a n t ib io t ic s e n s it iv it y p r o f il e o f e n t e r ic b a c t e r iaISOLATED FROM SOIL SAMPLES AROUND KEMRI (CMR) AND ITS

ENVIRONS IN NAIROBI KENYA

By:Peter Shigoli Mashedi

(BSc Medical Microbiology, JKUAT)

\ thesis submitted in partial fulfillment of the requirements for the award of the degree of Master of Science in Microbiology of the University of Nairobi.

November, 2012

i

pec I a ration

flie work described herein is my original work as part of ongoing project. Articles and texts hed have been acknowledged. The contents in this thesis have not been submitted previously to ly other Univeristy in whole or part, for the award of any degree or academic titles

>tter Shigoli Mashedi

lture..

te.... 3o

his thesis has been submitted with the approval as the University supervisor. 'Jr Miriam M. Jumba Jchool of Biological Sciences Jniversity of Nairobi Signature . . . ^)ate............... ./ .10 Is3L Q J 5 -

'his thesis has been submitted with the approval as the External supervisor.

)r. Christine Bii

-enter for Microbiology Research (CMR) Kenya Medical Research Institute, Nairobi * ) Box 54840-00200 NAIROBI.

Signature....... ~ ...................................

pate....................... ...................................................................

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Dedication

I dedicate this study to my family members.

Acknowledgment

I would like to thank the Almighty God lor giving me the strength and guidance to have completed this project. I also thank my supervisors Dr. Miriam Jumba and Dr. Christine Bii for guiding me with the project and also the entire stall at Opportunistic Infections Laboratory in KEMRI Nairobi, for the assistance they gave me.

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AbstractSoil is able to contain enteric bacteria and other pathogens in great concentrations, as it is normally a recipient of solid and liquid waste materials frequently. Recent studies elicit that soil may have a greater role in the transmission of enteric diseases than previously expected, even though its role as a reservoir of certain bacterial pathogens is not in doubt. Enteric bacteria are responsible for causing most gastrointestinal infections, for example salmonellosis, dysentery, typhoid fever and other infections caused by Yersinia sp. and Escherichia coli 0157:H7 and many other strains. The study was aimed at determining the prevalence of enteric bacteria from various soil samples collected around Nairobi, and to compare their drug susceptibility profile with those from clinical samples. The soil samples were collected from various locations in Nairobi within a radius of 30km from Kenya Medical Research Institute, Centre for Microbiology Research in Nairobi, with their Global Position System (GPS) location recorded down, then transported to the laboratory. Ten grams of each o f the soil samples were serially diluted then plated on Mueller-I Iinton agar and incubated at 30°C overnight, the colonies were Gram stained and the Gram-negative colonies inoculated on Analytic Profile Index kit (API 20E) for further identification. Antibiotic sensitivity testing was done using Disc Diffusion method and then compared with clinical isolates. Out of the soil samples (n=236) inoculated onto Mueller- Hinton agar, 17 were positive for Proteus salmonicida, which represents a prevalence of 7.2% of enteric bacteria in the soil. The other isolated Gram negative bacteria were Myroides spp, Pseudomonas putida. Pseudomonas aeruginosa, Stenotrophomonas maltophila and Alcaligenes spp. Proteus salmonicida showed a higher sensitivity to the antibiotics compared to the clinical Proteus except for Cefotaxime antibiotic which was resistant to it. In conclusion, soil may be a significant a reservoir for the enteric bacteria contributing to antibiotic resistance as indicated by Proteus salmonicida with resistance to Cefotaxime antibiotic, compared to Proteus species from the clinical sources which was sensitive to the same antibiotic.

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T A BL E O F C O N T E N T SDeclaration.......................................................................................................................................... iiDedication...........................................................................................................................................iiiAcknowledgement.............................................................................................................................. ivAbstract.............................................................................................................................................. v

1.1 Introduction................................................................................................................................ 12.0 CHAPTER TWO.........................................................................................................................52.1 Literature review............................... !...................................................................................... 5

2.1.1 Enteric bacteria.....................................................................................................................52.1.2 Isolation, identification and drug susceptibility testing of enteric bacteria.......................62.1.3 Sources of soil contamination by enteric pathogens.......................................................... 72.1.4 Fate of enteric bacteria in the soil....................................................................................... 92.1.5 The role of soil as a reservoir for bacteria in contributing to antibtiobitc resistance ... 10

2.2 Justification............................................................................................................................... 122.3 Hypothesis................................................................................................................................. 132.4 Objectives.......................................................... ......................................................................13

2.4.1 Overall objective................................................................................................................. 132.4.2 Specific objectives.................................. ;.........................................................................13

3.0 CHAPTER THREE.................................................................................................................. 143.1 Materials and methods..............................................................................................................14

3.1.1 Study site ................................................................................................ 143.1.2 Sampling.............................................................................................................................153.1.3 Sample size and its justification........................................................................................153.1.4 Sample collection................................................................................................................15

3.2 Laboratory methodology.......................................................................................................... 163.2.1 Sample processing and identification of enteric bacteria................................................163.2.2 Antibiotic susceptibility testing for the enteric bacteria................................................... 163.2.4 Variables................................................. ;.........................................................................17

3.3 Experimental design.................................................................................................................. 17VI

4.0 CHAPTER FOUR........................................................................................................................184.1 Results........................................................................................................................................18

4.1.3 Colour reactions on API 20E kit after incubation and addition of reagents...................224.1.4 Distribution of the enteric soil isolates............................................................................... 24

4.1.5 Antibiotic sensitivity profile of the soil and clinical isolates........................................ 254.2 Data analysis.............................................................................................................................29

5.0 CHAPTER FIVE....................... ............................................................................................... 305.1 Discussion.....................................................................................................i......................... 305.2 Conclusion.................................................................................................................................335.3 Recommendations.................................................................................................................... 33

6.0 REFERENCES.......................................................................................................................34APPENDIX....................................................................... 40

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LIST OF TABLES1: Gram stain reactions of isolates 192: Antibiotic sensitivity profile of Proteus salmonicida from soil 253: Antibiotic sensitivity profile of Proteus species from clinical isolate 264: Antibiotic amounts impregnated on disc and breakpoints for enteric bacteria 26

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LIST OF FIGURES1: Map of Nairobi 142: Gram stain 213: Gram stain 214: Various API reactions 235: Various API reactions 236: Distribution of enteric soil isolates 247: Antibiotic sensitivity 278: Comparison of inhibition zones of Proteus species from the soil and clinical samples 28

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1.0 C H A P T E R O N E1.1 IntroductionHumans are in contact with soil constantly, either directly or indirectly via food, water and air and thus soil may act as a vector and source of important human disease causing agents. Although many of the diseases associated with soils have been well characterized and studied, enteric diseases and their link to soil have been understudied and possibly underestimated. In order to clarify this connection, diseases associated with soil have been classified depending on the origin of the etiological agent as follows (Toze, 1997): (a) soil-associated diseases which are caused by opportunistic or emerging pathogens that belong to the normal soil microbiota (e.g. Aspergillus fumigatus is a very common fungus occurring in soils and can infect the lungs via inhalation of spores), (b) soil-related diseases, which result in intoxication from the ingestion of food contaminated with entero- or neurotoxins (Clostridium botulinum, C. perfrigens and Bacillus cereus), (c) soil-based diseases caused by pathogens indigenous to soil (which include C. tetani, B. anthracis, and C. perfringens) and (d) soil-borne diseases caused by enteric pathogens which get into soil by means of human or animal excreta. Enteric pathogens transmitted by the fecal-oral route are bacteria, viruses, protozoa and helminths.

Gastrointestinal infections are the most common diseases caused by enteric bacteria. Some examples are salmonellosis {Salmonella sp), cholera {Vibrio cholerae), dysentery {Shigella sp.) and other infections caused by Campylobacter jejuni. Yersinia sp. and Escherichia coli 0157:H7 among others. E. coli 0157:117 successfully causes infections because of its low infectious dose (ID), which can be as few as ten cells. (Rosen. 2000). Annual summaries of food-borne and water-borne disease outbreaks published by the Centers for Disease Control show that, in the early 2000, there was an increase in food-borne and water-borne outbreaks caused by enteric pathogens. It is possible that the water and food contaminations were related to the practices mentioned above. For example, in the United States, water-borne diseases caused by contaminated ground water increased in the 1990s (Plym-Forshell and Ekesbo, 1993; Craun and Caldron, 1996). Fruits and vegetables frequently come in contact with soil post-harvest and thus may become contaminated with soil enteric bacteria present in sewage sludge or manure spread. One of the first cases of infection with E. coli 0157:H7 linked to the use of animal excreta as manure was with an ovo-vegetarian woman. The woman consumed almost exclusively the food

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produced in her garden, in which she used the manure from her own cow as a fertilizer (OMRI, 1998). In 1970, an outbreak occurred as a result of the ingestion of vegetables irrigated with wastewater. Further studies indicated that Vibrio cholerae was present in the irrigated soils (Shuval et al., 1986). Fruit juice and cider may become contaminated as a result of the fruit falling to the ground and coming in contact with soil which may contain pathogens from animal excreta or sewage sludge used as fertilizer. Unpasteurized juice has been associated with at least 15 food-borne illness outbreaks since 1900 (Parish, 1997).

Bacterial antibiotic resistance is a serious public health concern due to the reduced potency of antimicrobial agents used in the treatment of infectious diseases (Martinez and Baquero, 2002). Enteric bacteria are present in large numbers in the human and animal gut, are medically important as infectious agents and exhibit antibiotic resistance (Paterson, 2002). Antibiotics are extensively used in human and veterinary medicine, and in agriculture, for the treatment of infections, growth enhancement, and prophylaxis in food animals. This leads to selection of drug- and multidrug-resistant bacteria (Barbosa and Levy, 2000). Antibiotic-producing microorganisms are found naturally in soil. This suggests intrinsic chromosomal antibiotic resistance originated in the soil in response to harsh environments generated by such antibiotic- producing microorganisms (Randal and Woodward, 2001). Whether naturally occurring or commercially made, stable antibiotics accumulate in soil inhabited by food animals and where antibiotics are used. This leads to selection for multidrug resistance, which can be chromosomally (intrinsic) or plasmid-encoded acquired (Owens et al., 2001).

Bacterial antibiotic resistance is a serious public health concern due to the reduced potency of antimicrobial agents used in the treatment of infectious diseases (Martinez and Baquero, 2002). Enteric bacteria are present in large numbers in the human and animal gut, are medically important as infectious agents and exhibit antibiotic resistance (Paterson, 2002). Antibiotics are extensively used in human and veterinary medicine, and in agriculture, for the treatment of infections, growth enhancement, and prophylaxis in food animals. This leads to selection of drug- and multidrug-resistant bacteria (Barbosa and Levy, 2000}. Antibiotic-producing microorganisms are found naturally in- soil. This suggests intrinsic chromosomal antibiotic resistance originated in the soil in response to harsh environments generated by such antibiotic- producing microorganisms (Randal and Woodward, 2001). Whether naturally occurring or

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commercially made, stable antibiotics accumulate in soil inhabited by food animals and where antibiotics are used. This leads to selection for multidrug resistance, which can be chromosomally (intrinsic) or plasmid-encoded acquired (Owens et al., 2001).

Antibiotic sensitivity testing can be used to determine whether soil bacteria have acquired resistance to a particular antibiotic. Several methods for antibiotic sensitivity testing can be used: Minimum Inhibitor) Concentration; Automated and Disc diffusion methods. Minimum Inhibitory Concentration (MIC) is used to determine the lowest concentration of antibiotic at which an isolate cannot produce visible growth after overnight incubation. The broth dilution method is one variation of MIC of which an isolate is inoculated in broth media at a specific inoculum density (in tubes or microtitre plates) containing antibiotics at varying levels (Bannerman, 2003). Doubling dilutions are used and after incubation, turbidity is recorded either with an automated reader or visually, and the breakpoint concentration established. Microtitre plates or ready-to-use strips are commercially available with antibiotics impregnated in the wells. Agar dilution method is a variation on this approach. In this method, a small volume of suspension is inoculated onto agar containing a particular concentration of antibiotic, and incubated (Bannerman, 2003). Thereafter, it is examined for zones of growth.

Automated methods are meant to reduce technical errors and lengthy preparation times (Bannerman, 2003). These methods entail the use of formatted microdilution panels as well as instrumentation and automated reading of plates. Most automated antimicrobial susceptibility testing systems provide automated inoculation, reading and interpretation. These systems have the advantage of being rapid and convenient, but one major limitation for most laboratories is the cost entailed in initial purchase, maintenance and operation of the machinery. According to Antibiotic resistance learning site for vet students (2012). examples ol these machines these include: Vitek System (bioMerieux, France), Micronaut (Merlin. Bornheim-Hesel, Germany). Walk-Away System (Dade International. Sacramento. Calif.), Phoenix (BD Biosciences, Maryland), Avantage Test System (Abbott Laboratories, Irving, Texas), , Sensitive ARIS ( I rek Diagnostic Systems, East Grinstead. UK), Phoenix (BD Biosciences, Maryland) and many others which are being developed.

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In this study, the Disc diffusion or the Kirby-Bauer test is another method used to determine antibiotic sensitivity. A growth medium, usually Mueller-Hinton agar, is first evenly inoculated throughout the plate with the isolate of interest that has been diluted at a standard Mac-Farland concentration. Commercially prepared disks, containing pre-impregnated standard concentration of a particular antibiotic, are then evenly dispensed onto the agar surface (Ryan and Ray, 2004). After an overnight incubation, the bacterial growth around each disc is observed and if the test isolate is susceptible to a particular antibiotic, a clear area where bacteria did not grow, will be observed around that particular disk. This clear zone around an antibiotic disk that has no growth is referred to as the zone of inhibition since this approximates the minimum antibiotic concentration sufficient to prevent growth of the test isolate (Ryan and Ray, 2004). This zone is then measured in millimeters and compared to a standard interpretation chart used to categorize the isolate as susceptible, intermediate or resistant. A variation on this approach is to use a strip impregnated along its length with a gradient of different concentrations of antimicrobial. After incubation this creates an ellipse shaped zone of no growth, the MIC can be read from the concentration markings on the strip (Ryan and Ray, 2004). In this method, no tables need to be referred to get an MIC value and the test requires less manipulations, as one strip will cover the whole concentration range. However it is an expensive method since one strip usually contains one antibiotic only.

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2.0 C H A P T E R T W O2.1 Literature review2.1.1 Enteric bacteria

Enteric bacteria refer to bacteria that are mainly found in the gastrointestinal tract of humans and animals. They belong to Enterobacteriaceae family (Graham el a/., 2000) and are Gram­negative. rod-shaped lacultative anaerobic bacteria, most of which are motile with peritrichous flagella, oxidase negative and have relatively simple growth requirements (Graham et al., 2000). Enterobacteriaceae are widely distributed in nature in plants and animals, and are important pathogens and they are part of the intestinal flora, while others are found in water or soil, or are parasites on a variety of different animals and plants (Knight and Girling, 2003).

This family may be classified into tribes, genera and species by their cultural and biochemical characteristics. The species are further classified into biotypes, serotypes, bacteriophage types and colicin types (Satish. 1999).

The five tribes are as outlined below:

Tribe 1 EscherichiaGenus: Escherichia, Eciwardsiella, Citrohacter, Salmonella, Shigella.

Tribe 2. KlebsiellaeGenus: Klebsiella, Enterobacter, Hafnia, Serratia.

Tribe 3 ProteaeGenus: Proteus

Tribe 4: ErwiniaeGenus: Erwinia

Tribe 5: YersinaeGenus: Yersinia

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Enteric bacteria occurring as normal tlora of the intestines have benefits such as synthesis and excretion of vitamins that can be absorbed by the human host as nutrients. Some of the vitamins secreted are vitamin K and B12 (Guarner and Malagelada. 2003). They also prevent colonization by pathogens competing for essential nutrients and attachment on the mucosa - thought to be the most beneficial effect in the intestines. Enteric bacteria have also been shown to stimulate the development of lymphatic tissue in the gastro intestinal tract, and also stimulate production of cross reactive antibodies since they act as antigens in an animal (Guarner and Malagelada. 2003). Enteric bacteria occur as pathogens when they invade the tissues of the gastrointestinal tract or due to secretion of exotoxin or enterotoxin. They cause diseases such as gastroenteritis, typhoid fever, paratyphiod fever, bacillary dysentery, cholera among others (Venkatasen, 2001).

2.1.2 Isolation, identification and drug susceptibility testing of enteric bacteria

Enteric bacteria are isolated in the laboratory using differential, enrichment or selective media. Differential media are media that aid in the presumptive identification of bacteria based on the appearance of the colonies on the medium, for example MacConkey and Xylose-Lactose- Desoxycholate (XLD) agar (Bannerman, 2003). Enrichment media allows fastidious organisms to grow because of the specific nutrients additives such as haemin. An example is Selenite-F broth for isolation of Salmonella sp. (Bannerman. 2003). Selective media are media that contain additives that enhance the presence of the desired organism by inhibiting other organisms. Most commonly the selection is attained with a dye or added antibiotic. An example is MacConkey agar that contains crystal violet that inhibits most Gram-positive organisms (Bannerman, 2003).

Various biochemical tests can also be used to further identify the enteric pathogens. Triple Sugar Iron test (TSI) is used to identify Hydrogen Sulphide producers e.g. Salmonella s p by changes in the media after incubation. Other media for biochemical identification include Indole, Vorges -Prosker and lysine indole mortality media.

The API-20E test kit for the identification of enteric bacteria provides an easy way to inoculate and read tests relevant to members of the.Family Enterobacteriaceae (Willey et al., 2008). A

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plastic strip holding twenty mini-test tubes is inoculated with a saline suspension of a pure culture (as per manufacturer's directions). This process also rehydrates the dessicated medium in each tube. A few tubes are completely filled, and some tubes are overlaid with mineral oil such that anaerobic reactions can be carried out. After incubation in a humidity chamber for 18-24 hours at 37°C. the color reactions are read (some with the aid of added reagents), and the reactions (plus the oxidase reaction done separately) are converted to a seven-digit code which is called the Analytical Profile Index (API). I he code can be fed into the manufacturer's database given as a chart to identify the corresponding enteric bacteria (Willey et al., 2008).

Gram staining can also be used to identify enteric bacteria. It is the most commonly used differential staining procedure for bacterial colonies because of its broad staining spectrum (Ryan et al., 2004). Gram positive retain the crystal violet dye because of increased number of cross linked teichoic acid and decreased permeability of the cell walls to organic solvents as they contain little lipids (Ryan et al.. 2004). Enteric bacteria are Gram-negative since their walls have increased permeability to the decolourizers. since they have a higher lipid content the tend to lose crystal violet stain and pick the counterstain, hence appearing red under a microscope (Ryan et al., 2004).

According to Clinical and Laboratory Standard Institute (2007), the antibiotics used for susceptibility testing for Enterobacteriaceae. are Ampicillin. Cefuroxime. Gentamycin. Ciprofloxacin, Gerttamycin, Naldixic, Co-trimoxazole, Chloramphenical, Cefotaxime and Erythromcin.

2.1.3 Sources of soil contamination by enteric pathogens

1 here is a concern about a possible increase in soil-borne diseases in human populations, given the successful land disposal practices of sewage and sewage sludges that result from wastewater treatment. 1 hese practices may favour the entry of considerable concentrations of enteric pathogens into soil, because large amounts of these solids are applied to lands or disposed of in landfills. A variety of treatment methods, such as composting, aerobic and anaerobic digestion, alkaline stabilization, conditioning, dewatering and heat drying, are used in waste-water

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treatment plants to reduce pollutants and to destroy pathogens (Keswick, 1984). Sludge, is the first product of this treatment and, if additional treatment is given in order to reduce the pathogen concentrations to specific levels, the material becomes a biosolid (Keswick, 1984). Biosolids are classified as either class A or class B. in categories established by the Environmental Protection Agency (EPA) in 2002, based on the following microbiology criteria: Class A biosolids must have a concentration of thermotolerant coliforms below 1.000 colony-forming units (CFU)/g dry weight (dw) by the most probable number (MPN) method, a Salmonella concentration of less than 4 CFU/g dw, an enteric virus concentration of less than four plaque-forming units/g dw and less than four viable helminth eggs/g dw. Class A biosolids can be applied to lawns and home gardens and given away to the public in bags or other containers. In general, they are used like any commercial fertilizer (EPA, 2002).

Class B biosolids may contain Escherichia coli, Salmonella, Shigella, Campylobacter, Cryptosporidium, Giardia, Norwalk virus and enteroviruses (EPA, 2002). Its use is restricted to land application, forest lands, reclasmation sites and for a period of time, access is limited, to the public and to livestock grassing and the harvest schedule is controlled. This time period allows for the natural die-off of pathogens in the biosolids.

Ihere is concern about the effect that the disposal of these solids may have on public health because (a) the fate of these enteric microorganisms in the soil is not well understood and thus they may be a contamination source for food or surface- and groundwater, (b) the infectious dose of some pathogens is low and this could imply a high risk, especially in special populations, such as the immune-compromised and the elderly, (c) there is a possibility of re-growth of pathogenic bacteria (Yeager and Ward 1981; Hay, 1996), (d) the presence of indicator bacteria, such as coliforms, which is used as an index of safety, does not accurately predict the presence of pathogens and (e) many diseases may be due to unknown agents and the methods for their detection have not yet been developed (Morbidity and mortality, 2000).

In developing countries, untreated domestic wastewater is an important source of enteric pathogens to soil because it is used in agricultural irrigation. This presents a high risk to farm workers and to consumers of food products irrigated with wastewater (Strauss, 1994).

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Other practices that favor the entry of considerable amounts of enteric bacteria into the soil environment are the use of human and animal excreta as manure and the inadequate disposal of human excreta in national parks and in general in areas where toilets are not provided (Cilimburg el al., 2000). Feachem el al., (1983) showed that the survival times of some excreted pathogens in soil and on crop surfaces were: Enteroviruses, thermotolerant coliforms and Salmonella spp persist less than 20 days, Vibrio cholerae persists less than ten days and helminth eggs may persist for several months.

Municipal or City solid waste may be another source of enteric pathogens to soil. Enteric pathogens may come from the excreta present in disposable diapers, pet feces, food waste and sewage sludge (Gerba. 1996). On-site soil disposal systems (OSDSs) treat domestic water for 20% of the United States population and could also result in soil, and consequently groundwater contamination (Scandura and Sobsey, 1997).

2.1.4 Fate of enteric bacteria in the soil

Soil moisture favors the survival of bacteria. Reductions in bacterial population densities are observed under dry soil conditions. Clays favor the adsorption of microorganisms to soil particles and this further reduces the die-off rates (Gerba and Bitton, 1984). Clays protect bacterial cells, by creating a barrier against microbial predators and parasites (Yeager and O'Brien, 1979). Hence, the rates of enteric bacteria survival are lower in sandy soils with a low water-holding capacity. pH affects the adsorption characteristics of cells, so inactivation rates in acidic soils are lower (Yeager and O'Brien. 1979). Increases in cation concentrations also result in increased adsorption rates, consequently affecting microbial survival. Soluble organic compounds increase survival and, in the case of bacteria, may favour their re-growth when degradable organic matter is present (Yeager and Ward. 1981).

Microbial movement in soils is dependent on the water saturation state. According to Sinton (1986), microorganisms move rapidly under saturated conditions, but only for a few centimeters, because microorganisms are in close contact with soil particles, promoting the adsorption of microorganisms onto the soil particles. When soil is saturated, all pores are filled with water,

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allowing microorganisms to pass through the soil hence, soil texture controls in part, the movement of microorganisms, because fine-grained soils avoid movement while coarse-grained soils promote the movement (Sinton, 1986). Another important environmental factor affecting microbial movement is rainfall. It can result in pathogen spread by runoff from places where manure or biosolids have been applied or by leaching through the soil profile (Gerba and Bitton, 1984).

2.1.5 The role of soil as a reservoir for bacteria in contributing to antihtiobitc resistance

Bacterial antibiotic resistance has become a serious public health concern because the causative agents of infections in humans and animals are becoming less receptive to the healing aspect of antibiotics due to the reduced potency of antimicrobial agents used in the treatment of infectious diseases (Martinez and Baquero, 2002. Since soil-dwelling bacteria not only produce antibiotics but also are exposed to a myriad of antibiotics produced by surrounding strains, they must develop multiple tactics to survive. According to RxPG news (201OX Researchers, from McMaster University say that study of bacteria found in soil may be critical in identifying how and why antibiotic resistance happens in bacteria that infect people. The researchers led by Professor Gerry Wright, screened 480 strains of soil bacteria isolated from diverse locations for resistance to 21 clinically relevant antibiotics. The study established that out that bacteria showed resistance to major classes of antibiotics, and that the method of Vancomycin resistance was similar to the resistance found in clinical isolates. The same study_also uncovered bacteria that produce enzymes capable of rendering antibiotics inactive, breaking down or modifying them. I heir study hence suggests that the soil serves as an under-recognized source of resistance that has the potential to reach clinically isolated bacteria.Clinically, aminoglycosides play an important role in the treatment of severe sepsis due to enterobacterial infection, as well as infections caused by selected gram-positive aerobic bacilli. They are naturally occurring antibiotics that bind to the 16S ribosomal RNA of the 30S ribosome in the aminoacyl-transfer RNA site (A-site), causing misreading and consequently inhibition of translation (Davies and Benveniste, 1973). The most common mechanism of clinical resistance

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to aminoglycosides is mediated by antibiotic-moditying enzymes such as kinases that confer a

high level o f resistance.

However, aminoglycoside kinases, enzymes that modify the antibiotic by the transfer o f a

phosphate group from A T P (adenosine triphosphate), have also been identified in soil-dwelling

antibiotic-producing actinomycctes with sequence homology to the enzymes found in clinical

pathogens (Davies. 1994). fo r these antibiotic producers, resistance likely evolved as a means o f

protection.

I he use of antibiotics in therapeutic treatments or as growth promoters and field cultivation o f

some gcncticallv modi lied plants are suspected to increase the risk o f antibiotic resistance gene

dissemination (Sandrinc el a l., 2008). Several commercial genetically modified plants contain

antibiotic resistance genes that are still under the control of bacterial promoters as remnants o f

i !k bacterial vectors used to construct the G M Ps. Ihese former bacterial genes could be

transferred more easily than other plant genes to soil bacteria because o f a high degree o f

homology facilitating recombination in potential bacterial recipients (W ilke cl al., 2005).

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2.2 Ju s tif ic a tio n

Studies are needed to determine the true risk of enteric bacterial infections related to soil. Among the studies that need to be carried out are: the survival of enteric microorganisms in different types of soil, the ability of different types of soils to either protect or inactivate pathogenic microorganisms, and the development of methods for the detection and quantification of enteric bacteria in soils and risk assessment. According to Santamaria and Toranzos (2003), data concerning the role of soil as a vector or reservoir of enteric bacterial infections for humans and animals are not readily available and in the absence of the data, it would be challenging to carry out risk assessment studies to determine the danger of the presence of enteric microorganisms in soil. Microorganisms present in soil, may eventually end up in the water or air as a result of run­off and wind, hence the role of soil when carrying out studies on enteric diseases cannot be overlooked. Most studies focus on the role of either water or food as the source of the pathogens during enteric disease outbreaks (Santamaria and Toranzos, 2003). However, it may be easier to detect the pathogens when standardized methods are developed, if soil is in fact an important source of microorganisms. It is also important to compare the drug sensitivity profile of these enteric soil bacteria, with those of clinical isolates in order to assess the risk of antimicrobial resistance and to find out whether environmental pressure caused them to be resistant to the antibiotics.

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2.3 H ypo thesis

There is a difference in antibiotic susceptibility of environmental and clinical bacterial pathogens in Nairobi.

2.4 Objectives2.4.1 Overall objective

• To isolate and determine antibiotic sensitivity of the enteric bacteria from soil samples collected around Nairobi, compared to clinical isolates.

2.4.2 Specific objectives

1. To isolate and identify enteric bacteria from the soil samples in Nairobi and its environs2. To determine the antibiotic sensitivity of the isolated enteric bacteria3. To compare the antibiotic sensitivity profile of the isolated enteric bacteria with those

from clinical isolates from the laboratory database.

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3.0 C H A P T E R T H R E E

3.1 Materials and methods3.1.1 Study site

The study was carried out at the Kenya Medical Research Institute at the Center for Microbiology Research (CMR) at the Mycology Opportunistic Infections laboratory in Nairobi. The soil samples were drawn from a 30 kilometer diameter circular area centered at the Kenya Medical Research Institute in Nairobi, Kenya. The study was expected to provide samples that are representative of Nairobi metropolitan and its environs. The study site is a high altitude area of around 1660 meters above sea level and it has a moderate climate with maximum temperature of 28° Centigrade and minimum temperature of 11° Centigrade (World Travel, 2012). It receives rainfall ranging from 15 millimeters to 200 millimeters annually (World Travel, 2012). It is located in Latitude 1° 17’S and Longitude 36°48'E (Maps of world, 2012).

Figure 1: Map of Nairobi.-the star on the map shows the location of KEMRI, CMR. (Courtesy of Maps of world, 2012)

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3.1.2 Sam pling

The GPS coordinates were awarded random numbers then simple random sampling method was used to select the random sites. In the event that the random site was unavailable due to access restrictions or lack of an appropriate site (example stones), the closest available site was used and the GPS coordinates recorded. Simple random sampling system was also used to select the four drugs, from the drugs available for susceptibility testing for Enterobacteriaceae i.e. Ampicillin, Cefuroxime, Gentamycin, Ciprofloxacin, Gentamycin, Naldixic, Co-trimoxazole, Chloramphenical, Cefotaxime and Ervthromcin. The scientific calculator was used to generate the random numbers that were used to sample the sites and the drugs, using the kRan #’ function found on the calculator. The drugs selected were Ciprofloxacin, Gentamycin, Cefotaxime and Chloramphenical.

3.1.3 Sample size and its justification

A total of 236 soil samples were collected in this study (an average of one sample per four square kilometers). A 30- kilometer diameter region centered at KEMRI, CMR Nairobi was sampled. This region encompasses 900 square kilometers. In order to provide an accurate representation of this area, one sample every 5 square kilometers - or at least 180 soil samples was needed. In order to make sure of an appropriate number of samples, 236 soil samples were collected to cater for some site that will not be accessible, such as the airport or military bases.

3.1.4 Sample collection

Once at the sample site, the GPS coordinates was recorded and sterile metal scoop was used to scoop some soil from the randomly selected site, and then placed in sterile paper bag and placed in a transportation box to the laboratory. The samples were then named starting with the zone (similar region) from which a samples was gotten followed by the sample number from that zone. For example sample 79.2/2 represents zone 79.2 and 2nd sample from that zone.

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3.2 Laboratory methodology3.2.1 Sample processing and identification of enteric bacteria

Ten grams of each of the soil samples was serially diluted in sterile distilled water in ratio 1:10, and then incubated for around two hours at 30° Centigrade. After incubation, a loopful of the diluted soil sample was inoculated on Muller-Hinton agar (HiMcdia Lab, India) plates, and then incubated overnight at 30° Centigrade. After incubation, the colony characteristics was observed so that the yeast colonies were avoided, and then the selected colonies were Gram stained and observed under the microscope under high power objective lens with aid of oil emulsion. The rod shaped Gram negative bacteria were sub-cultured on Muller-Hinton agar to get purity isolates. The Gram negative bacteria were most likely represent the enteric bacteria and their corresponding colonies were be inoculated onto Analytic Profile Index Kit (API 20E, BioMereiux, France), and then incubated overnight at 30°Centigrade. After incubation, the API 20E was scored based on various biochemical reactions such as Hydrogen Sulphide production, various amino acid and carbohydrate reaction among others, that aided in identification of the enteric bacteria. The color reactions were observed (some with the aid of added reagents such as JAMES reagent), and the reactions (plus the oxidase reaction done separately) were converted to a seven-digit code which were read on the manufacturer's manual to show the corresponding enteric bacteria species.

3.2.2 Antibiotic susceptibility testing for the enteric bacteria

The identified enteric bacteria was be streaked onto a new Mueller-Hinton agar plate to get a purity plate. 4-5 isolated colonies were picked and inoculated into 5mL normal saline and emulsified uniformly. The turbidity was adjusted to that of 0.5 McFarland (Approximately 1.5 *10xCFU/mL) and the inoculum swabbed on the entire surface of Muller-Hinton agar (HiMedia Lab. India) plates three times while rotating the plate 60° between streaking to obtain uniform inoculation. The plates were then allowed to stand at room temperature for about three minutes

16

to allow any surface moisture to be absorbed before application of the drug discs. The antibiotic discs were applied on the surface of the plates using a sterile forceps, and then incubated at 30° Centigrade overnight. The antibiotics tested and their concentrations are indicated in table 1.

The zones of inhibition were then compared to those of enteric bacteria isolated from clinical samples using the data from tests done in the laboratory, during the same duration of the study. Antibiotic susceptibility testing was done according Clinical and Laboratory Standards of 2007.

Sterile water with no soil sample was used as a method of negative control in the study. Also Escherichia coli ATCC 25922, was used as standard control strain to check for the efficacy of the antibiotics.

3.2.4 Variables

The characteristics to be measured in the study were measured numerically such as zones of inhibition in millimeters, and number of enteric bacteria isolated. The dependent variables were the zones of inhibition, and the number of isolated enteric bacteria. The independent variables included location of the soil and the drugs used in susceptibility testing.

3.3 Experimental design

Randomized Block design was used in the study where, the sites that were homogenous in nature (for example if they are close to each other according to their coordinates) was grouped together to form the blocks. The four randomly chosen drugs formed the treatments. Within each block, after only with control randomized experiment was applied.

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4.0 C H A P T E R FO U R4.1 Results

A total of 236 soil samples were inoculated on Mueller-Hinton agar plates and 52/236 [22%] of the inoculated samples were positive for bacteria. Of the 52 samples that grew bacteria, 38 [73%] were Gram negative bacteria of which they were inoculated onto the Analytic Profile Index (API 20E) kit for identification. Of the 38 Gram negative bacteria isolated. 17 [44%] were identified as Proteus salmonicida, 11 [28%] were Myroides spp, 5 [ 17%] were Pseudomonas putida. Pseudomonas aeruginosa and Stenotrophomonas maltophila were two each [5% each], and one Alcaligenes spp [2%] was identified using the API 20E kit. ). The other isolated Gram negative bacteria (Myroides spp, Pseudomonas putida, Pseudomonas aeruginosa,Stenotrophomonas maltophila and Alcaligenes spp) are not members o f the Enterobacteriaceae family though some maybe isolated from the intestinal tract such as Pseudomonas and Alcaligenes spp.

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4.1.2 Gram stain reactions of isolates that grew on Mucller-Hinton agar plates

In the growth on Mueller- Hinton agar, 52 isolates were bacteria out of the 236 soil inoculations, representing 22% prevalence of bacteria in the soil. When the 52 isolates were Gram stained, most of them were Gram negative rods (n=38) as shown in table one below.

Table 1: Gram stain reactions of isolatesIsolate Gram Cell shape58.3/5 - Cocci10.2/5 - Cocci1.9/20 +• Cocci60.1/48 - Rod1.9/19 - Cocci60.1/13 - Cocci93.1/1 - Cocci60.1/57 - • Rod93.2/3 - Cocci60.1/51 - Rod60.1/66 - . Rod60.1/64 - Rod50.3/60 + Cocci1.9/20 - Rod79.3/3 - Rod69.2/6 Rod71.2/10 - Rod79.3/4 - Rod62.3/5 - Rod69.2/8 - Rod2.3/15 - Rod62.3/2 • Rod71.2/10 - Rod

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Table 1 continued2.3/8 Rod12.1/8 - Rod12.1/2 - RodIMIS - Rod79.:2/17 - Rod79.2/6 - Rod9.2/1 - Rod74b.2/15 - Rod60.1/16 - Rod60.1/1 - Rod40.1/3 - Rod68.3/12 - Rod29.2/10 - Rod40/3 - Rod76.1/17 - Rod68.3/12 - Rod4773 - Rod53.2/34 + Cocci53.2/15 + Rod80.3/1 + Cocci49. lb/13 - Rod53.2/31 + Rod53.2/33 + Rod78.3/12 + Rod5.1/30 + Rod5.3/5 + Rod60.1/5 Rod53.5/5 + Rod5.2/9 + Rod

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Some isolates were a mixture of Gram positive rods and cocci as shown in figure two below. Some Gram negative rods (figure three) appeared large with terminal endospores, a survival tool in soil bacteria, while some were tiny rods. Other Gram positive rods appeared in chains, a characteristic of some soil bacteria such as Clostridium spp.

Figure 2: Gram stain - Gram positive cocci and rods (xlOO oil immersion)

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The Analytic Profile Index 20E kit was used to identify the bacterial colonies. The plates that gave the Gram negative reaction were traced back and a pure colony on each plate was inoculated on the API 20E kit. The API kit gave various biochemical reactions (as shown in figures four and five) such Hydrogen Sulphide reaction that gave a black color.

NB:

The biochemical reactions on API kit are:

Amino acids arginine (ADH), lysine (LDC) and ornithine (ODC). Decarboxylation is is shown by an alkaline reaction (red color of the particular pH indicator used)

Carbohydrates glucose, mannitol, inositol, sorbitol, rhamnose, sucrose, melibiose, amygdalin and arabinose. Fermentation is shown by an acid reaction (yellow color of indicator).

Tryptophan deaminase (TDA) gives a deep brown color with the addition of ferric chloride; positive results for this test correlate with positive phenylalanine and lysine deaminase reactions which are characteristic of Proteus, Morganella and Providencia.

Urea (URE) reaction gives an orange-red color.

ONPG (Orth-nitrophenyl-b-D-galactopyranoside) positive reaction gives a yellow colouration, while citrate (CIT) reaction gives a blue color.

Voges Proskauer (VP) gives a pink coloration within ten seconds of adding VP1 and VP2 reagents if the reaction is positive.

Hydrogen sulfide production (H2S) and gelatin hydrolysis (GEL) result in a black color throughout the tube (Willey et al., 2008).

4.1.3 Colour reactions on API 20E kit after incubation and addition o f reagents

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r

Figure 4: Various API reactions - positive reactions on ONPG (Orth-nitrophenyl-b-D- galactopyranoside), CIT (Citrate), URE (Urea) and TDA (Tryptophane DiAminase) on Pseudomonas aeruginosa.

f mmA

FiRure5: Various API reactions - positive reactions on ADH (Arginine DiHydroiase), TDA, VP(̂ °ges Proskauer) and GEL (Gelatinase) on Proteus salmonicida .NB:

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4.1.4 Distribution of the enteric soil isolates

Most of the isolates that yielded Proteus salmonicida bacteria were found to occur at the North

Western and South Eastern part o f Nairobi, as shown on the map below.

Figure 6: Distribution o f enteric soil isolates- the red stars indicate the soil sample zones that

yielded P. samlonicida.

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Most of the isolates from soil samples demonstrated the least resistance to Chloramphenicol, with zones of inhibition ranging from 16 mm to 41 mm. 1 ligh resistance levels were observed in Cefotaxime with zones if inhibition at 6mm at shown in table two. The zones of inhibiton observed in Ciprofloxacin and Gentamycin, ranged from 22 mm to 27 and 24 mm to 30 mm respectively. The Proteus species from clinical isolate (otitis-ear infection) demonstrated the least resistance to Cefuroxime, while it was most resistant to Chloramphenicol antibiotic as shown in table three.

Table 2: Antibiotic sensitivity profile of Proteus salmonicicla from soil

4.1.5 Antibiotic sensitivity profile of the soil and clinical isolates

Isolate Zones of inhibition in millimeters

C iprofloxacin Gentam ycin Cefotaxime Chloram phenicol

40.1/3 26 27 6 1679.2 27 28 6 362.3/8 24 25 6 3629.2/10 24 28 6 3047.3/3 24 26 6 3474B.2/2 23 25 6 3474B.2/15 26 30 6 2879.2/6 25 28 6 2062.3/2 27 30 6 409.2/9 22 25 6 3379.3/12 26 30 6 3079.3/7 26 29 6 372.3/15 25 28 6 4179.2/9 33 30 6 4179.3/1 24 24 6 369.2/1 25 26 6 3271.2/5 22 25 6 37

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Table 3: Antibiotic sensitivity profile of Proteus species from clinical isolate (otitis)Drug Zones o f inhibition in mm

Ampicillin 13

Cefiiroxime 24

Gentamycin 12

Ciprofloxacin 21

Co-trimoxazole 14

Chloramphenical 11

Cefotaxime 18

Erythromycin 18

The zones of inhibition were compared to the Clinical Laboratory Standard Institute breakpoints of 2007, as shown in table four below.

Table 4: Antibiotic amounts impregnated on disc and breakpoints for enteric bacteria

Drug Am ount impregnated on disc (pg) Breakpoints(m m )

Ampicillin 2 14-16 I

Cefuroxime 30 15-22 I

Gentamycin 10 13-14 1

Ciprofloxacin 5 16-20 1

Nalidixic 30 14-18 I

Co-trimoxazole 50 11-15 I

Chloramphenical 50 13-171

Cefotaxime 30 15-17 1

Erythromycin 15 >22 R

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Proteus species isolated from otitis was used to compare the antibitiotic sensitivity profile with the Proteus isolated from the soil samples. Proteus salmonicida showed to be generally more sensitive to the antibiotics, compared to the Proteus from otitis as shown on the graph in figure eight. However P. salmonicida was resistant to Cefotaxime antibiotic while the Proteus from clinical specimen was sensitive to the same antibiotic.

■ otitis

■ soil

Antibiotics

Figure 8: comparison of inhibition zones of Proteus species from the soil and clinical samples - the graph shows higher zones of inhibition of soil samples than of the clinical sample (otitis). NB: CIP (Ciprofloxacin), GM (Gentamycin), CTX (Cefotaxime), CHL (Chloramphenical).

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4.2 D ata a n a l y s i s

Data were entered in Excel spread sheets. Analysis of Variance (ANOVA) was used to test the effect of the antibiotics on the enteric bacteria. Chi-square test was used to test the association between the number of enteric bacteria isolated and the sample site zones. It was also be used to test any association between the number of enteric bacteria isolated and the seasons. Student T- test was used to test if there was difference between the zones of inhibition of the four antibiotics, between the soil samples and the clinical samples of the enteric bacteria. All were tested at p< 0.05. SPSSR version 16.0.2 software was used for the statistical analysis

One-way Analysis of Variance was used to test the effects of the antibiotics on P. scilmonicida. Significant effect was observed overall (F=2.729. d.f=3 at p<0.005) hence the null hypothesis of was rejected. In Cefotaxime however, no effect was observed in since there was no variation of the zones of inhibition from the mean. Ciprofloxacin showed the most effect among the four antibiotics since it had the greatest variation of zones of inhibition from its mean from its mean.

Student's t-test was used to compare the zones of inhibition between the soil and clinical isolates. Significant results were also observed (t=3.873, d.f=3 at p<0.()5). The null hypothesis of was rejected meaning there was a difference in the zones of inhibition between the two sources ofisolates.

Chi-Square test was used to test association between the soil samples zones and number of enteric bacteria isolated: X2=2.364, df=2, p<0.05. The results show a higher significance value at p<0.05, hence the null hypothesis was rejected meaning there was association between the number of enteric bacteria isolated and the soil sample zones.

29

J0 CHAPTER FIV E

(i pMtntwn

xdmi ulmonmJa was the only enteric bacteria isolated out of the „x tifBm ^ . - n ,* * * * * * ™ Analytical Profile Index (API 20E). U th o u c ,.... . can he a«,u.r*J tnan

contaminated fish It is pathogenic lor fish, causing disease known , , „ mWK>1,,.........„„d tt toil temples (n=236) inoculated onto Mueller II,..ton agar. 17 grew /• gfcsum. which represents a prev alence of 7.2% o f enteric bacteria in the v.,1 I hi. k>w *ulcnce compares well with the study done by Ntabo el at.. (2010) who were analcred wenafrom soil samples in Juja and Kakamega forest Out of the 137 pure iv»lntc% ihcv •palatesofSerratia marcescens were the only enteric bacteria, which represent! a pccval. ru.

Previous studies show that enteric bacteria find it difficult to compete with tin n.itm.i

micro-biota due to the low am ount o f nutrients in that ecosystem (Burton et a! I 7 |

xx results contrast with the study done by Burgos et al.% (2004) investigating the pcescruc «*i

* :njg-resistant enteric bacteria in dairy farm topsoil. I hey isolated 102 enteric KkI. ru lr - inn tarm top soils and 9 isolates were obtained from adjacent roadsides (non-dairy voil)

v Jrgc number of enteric bacteria isolates could be attributed to the cow dung that wa prevent

7c dairy farms. The enteric bacteria they isolated were: ( 'itrobacter hraakn (7 trohm ter ‘̂ n ( itrobacter koseri, Enterobacter gergoviae, Enterohacter laylorae. Escherichia o>h

dla pneumonia, Proteus mirabilis, Proteus vulgaris. Pseudomonas aeruginosa ’•d»monas fluorescens, Shigella spp. and Serratia plymuthic

Profile Index kit (A P I 20E ) provided a direct way o f identifying enteric bacteria and

c wtidious Gram negative bacteria. O f the 38 Ciram negative isolates inoculated

‘ < provided identities as Proteus salmonicida. M yroidcs spp Pseudomonas puttda * *onas aeruginosa, Stenotrophom onas maltophila and Alcaligenes spp O f Ihcsc Gram

**** bacteria, Proteus salm onicida was the only enteric bacteria targeted. The enteric

^ gave positive reactions on A D H (A rg in ine D iHydrolase), I D A ( I ryptophanc

~ VP (Voges Proskauer) and G E L (Gelatinase). According to W illey et aU ' 3,1 enzyme present in the enteric bacteria that is responsible f«'i releasing

Canine during the urea cycle. T D A is an enzyme used by bacteria to produce X1M30

deaminaton of Trypyophan. Positive VP test meant that the enteric bacterium was able to produce acetion in the culture (Willey et cil., 2008). Gelatinase is an enzyme used by the enteric bacteria to break down protein gelatin from collagen (Ryan and Ray, 2004). However, it is noted that Escherichia coli a typical enteric bacteria, was not isolated in this study. It may have been present in the soil samples but in a lower concentration than the isolated bacteria.

Antibiotic sensitivity profile was also performed on the enteric bacteria. Disc diffusion method was used according to Clinical and Laboratory Standards (CLSI) o f 2007. Four randomly chosen antibiotics were used (Chloramphenicol, Cefotaxime, Ciprofloxacin and Gentamycin) and P. salmonicida was most sensitive to Chloramphenicol while resistant to Cefotaxime antibiotics. This could mean the bacteria had developed resistance to the antibiotic due to environmental exposure, as the Cefotaxime antibiotic showed sensitivity to standard Escherichia coli (ATCC 25922) that was used as a method of control to show the efficacy of the four antibiotics used. Soil bacteria have been shown to develop antibiotic resistance to some of the antibiotics, as a means of survival since they are exposed to a myriad of other antibiotic producing bacteria e.g. Actinomycetes (Davies, 1994). Although the exact method of resistance could not be established in this study, resistant gene transfer through plasmids may be implicated considering the prevalence of plasmids in soil bacteria (Wilke et al., 2005). Other mechanisms ofbacterial resistance such as efflux pumps and porin mutations could also have occurred. . In the study done by Burgos et al., (2004) they used 22 isolates for further study based on medical importance or high frequency of occurrence. They used Minimal Inhibitory Concentration method (MIC) to performed antibiotic sensitivity profile on the 22 isolates. The antibiotics they used were Chloramphenicol, Penicillin G, Nalidixic acid and Tetracycline. Most of their isolates showed higher resistant levels to Chloramphenicol which was contrary to this study, as the same antibiotic showed highest sensitivity. Most of the isolates demonstrated the least resistance to Nalidixic acid and Tetracycline. The isolates also showed highest resistance to Penicillin G.

In data analysis, one-way Analysis Of Variance (ANOVA) was used to test the effect of the treatment (the four antibiotics) on the replicates/subjects (isolates). Ciprofloxacin was found to have greatest an effect within and between groups. This means it was the most effective drug among the four antibiotics chosen. Chloramphenicol showed the highest zones of inhibition on paper but upon analysis it was not as effective as Ciprofloxacin, since its zones of inhibition

31

were not that varying from the mean. Cefotaxime did not show any effect within and between the groups as the antibiotic gave the same zones of inhibition of 6 millimeters hence there was no difference from the mean. Thus the antibitotic was the least active among the four chosen. In summary, the most effective antibiotics were Ciprofloxacin followed by Chloramphenicol. Gentamycin and Cefotaxime was the least effective one. Burgos et al., (2004) used the non- parametric one-tail Wilcoxon-Paired sample test to test the effects of salicylate on the antibiotic resistance to Chloramphenicol, Nalidixic acid, Penicillin, and Tetracycline. The one-tailed test was used, as the priori hypothesis was that salicylate increased antibiotic resistance in the isolates. Results were considered significant at P <0.05.

Some of the challenges encountered during the study were expenses in terms of the API 20E kit and its reagents were quite costly. Also we encountered a challenge in the incubation temperature of the soil isolates, in that at first the inoculums was not showing any growth on the Mueller-Hinton agar, which was due to the set temperature of 37°C. Since the target isolates were environmental in nature, they required a lower incubation temperature of about 30°C. Also delays were encountered in the supply of the materials to be used in the study.

32

5.2 Conclusion

Soil may be a significant a reservoir for the enteric bacteria contributing to antibiotic resistance as indicated by Proteus salmonicida with resistance to Cefotaxime antibiotic, compared to Proteus species from the clinical sources which was sensitive. This study suggests that the soil serves as an under-recognized source of resistance with the potential to reach clinical isolates of the bacteria. The study however showed a low prevalence of enteric bacteria from the soil sampled.

5.3 Recommendations

Cefotaxime antibiotic should not be used for antibiotic sensitivity testing of Proteus salmonicida isolated from soil samples. However Chloramphenicol, Ciprofloxacin and Gentamycin can form good antibiotic regiments against the bacterium, as it is quite sensitive to the antibiotics. There is a possibility that low levels of antibiotic resistance might persist on topsoil, suggesting the need for topsoil analysis of antibiotic residues as well as the establishment of surveillance programs for antibiotic resistant bacteria in soil. Other antibiotics other than the ones used in this study, could be tested against enteric soil bacteria to determine if any resistance could be exhibited.

33

6.0 REFERENCES

Bannerman T.L. (2003). Manual o f C linical Microbiology. 8lh ed. American Society for Microbiology Press.

Barbosa T., and Levy S.B. (2000). The impact of antibiotic use on resistance development and persistence. Drug Resistance, 3:303-311

BioMeriex France (2010). Analytic Profile Index 20E.

Burgos J.M., Ellington B.A., Varela M.F. (2004). Presence of Multidrug-Resistant Enteric Bacteria in Dairy Farm Topsoil. Journal o f Dairy Science. New Mexico, 88:1391-1398

Cilimburg A., Monz C., Kehoe S. (2000). Wildland recreation and human waste: a review of problems, practices, and concerns. Environmental Management. 25:587-598

Clinical and Laboratory Standard Institute (2007). Performance standards for antimicrobial susceptibility testing. CLSI approved standard M100-S17. Clinical and Laboratory Standards Institute. Wayne, PA.

Davies J. (1994). Inactivation of antibiotics and the dissemination of resistance genes. Science, 264:375-382.

Davies J. and Benveniste R. (1973). Aminoglycoside antibiotic-inactivating enzymes in actinomycetes similar to those present in clinical isolates of antibiotic-resistant bacteria. National Academic Science, 70:2276-2280.

34

Craun G.F. and Calderon R.L. (1996). Microbial risk in groundwater systems: epidemiology of waterborne outbreaks. Groundwater Foundations. Boston.

Environmental Protection Agency (2002). Land application of biosolids. (Office of inspector general status report 2002-S-000004) http://www.epa.gov/oigearth/eroom.htm [accessed June, 2011].

Feachem G.G., Bradley D.J., Garelick H., Mara D.D. (1983). Sanitation and disease—health aspects o f excreta and wastewater management. (World bank studies in water supply and sanitation 3) Wiley, Chichester.

Gerba C.P. (1996). Microbial pathogens in municipal solid waste. Microbiology o f solid waste. CRC Press, New York, pp 155-173.

Gerba C.P. and Bitton G. (1984). Microbial pollutants: their survival and transport pattern to groundwater. Groundwater pollution microbiology pp 39-54.

Graham S.M., Molynex E.M., Walsh L.A.. Cheesebrough J.S., Malcom E.M., Hart H.A. (2000). Non Typhoidal Salmonella Infections of Children in Tropical Africa. Pediatric Disease Journal, 19:1189-1196.

Guarner F. and Malagelada J.R. (2003). Gut flora in health and disease. Lancet, 361: 512-519.

Hay C.J. (1996). Pathogen destruction and biosolids composting. 2nd ed. BioCycle.

35

JohannaS., Gary A. (2002). Enteric pathogens and soil: a short review. International Microbiology, 6:5-9.

Kelly Struble (2002). Resident Physician, Department of Internal Medicine, University of Oklahoma College o f Medicine.

Keswick B.H. (1984). Sources of ground water pollution. Groundwater pollution microbiology. Wiley, New York, pp 39-54.

Knight D.J. and Girling K.J. (2003). Gut flora in health and disease. Lancet, 361:1831.

Martinez, J.L. and Baquero F. (2002). Mutation frequencies and antibiotic resistance. Antimicrobiol Agents Chemotherapy, 44: 1771-1777.

Morbidity and mortality (1997). Outbreaks of Escherichia coli 0157:H7 and cryptosporidiosis associated with drinking unpasteurized apple cider. Morbid Mortal Wkly Rep, 46:4-8.

Ntabo R., Boga H., Muigai A., Mwirichia R. (2010). Isolation and characterization of bacteria isolates from soil feeding termites and soil from Juja and Kakamega forest in Kenya. http://elearning.jkuat.ac.ke/journals/ojs/index.php/jscp/article/view/675 [accessed July, 2012].

Organic Materials Review Institute (1998). Use of manure, compost and sewage sludge in the December 1997 proposed national organic program. http://www.OMRI.org/USDA.html [accessed January, 2011].

36

Owens W.E., Nickerson S.C., Boddie R.L., Tomita G.M., Ray C.H. (2001). Prevalence of mastitis in dairy heifers and effectiveness of antibiotic therapy. Journal o f Dairy Science, 84: 814-817.

Paterson D.L. (2002). Serious infections caused by enteric gram-negative bacilli—mechanisms of antibiotic resistance and implications for therapy of gram-negative sepsis in the transplanted patient. Journal o f Respiratory Infection, 17: 260-264.

Parish M.E. (1997). Public health and non-pasteurized fruit juices. Critical Review Microbiology, 23:109-119.

Plym-Forshell L. and Ekesbo I. (1993). Survival of Salmonella in composted and not composted animal manure. Journal o f Veterinary Medicine, 40:654-658.

Randal L. and Woodward M.J. (2001). Multiple antibiotic resistance {mar) locus in Salmonella enterica Serovar Typhimurium DTI04. Applied Environmental Microbiology, 67:1190- 1197.

Rosen B.H. (2000). Waterborne pathogens in agricultural watersheds. USDA-Natural Resources Conservation Service, http://www.wcc.nrcs.usda.gov/watershed/projects [accessed January, 2012].

Ryan K.J. and Ray C.G. (2004). Sherris Medical Microbiology. 4th ed. McGraw Hill. ISBN 0 8385-8529-9.

37

RxPG news (2010). “Dirt may be a key to how bacteria that infect humans develop a resistance to anitbitic drugs”, www.mcmaster.ca [accessed April, 2011].

Sandrine D., Herve S., John P., Elisabeth N., Dominique B., Pascal S. (2008). Antibiotic resistant soil bacteria in transgenic plant fields. PNAS, 105:3957-3962

Santamaria J. and Toranzos G.A. (2003). Enteric pathogens and soil: a short review. International Microbiology. 6:5-9.

Satish Gupte (1999). Short Text Book o f Medical Microbiology. 7th ed. Jaypee Publishers, India, p. 628-931.

Scandura J.E. and Sobsey MD. (1997) Viral and bacterial contamination of ground water from onsite sewage treatment systems. Water Science Technology. 35:141-146.

Shanahan F. (2002). Best Practice & Research Clinical Gastroenterology. Vol. 16, p. 915-931.

Shuval HI., Adin A., Fattal B„ Rawitz E., Yekutiel P. (1986). Wastewater irrigation in developing countries, health effects and technical solutions. UNDP project management report 6. World Bank, Washington, D.C., pp 27-57.

Sinton L.W. (1986). Microbial contamination of alluvial gravel aquifers by septic tank effluent. Water Air Soil Pollution. 28:407-425.

38

Strauss M. (1994). Health implications of excreta and wastewater use. Hubei environmental sanitation study, 2nd workshop. Hubei, Wuhan.

Toze S. (1997). Microbial pathogens in wastewater. Literature review’ for urban water systems. CSIRO land and water technical report, http://www.clw.csiro.au/publications/technical/trl 97 [accessed June, 2011].

Venkatesan. P. (2001). Essentials o f Medical Laboratory Technology. 2nd Ed. Swathi Publishers, India, p. 13-15.

Wilke M.S., Lovering A.L., Strynadka N.C. (2005). beta-Lactam antibiotic resistance: a current structural perspective. Currrent Opinion Microbiology, 8:525-533.

Willey J., Sherwood L„ Woolverton C. (2008). Prescott. Harley & Klein Microbiology, l'" Ed. McGraw Hill.

Yeager J.G. and O'Brien RT. (1979). Enterovirus inactivation in soil. Applied Environmental Microbiology, pp. 694-701.

Yeager J.G. and Ward RL. (1981). Effects of moisture content on long term survival and regrowth of bacteria in wastewater sludge. Applied Environmental Microbiology,41:1117-1122

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