Project Abstract

<p> </p><h3>                ABSTRACT</h3><h3>Background</h3><p>Plants are rich in secondary metabolites and are being used for the treatment of various ailments in the indigenous system of medicine. Many developing countries are facing illnesses, and deaths among children are caused by germs, which get into the mouth via water and food. In addition, it has been estimated that up to 80% of all disease and sickness in the world is caused by inadequate sanitation, polluted water or unavailability of water. Thus, this study investigates the water purifying property of <i>Moringa oleifera</i>&nbsp;seed powder and determines the role of seed extracts against a few bacterial growths.</p><h3>Methods</h3><p>Water samples were obtained randomly during January and February, 2015, from the Angereb and Shinta rivers, Gondar, Ethiopia. Both sites of water samples were subjected for purification studies and treated with dried seed powder. Treated water samples were subjected to bacteriological analysis using most probable number technique.</p><h3>Results</h3><p>Addition of aluminum sulfate as a coagulant lowered the water pH from 7.2 to 3.66, while the seed extract water pH remained the same. Treatment of 0.016 g/L of <i>M. oleifera</i>&nbsp;decreased water turbidity from 208.3 nephelometric turbidity units (NTU) to 33.66 NTU (83.84%) and from 129 NTU to 16.8 NTU (86.98%) for the Shinta and Angereb river water samples, respectively. The highest microbial load reduction was found with the Angereb (97.17%) and Shinta (97.50%) rivers. The acetone extracts showed maximum antibacterial activity with 19.00 mm against <i>Salmonella typhii</i>&nbsp;(clinical isolate), while <i>Shigella dysenteriae</i>&nbsp;(clinical isolate) was the least sensitive with 7.66 mm on the aqueous extract. The most frequent MIC value was 6.25 mg/mL followed by 12.5 mg/mL. The acetone extract is the most potent in inhibiting and killing the test organisms at a very low concentration for <i>Shigella typhii</i>.</p><h3>Conclusion</h3><p>Taken together, the seed powder exhibits a remarkable reduction in turbidity and coliform count which makes the seed powder a good source for water purification. The acetone extract of seed had a strong antibacterial activity. It reveals that the seed powder and its extract can control and reduce waterborne bacterial diseases. This investigation facilitates benefits to those who cannot afford and or have access to clean drinking water in Ethiopia and elsewhere.</p> <br><p></p>

Project Overview

<p> </p><div><p><b>1.0 INTRODUCTION</b><br></p><p><b>1.2 BACKGROUND STUDY</b></p><p>Ethiopia is a tropical country which is endowed with rich plant biodiversity, and there are many plant species found in this country for medicinal purposes, and it is often reported and explored [1]. Many local healers in developing countries still depend on traditional medicines for their primary healthcare. Therefore, such plants should be investigated to understand their properties, safety and efficiency for various types of exploitation. Different plant parts are rich in a wide variety of secondary metabolites such as tannins, terpenoids, alkaloids, flavonoids, glycosides, which have been found to have antimicrobial properties [2,3,4]. In recent days, these plant metabolites are also used in the fabrication of metal and metal oxide nanoparticles, which have shown effective antimicrobial properties [5.6.7.8.10]. <i>Moringa oleifera</i>&nbsp;Lam. belongs to the family Moringaceae and is a valuable plant, found in many countries of the tropics and subtropics. Its leaves, fruit, flowers and immature pods are used as a highly nutritive vegetable in many countries, particularly in India, Pakistan, Philippines, Hawaii and many parts of Africa [3]. Seed extract is observed to have a protective effect by decreasing liver lipid peroxides and is antihypertensive [11-13]. <i>M. oleifera</i>&nbsp;roots, leaves, seed, fruit, flowers, bark and immature pods are used as cardiac and circulatory stimulants, contain antipyretic, antiepileptic, antitumor, antiinflammatory, antiulcer, diuretic, antihypertensive, cholesterol lowering, antispasmodic, antidiabetic, hepatoprotective, antioxidant, antibacterial and antifungal activities, and are being used for the treatment of various ailments in the indigenous system of medicine [3].</p><p>Water is used for several purposes by humans, but the level of purity of the water being consumed is very crucial since it has a direct effect on health. More than half of all illnesses and deaths among children are caused by germs, which get into the mouth via water and food. The World Health Organization has estimated that up to 80% of all disease and sickness in the world is caused by inadequate sanitation, polluted water or unavailability of water [13]. Every day 2 million tonnes of sewage, industrial and agricultural waste is discharged into the world’s water [14], the equivalent of the weight of the entire human population of 6.8 billion people. According to UN estimates, the amount of wastewater produced annually is about 1500 km3, six times more than that exists in all the rivers of the world [14].</p><p> Lake Tana occupies a wide depression in the Ethiopian plateau and is the largest lake in the country. The lake is the source of the Blue Nile river with a total surface area of 3600 km2, a volume of 28 km3&nbsp;and an average elevation of 1911 m above the sea level. More than 40 small and big rivers are reported to feed Lake Tana. It is located at a latitude of 12 (12°0′0N) and a longitude of 37.33 (37°19′60E). It was formed by a volcanic blockage that reversed the previously north-flowing Blue Nile river and created one of Africa’s greatest waterfalls. The majority of people living around Lake Tana and its rivers are still utilizing water for drinking and other routine activities without any purification. It has been found that the dry seeds of <i>M. oleifera</i>&nbsp;are used in place of aluminum sulfate by rural women to treat highly turbid Nile water. It has also been reported that when the crushed seeds are added to raw water, the proteins produce positive charges acting like magnets and attracting the predominantly negatively charged particles (such as clay, silt, bacteria and other toxic particles in water) [15]. Crushed seeds are also capable of attracting and sticking fast to bacteria and viruses that are found in contaminated and turbid water [16]&nbsp;A comparative study has also been carried out with harvesting surface rainwater and its purification by using seed extracts and aluminum sulfate [17]. Seeds have also shown antimicrobial activity [18,19]. Most of the workers have reported the use of leaf and or bark extracts elsewhere, while the scientific and effective use of <i>M. oleifera</i>&nbsp;dried seed powder for water purification and antibacterial efficacy under Gondar, Ethiopia agro-climatic zone is not reported so far. Therefore, the present study was designed. In this study, the water purifying property of the seed powder specifically for the Angereb and Shinta rivers and the role of seed extracts against bacterial growth namely <i>Escherichia coli</i>&nbsp;(ATCC2592), <i>E. coli</i>&nbsp;(clinical isolate), <i>Salmonella typhii</i>&nbsp;(clinical isolate) and <i>Shigella dysenteriae</i>&nbsp;(clinical isolate) were examined</p><div><h3>1. 2 METHOD</h3><p><b><i>1.2.1 Study area</i></b><br></p><p>Gondar is bounded by 120 35′ 07″ North latitude and 370 26′ 08″ East longitude, and it has a narrow range of altitude, i.e., 2000–2200 m above the sea level, which is north of Lake Tana and southwest of the Simien Mountains. The Angereb river is a river of Ethiopia and eastern Sudan and one of the sources of the Nile river found in Gondar town. It flows west to join the Atbarah river [21]. The other river flowing in Gondar is the Shinta. The Shinta contributes to the Angereb river which is a part of the tributaries of Lake Tana. The Shinta river drains from north to south, and it serves as natural sewerage lines for domestic and industrial wastes [21]. All samples were labeled and transported to the Tewodros campus Microbiology Laboratory, Department of Biology, University of Gondar, Gondar, Ethiopia, and stored at 4 °C for further studies.</p></div> <h3>Plant material collection, identification and treatments</h3><p>Seeds of <i>Moringa oleifera</i>&nbsp;used in this study were obtained from the Agricultural and Forestry office, Adirkay, Gondar. Adirkay district has a latitude and longitude of 13°29′ 50°60′N38°03′ 25.96°E with an elevation of 2025 m above the sea level. The area is predominately rural and most residents live in villages as agriculturists. The experiment was carried out during October 2014 to May 2015 in the Microbiology Laboratory, Department of Biology, University of Gondar, Ethiopia. The treatments given were the varying concentrations of powder produced from <i>M. oleifera</i>&nbsp;seeds and the positive and negative control (aluminum sulfate and no seed powder, respectively) for water purification. Further, the varying concentration of extracts with different solvents was produced from <i>M. oleifera</i>&nbsp;seeds and the positive control (ciprofloxacin) was used for antibacterial test.</p><p> </p><h3>Extraction of Moringa seed powder and antimicrobial test organisms</h3><p>Mature seeds of <i>M. oleifera</i>&nbsp;were chosen from dry cracked fruits. The plucked fruits were cracked to obtain the seeds which were air-dried for 2 days. The shells surrounding the seed kernels were removed using a knife, and the kernels were powdered using a laboratory mortar and pestle and sieved using a sieve with a pore size of 2.5 mm2 to obtain a fine powder. The powder was stored in a sterile bottle at room temperature in a dark place. The powdered sample was successively extracted with methanol, acetone and aqueous in increasing polarity. In this procedure, 50 g of <i>M. oleifera</i>&nbsp;powdered seeds was soaked in 250 mL of each of the solvents which were acetone, methanol and aqueous and in all cases equal volumes of solvents were used. They were left shaking on a horizontal shaker for 3 days. Then, the extracts were filtered separately through Whatman no.1 filter paper. The filtrates were then centrifuged at 5000 rpm for 15 min. The supernatant of each extract was evaporated by using Rota vapor (Laborator 4000-efficient, Heidolph, Germany). The crude extracts were stored at 4 °C. The yields of acetone, methanol and water extracts weighed 16, 15 and 13%, respectively. Each test was replicated three times [<a target="_blank" rel="nofollow" href="https://agricultureandfoodsecurity.biomedcentral.com/articles/10.1186/s40066-018-0177-1#ref-CR22">22</a>]. The antibacterial properties of the crude extracts were tested against the test organisms. Pure cultures of <i>Escherichia coli</i>&nbsp;(ATCC 2592) were taken from the microbiology laboratory, Department of Biology, while <i>E. coli</i>&nbsp;(clinical isolate), <i>Salmonella typhii</i>&nbsp;(clinical isolate) and <i>Shigella dysenteriae</i>&nbsp;(clinical isolate) were obtained from the Gondar College of Medical Science, University of Gondar, Ethiopia. Organisms were chosen based on reports of their human and livestock pathogenicity in water.</p><p> </p><h3>Sample preparation and laboratory analyses</h3><p>Forty liters of raw water samples were fetched from the Shinta and Angereb rivers. Thereafter, dispensed into six beakers containing 500 mL each for <i>M. oleifera</i>&nbsp;and aluminum sulfate. Five different concentrations of the stock solution for the loading dose were prepared by weighing 2.0, 4.0, 6.0, 8.0 and 10.0 g of <i>M. oleifera</i>&nbsp;seed powder and aluminum sulfate each separately into a beaker containing 500 ml of distilled water. The mixtures in the beakers were stirred using a glass rod to obtain a clear solution. A 500 ml of distilled water with no <i>M. oleifera</i>&nbsp;seed powder extract was kept as negative control [<a target="_blank" rel="nofollow" href="https://agricultureandfoodsecurity.biomedcentral.com/articles/10.1186/s40066-018-0177-1#ref-CR23">23</a>]. Two ml of the various concentrations including the control of all the loading dosages prepared was measured into a beaker containing 500 ml of the sample river water. The solutions were mixed rapidly for 2 min, followed by 10 min of gentle mixing using a sterile glass rod to aid in coagulant formation. The suspensions were left to stand without disturbance for 1 h. This is the method adopted since there is no standard method for conducting the jar test [24]. The supernatants formed were recovered and subjected to total coliforms count, pH and turbidity measurements.</p> <h3>Total coliform using most probable number (MPN) test</h3><p>In determining the MPN of coliforms that were present in each of the treated water samples, the multiple tube fermentation method was adopted. Lauryl tryptose broth (LTB) tubes were used for the bacteria growth. Two types of the LTB tubes were prepared. These were the single-strength lauryl tryptose broth (SSLTB) and the double-strength lauryl tryptose broth (DSLTB). In the single strength, 6.5 g of the LTB powder was weighed and dissolved in 500 ml of distilled water. The double strength was prepared using exactly twice the weights of the reagents used and stirred gently for 10 min as it was done in the single-strength LTB preparation [25]. An estimate of the number of the coliforms (most probable number) was done in the presumptive test. In this procedure, 15 test tubes with 15 ml of lauryl tryptose broth were inoculated with 10 mL of treated water samples and the control at different intervals. Five tubes received 10 ml of water, the LTB used was double strength for this case, other five tubes received 1 ml of water, and the last five tubes received 0.1 ml of water. Here, the last 10 tubes contained single-strength LTB. The test tubes were then incubated for 24–48 h at 37 °C. For confirmatory test, from each positive presumptive test tube three loopful of samples were transferred to test tubes containing brilliant green lactose bile broth (BGLB), further incubated at 37 °C for 24 h and inspected for the presence of gas. The number of tubes showing gas production was counted, and the figure was compared to a table developed by the American Public Health Association [26]. The number was the MPN of coliform per 100 ml of the water sample [27].</p><h3>Turbidity and pH measurement</h3><p>Turbidity was determined by the nephelometric method using turbidimeter on water samples on the jar tests. The turbidity level of the water samples was measured before and after treatment of the sample with different doses of <i>M. oleifera</i>&nbsp;seed powder and aluminum sulfate using turbidity meter. A test tube was filled with the sample to the 10 ml mark, and turbidity meter reading was taken against the blank tube. The result was read directly from the turbidity meter display and reported as nephelometric turbidity units (NTU) [28]. A buffer solution calibrated pH meter was used to measure the hydrogen ion concentration of the water before and after treatment with Moringa seed powder and aluminum sulfate. Accordingly, the result was read from the display [29]. In addition, McFarland standard was comparable to a bacterial suspension of 1.5 × 108 cell/mL [30].</p><h3>Antimicrobial tests using different methods</h3><p>The isolates were preserved on nutrient agar plate and incubated at 37 °C for 24 h according to the manufacturer’s specification. The isolates were preserved on agar slant and stored at 4 °C until further use. A loopful of the test organisms was inoculated into sterile normal saline. The bacterial suspension was compared to the 0.5 McFarland standards [31]. The antibacterial assay was performed by agar well diffusion technique, and MIC and MBC were determined.</p><h3>Agar well diffusion</h3><p>Bacterial broth culture was prepared to a density of 108 cells mL−1 of 0.5 McFarland standards. The aliquot was spread evenly onto Mueller–Hinton agar using a sterilized cotton swab. Then, the plated medium was allowed to dry at room temperature for 30 min [32]. On each plate, equidistant wells were prepared with a 6-mm-diameter sterilized, cork borer, which were 2 mm from the edge of the plate. Fifty microliters of each extract (50 mg/mL) was aseptically introduced into a respective agar well. Ciprofloxacin (25 µg/mL) was used as standard (positive) control and sample free solutions as blank control. This was followed by allowing the agar plate on the bench for 40 min pre-diffusion followed by incubation at 37 °C for 24–48 h. The formation of clear inhibition zone of  ≥ 7 mm diameters around the wells was regarded as significant susceptibility of the organisms to the extract [33]. The experiment was performed in triplicates.</p><h3>Determination of minimum inhibitory concentration (MIC)</h3><p>The MIC was determined for extracts that showed ≥ 7 mm diameter growth inhibition zone. The test was performed using agar dilution method. In this technique, a serial of twofold dilution of the extract was prepared in Mueller–Hinton agar. The bacterial inoculum which was standardized according to the McFarland standard was inoculated on the surface of the agar. The extract solution (50 mg/mL) was serially diluted as 1:2, 1:4 and 1:8 to bring 25, 12.5 and 6.25 mg/mL concentrations, respectively [22]. The extracts were then aseptically introduced as described above. The inhibition of growth was assessed after 24 h incubation at 37 °C, and the minimum concentration that inhibited growth was considered as MIC value of the extract.</p><h3>Determination of the minimum bactericidal concentration (MBC)</h3><p>The MBC of the plant extracts was determined by the modified method of Spencer and Spencer [34]. The plates that showed no growth after incubation of the batch of agar plates on MIC were sub-cultured on Mueller–Hinton agar plates and incubated at 37 °C for 24 h. The highest dilution (least concentration) that yielded no single bacterial colony and showed no visible growth after incubation was taken as the MBC.</p><h3>Statistical analysis</h3><p>Data were analyzed using SPSS (version 16) statistical software. Mean coliform and turbidity reduction were calculated. One-way ANOVA was used to test existence of statistically significant difference between mean zones of inhibition. A significance level of <i>p</i>&nbsp;value less than 0.05 was used.</p> <br><p></p> <br><p></p><br> <br><p></p><br> </div><br><p></p>