ANTIBACTERIAL ACTIVITIES OF DIFFERENT … of S. aureus colonies included test for catalase, slide coagulase and DNase (Koneman et al., 1997), as well as Api-Staph strip system (API

Egypt. J. Exp. Biol. (Bot.), 10(1): 75 – 85 (2014) © The Egyptian Society of Experimental Biology

ISSN: 1687-7497 On Line ISSN: 2090 - 0503 http://www.egyseb.org

R E S E A R C H A R T I C L E

Mosta fa M. E l-Sheekh Shymaa M. E l-Shafaay At ef M. Abo u-Shad y Enas M. E l-Bal lat

ANTIBACTERIAL ACTIVIT IES OF DIFFERENT EXTRACTS OF SOME FRESH AND MARINE ALGAE

ABSTRACT: The in v it ro antimicrobial act ivity of algal extracts belonging to two species of microalgae; Chlorella vulgaris, Spirulina p latensis and two species of seaweeds; Sargassum vulgare and Sargassum wight ii were tested against methici ll in-resistant Staphylococcus aureus (MRSA) and methicil l in-sensit ive S. aureus (MSSA) clinical isolates. In this study, microt iter plate reader assay method was used to elucidate the effects. The ext racts were prepared in four different solvents, out of which methanol extracts showed better and promising results. The results also confirmed the potential use of algal extracts as a source of antimicrobial compounds.

KEY WORDS: Antimicrobial activi ty, Algae, Bacteria, Microt iter plate assay

CORRESPONDENCE: Mostafa Moham ed El-Sheekh Botant Department, Faculty of Science, Tanta Universi ty, Egypt. E-mail: [email protected]

Shymaa Moham ed E l-Shafaay At ef Mohamed Abo u-S hady Enas Mosta fa E l-Ba lla t Botany Department, Faculty of Science, Tanta University, Egypt ARTICLE CODE: 07.02.14

INTRODUCTION: Bacterial resistance has been the main

factor responsible for the increase of morbidity, mortality and health care costs of bacterial infections. The defense mechanism against antibiotics is widely present in bacteria and became a world health problem. The increasing prevalence of multidrug resistance strains of bacteria and the recent appearance of strains with reduced susceptibi lity to antibiotics raises the specter of untreatable bacterial infections and adds urgency to the search for new infection-fight ing strategies to control microbial infections (Mala et al. , 2009).

It is not only the resistance but also the cost of synthet ic chemicals lead to search for al ternate medicine such as antimicrobial compounds from natural sources. Algae derived natural products and antibiotics are found to be the effective alternative recognized from natural environmental resources. The f i rst invest igation on antibiotic activity of algae was carried out by Pratt et al. (1944). One of the potential groups of natural resource i s algae which are known to possess promising novel bioactive substances (Vadas, 1979; Metzger et al., 2002). Since algae have been used in traditional medicine for a long time and also some algae have bacteriostatic, bactericidal , antifungal, antivi ral and antitumor act ivity, they have been extensively studied by several researchers (Justo et al. , 2001). Many investigators have reported antibacterial activit ies of microalgae as due to fatty acids (Cooper et. al. , 1983; Findlay and Patil, 1984). Compounds with cytostatic, antiviral, anthelmintic, antifungal, and antibacterial activities have been detected in green, brown and red algae (Lindequist and Schweder, 2001; Newman et al., 2003).

Cyanobacteria are potential sources of high value chemicals and pharmaceuticals. Spirulina i s one of the most potential cyanobacteria used in medicine. Spirulina i s safe for human consumption as medicine, because i t is free of microcystin toxin and the long term dietary supplementation of up to 5%

Egypt. J. Exp. Biol. (Bot.), 10(1): 75 – 85 (2014)


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of the Spiru lina may be consumed without evident toxic side effects (Yang et al. , 2011). It is known to produce intracel lular and extracellular metabol ites with diverse biological activit ies such as antifungal (Mac Millan et al. , 2002), antiviral (Hayashi et al. , 1996), and antibacterial activit ies (Kaushik and Chauhan, 2008).

The ability of seaweeds to produce secondary metabolites of potential interest has been extensively documented (Faulkner, 1993). There are numerous reports of compounds derived from macroalgae with a broad range of biological activities, such as antibiotics (antibacterial and ant ifungal properties), antiviral diseases (Trono, 1999), anti tumor and anti-inflammatory (Scheuer, 1990), as well as neurotoxins (Kobashi, 1989). Chemical structure types include sterols (Ahmad et al. , 1993), i soprenoids amino acids, terpenoids, phlorotannins, steroids, phenolic compounds, halogenated ketones and alkanes, cyclic polysulphides, fatty acids and acrylic acid can be counted (Mtolera and Semesi, 1996). The aim of thi s research was to study the antibacterial activi ties of dif ferent extracts of marine macroalgae and fresh water microalgae.

MATERIAL AND METHODS: Organisms and culture conditions: Microalgae:

The two microalgal strains were obtained from the culture collection of the Botany Department, Faculty of Science, Mansoura University, Egypt. Zarrouk's (1966) and Kuhl 's (1962) media were used for cul tivation of S. platensis and C. vulgaris, respectively. The culture flasks were aerated with air mixed with 3% CO2 to accelerate algal growth and incubated at 30ºC under continuous illumination provided from day light fluorescent tubes giving l ight intensity of 80 µEm - 2s- 1 for C. vulgaris and 2500 Lux for S. platensis. Before bubbl ing into the culture, the mixture was allowed to pass through bacterial filter (0.2 µm diameter). S. platensis was grown until the late exponential phase of the growth (the 10 th day), while C. vulgaris was grown and harvested til l (the 12 th day) and the cultures were harvested by centrifugation at 3500 rpm for 15 minutes. The pellet was rinsed three t imes and resuspended in sterilized distil led water to remove traces of growth medium. The col lected biomass was dried in an oven at (40-60ºC) and then powdered by manual morter. Macroalgae (seaweeds):

In this study, two species of seaweeds were collected from the rocky areas in few meters below the water surface in Red sea, Seuz beach, Egypt during November 2012. The seaweeds were brought to the laboratory

in plastic bags containing sea water to prevent evaporation. Algae were then cleaned from epiphytes and rock debris and given a quick fresh water rinse to remove surface salts. The seaweeds were identi fied following Abbott and Hollenberg (1976), Aleem (1993), and Taylor (1960). The samples were air dried in the shade at room temperature 25-30ºC on absorbent paper, cut into small pieces and grounded to fine powder. These powdered samples were stored in light plastic bags. Tested bacteria:

The samples were collected from di fferent patients cl inically diagnosed to have bacterial infection (impetigo). This was achieved by visi ting the outpat ient cli nic of Dermatology Department at Tanta University Hospital. The samples included different isolates of Staphylococcus aureus. The best purifi ed isolates were used in the study. The patient samples were transferred in 2 ml phosphate–buffered saline (PBS; NaCl, 8 g/L; KCl, 0.2 g/L; Na2HPO4, 1.15 g/L; KH2PO4, 0.2 g/L) and were forwarded to the Bacteriology Laboratory in Botany Department, Faculty of Science, Tanta University. All cul ture swabs were processed in the same day that they were collected. Each specimen was plated to a mannitol salt agar media. Culture plates were incubated up to 24 h at 37°C, and then examined for colony morphology consistent with S. aureus. Identification of S. aureus colonies included test for catalase, slide coagulase and DNase (Koneman et al. , 1997), as well as Api-Staph strip system (API System S.A., Montalïeu-Vercieu, France).

A positive bacterial growth (basel ine) on nutrient agar, the diagnosis of skin lesions in clinical samples, was based on the presence of > 105 colony forming unit (CFU) of bacteria/ml in the culture (Wil liams et al. , 1990). Methods of determination of the physiological and biochemical characteristics of bacterial isolates: A. Growth on Mannitol Salt Agar (MSA):

This type of medium is both select ive and differential. The MSA is be selected for organisms such as Staphylococcus species which can l ive in high salt concentration. The di fferential ingredient in MSA is the sugar mannitol. Organisms which are capable of using mannitol as a food source wi ll produce acidic byproducts of fermentation that wi ll lower the pH of the media. The acidi ty of the media will cause the pH indicator, phenol red, to turn yel low. Staphylococcus aureus is capable of fermenting mannitol and covert the color of the media from pink into yellow. B. Catalase test:

A few drops of 3% (v/v) hydrogen peroxide were added to the bacterial growth appeared on the surface of nutrient agar

E l- Sh e ek h et a l . , A nt ib ac t e r i a l A c t i v i t i es o f D i f f e r en t E xt r act s o f S om e Fr es h an d M ar i ne A lga e

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plates. Gas bubbles either instantly or after 5 minutes if the bacteria were catalase positive. C. Deoxyribonuclease (DNase) test:

DNase agar (oxoid) was inoculated by the i solated bacteria for 2 days. At the end of incubation period, the plates were treated with 1% HCl. The DNA molecules hydrolyzed to a mixture of mono and polynucleotide by the action of enzyme DNase produced by the organism. HCl reacted with the nucleotides in the medium forming a cloudy precipi tate and a clear area contained the nucleotide fractions which were not precipi tated by the acid. D. Coagulase test:

Coagulase is a protein enzyme produced by several microorganisms that enables the conversion of fibrinogen to fibrin. In laboratory, it is used to distinguish between different types of Staphylococcus isolates. Antibiotic susceptibility of the tested isolates:

The susceptibi lity of the tested S. aureus isolates to 9 antimicrobial agents was performed by modified Kirby-Bauer single-disk diffusion technique on Muller Hinton agar (Robert et al. , 2003). The used ant imicrobials were amoxicil lin, oxacillin, imipenem, gentamicin, chloramphenicol , vancomycin, deoxycycline, clindamycin and nitrofurantoin. The results of the susceptibili ty tests were interpreted according to the criteria establ ished by the Clinical and Laboratory Standards Institute (CLSI, 2010). Methicill in-resistance was tested by using 1 mg oxacill in disk (Oxoid). Determination of the suitable extraction time of antimicrobial material:

A sample of Chlorella vulgaris was extracted by soaking in 70% methanol as an example for solvents in a conical flask at room temperature 25°C-30°C for different periods 24h, 48h, 72h and 96h (1:15 v/v) (Rosell and Srivastava, 1987). The extracts were filtered and evaporated under reduced pressure. The residues were concentrated to constant concentration and the antimicrobial activi ty against Staphylococcus aureus was assayed using hol low well technique by making five well s of 7 mm diameter cut with steri lized cork borer and 50 µL of algal extract and methanol as control . The diameter of inhibit ion zones was calculated at the end of incubation period and the results were compared with each other and the best extraction time was used in our study. Determination of the best algal extract with the best solvent using microtiter plates assay method:

The extraction was carried out with different solvents (i.e., 70% ethanol, 70% methanol, 70% ethyl acetate and 70% chloroform) by soaking all algal samples in the respective solvents. Ant ibacterial activi ty assay of the algal extracts was conducted

using microti ter plates reader assay method according to Bechert et al. (2000) with some modifications. Aliquot of 100 µl of bacterial isolate (106 CFU/ml) in Muller Hinton Broth media was transferred to each well of 96 well plates, 50 µl of extracts were added to each well in repl ica. The plates were incubated under microaerophi lic conditions at 37ºC for 24h. After incubation, the absorbance of the samples was determined using automated ELIZA microplate reader adjusted at 630 nm. The inhibi tion percentage of algal extracts was calculated according to the following equation (Mulyono et al. , 2012). The test was carried out in triplicate (in same 96-wel l plate) and repeated twice for each strain and each tested agent.

Inhibition percentage (%) = 100 – (A bacteria + NB + extract – A NB + extract) / (A bacteria

+ NB + solvent - A NB + solvent) x 100 Where: *A= Absorbance; *NB= Nutrient Broth media; Treatment = Abac te r ia + NB + ext rac t ; (-Ve) control = Ab ac te r ia + NB + solven t; Blank' = ANB + ex trac t ; Blank'' = ANB + sol ven t

Fractionation and characterization of antimicrobial crude extract using column chromatography:

Selected active crude extracts (4g) were fractionated by column chromatography on silica gel G (EDWC, 60-120 mesh). Column (2 cm x 40 cm) was set up in chloroform with silica gel (30-40 g), eluted with gradients of solvents from 7: 3% of chloroform: methanol to 3: 7% chloroform: methanol and the collected fractions were evaporated to dryness with the rotary evaporator. The dried samples were dissolved in pure methanol and assayed for their antimicrobial activit y. The maximum absorpt ion of the active fractions was measured by spectrophotometer (UV 2101/ pc) using quartz cuvette containing the di fferent fractions. After detection of the material which has antagonistic activity against bacteria, it is lyophilized and was subjected to the following analysis in order to reveal its structure as far as possible. Agar diffusion method for determining the minimum inhibitory concentration (MIC) for bacteria:

MIC was determined by agar method (Chattopadhyay et al. , 1998). Serial dilutions (200, 100, 50, 25, 12.5, 6.25, 3, and 1 mg/ml) of the antimicrobial material were tested against the most resistant pathogenic strain of Staphylococcus aureus isolate with concentration (108 CFU). The inoculated plates were then incubated at 37°C for 24h. The lowest concentration which did not show any visible growth was considered as the MIC. Statistical analysis:

Results are presented as mean ± SD (standard deviat ion) for three replicates. The



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stat ist ical analysis were carried out using SAS and SYSTAT statistical software packages, the degree of signifi cance of the obtained results was tested. One and three way analyses of variance were carried out.

RESULTS: Determination of the suitable extraction time of antimicrobial material:

This experiment was carried out in order to determine the sui table extraction t ime of the antimicrobial material using Chlorella vulgar is as an example for algae extracted by soaking in 70% methanol at room temperature. The antimicrobial activity against a bacterial strain (Staphylococcus aureus) was determined as a diameter of inhibition zone using agar di ffusion method. The obtained results indicated that the inhibition zone of Staphylococcus aureus at dif ferent extraction times (24h, 48h, 72h, and 96h) were 1.5, 1.7, 1.9, and 1.9 mm, respectively indicating that the 72h ext raction time would be sufficient to extract the antimicrobial substance (Fig. 1). Therefore, we used it in the rest of our study.

Fig. 1. Effect of different extraction t imes at room

temperature for extracting the ant imicrobial substance from Chlorella vulgaris using 70% methanol against Staphylococcus aureus; 1 = 24h; 2 = 48h; 3 = 72h; 4 = 96h; c = Control (70% methanol).

Determination of the physiological and biochemical characteristics of bacterial isolates: Growth on Mannitol Salt Agar (MSA):

MSA was selected for organisms such as Staphylococcus species which can live in areas of high salt concentration. The differential ingredient in MSA is the sugar mannitol . Organisms capable of using mannitol as a food source would produce acidic byproducts of fermentation that would lower the pH of the media. The acidi ty of the media would cause the pH indicator, phenol red, to turn yellow. Staphylococcus aureus was capable of fermenting mannitol

and coverting the color of the media from pink into yel low. All the recovered Staphylococcus aureus isolates were shown positive results (Fig. 2).

Fig. 2. Photo 2. The effect of Staphylococcus aureus

on mannitol salt agar medium.

Catalase test: A few drops of 3% (v/v) hydrogen

peroxide were added to the bacterial growth appeared on the surface of nutrient agar plates. Gas bubbles were appeared instantly. All the Staphylococcus aureus isolates were catalase posi tive (Fig. 3).

Fig. 3. Formation of gas bubbles when drops of hydrogen peroxide were added to the Staphylococcus aureus growth appeared on the surface of nutr ient agar plates

Deoxyribonuclease (DNase) test: DNase agar was inoculated by the

isolated bacteria for 2 days. At the end of incubation period, the plates were treated with 1% HCl. The DNA molecules hydrolyzed to a mixture of mono and polynucleotide by the action of enzyme DNase produced by the Staphylococcus aureus. HCl reacted with the nucleotides in the medium forming a cloudy precipi tate and a clear area contained the nucleotide fract ions which were not



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precipitated by the acid. All the Staphylococcus aureus isolates were DNase positive (Fig. 4).

Fig. 4. Formation of a cloudy prec ipitate and a clear area contained the nucleotide fractions which were not prec ipitated by the HC l.

Coagulase test: Coagulase is a protein enzyme produced

by several microorganisms that enables the conversion of fibrinogen to fibrin. In the laboratory, it is used to distinguish between different types of Staphylococcus isolates. All the recovered Staphylococcus aureus isolates were coagulase positive (Fig. 5).

Susceptibil ity of S. aureus isolates to different antibacterial agents:

The susceptibility of S. aureus isolates to 9 different antimicrobial agents representing 9 classes was conducted using disk diffusion method. The used antimicrobial discs were as follow: Amoxicillin (AMX), Oxacillin (OX), Imipenem (IPM), Gentamicin (GEN), Chloramphenicol (CL), Vancomycin (VA), Deoxycycline (DO), Clindamycin (DA) and Nitrofurantoin (F). The sizes of the inhibition zones were interpreted by referring to Clinical and Laboratory Standards Institute standards and the results were reported as being susceptible, intermediate or resistant to the antibiotics that were tested. The obtained data showing the incidence of antimicrobial resistance among isolates are shown in table 1. Isolates

with resistant phenotypes included those that were classified as intermediate resistance. Table 1. Inc idence of antibacter ial res istance

among S. aureus isolates.

Antibacterial agent No. (%) of resistant isolates AminoPenicillins Amoxicillin (AMX) 8 (100 %) Penicillins Oxacillin (OX) 7 (87.5 %) Carbapenems Imipenem (IPM) 1 (12.5 %) Aminoglycosides Gentamicin (GEN) 5 (62.5 %) Phenicols Chloramphenicol (CL) 6 (75 %) Glycopeptides Vancomycin (VA) 5 (62.5 %) Tetracyclines Deoxycycline (DO) 4 (50 %) Lincosamides Clindamycin (DA) 8 (100 %) Nitrofurantions Nitrofurantoin (F) 4 (50 %)

As shown in table 1, the incidence of resistance to different tested antibiotics ranged between 12.5% (imipenem) and 100% (amoxicill in and clindamycin). For pencillins, the incidence of resistance to oxacillin was 87.5% but for aminopenicill in was 100%. In case of aminoglycosides, phenicols, glycopeptides, tetracyclines and nitrofurantions, there was moderate incidence of resistance; gentamicin (62.5%), chloramphenicol (75%), vancomycin (62.5%), deoxycycline (50%) and ni trofurantion (50%), respectively. The only class of antibiotics showed very low incidence of resistance with most isolates was carbapenems. Determination of the best solvent for extracting the antibacterial material from different micro and macroalgae using microtiter plates:

This experiment was carried out in order to determine the best solvent (70% ethanol, 70% methanol, 70% ethyl acetate and 70% chloroform) for extracting the antibacterial material from different tested algae (Chlorella Vulgaris, Spirulina platensis, Sargassum vulgare and Sargassum wight ii) against methicillin-resistant S. aureus (MRSA) and methicillin-sensi tive S. aureus (MSSA) clinical isolates.

Antibacterial act ivity assay of the algal extracts was conducted using microti ter plates reader assay method. The results presented in table 2 show that ethanol, methanol, ethyl acetate and chloroform extracts of different algae possessed antibacterial activity against the bacteria tested. The blind control (ethanol, methanol, ethyl acetate and chloroform) did not inhibit the tested bacteria. 70% methanol extracts showed the strongest inhibition against the tested bacteria with inhibition activity percentage* on 26.92%, followed by 70% chloroform extracts with inhibition act ivity percentage 25.54%,

Fig. 5. Clumps were not mixed uniformly into coagulase plasma, represent ing a positive slide coagulase test which is indicative of Staphylococcus aureus .



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followed by 70% ethyl acetate extracts with inhibit ion activity percentage 24.86%, whereas 70% ethanol showed the weakest inhibit ion with inhibition activity percentage 22.68% of all algae against all tested

bacteria. Therefore, we use 70% methanol as solvent for extractions. *Inhibition activity percentage = (average diameter of inhibition zone of each solvent/ average diameter of inhibition zone of all solvents) x 100

Table 2. One way analys is of variance (ANOVA) of different algal extracts against different strains of Staphylococcus aureus using different solvents

Staphylococcus aureus Algae Solvent

(70%) Isolate (E)

Isolate (C)

Isolate (D)

Isolate (H)

Isolate (F)

Isolate (C')

Isolate (A)

Isolate (R)

Ethanol 99.60 ± 0.02 99.98 ± 0.01 99.95 ± 0.01 96.83 ± 0.01 99.56 ± 0.01 99.04 ± 0.01 95.90 ± 0.02 97.08 ± 0.01

Methanol 96.17 ± 0.01 93.37 ± 0.02 95.02 ± 0.01 92.85 ± 0.02 89.71 ± 0.01 96.06 ± 0.02 92.03 ± 0.02 92.71 ± 0.02

Ethyl acetate 89.63 ± 0.02 80.05 ± 0.02 83.54 ± 0.02 50.50 ± 0.0 62.63 ± 0.02 53.82 ± 0.02 74.84 ± 0.03 95.40 ± 0.0 Spirulina platensis

Chloroform 92.71 ± 0.01 88.47 ± 0.02 75.51 ± 0.02 79.97 ± 0.02 87.56 ± 0.02 90.54 ± 0.01 81.85 ± 0.02 70.30 ± 0.01

F-Value 1.33** 0.10(ns) 1.33** 3.00* 1.33** 1.33** 0.10( ns) 2.67*

Ethanol 72.74 ± 0.01 82.78 ± 0.02 69.38 ± 0.0 99.84 ± 0.02 58.76 ± 0.0 48.62 ± 0.02 94.81 ± 0.01 89.40 ± 0.0

Methanol 99.85 ± 0.01 99.95 ± 0.01 99.94 ± 0.06 99.97 ± 0.02 99.85 ± 0.02 99.98 ± 0.01 99.99 ± 0.0 99.98 ± 0.01

Ethyl acetate 99.28 ± 0.01 99.95 ± 0.01 99.32 ± 0.02 99.81 ± 0.01 99.96 ± 0.02 99.82 ± 0.01 99.97 ± 0.02 99.96 ± 0.01 Chlorella vulgaris

Chloroform 99.32 ± 0.0 99.96 ± 0.01 99.32 ± 0.02 99.06 ± 0.01 99.42 ± 0.02 99.67 ± 0.02 99.63 ± 0.55 99.90±0.07

F-Value 2.00* 6.00* 3.00* 1.50* 3.00* 1.33** 2.67* 2.67*

Ethanol 74.81 ± 0.01 72.22 ± 0.02 97.63 ± 0.01 80.72 ±0.01 67.04 ± 0.0 54.25 ± 0.02 78.27 ± 0.01 99.80 ± 0.0

Methanol 99.96 ± 0.03 99.32 ± 0.03 94.80 ± 0.01 99.96 ± 0.02 99.96 ± 0.0 99.92 ± 0.02 99.83 ± 0.02 96.80 ± 0.0

Ethyl acetate 75.72 ± 0.01 74.16 ± 0.01 70.76 ± 0.01 99.93 ± 0.02 99.95 ± 0.05 99.54 ± 0.01 84.62 ± 0.02 99.60 ± 0.0 Sargassum vulgare

Chloroform 59.71 ± 0.0 83.40 ± 0.01 94.06 ± 0.01 99.01 ± 0.01 98.86 ± 0.02 99.03 ± 0.0 99.07 ± 0.02 99.13 ± 0.01

F-Value 2.67* 1.50* 1.33** 2.67* 4.00* 2.68* 1.33** 1.33**

Ethanol 85.46 ± 0.01 42.45 ± 0.01 50.00 ± 0.0 91.36 ± 0.03 87.83 ± 0.06 97.75 ± 0.01 64.00 ± 0.0 83.20 ± 0.0

Methanol 89.22 ± 0.02 97.92 ± 0.02 99.28 ± 0.01 99.82 ± 0.02 99.60 ± 0.01 99.95 ± 0.01 99.93 ± 0.01 99.96 ± 0.01

Ethyl acetate 93.86 ± 0.01 98.37 ± 0.02 99.57 ± 0.02 99.71 ± 0.02 99.95 ± 0.02 99.94 ± 0.04 99.86 ± 0.02 99.93 ± 0.01 Sargassum wightii

Chloroform 95.34 ± 0.01 99.00 ± 0.0 98.56 ± 0.01 95.59 ± 0.01 95.42 ± 0.02 95.27 ± 0.01 96.96 ± 0.01 92.28 ± 0.02

F-Value 1.33** 2.67* 3.00* 1.50* 1.50* 1.50* 2.00* 3.00*

(±) standard deviation of the means (n=3); * Signif icant at P ≤ 0. 01,** Signif icant at P ≤ 0. 001, and (ns) Non-s ignif icant at P ≥ 0.01

One way ANOVA analysis confirmed that the variation in antibacterial activit ies in relat ion to algae and di fferent bacterial isolates were significant at p ≤ 0. 01 for almost treatments except Spirulina p latensis against isolates (E, D, F, and C'), Chlorella vulgar is against isolate (C'), Sargassum vulgare against isolates (D, A, and R) and Sargassum wight ii against isolate (E) were significant at P ≤ 0. 001. On the other hand, Spiru lina p latensis against isolates (C and A) were non-signif icant at p ≥ 0.01 (Table 2). However three-way ANOVA analysis confirmed that the variation in the antibacterial activity in relat ion to algae, bacteria, solvents and their interactions on antibacterial activity were significant at p ≤ 0.0001 for al l treatments (Table 3). Table 3. Three-way analys is of variance (ANOVA)

of different algal extracts against different strains of Staphylococcus aureus using different solvents

Source DF F-Value P-Value R2 Algae 3 99999.99 0.0001 Solvent 3 99999.99 0.0001 Bacteria 7 88665.46 0.0001 Algae*Solvent 9 99999.99 0.0001 Algae*Bacteria 21 99999.99 0.0001 Solvent*Bacteria 21 80146.77 0.0001 Algae*Solvent*Bacteria 63 99999.99 0.0001

99.9%

Fractionation and characterization of the antimicrobial crude extract using column chromatography:

The ant ibacterial material i solated from Chlore lla vulgar is was transferred to a column containing Sil ica gel (EDWC, 60-120 mesh). The material was eluted using gradients of solvents from 7:3% of chloroform: methanol to 3:7% chloroform: methanol. The different fractions were collected every 5 min and examined for antimicrobial activity using agar di ffusion assay method. The results showed that 22 fract ions were collected and only 3 fractions had ant imicrobial activity. UV absorption spectrum of these fractions was determined using spectrophotometer (UV 2101/ pc) at range of 250-800 nm. The results showed that the 3 fractions had the same absorption peaks (2 peaks at 410 and 664 nm) (Fig. 6). The different pigments and impuriti es were removed by filtration using charcoal. The UV spectrum of the puri fied antimicrobial material was then carried out in pure methanol. This spectrum shows one absorption peak at 410 nm, indicat ing the presence of an aromatic compound (Fig. 7). The obtained compound was examined for antimicrobial act ivity. The results showed that the compound sti ll has antimicrobial activity (Fig. 8), indicat ing that the compound having



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a peak at 664 nm does not have any antimicrobial activity.

Fig. 6. UV spectrophotometer scanning of dif ferent active f ract ions separated by column chromatography; A=fraction 1, B=f raction 2, C=f raction 3. *Abs. =Absorption

Fig. 7. UV spectrum of the ant imicrobial material isolated from Chlorella vulgaris af ter

purif ication by charcoal. *Abs. =Absorption

Fig. 8. The antimicrobial activity of the act ive

fraction before and after purif ication by charcoal; A=active fraction after purif ication, B=act ive fraction before purif ication, and C=control (Methanol).

Determination of minimum inhibitory concentration (MIC) of the purified antagonistic antimicrobial material extracted from Chlorella vulgaris:

MIC was determined by agar diffusion methods. Serial dilutions (200, 100, 50, 25, 12.5, 6.25, 3, and 1 mg/ml) of the antimicrobial material extracted from Chlorella vulgar is were tested against the most resistant isolate of bacteria (Staphylococcus aureus) as Gram-positive bacteria. The concentration of the tested Staphylococcus aureus was 108 CFU. The plates were inoculated and incubated at 37°C for 24h. The lowest concentrat ion of the antimicrobial material, which did not show any visible growth, was considered as the MIC. The results presented in (Fig. 9), showed the MIC value for Staphylococcus aureus is (6.25 mg/ml).

Fig. 9. Minimum inhibitory concentration (MIC) of the

purif ied antagonistic material extracted from Chlorella vulgaris extracted by 70% methanol: A) 200mg/ml; B) 100mg/ml; C) 50mg/ml; D) 25mg/ml; E) 12.5 mg/ml; F) 6.25 mg/ml; G) 3 mg/ml; H) 1 mg/ml.

B

A C



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DISCUSSION: This study depended on the phenotypic

methods commonly used in the identification of Staphylococcus aureus. Firstly, mannitol sal t agar (MSA) was used for identi fication of S. aureus clinical samples. Mannitol Salt Agar (MSA) is both a selecti ve and differential medium used in the isolat ion of staphylococci. Early work by Gordon indicated that the fermentation of mannitol could be used as a means of differentiating pathogenic staphylococci from nonpathogenic staphylococci (Gordon, 1904). Although his work was confi rmed and expanded upon by several i nvestigators (Dudgeon, 1908), mannitol fermentation was not used routinely as an ident ificat ion tool .

The results showed that al l the isolates are positive reaction for manitol salt agar. This is because Staphylococci can withstand the osmotic pressure created by 7.5% NaCl, while this concentrat ion wil l inhibit the growth of most other gram-positive and gram-negati ve bacteria (Koch, 1942). Additionally, MSA contains mannitol and uses phenol red as a pH indicator (pH = 7.8) in the medium. When mannitol is fermented by a bacterium, acid i s produced, which lowers the pH and results in the formation of a yellow area surrounding an isolated colony on MSA.

The second test for ident ification of S. aureus was catalase test which showed that all clinical isolates are catalase posi tive. These results agree with that reported by Amini et al. (2012). Staphylococci have the abil ity to produce enzymes such as catalase considered to be virulence determinants. This enzyme allows bacteria to better resist intra- and extra-cellular ki lling by hydrogen peroxide (Grüner, 2007). Species of the genus Staphylococcus are characterized by the production of catalase. Among them, only two species, Staphylococcus saccharolyt icus and Staphylococcus aureus subsp. anaerobius , are not able to produce catalase (Kloos and Bannerman, 1999; Sanz et al. , 2000).

In order to support the previous biochemical tests, DNase tests and slide coagulase tests were carried out in our present study. The combination of all the biochemical tests increased the sensi tivity to identify the S. aureus among the bacterial isolates. DNase and tube coagulase tests were carr ied out and the results were positive for both. These results agree with that reported by Amini et al. (2012) and Koneman et al. (1997).

The posi tive results of S. aureus in case of DNase test might be due to the abi lity of these bacteria to produce DNase enzymes (deoxyribonuclease) which breaks down the DNA by adding HCl which reacts with the nucleotides in the medium forming a cloudy

precipi tate and a clear area that contain the nucleotide fract ions which were not precipi tated by the acid.

The organic extraction solvents we used were ethanol, methanol, ethyl acetate and chloroform. This is because organic solvent always provides a high efficiency in extracting compounds for antimicrobial acti vi ties compared to water-based methods (Parekh and Chanda, 2007; Axelsson and Gentil i, 2014). Lipid-soluble ext racts from fresh water microalgae and marine macroalgae have been invest igated as a source of substances with pharmacological propert ies. Moreover, several dif ferent organic solvents have been used for screening algae for antimicrobial activity. Sastry and Rao (1994) showed antibacterial activity against Gram-posit ive and Gram-negative pathogenic strains after successive extraction with benzene, chloroform, and methanol. Likewise, Mahasneh et al. (1995) showed antibiotic activity in organic extracts of six species of marine algae against mult i-antibiotic resistant bacteria.

Regarding the extraction time, Rosell and Srivastava (1987) used different lengths of time 24h, 48h and 72h for extracting the components of algae at room temperature. They found that no signif icant difference among the di fferent extraction periods. In contrast, our present study showed that 72h extract ion of antimicrobial material has been found suff icient. This might be due to that the extract ion of antimicrobial material increased with soaking t ime unti l saturation. Our results agreed with that reported by Elshobary (2010) who reported that the most active ext ract was at 72h.

Many in vit ro methods have been used to detect the antimicrobial activity of algal material. The most commonly used methods are the agar disk/wel l diffusion assay and the agar/broth dilution assay (Patton et al. , 2006; Elshobary, 2010). Studies of date have used the popular agar disk/wel l diffusion method (Almajano et al. , 2008; Gutierrez et al., 2008) despite its MIC data being potentially subjective due to the different measurement methods of the inhibiti on zone. There has been an increase in the use of the broth di lut ion assay and, in particular, the 96-well microplate assay in recent years (Eloff, 1998; Casey et al. , 2004; Turcotte et al. , 2004; Patton et al. , 2006; Rufián-Henares and Morales 2006). This assay has been identified as being more accurate than the agar/disk di ffusion, less tedious and can be automated. This becomes very attractive as the throughput of samples is high and the increased accuracy of the method al lows results to be easily compared since the MICs can be determined. On contrast, Ostrosky et al. (2008) and Chiheb et al. (2009) reported that agar dilution is the most widely used



83

method due to i ts simplicit y of implementation and low cost.

The obtained results in our study revealed that the strongest antibacterial activi ties were exhibited using 70% methanol. These results were in accordance with de Cano et al. (1990) who reported that Nostoc muscorum extract produced using methanol showed the highest antibacterial and anti fungal activit ies against two human pathogens; staphylococcus aureus and Candida albicans. Pandian et al. (2011) also tested petroleum-ether, chloroform and methanol of Acanthaphora spicifera in vit ro for thei r antibacterial and antifungal activi ty against Escherich ia coli, Bacillus subt ilis, Bacillus palmitus, Pseudomonas aeruginosa, and against Candida albicans, Microsporum gypseum, Aspergillus n iger , respect ively by disc di ffusion techniques. The methanolic extract of Acanthaphora spicifera showed

higher antibacterial and anti fungal activity compared to the other two extracts. Simic et al. (2012) demonstrated that the methanolic extract of green microalgae, Trentepohlia umbrina, has a strong antimicrobial activity against Staphylococcus aureus, Escherichia coli, Klebsie lla pneumonia , Pseudomonas aeruginosa, Enterococcus faecalis, Aspergillus n iger , Candida a lb icans, Fusarium oxysporum, Penicill ium purpurescens and Trichoderma harsianum .

Finally, we conclude that algae are potential source of bioacti ve compounds especial ly green algae (Chlorella vu lgar is) . Thus, the lowest concentrations of the algal extracts show a great effect against the most resistant isolates of Staphylococcus aureus. So, further studies can be done to evaluate the in v ivo ef fect of these extracts and introduce these extracts to the pharmaceutical uses.

REFERENCES: Abbott IA, Hollenberg IG. 1976. Marine algae of

California Stanford University press. Stanf ord, CA, pp. 844.

Ahmad VU, Aliya R, Perveen S, Shameel M. 1993. Sterols from marine green alga Codium decortacatum. Phytochemistry, 33(5): 1189-1192.

Aleem AA. 1993. The mar ine Algae of Alexandria, Faculty of science, University of Alexandria, Egypt. Privately published, pp. 139.

Almajano MP, Carbó R, Jiménez AJL, Gordon MH. 2008. Antioxidant and antimicrobial act ivit ies of tea infusions. Food Chem., 108(1): 55–63.

Amini R, Abdulamir AS, Chung C, Jahanshir i F, W ong CB, Poyling B, Hematian A, Sekawi Z, Zargar M, Jali lian F. 2012. Circulation and transmission of methic il lin-res istant Staphylococcus aureus among college students in Malays ia (cell phones as reservoir); As ian Biomed., 6(5): 659-673.

Axelsson A, Gent il i G. 2014. A s ingle-step method for rapid extraction of total l ipids f rom green microalgae. PLoS ONE 9(2): e89643.

Bechert T, Steinrücke P, Guggenbichler JP. 2000. A new method for screening ant i-inf ective biomaterials. Nat. Med., 6(9): 1053-1056.

Casey JT, O'Cleir igh C, Walsh PK, O'Shea DG. 2004. Development of a robust microplate plate-based assay method for assessment of bioactivity. J. Microbiol. Methods , 58(3): 327-334.

Chattopadhyay, D., Sinha, B. and Vaid, L.K. 1998. Antibacterial activity of Syzygium species: A report. Fitoterapia, 69(4): 365-367.

Chiheb I, Riadi H, Martinez-Lopez J, Dominguez SJF, Gomez VJA, Bouziane H, Kadir i M. 2009. Screening of antibacterial activity in mar ine green and brown macroalgae from the coast of Morocco. Af r. J. Biotechnol., 8(7): 1258–1262.

CLSI 2010. Performance standards for ant imicrobial susceptibilit y testing; twent ieth informational supplement, CLSI document M100-S20, W ayne, PA: Clinical and Laboratory Standards Institute.

Cooper S, Battat A, Marot P, Sylvester M. 1983.

Production of antibacterial act ivit ies by two bacillar iophyceae grown in dialys is culture. Can. J. Microbiol., 29(3): 338-341.

de Cano MMS, de Mulé MCr, de Caire GZ, de Halperin DR. 1990. Inhibit ion of Candida albicans and Staphylococcus aureus by phenolic compounds from the terrestr ial cyanobacterium Nostoc muscorum. J. Appl. Phycol., 2(1): 79-82.

Dudgeon LS. 1908. The differentiat ion of the staphylococc i. J. Pathol. Bacteriol., 12(2): 242-257.

Elof f JN. 1998. A sensit ive and quick microplate method to determine the minimal inhibitory concentration of plant extracts f rom bacteria. Planta Med., 64(8): 711–713.

Elshobary ME. 2010. The antimicrobial act ivit ies of some seaweed collected from Alexandria coast. M.Sc. Thesis, Tanta Univers ity, Egypt.

Faulkner DJ. 1993. Mar ine natural products. Nat. Prod. Rep., 19(1):1-48.

Findlay JA, Pat il AD. 1984. Ant ibacterial constituents of the diatom Navicula delognei. J. Nat. prod., 47(5): 815-818.

Gordon MH. 1904. Report of some characters by which various streptococci and staphylococci may be dif ferentiated and identif ied. Ann. Rep t. of the Local Govt. Board Supp., 33: 388-430.

Grüner BM, Han SR, Meyer HG, Wulf U, Bhakdi S, Siegel EK. 2007. Characterization of a catalase-negative methic il lin-res istant Staphylococcus aureus strain. J. Clin. Microbiol., 45(8): 2684–2685.

Gutierrez J, Barry-Ryan C, Bourke P. 2008. The antimicrobial ef f icacy of plant essential oil combinations and interactions with food ingredients. Int. J. Food Microbiol., 124(1): 91-97.

Hayashi T, Hayashi K, Maeda M, Kojima I. 1996. Calcium spirulan, an inhibitor of enveloped virus replication, from a blue green alga Spirulina platens is. J. Nat. Prod., 59(1): 83-87.



84

Justo GZ, Silva MR, Queiroz ML. 2001. Effects of green algae chlorella vulgaris on the response of the host hematopoietic system to intraperitoneal ehrl ich asc ites tumour transplantat ion in mice. Immunopharmacol. Immunotoxicol., 23(1): 199-131.

Kaushik P, Chauhan A. 2008. In v itro antibacterial activity of laboratory grown culture of Spirulina platens is. Indian J. Microbiol., 48(3): 348-352.

Kloos WE, Bannerman TL. 1999. Staphylococcus and Micrococcus. In: “Manual of Clinical Microbiology. (Murray PR. Ed.)”. W ashington DC, Am. Soc. Microbiol., 7 t h ed., pp. 2113.

Kobashi K. 1989. Pharmacologically act ive metabolites from symbiotic microalgae in Okinawan marine invertebrates. J. Nat. Prod., 52(2): 225-238.

Koch FE. 1942. Electivnährboden für Staphylokokken. Zentr. Bakt. Paras itenk. I Orig., 149: 122–124.

Koneman EW, Allen SD, Janada W M, Schreckenberger PC, Winn W. 1997. Color Atlas and Textbook of Diagnostic Microbiology, JB Lippincott, Philadelphia, pp. 1395.

Kuhl A. 1962. Zur Physiologie der Speicherung konders ierter anor-ganischerPhosphatein Chlorella. Vortr. Botan. hrsg. Deut. Botan. Ges. (N.F.), 1: 157–166

Lindequist U, Schweder T. 2001. Marine biotechnology. In: “Biotechnology. (Rehm HJ, Reed G. Eds)”, vol. 10. Wiley-VCH, Weinheim, pp. 441-484.

Mac Millan JB, Ernst-Russell MA, De Ropp JS, Molinski TF. 2002. Lobocyclamides A-C, l ipopeptides from a crypt ic cyanobacterium mat containing Lyngbya confervoides . J. Org. Chem., 67(23): 8210-8215.

Mahasneh I, Jamal M, Kashashneh M, Zibdeh M. 1995. Antibiotic act ivity of marine algae against multi-ant ibiotic res istant bacteria. Microbios, 83(334): 23-26.

Mala R, Sarojini M, Saravanababu S, Umadevi N. 2009. Screening for antim icrobial activity of crude extracts of Spirulina Platensis . J. Cell Tissue Res. , 9(3): 1951-1955.

Metzger P, Rager MN, Largeau C. 2002. Botryolins A and B, two tetramethyl sequalene tr iethers f rom the green microalga Botryococcus braunii. Phytochemistry, 59(8): 839-843.

Mtolera MSP, Semesi AK. 1996. Ant imicrobial activity of extracts from s ix green algae from Tanzania. In: “Current trends in mar ine botanical research in the East Afr ican region (Bjork M, Semesi AK, Pedersen M, Bergman B. eds)”. Sida, Marine science Program, Department for research cooperation, SAREC, Stockholm, pp. 211–217.

Mulyono N, Lay BW , Rahayu S, Yaprianti I. 2012. Antibacterial activity of petung bamboo (Dendrocalamus Asper) leaf extract against pathogenic Escherichia coli and their chemical identif ication. Int. J. Pharm. Biol. Arch., 3(4): 770-778.

Newman DJ, Cragg GM, Snader KM. 2003. Natural products as source of new drugs over the period 1981-2002. J. Nat. prod., 66(7): 1022-1037.

Ostrosky EA, Mizumoto MK, Lima ME, Kaneko TM, Nishikawa SO, Freitas BR. 2008. Methods for

evaluation of the antimicrobial activity and determination of Minimum Inhibitory Concentration (MIC) of plant extracts. Rev. Bras. Farmacogn., 18(2): 301-307.

Pandian P, Selvamuthukumar S, Manavalan R, Parthasarathy V. 2011. Screening of antibacterial and ant ifungal activit ies of red marine algae Acanthaphora spic ifera (Rhodophyceae). J. Biomed. Sci. Res., 3(3): 444- 448.

Parekh J, Chanda S. 2007. In v itro antibacterial activity of crude methanol extract of W oodfordia fruticosa Kurz f lower (Lythacease). Braz. J. Microbiol., 38(2): 230-239

Patton T, Barrett J, Brennan J, Moran N. 2006. Use of a spectrophotometer ic bioassay for determination of microbial sensitivity to manuka honey. J. Microbiol. Meth., 64(1): 84-95.

Pratt R, Daniels TC, Eiler JJ, Gunnison JB, Kumler WD, Oneto JF, Strait LA, Spoehr HA, Hardin GJ, Milner HW, Smith JH, Strain HH. 1944. Chlorell in. An antibacterial substance from Chrole lla. Science, 99(2574): 351-352.

Robert S, Anders RL, Niels F, Frabk E. 2003. Evaluation of different disk diffusion/media for detection of methic il lin resistance in Staphylococcus aureus and coagulase-negative staphylococc i. APMIS, 111(9): 905-914.

Rosell K, Srivastava LM. 1987. Fatty ac ids as antimicrobial substances in brown algae. Hydrobiologia, 151-152(1): 471-475.

Rufián-Henares JA, Morales FJ. 2006. A new applicat ion of a commerc ial microtiter plate based assay for assessing the antimicrobial activity of Mail lard reaction products. Food Res. Int ., 39(1): 33–39.

Sanz R, Marín I, Ruiz-Santa-Quiter ia JA, Orden JA, Cid D, Diez RM, Silhadi KS, Amils R, de la Fuente R. 2000. Catalase def iciency in Staphylococcus aureus subsp. anaerobius is associated with natural loss-of-function mutations within the structural gene. Microbiology, 146(pt 2): 465–475.

Sastry VMVS, Rao GRK. 1994. Antibacterial substances from marine algae: successive extraction us ing benzene, chloroform and methanol. Bot. Mar., 37(4): 357-360.

Scheuer PJ. 1990. Some marine ecological phenomena: chemical bas is and biomedical potential. Sc ience, 248 (4952): 173-177.

Simic S, Kosanic M, Rankovic B. 2012. Evaluation of In Vitro Antioxidant and Ant imicrobial Activit ies of Green Microalgae Trentepohlia umbrina. Not. Bot. Hort i. Agrobo., 40(2): 86-91

Taylor W R. 1960. Marine Algae of the eastern tropical and subtropical coasts of the Americas. Univers ity of Michigan Press, Ann Arbor, Michigan, pp. 870.

Trono Jr GC. 1999. D iversity of the seaweed f lora of the Philippines and its util ization. Hydrobiologia, 398-399(0): 1-6.

Turcotte C, Lacroix C, Kheadr E, Grignon L, Fliss I. 2004. A rapid turbidometr ic microplate bioassay f or accurate quant if icat ion of lactic ac id bacteria bacteriocins. Int. J. Food Microbiol. , 90(3): 283–293.



85

Vadas RL. 1979. Seaweeds: An overview, ecological and economic importance. Experientia, 35(4): 429-570.

Williams RE, Gibson AG, Aitchison TC, Lever R, Mackie RM. 1990. Assessment of contact-plate sampling technique and subsequent quantitat ive bacterial studies in atopic dermatit is. Br. J. Dermatol., 123(4): 493-501.

Yang Y, Park Y, Cassada DA, Snow DD, Rogers DG, Lee J. 2011. In vitro and in v ivo saf ety

assessment of edible blue-green algae, Nostoc commune var. sphaeroides kütz ing and Spirulina platens is . Food Chem. Toxicol., 49(7): 1560-1564.

Zarrouk C. 1966. Contribution à l’étude d’une cyanophycée. Inf luence de divers’ facteurs physiques et chimiques sur la croissance et la photosynthèse de Spirulina maxima. Ph.D. Thesis, Université de Paris, Paris.

األنشطة المضادة للبكتريا للمستخلصات المختلفة من بعض الطحالب

إيناس مصطفى البالطعاطف محمد أبوشادي، مصطفى محمد الشیخ، شیماء محمد الشافعي،

قسم النبات، كلیة العلوم، جامعة طنطا، مصر

تھدف ھذه الدراسة إلى دراسة النشاط ضد میكروبي نوعین من الطحالب لمستخلصات الطحالب التى تنتمى إلى

) كلوريلال فولجاريز وسبیرولینا بالتینسیس(الدقیقة مثل سراجسیوم فولجار (ونوعین من األعشاب البحرية مثل

ضد سالالت من البكتريا ) وسراجسیوم ويجتاىّ عزلھا) استافیلوكوكس أوريوس( وقد قمنا بإستخدام . تم

بى وتمّ طريقة أطباق المیركوتیتر لتقییم النشاط الضد میكرو وأوضحت . تحضیر المستخلصات بإستخدام أربع مذيبات مختلفة

ھذه الدراسة أن مستخلصات المیثانول أعطت أفضل نتائج وقد ّت نجاح استخدام مستخلصات الطحالب كمصدر للمركبات أكد

. الضد میكروبیة

:المحكمون عفت فھمي شبانه قسم النبات، علوم القاھرة .د.أ زينب خلیل إبراھیم قسم النبات، علوم القاھرة.د.أ

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ANTIBACTERIAL ACTIVITIES OF DIFFERENT … of S. aureus colonies included test for catalase, slide coagulase and DNase (Koneman et al., 1997), as well as Api-Staph strip system (API