Download Antibiotic Sensitivity Profile Of Enteric Bacteria Isolated From Soil

a n t ib io t ic s e n s it iv it y p r o f il e of e n t e r ic b a c t e r ia
ISOLATED 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 Ia 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.... 3 o
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 QJ5 '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....... ~
...................................
p ate....................... ...................................................................
II
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.
IV
Abstract
Soil 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 of 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.
v
TABLE OF CO NTENTS
Declaration..........................................................................................................................................ii
Dedication...........................................................................................................................................iii
Acknowledgement.............................................................................................................................. iv
Abstract.............................................................................................................................................. v
1.1 Introduction................................................................................................................................ 1
2.0 CHAPTER TWO.........................................................................................................................5
2.1 Literature review............................... !...................................................................................... 5
2.1.1 Enteric bacteria.....................................................................................................................5
2.1.2 Isolation, identification and drug susceptibilitytesting of enteric bacteria.......................6
2.1.3 Sources of soil contamination by enteric pathogens..........................................................7
2.1.4 Fate of enteric bacteria in the soil....................................................................................... 9
2.1.5 The role of soil as a reservoir for bacteria in contributing to antibtiobitc resistance ... 10
2.2 Justification............................................................................................................................... 12
2.3 Hypothesis.................................................................................................................................13
2.4 Objectives................................................................................................................................13
2.4.1 Overall objective.................................................................................................................13
2.4.2 Specific objectives..................................;.........................................................................13
3.0 CHAPTER THREE..................................................................................................................14
3.1 Materials and methods..............................................................................................................14
3.1.1 Study site................................................................................................
14
3.1.2 Sampling.............................................................................................................................15
3.1.3 Sample size and its justification........................................................................................15
3.1.4 Sample collection................................................................................................................15
3.2 Laboratory methodology.......................................................................................................... 16
3.2.1 Sample processing and identification of enteric bacteria................................................16
3.2.2 Antibiotic susceptibility testing forthe enteric bacteria...................................................16
3.2.4 Variables................................................. ;.........................................................................17
3.3 Experimental design..................................................................................................................17
VI
4.0 CHAPTER FOUR........................................................................................................................18
4.1 Results........................................................................................................................................18
4.1.3 Colour reactions onAPI 20E kitafter incubation and addition of reagents...................22
4.1.4 Distribution of the enteric soil isolates...............................................................................24
4.1.5 Antibiotic sensitivityprofile of the soil and clinical isolates........................................ 25
4.2 Data analysis.............................................................................................................................29
5.0 CHAPTER FIVE....................... ............................................................................................... 30
5.1 Discussion.....................................................................................................i......................... 30
5.2 Conclusion.................................................................................................................................33
5.3 Recommendations.................................................................................................................... 33
6.0 REFERENCES.......................................................................................................................34
APPENDIX.......................................................................
40
vii
LIST OF TABLES
1: Gram stain reactions of isolates
2: Antibiotic sensitivity profile of Proteus salmonicida from soil
19
25
3: Antibiotic sensitivity profile of Proteus species from clinical isolate
4: Antibiotic amounts impregnated on disc and breakpoints for enteric bacteria
26
26
viii .
*
LIST OF FIGURES
1: Map of Nairobi
2: Gram stain
3: Gram stain
4: Various API reactions
5: Various API reactions
6: Distribution of enteric soil isolates
7: Antibiotic sensitivity
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21
21
23
23
24
27
8: Comparison of inhibition zones of Proteus species from the soil and clinical samples 28
IX
1.0 C H A P T E R O N E
1.1 Introduction
Humans 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 antibioticproducing 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 antibioticproducing microorganisms (Randal and Woodward, 2001). Whether naturally occurring or
2
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.
3
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 TW O
2.1 Literature review
2.1.1 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 Gram­
negative. 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 Escherichia
Genus: Escherichia, Eciwardsiella, Citrohacter, Salmonella, Shigella.
Tribe 2. Klebsiellae
Genus: Klebsiella, Enterobacter, Hafnia, Serratia.
Tribe 3 Proteae
Genus: Proteus
Tribe 4: Erwiniae
Genus: Erwinia
Tribe 5: Yersinae
Genus: Yersinia
5
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-LactoseDesoxycholate (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
6
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
1here 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
7
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).
8
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,
9
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
10
to aminoglycosides is mediated by antibiotic-moditying enzymes such as kinases that confer a
high level o f resistance.
However, aminoglycoside kinases, enzymes that m odify 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 enzym es found in clinical
pathogens (Davies. 1994). f o 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
dissemination (Sandrinc
plants are suspected to increase the risk o f antibiotic resistance gene
el al ., 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 P s.
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
11
cl al.,
2005).
2.2 Justificatio 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 run­
off 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.
12
2.3 H ypothesis
There is a difference in antibiotic susceptibility of environmental and clinical bacterial pathogens
in Nairobi.
2.4 Objectives
2.4.1 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 environs
2. To determine the antibiotic sensitivity of the isolated enteric bacteria
3. To compare the antibiotic sensitivity profile of the isolated enteric bacteria with those
from clinical isolates from the laboratory database.
13 •
3.0 C H A P T E R T H R E E
3.1 Materials and methods
3.1.1 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)
14
3.1.2 Sampling
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.
15
3.2 Laboratory methodology
3.2.1 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.
17
4.0 C H A PT E R FO U R
4.1 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 of the Enterobacteriaceae
family though some maybe isolated from the intestinal tract such as Pseudomonas and
Alcaligenes spp.
18
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 isolates
Isolate
58.3/5
10.2/5
1.9/20
60.1/48
1.9/19
60.1/13
93.1/1
60.1/57
93.2/3
60.1/51
60.1/66
60.1/64
50.3/60
1.9/20
79.3/3
69.2/6
71.2/10
79.3/4
62.3/5
69.2/8
2.3/15
62.3/2
71.2/10
Gram
+•
•
.
+
-
Cell shape
Cocci
Cocci
Cocci
Rod
Cocci
Cocci
Cocci
Rod
Cocci
Rod
Rod
Rod
Cocci
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
•
19
Table 1 continued
2.3/8
12.1/8
12.1/2
IMIS
79.:2/17
79.2/6
9.2/1
74b.2/15
60.1/16
60.1/1
40.1/3
68.3/12
29.2/10
40/3
76.1/17
68.3/12
4773
53.2/34
53.2/15
80.3/1
49. lb/13
53.2/31
53.2/33
78.3/12
5.1/30
5.3/5
60.1/5
53.5/5
5.2/9
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Cocci
Rod
Cocci
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
Rod
+
+
+
+
+
+
+
+
+
+
20
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)
21
4.1.3 Colour reactions on API 20E kit after incubation and addition of reagents
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).
22
r
Figure 4: Various API reactions - positive reactions on ONPG (Orth-nitrophenyl-b-Dgalactopyranoside), CIT (Citrate), URE (Urea) and TDA (Tryptophane DiAminase) on
Pseudomonas aeruginosa.
f mm
A
FiRure5: Various API reactions - positive reactions on ADH (Arginine DiHydroiase), TDA, VP
(^°ges Proskauer) and GEL (Gelatinase) on Proteus salmonicida .
NB:
23
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.
24
4.1.5 Antibiotic sensitivity profile of the soil and clinical isolates
Most of the isolates from soil samples demonstrated the least resistance to Chloramphenicol,
with zones of inhibition ranging from 16 mm to 41 mm. 1ligh 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
Isolate
40.1/3
79.2
2.3/8
29.2/10
47.3/3
74B.2/2
74B.2/15
79.2/6
62.3/2
9.2/9
79.3/12
79.3/7
2.3/15
79.2/9
79.3/1
9.2/1
71.2/5
Z o n e s of in h ib ition in m illim eters
C ip ro flo x a c in
G e n ta m ycin
C efotaxim e
C h lo ra m p h e n ic o l
26
27
24
24
24
23
26
25
27
22
26
26
25
33
24
25
22
27
28
25
28
26
25
30
28
30
25
30
29
28
30
24
26
25
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
6
16
36
36
30
34
34
28
20
40
33
30
37
41
41
36
32
37
25
Table 3: Antibiotic sensitivity profile of Proteus species from clinical isolate (otitis)
Z o n e s o f in h ib ition in m m
Drug
Ampicillin
Cefiiroxime
Gentamycin
Ciprofloxacin
Co-trimoxazole
Chloramphenical
Cefotaxime
Erythromycin
13
24
12
21
14
11
18
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
Ampicillin
Cefuroxime
Gentamycin
Ciprofloxacin
Nalidixic
Co-trimoxazole
Chloramphenical
Cefotaxime
Erythromycin
A m o u n t im p regnated on disc (p g)
B re a k p o in ts(m m )
2
14-16 I
30
15-22 I
10
13-14 1
5
16-20 1
30
14-18 I
50
11-15 I
50
13-171
30
15-17 1
15
> 22 R
26
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).
28
4.2 Data a n a ly s is
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 Ttest 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. SPSSRversion 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 of
isolates.
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
J0CHAPTER FIVE
(i pMtntwn
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,,.........„„
dtt toil temples (n=236) inoculated onto Mueller II,..ton agar. 17 grew /•
gfc
sum. which represents a prev alence of 7.2% of 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 sh o w that enteric bacteria find it difficult to compete with tin n.itm.i
micro-biota due to the low am ou n t o f nutrients in that ecosystem (B u rton et a! I 7 |
xx results contrast with the study d o n e by B u rg o s et al.%(2004) investigating the pcescruc «*i
* :njg-resistant enteric bacteria in d a iry farm topsoil. I hey isolated 102 enteric Kk I. r u lr inn tarm top soils and 9 isolates were obtained from adjacent roadsides (non-dairy voil)
v Jrgc number of enteric bacteria isolate s could be attributed to the cow d u ng 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
xdmi
Profile Index kit ( A P I 2 0 E ) p rovid ed a direct w a y o f identifying enteric bacteria and
c wtidious Gram negative bacteria. O f the 38 Ciram negative isolates inoculated
‘ < provided identities as Proteus salmonicida. Myroidcs spp Pseudomonas puttda
* *onas aeruginosa, Stenotrophomonas maltophila and Alcaligenes spp O f Ihcsc Gram
**** bacteria, Proteus salmonicida w a s the o n ly enteric bacteria targeted. The enteric
gave positive reactions on A D H (A rg in in e D iH yd ro la se ), I D A ( I ryptophanc
~ VP (Voges Proskauer) and G E L (Gelatinase). A cco rd in g to W ille y 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 enzym e used by bacteria to produce
X1M
^
30
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) of 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 nonparametric 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 of Clinical 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 of 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 of 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 of 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, 67:11901197.
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
39