1155/2014/481508

Evaluating the Biodeterioration Enzymatic Activities of Fungal Contamination Isolated from Some Ancient Yemeni Mummies Preserved in the National Museum

Khalid Mohammed Naji,¹Qais Yusuf M. Abdullah,²Aida Qaseem M. AL-Zaqri,³and Saeed M. Alghalibi²

Academic Editor: Saad Tayyab

Received29 Jul 2014

Revised25 Sept 2014

Accepted16 Oct 2014

Published13 Nov 2014

Abstract

Sophisticated mummification using chemical preservation was prevalent in ancient Yemeni civilization as noted in the 4th century B.C. mummies of the National Museum of Yemen, Sana’a, used in this study. Five of these mummies were used to evaluate hydrolytic enzymes produced as a result of fungal contamination. Forty-seven fungal species were isolated, thereby reflecting a high degree of contamination which may have resulted from the poor ventilation and preservation system. Aspergillus was the most common genus isolated (48.9%). Fifteen isolates exhibited ability to produce cellulase (EC; 3.2.1.4), Aspergillus candidus being the highest cellulose-producer. Pectin lyase (PL, EC; 4.2.2.2) and pectin methyl esterase (PME, EC; 3.1.1.11) were produced by Trichoderma hamatum, whereas chitinase (EC; 3.2.1.14) was produced by Aspergillus niger. Protease activity was noted by only Cladosporium herbarum. The higher activities of these fungal hydrolytic enzymes represent the major threats of biodeterioration including deteriorating linen bandages as well as the mummy bodies. Therefore, it is recommended to improve the preservation system of the mummies at the National Museum to minimize the contamination up to the lowest level and protect the mummies from biodeterioration.

1. Introduction

Human mummies are extremely susceptible to damage from environmental factors [1]. There are a number of abiotic and biotic factors, such as pollution, light, humidity, temperature, microorganisms, and insects, that have deteriorating effects on museum materials [2]. The most serious damage on mummies occurs by microorganisms such as bacteria, actinomycetes, and fungi [1, 3, 4].

In an indoor environment, fungi are the most hazardous microorganisms responsible for the biocontamination of museum collections. Unlike bacteria whose growth in museums is limited by their need for water, fungi are able to grow at relatively low levels of temperature and humidity such as those usually maintained in the museums [5, 6]. Even in quite harsh conditions including dryness and low temperature, fungal spores are able to survive for a long time [7]. Fungi are usually present in display and storage conditions of museums [8, 9].

Fungal degradation is one of the highest risk factors for deterioration of ancient mummies [10, 11]. The deterioration by fungi seems to be a predominant feature in the museums of many countries all over the world [8, 12]. Fungi excrete enzymes that digest organic matter, thereby altering and weakening those materials. In addition, many molds contain colored substances that can cause stains and spots on textiles and decrease the strength of the fabric. Many fungi can be dangerous to people and in some cases pose a major health hazard and cause zoonotic superficial infections as a consequence of invading keratinized tissues of skin, hair, and nails [13, 14].

Fungal biodeterioration results in a change in the quality or value of a material and makes it less functional in utilization terms. A battery of extracellular hydrolytic enzymes excreted by fungi results in the formation of acidic products that cause chemical alteration of the material under attack [15]. Cellulases catalyze hydrolysis of cellulose, break internal bonds to disrupt its crystalline structure, and expose individual cellulose polysaccharide chains [16]. Pectinases catalyze hydrolysis of pectic substances readily soluble in water. Pectolytic enzymes are divided into two classes, namely, pectinesterases and pectin depolymerases. The former enzyme deesterifies pectin by the removal of methoxy residues and is called pectin methoxy esterase (PME), while the latter splits the main chain and is further classified as polygalacturonase (PG) and pectin lyases (PL) [17–19]. Chitinases are glycosyl hydrolases having the ability to degrade β-1, 4-linked insoluble linear polymers of N-acetylglucosamine (chitin) directly to low molecular weight chitooligomers [20]. Proteases, a complex group of hydrolytic enzymes, hydrolyze proteins into small peptides and amino acids by the addition of water across amide bonds. They have evolved different classes of proteases that perform the same reaction by using completely different catalytic mechanisms. Fungi are known to produce acid, neutral, alkaline, and metalloproteases. A single organism can produce more than one type of protease [21]. Fungal proteases are active over a wide pH range (pH 4 to 11) and exhibit broad substrate specificity [22].

The ancient Yemeni mummies discovered in 1983 in mountain caves near Shebam al-Giras were rescued by the Archeology Department Expedition of University of Sana’a [23]. The Yemeni mummies are different from those of Egypt. They are older and go back to the 4th century B.C. according to results of radiocarbon analysis of the leather and linen wrappings [24]. The dead bodies were treated different from the Egyptian ones. Local materials used in mummification to absorb moisture such as raá plant (Aerva javanica), henna (Lawsonia inermis), myrrh, and other aromatic plants were used to preserve the corpses. Oxides of iron and sulfur were used to fix proteins in the corpse. The process was completed with the corpses being wrapped in linen (Linum usitatissimum) bandages. The bandaging took place over an elongated period of days [25]. Most of the materials used in the mummification are natural organic substances. Hence, these mummies are liable to deterioration. In addition, the storage conditions of the mummified corpses in the museum display are in unsuitable environmental conditions. These mummies are of special importance as they depict ancient Yemeni civilization and its use of advanced medical and scientific knowledge in the preservation of the corpses.

This study aimed at screening of fungal contamination of five Ancient Yemeni mummies preserved in the National Museum of Yemen at Sana’a. The fungal hydrolytic enzymes assessed were cellulase, PL, PME, chitinase, and protease. This in turn will help in our understanding for the use of better preservative conservation procedures in order to safeguard the mummies from future damage.

2. Materials and Methods

2.1. Mummy Samples

Five mummies preserved in the National Museum of Yemen at Sana’a were used in this study for the collection of fungal contamination. The details of the mummies used are as described in Table 1.

2.2. Screening of Fungal Contamination

Samples were taken from different skin areas (face, hands, and feet) of the five mummies. Fungi were isolated using the conventional methods of swabbing and streaking [26] on Potato Dextrose Agar (PDA). Plates incubated at °C for 7 days. Three replicates were prepared for each sample. The resultant colonies were isolated, purified, and identified. Pure cultures of the identified fungi were transferred to a slant and used for physiological studies. Fungi were identified based on their morphology according to Raper and Fennell [27], Ellis [28], and Barnett and Hunter [29].

2.3. Screening the Hydrolytic Enzymatic Activity of Isolated Fungi

Twenty-one isolates were examined for their ability to produce common hydrolytic enzymes. Standard methods were used for assay of the following enzyme activities: cellulase [30], pectinase [31], chitinase [32], and protease [33].

2.4. Production of Enzymes in Liquid Nutrient

The isolates that were found to exhibit highest activity of each enzyme on solid media were grown in liquid media for further analysis.

2.5. Growth Condition

The fungi were grown in Erlenmeyer flasks (100 mL) containing 25 mL of the liquid growth medium. After sterilization, the flasks were inoculated with a single 6 mm disc cut out from the margin of 5 d colony of the fungus grown on Czapek’s agar medium. The inoculated flasks were incubated as stationary culture at 30°C for 7, 14, and 21 days [34]. The experiments were carried out in triplicate.

2.6. Growth Medium

Cellulase. 100 mL of the liquid medium contained cellulose, 1 g; (NH₄)₂SO₄, 0.14 g; K₂HPO₄, 0.6 g; KH₂PO₄, 0.20 g; and MgSO₄7H₂O, 0.01 g in distilled water [35]. An isolate of Aspergillus candidus was inoculated into sterile media for the production of cellulase.

Pectinase. Trichoderma hamatum isolate was used for the production of PL and PG enzymes using the medium described by Eggins and Pugh [36]. The composition per 1000 mL was pectin, 10 g; L-asparagine, 0.5 g; (NH₄)₂SO₄, 0.5 g; KH₂PO₄, 1.0 g; KCl, 0.5 g; CaCl₂, 0.2 g; yeast extract, 0.5 g; MgSO₄7H₂O, 0.2 g in distilled water. The pH was adjusted to 5 using acetate buffer.

Chitinase. Aspergillus niger isolate was used for the production of chitinase using the medium described by Hsu and Lockwood [32]. 100 mL of liquid growth medium contained NaNO₃, 0.5 g; KH₂PO₄, 0.2 g; KCl, 0.1 g; yeast extract, 0.05 g; MgSO₄ 7H₂O, 0.05 g in distilled water. The pH was adjusted to 5 using acetate buffer.

Protease. Cladosporium herbarum isolate was used for protease production in liquid medium as described by Ali [37]. 1000 mL of liquid growth medium contained K₂HPO₄, 1 g; MgSO₄7H₂O, 0.5 g; KCl, 0.5 g; FeSO₄7H₂O, 0.01 g; sucrose, 30 g; casein, 1% (w/v) in distilled water. The pH was adjusted to 5 using acetate buffer.

2.7. Enzymes Assay

The culture filtrates were used as crude enzyme preparations for all mentioned enzymes.

Cellulase activity was measured by estimating the liberated amount of reducing sugar using dinitro salicylic acid (DNS), according to Miller [38] with the modification of Bernfeld [39] using glucose for the standard curve. The absorbance was read at 540 nm using boiled enzyme as control. One unit of cellulase activity is defined as the quantity of enzyme that catalyzes the liberation of 1 μmol/min of reducing sugar measured using glucose as standard.

Pectinase. Two enzymes of pectinase family were estimated.(a)Pectin lyase (PL) activity was assayed spectrophotometrically by determining uronide at 235 nm [40]. 3 mL of reaction mixture contained 0.1% pectin in 0.05 M of Tris-HCl buffer (pH 8) and 0.5 mL of culture filtrate. The reaction mixture was incubated at 30°C for 3 hours. Then 3 mL of 0.01 N HCl and 1 mL of reaction mixture were mixed and the optical density read. One unit of PL activity is defined as the amount of enzyme which causes an increase in absorbance of 0.01 in 30 min.(b)Pectin methyl esterase (PME) activity was assayed based on the method described by Kertesz [41] and modified by Olutiola and Akintunde [42]. In sterilized 25 mL conical flask, 2 mL of culture filtrate was mixed with 5 mL of 1.2% pectin (in 0.05 M Tris-HCl buffer containing 5 mM sodium carbonate, pH 8). The flask was incubated in water bath at 30°C for 3 h followed by titration with 0.02 N NaOH to adjust pH to 8. One unit of PME activity was defined as the amount of enzyme required for addition of 1 μ equivalent of NaOH per hour to maintain the reaction at pH 8.

Chitinase activity was estimated according to Monreal and Reese [43]. 5 mL reaction mixture contained 1% colloidal chitin in phosphate buffer (pH 5) and 1 mL of culture filtrate. The mixture was incubated at 37°C for 1 h, reaction arrested by adding 0.5 mL of DNS, boiled for 15 min in a water bath, and cooled to room temperature. Optical density was then measured at 540 nm against a suitable blank. The reducing sugar released was determined from the standard curve of glucose. One unit of chitinase activity is defined as the amount of enzyme, which produced 1 μmol of reducing sugar min⁻¹ under assay conditions.

Protease activity was estimated by the method of Folin and Ciocalteu [44]. The reaction mixture containing 5 mL of 0.65% casein (pH 7.5) and 1 mL of enzyme extract was mixed well by swirling and incubated at 37°C for 10 min. The reaction was arrested by adding 5 mL of cold 10% trichloroacetic acid and incubated at 37°C for 30 min. It was filtered through Whatman filter paper and the filtrate thus obtained was used for color development. 5 mL of Na₂CO₃ and 1 mL of Folin-Ciocalteu’s (FC) reagent were added to an aliquot of the filtrate and incubated at 37°C for 30 min and absorbance was read at 660 nm. The amount of amino acids released was calculated using standard curve of tyrosine. One unit of protease activity is defined as the amount of enzyme that produced 1 μmol of tyrosine min⁻¹ under assay conditions.

Protein Content. Soluble protein was estimated according to Lowry et al. method [45], using bovine serum albumin (BSA) as standard.

2.8. Determination of Dry Weight

Fungal mycelia were separated from the growth medium by filtration through Whatman no. 1 filter paper. The mycelia were then washed and dried in an oven at 60°C for 24 h. The dry weight of the mycelia was recorded.

2.9. Statistical Analysis

Two-way analysis of variance (ANOVA) was used for statistical evaluation of enzymes activities and dry weight by using GraphPad Prism version 6. Data were shown as mean ± SD. The value of was considered statistically significant.

3. Results and Discussion

Fifteen species belonging to 10 genera of fungi were isolated from five ancient Yemeni mummies on PDA. The total count of fungi isolated from all samples was 47 (Table 2). Mummy 1 (M1) had the lowest fungal contamination, that is, only two species of Aspergillus (Figure 1). In contrast, M4 showed the highest contamination with 12 isolates of 9 species (Figure 1). The other mummies showed moderate contamination. According to the numbers of isolated fungi from the five mummies, the order from higher to lower contamination is as follows: M4 > M2 = M5 > M3 > M1 (Figure 1).

The diversity of fungal profile of each mummy, although preserved in the same museum, may have resulted from environmental conditions, which differ in each room inside the museum due to several factors including the construction and design of the building. Also, the place from which the mummies were unearthed varies. Each mummy was preserved in a separate room, and preservation conditions vary from room to room in terms of ventilation, humidity, and temperature (data not shown). In addition to that, some mummies were treated by disinfectant whereas others were not.

Aspergillus was the most common genera isolated from the mummy samples with high frequency of 48.94% of the total fungal count, the most common species being A. niger (25.53%) followed by A. flavus (10.63%), while A. fumigatus, A. candidus, and A. ustus were isolated in low frequency. Cladosporium and Penicillium were isolated in moderate frequency representing 14.89% and 12.76%, respectively. Aureobasidium pullulans, Chaetomium thermophilum, Mucor circinelloides, Scopulariopsis koningii, Stachybotrys chartarum, Trichoderma hamatum, and Ulocladium chartarum were isolated in rare incidence from mummy samples.

Our results exhibited similar fungal contamination when compared to isolates from Egyptian mummies. These included Alternaria tenuis, Aspergillus humicola, Aspergillus niger, Chaetomella horrid, Chaetomium globosum, Hormodendrum viride, and Penicillium corylophilum [11]. Twenty-three mummies from cemeteries and caves of northern Mexico were found to have mainly Penicillium, Aspergillus, Cladosporium, Alternaria, Candida tropicalis, and Candida albicans [46]. Also three species of Aspergillus and one of Alternaria were found to be associated with open deteriogens skeleton of giraffe in zoological museum of Punjab University in Lahore India [47].

Twenty-four isolates were examined for their ability to produce cellulase, pectinase, chitinase, and protease. Among these, 15 isolates exhibited enzymatic potential (Table 3). Cellulase enzymes degrade cellulose to disaccharides, while endoglucanases cut the cellulose chain in a random fashion and exoglucanases successively remove single cellobiose or glucose units from the nonreducing end of the cellulose chain [10]. In this study Aspergillus candidus, Aureobasidium pullulans, Ulocladium chartarum, Cladosporium cladosporioides, A. ustus, and Penicillium sp., respectively, exhibited high ability to produce cellulolytic activity based on the zone area. Similar results showed that Penicillium sp. produced high activity of cellulose [48]. On the other hand, enzyme extract obtained from A. niger and A. terreus was found to be rich in β-glucosidase [49]. Aspergillus candidus was prepared for quantitative estimation of cellulase; it showed significant reduction in cellulase activity with time of incubation period (Figure 2(a)). Ancient Yemeni mummies are 2000 years old, and their mummification procedure contains many botanical materials. Plenty of species able to produce cellulase suggest that this enzyme is the main cause of degradation of linen bandages of mummies and other materials.

(a)

(b)

(c)

(d)

(e)

Figure 2

Levels of specific activity (SA) (U/mg protein) during growth incubation at 28°C (left -axis) for (a) cellulase from Aspergillus candidus, (b) chitinase from Aspergillus niger, (c) protease from Cladosporium herbarum, (d) pectin lyase (PL) from Trichoderma hamatum, (e) pectin methyl esterase (PME) from Trichoderma hamatum. Right -axis indicates the effect of incubation period on dry weight (DW) of each fungus growth. Data presented in these graphs are mean ± standard division for double measures of three separate repeats.

Pectinolytic activity is one of the most important extracellular enzymes of fungi which hydrolyzes pectic substance commonly present in plant cell walls [50]. Our results revealed that, among the tested fungi, Penicillium echinulatum and Trichoderma hamatum showed the highest pectolytic activity for both pectinases enzymes, pectin methyl esterase (PME) and pectin lyase (PL). These results are in agreement with many studies that have shown Trichoderma sp. [51] and Pencillium sp. to be the most common pectinase producers [50]. In contrast, Aspergillus sp. showed the highest pectinolytic activity [52] followed by Scopulariopsis koningii and Mucor circinelloides. Activities of PL and PME from Trichoderma hamatum did not show any difference in the specific activity of both enzymes in the second week of incubation (Figures 2(b) and 2(c)), but in the third week both showed significant elevation (Table 4). This suggests that pectinase enzymes are involved in fungi metabolism and may help in its growth adaption. These enzymes play an important role in deteriorating linen bandages as well as the mummy.

Results of our study show that A. niger and P. citrinum were able to produce chitinase enzyme. These results are similar to high chitinolytic activity of A. niger [53]. However, Trichoderma viride showed the maximum chitinolytic activity from eight fungal species belonging to 5 genera isolated from agricultural soil sample [54]. The activity of chitinase produced by A. niger showed acute reduction in the second week of incubation time when compared to the first week (Figure 2(b)). This suggests that chitinase enzyme of A. niger isolated from mummy skin is not involved in the growth adaptation of the isolates.

Proteolytic enzymes are a large group of enzymes that cleave the peptide bonds of proteins to small fragments and amino acids. Fungi cause proteolysis of collagen which is dependent on many factors such as storage and environmental conditions, in addition to presence of certain substances that reside on the mummy’s skin [10]. In our results, only one fungal isolate was able to produce proteolytic activity (Table 2). This isolate was C. herbarium. The proteolytic activity was found to reduce after 2 weeks of incubation whereas, after 3 weeks, it was elevated by 2-fold (Figure 2(c)). In contrast, the fungi from different source than mummy showed higher proteolytic activity. 72.2% of 54 fungal isolates showed proteolytic activity [55].

Identifying the fungal species and the factors enhancing their growth is an essential step for the success in establishing a proper and effective strategy for preservation of the collection of mummies in museums. The fungal problem should be addressed either by controlling physical conditions of the surrounding environment in which they are stored/displayed or by the treatment with biocides. Control of the environmental conditions is the best means of protecting mummies.

4. Conclusion

It can be concluded that the higher degree of contamination of the ancient Yemeni mummies preserved in the National Museum of Yemen may result from the poor level of preservation conditions. Also the higher activity of hydrolytic enzymes including cellulase, PL, PME, chitinase, and protease from the isolated fungi species represents the major threat that can cause biodeterioration of the ancient Yemeni mummies. Therefore, it is recommended to improve the preservation systems of the mummies at the National Museum to minimize the contamination to the lowest level, which in turn will protect the mummies from biodeterioration.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.

References

A. Arya, A. R. Shah, and S. Sadasivan, “Indoor aeromycoflora of Baroda museum and deterioration of Egyptian mummy,” Current Science, vol. 81, no. 7, pp. 793–799, 2001.
View at: Google Scholar
A. E. David, “Conservation of mummified Egyptian remains,” in Science in Egyptology: Proceedings of the Science in Egyptology Symposia, A. R. David, Ed., Manchester University Press, Manchester, UK, 1986.
View at: Google Scholar
M. L. E. Florian, “Conidial fungi (mould, mildew) biology: a basis for logical prevention, eradication and treatment for museum and archival collections,” Leather Conservation News, vol. 10, no. 1, pp. 1–29, 1994.
View at: Google Scholar
O. Abdel-Kareem, J. Szostak-Kotowa, W. Barabasz, I. Paśmionka, and A. Galus, “Fungal biodeterioration of ancient Egyptian textiles, part I: survaying study for the most dominant fungi on ancient Egyptian textiles,” in Drobnousreoje W Środowisku Występowanie, Aktywność i Znaczenie, Wyd, pp. 279–290, AR Kraków, 1997.
View at: Google Scholar
W. P. Lull and P. N. Banks, Conservation Environment Guidelines for Libraries and Archives, vol. 102, Canadian Council of Archives (CCA), Ottawa, Ontario, Canada, 1995.
S. Ogden, “Temperature, relative humidity, light and air quality: basic guidelines for preservation,” in Preservation of Library and Archival Materials: A Manual, S. Ogden, Ed., p. 412, Northeast Document Conservation Center (NEDCC), Andover, Mass, USA, 3rd edition, 1999.
View at: Google Scholar
M. L. Florian, Fungal Facts: Solving Fungal Problems in Heritage Collections, Archetype Publications, London, UK, 2002.
M. R. Giuliani and M. P. Nugari, “A case of fungal biodeterioration on an ancient textile,” in Proceedings of the 9th Triennial Meeting of the ICOM Committee for Conservation, pp. 305–307, Washington, DC, USA, 1993.
View at: Google Scholar
M. L. Florian, Fungal Facts, Solving Fungal Problems in Heritage Collections, Archetype Publications, London, UK, 2004.
R. Martínez, F. Hernández, B. Millán, P. Manzano, and J. Méndez, “Effectiveness of imazalil to control the effect of fungal deterioration on mummies at the Mexico City Museum ‘El Carmen’,” Revista Iberoamericana de Micología, vol. 24, no. 4, pp. 283–288, 2007.
View at: Publisher Site | Google Scholar
A. Elnaggar, A. Sahab, S. Ismail, G. Mahgoub, and M. Abdelhady, “Microbial study of Egyptian mummies: an assessment of enzyme activity, fungicides and some mummification materials for the inhibition of microbial deterioration,” e-Conservation Magazine, no. 16, pp. 39–49, 2010.
View at: Google Scholar
N. Valentin, “Microorganisms in museum collections,” Coalition, vol. 19, pp. 1579–8410, 2010.
View at: Google Scholar
R. Kowalik, “Microbiodeterioration of library materials. Part 2. Microbiodecomposition of basic organic library materials,” Restaurator, vol. 4, no. 3-4, pp. 135–219, 1980.
View at: Google Scholar
I. Weitzman and R. C. Summerbell, “The dermatophytes,” Clinical Microbiology Reviews, vol. 8, no. 2, pp. 240–259, 1995.
View at: Google Scholar
R. J. Koestler, “Polymers and resins as food for microbes,” in Of Microbes and Art: The Role of Microbial Communities in the Degradation and Protection of Cultural Heritage, O. Ciferri, P. Tiano, and G. Mastromei, Eds., p. 250, Kluwer Academic/Plenum Publisher Company, New York, NY, USA, 2000, (International Conference on Microbiology and Conservation (ICMC), Florence, Italy, pp. 153–167).
View at: Google Scholar
M. K. Bhat and S. Bhat, “Cellulose degrading enzymes and their potential industrial applications,” Biotechnology Advances, vol. 15, no. 3-4, pp. 583–620, 1997.
View at: Publisher Site | Google Scholar
D. R. Kashyap, P. K. Vohra, S. Chopra, and R. Tewari, “Applications of pectinases in the commercial sector: a review,” Bioresource Technology, vol. 77, no. 3, pp. 215–227, 2001.
View at: Publisher Site | Google Scholar
S. N. Gummadi and T. Panda, “Purification and biochemical properties of microbial pectinases—a review,” Process Biochemistry, vol. 38, no. 7, pp. 987–996, 2003.
View at: Publisher Site | Google Scholar
D. Silva, K. Tokuioshi, E. da Silva Martins, R. da Silva, and E. Gomes, “Production of pectinase by solid-state fermentation with Penicillium viridicatum RFC3,” Process Biochemistry, vol. 40, no. 8, pp. 2885–2889, 2005.
View at: Publisher Site | Google Scholar
P. E. Yuli, M. T. Suhartono, Y. Rukayadi, J. K. Hwang, and Y. R. Pyun, “Characteristics of thermostable chitinase enzymes from the indonesian Bacillus sp.13.26,” Enzyme and Microbial Technology, vol. 35, no. 2-3, pp. 147–153, 2004.
View at: Publisher Site | Google Scholar
R. A. Lindberg, W. G. Rhodes, L. D. Eirich, and H. Drucker, “Extracellular acid proteases from Neurospora crassa,” Journal of Bacteriology, vol. 150, no. 3, pp. 1103–1108, 1982.
View at: Google Scholar
M. B. Rao, A. M. Tanksale, M. S. Ghatge, and V. V. Deshpande, “Molecular and biotechnological aspects of microbial proteases,” Microbiology and Molecular Biology Reviews, vol. 62, no. 3, pp. 597–635, 1998.
View at: Google Scholar
W. D. Glanzman, “Intentionally mummified remains of the pre-Islamic period in South Arabia,” Canadian Association for Physical Anthropology Newsletter, no. 1, pp. 2–7, 2005.
View at: Google Scholar
NEWSBRIEFS, “Mummified bodies found in North Yemen,” Archaeology, vol. 37, p. 83, 1984.
View at: Google Scholar
Y. M. Abdullah, Papers in History and Archaeology of Yemen, 1986, (Arabic).
R. Boano, E. Fulcheri, R. Grilletto, E. Leospo, and E. Rabino Massa, “Histological analysis and staining techniques modified and verified on ancient mummified tissues to study microorganism infestations,” Bollettino Della Societa Italiana di Biologia Sperimentale, vol. 75, no. 7-8, pp. 39–45, 1999.
View at: Google Scholar
K. B. Raper and D. J. Fennell, The Genus Aspergillus, Williams and Wilkins, Baltimore, Md, USA, 1965.
M. B. Ellis, Dematiacious Hypomycetes, Commonwealth Mycological Institute, Kew, UK, 1971.
H. Barnett and B. Hunter, Illustrated Genera of Imperfect Fungi, The American Phytopathological Society, APS, St. Paul, Minn, USA, 4th edition, 1999.
R. M. Teather and P. J. Wood, “Use of Congo red-polysaccharide interactions in enumeration and characterization of cellulolytic bacteria from the bovine rumen,” Applied and Environmental Microbiology, vol. 43, no. 4, pp. 777–780, 1982.
View at: Google Scholar
L. Hankin, M. Zucker, and D. C. Sands, “Improved solid medium for the detection and enumeration of pectolytic bacteria,” Applied Microbiology, vol. 22, no. 2, pp. 205–209, 1971.
View at: Google Scholar
S. C. Hsu and J. L. Lockwood, “Powdered chitin agar as a selective medium for enumeration of actinomycetes in water and soil,” American Society for Microbiology, vol. 29, no. 3, pp. 422–426, 1975.
View at: Google Scholar
R. R. Paterson and P. D. Bridge, Biochemical Techniques for Filamentous Fungi, International Mycological Institute, CAB International, Surrey, UK, 1994.
S. B. Oyeleke, E. C. Egwim, and S. H. Auta, “Screening of Aspergillus flavus and Aspergillus fumigatus strains for extracellular protease enzyme production,” Journal of Microbiology and Antimicrobials, vol. 2, no. 7, pp. 83–87, 2010.
View at: Google Scholar
M. Ghraib, K. A. Youssef, and M. M. Nour EL-Deen, “Biodegradation of some cellulose wastes,” Faculty of Education, Ain Shams University, vol. 16, pp. 87–105, 1992.
View at: Google Scholar
H. O. Eggins and G. J. Pugh, “Isolation of cellulose-decomposing fungi from the soil,” Nature, vol. 193, no. 4810, pp. 94–95, 1962.
View at: Publisher Site | Google Scholar
O. A. G. Ali, “Formation of protease by Aspergillus fumigatus and Pencillium sp,” King Saud University–Science, vol. 4, no. 2, pp. 127–136, 1992.
View at: Google Scholar
G. L. Miller, “Use of dinitrosalicylic acid reagent for determination of reducing sugar,” Analytical Chemistry, vol. 31, no. 3, pp. 426–428, 1959.
View at: Publisher Site | Google Scholar
P. Bernfeld, “Enzymes of starch degradation and synthesis,” Advances in Enzymology and Related Subjects of Biochemistry, vol. 12, pp. 379–428, 1951.
View at: Google Scholar
R. T. Sherwood, “Pectin lyase and polygalacturonase production by Rhizoctonia solani and other fungi,” Phytopathology Journal, vol. 56, pp. 279–286, 1966.
View at: Google Scholar
Z. I. Kertesz, The Pectic Substances, Interscience, New York, NY, USA, 1951.
P. O. Olutiola and O. A. Akintund, “Pectin lyase and methylesterase production by Penicillium citrinum,” Transactions of the British Mycological Society, vol. 72, pp. 49–55, 1979.
View at: Publisher Site | Google Scholar
J. Monreal and E. Reese, “The chitinase of Serratia marcescens,” Canadian Journal of Microbiology, vol. 15, no. 7, pp. 689–696, 1969.
View at: Publisher Site | Google Scholar
O. Folin and V. Ciocalteu, “On tyrosine and tryptophane determinations in proteins,” The Journal of Biological Chemistry, vol. 73, no. 2, pp. 627–650, 1927.
View at: Google Scholar
O. Lowry, N. J. Rosebrouph, A. L. Farr, and R. J. Randall, “Protein measurement with the Folin phenol reagent,” The Journal of Biological Chemistry, vol. 193, no. 1, pp. 265–275, 1951.
View at: Google Scholar
F. Rojo-Pérez, Riesgos Ocupacionales por Manejo de Restos Momificados. Tesis de Licenciatura en Antropología Física, Escuela Nacional del Antropología e Historia, UNAM, Mexico City, Mexico, 1998.
M. A. Shabbir, A. M. Khan, M. Khan, M. Iqbal, and F. Ahmad, “Fungal biodeterioration: a case study in the Zoological museum of the Punjab University,” Journal of Animal and Plant Sciences, vol. 17, pp. 3–7, 2007.
View at: Google Scholar
P. F. Hamlyn, Fungal Biotechnology, British Mycological Society, Newsletter, 1998.
L. V. Ray, A. W. Pal, A. K. Ghosh, and P. C. Chattodyay, “Cellulase and $β$ -glocosidase from Aspergillus niger and sacchrification of some cellulose wastes,” Journal of Microbiology and Biotechnology, vol. 8, pp. 85–94, 1993.
View at: Google Scholar
D. Silva, E. da Silva Martins, R. da Silva, and E. Gomes, “Pectinase production by Penicillium viridicatum RFC3 by solid state fermentation using agricultural wastes and agro-industrial by-products,” Brazilian Journal of Microbiology, vol. 33, no. 4, pp. 318–324, 2002.
View at: Google Scholar
E. S. Martins, D. Silva, R. Da Silva, and E. Gomes, “Solid state production of thermostable pectinases from thermophilic Thermoascus aurantiacus,” Process Biochemistry, vol. 37, no. 9, pp. 949–954, 2002.
View at: Publisher Site | Google Scholar
S. R. Patil and A. Dayanand, “Optimization of process for the production of fungal pectinases from deseeded sunflower head in submerged and solid-state conditions,” Bioresource Technology, vol. 97, no. 18, pp. 2340–2344, 2006.
View at: Publisher Site | Google Scholar
E. F. Sharaf, “A potent chitinolytic activity of Alternaria alternata isolated from Egyptian black sand,” Polish Journal of Microbiology, vol. 54, no. 2, pp. 145–151, 2005.
View at: Google Scholar
E. F. Sharaf, A. Q. El-Sarrany, and E. M. El-Deeb, “Biorecycling of shrimp shell by Trichoderma viride for production of antifungal chitinase,” African Journal of Microbiology Research, vol. 6, no. 21, pp. 4538–4545, 2012.
View at: Publisher Site | Google Scholar
A. A. Mohammed and N. A. Hussein, “Proteolytic and lipolytic activity of fungi isolated from luncheon meat and poultry in Assiut city,” Assiut Veterinary Medical Journal, vol. 50, no. 100, pp. 100–113, 2004.
View at: Google Scholar

Copyright

Copyright © 2014 Khalid Mohammed Naji et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

PDF Download Citation

Download other formats

Order printed copies

Views

2695

Downloads

1790

Citations