OKWUBIE, LAMBERT
(PG / M.PHARM / 10/ 52771)
EVALUATION OF OXIDATIVE STRESS INDICATORS ON FIVE
MEDICINALPLANTS FROM CRUDE OIL POLLUTED ENVIRONMENT
DEPARTMENT OF PHARMACOGNOSY AND
ENVIRONMENTAL MEDICINE
FACULTY OF PHARMACEUTICAL SCIENCES
Digitally Signed by: Content manager’s Name
DN : CN = Webmaster’s name Ameh Joseph Jnr O= University of Nigeria, Nsukka
OU = Innovation Centre
EVALUATION OF OXIDATIVE STRESS INDICATORS ON FIVE
MEDICINALPLANTS FROM CRUDE OIL POLLUTED ENVIRONMENT
BY
OKWUBIE, LAMBERT
(PG / M.PHARM / 10/ 52771)
PROFESSOR S. I. INYA AGHA
(SUPERVISOR)
DEPARTMENT OF PHARMACOGNOSY AND
ENVIRONMENTAL MEDICINE,
FACULTY OF PHARMACEUTICAL SCIENCES,
UNIVERSITY OF NIGERIA, NSUKKA.
JANUARY 2014
i
EVALUATION OF OXIDATIVE STRESS INDICATORS ON FIVE
MEDICINAL PLANTS FROM CRUDE OIL POLLUTED ENVIRONMENT
BY
OKWUBIE, LAMBERT
(PG / M.PHARM / 10/ 52771)
BEING A PROJECT REPORT SUBMITTED TO THE DEPARTMENT
OF PHARMACOGNOSY AND ENVIRONMENTAL MEDICINE,
FACULTY OF PHARMACEUTICAL SCIENCES, UNIVERSITY OF NIGERIA,
NSUKKA IN PARTIAL FULFILMENT OF THE REQUIREMENT FOR THE AWARD
OF MASTER OF PHARMACY DEGREE.
PROFESSOR S. I. INYA AGHA
(SUPERVISOR)
DEPARTMENT OF PHARMACOGNOSY AND ENVIRONMENTAL MEDICINE,
UNIVERSITY OF NIGERIA, NSUKKA.
JANUARY 2014
ii
CERTIFICATION
Okwubie, Lambert, a postgraduate student in the department of pharmacognosy and environmental medicine; with Reg. No. PG/M.PHARM/10/52771 has satisfactorily completed the requirement for the course work and research for the Degree of Master of Pharmacy (M.Pharm) in Pharmacognosy and Environmental Medicines. The work embodied in this project is original and has not been submitted in part or full for any other Diploma or Degree of this or any other University
______
Prof S.I. Inya Agha Dr. Mrs. U.E. Odoh
Supervisor Head of Department
______
Date Date
iii
DEDICATION
This work is dedicated to GOD Almighty, my beloved wife, and sons.
iv
ACKNOWLEDGEMENT
First and foremost, I thank the LORD GOD Almighty who gave me the ability to run this program. To HIM be all the glory in JESUS name Amen. I want to say a big thank you to my supervisor, Professor S.I. Inya Agha who gave her professional support and motherly advice, and Dr, Mrs. E.U Odoh for her wise contributions. I also appreciate the support of my wife and children, the management and staff of stratech laboratory,
Enugu for their immense contribution to this work. To the following, Pharm. Elechi , Mrs
D. Okoroafor of University of Port Harcourt , and Mr. and Mrs. Gilbert Nwaigwe, I say thanks to you all.
v
TABLE OF CONTENT
Title page………………………………………………………………………………………..i
Approval Page………………………………………………………………………………….ii
Dedication………………………………………………………………………………………iii
Acknowledgement……………………………………………………………………………..iv
Table of contents………………………………………………………………………………..v
List of tables……..……………………………………………………………………………..viii
List of Figures……………………………………………………………………………….…..ix
Abstract………………………………………………………………………………………..…x
CHAPTER ONE 1
1.1 INTRODUCTION...... 1
1.2 Aim and Objective…………………………………………………………………………..4
1.3 Significance of the study…………………………………………………………………..4
CHAPTER TWO 5
2.1 literature review of crude oil pollution of environment in Nigeria……………………..5
2.2 What are medicinal plants?...... 11
2.3 Literature review of medicinal plants use for this investigation………………………12 2.3 Effects of crude oil pollution on the environment………………………………………17
2.3 Crude oil spills; impacts on plants……………………………………………………….24
2.4 Oxidative stress; Phytopathology, Phytopathogenesis………………………………..26
vi
2.5 Environmental stress features…………………………………………………………...27
2.6 Biochemical feature in Oxidative stress formation…………………………………….28
2.4 The medicinal plants under study……………………………………………………….30
2.5 Antioxidant properties of the medicinal plants under study…………………………..48
CHAPTER THREE EXPERIMENTAL 49
3.1 materials and apparatus …………………………………………………………………49
3.2 Collection of test and control samples of medicinal plants……………………………49
3.3 Quantitative determination of ascorbic acid contents of the samples……………….50
3.4 Quantitative determination of the total phenolic compounds in the samples……….50
3.5 Quantitative determination of lipid peroxide content of the samples………………...51
3.6 Determination of the Ash values of the leaves of the samples……………………….51
3.7 Epidermal cells and stomata of the leaves of the samples…………………………...53
3.8 Phytochemical screening of the medicinal plants……………………………………..53
3.9 Statistical analysis…………………………………………………………………………56
CHAPTER FOUR: RESULTS 57
4.1 Quantitative determination of the ascorbic acid contents of the samples…….…….57
4.2 Quantitative determination of the Total phenolic compound of the samples……….58 4.3 Quantitative determination of lipid peroxide content of the samples………………...59
4.4 The Ash values of the leaves of the samples…………..………………………………60
4.5 Phytochemical screening of the medicinal plants……………..……………………….64
Vii
4.6 The epidermal examination………………………………………………………………65
CHAPTER FIVE: DISCUSSION AND CONCLUTION………………………………….70
REFERENCES…………………………………………………………………………………77
APPENDIX……………………………………………………………………………………..86
viii
LIST OF TABLES
1: Result for the ascorbic acid contents of the samples...... 57
2: Result for the total phenolic compounds in the samples...... 58
3: Result for lipid peroxide content of the samples………………………………………59 4: Result for Total ash of the samples…………………………………………………… 60
5: Result for the water soluble ash…………………………………………………………61
6: Result for the acid insoluble ash………………………………………………………...62
7: Result for the sulphated ash……………………………………………………………..63
8: Result for Quantitative screening of the phytochemical contents of
the medicinal plants………………………………………………………………………64
ix
LIST OF FIGURES
1: A section of Imo River polluted by oil…………………………………………………….10
2: Matured Chromolaena odorata with flowers…………………………………………….30
3: A section of Psidium guajava (guava) tree………………………………………………36
4: Aspilia africana plant with flowers…………………………………………..…………….40
5: Picture of Manihot esculenta (cassava) plant……………………………………………43
6: Gongronema latifolium (Utazi) plant………………………………………………………46
7a: photomicrograph of upper epidermis of test sample of Aspilia africana ...... 65
7b: photomicrograph of upper epidermis of control sample of Aspilia africana...... 65
8a: photomicrograph of lower Epidermis of test sample of Aspilia africana…………….65
8b: photomicrograph of lower epidermis of control sample of Aspilia africana…………65
9a: photomicrograph of upper epidermis of test sample of Chromolaena odorata……..66
9b: photomicrograph of upper epidermis of control sample of Chromolaena odorata…66
10: photomicrograph of lower epidermis of test sample of Chromolaena odorata……..66
11a photomicrograph of lower epidermis of test sample of Psidium guajava…………..67
11b photomicrograph of lower epidermis of control sample of Psidium guajava……….67
12a: Photomicrograph of upper epidermis of test sample of sample of
Psidium guajava…………………………………………………………………………67 12b: Photomicrograph of upper upper epidermis of control Sample of
Psidium guajava………………………………………………………………………….67
x
13a: Photomicrograph of test sample of Manihot esculenta lower epidermis…………..68
13b Photomicrograph of control sample of Manihot esculenta lower epidermis………..68
14a: Photomicrograph of test sample of Manihot esculenta upper epidermis…………..68
14b: Photomicrograph of control sample of Manihot esculenta upper epidermis………68
15a: Photomicrograph of test sample of Gongronema latifolium lower epidermis……..69
15b: Photomicrograph of control sample of Gongronema latifolium lower epidermis….69
16a: Photomicrograph of test sample of Gongronema latifolium upper epidermis……..69
16b: Photomicrograph of control sample of Gongronema latifolium upper epidermis…69
xi
ABSTRACT
In this work, evaluation of oxidative stress indicators on five medicinal plants
collected from crude oil polluted environment was carried out. The medicinal plants evaluated are Psidium guajava, (L) (Myrtaceae ), Chromolaena odorata , (L) king and H.E
Robins ( Asteraceae ), Aspilia africana , (Pers.) C.D.Adams, ( Asteraceae ), Manihot
esculenta . Crantz (Euphorbiaceae ), and Gongronema latifolium Benth (Asclepiadaceae )
Oil pollution affect both the physiological, biochemical and physical states of the plants in their natural environments, invariably some of these changes may be due to oxidative stress on the plants brought about by the oil pollution of their natural environment
The oxidative stresses in these five medicinal plants were evaluated using some oxidative stress indicators like ash values, effects on the epidermal cells and the stomata, the lipid peroxide, the total phenolic compounds contents and the ascorbic acid contents. Quantitative phytochemical screening was carried out, to examine the effect of the stress on the production of secondary metabolites by these medicinal plants.
From the results it was observed that there were indication of oxidative stress with respect to the ascorbic acid contents and ash values, but with respect to the effect on the epidermal cells and stomata lipid peroxide and total phenolic compounds, some of the test samples indicate oxidative stress while others did not. On the overall Psidium guajava gave indication of oxidative stress than the remaining four plants, i.e. A.
africana, G. latifolium, Chromolaena odorata and M. esculenta.
1
CHAPTER ONE
INTRODUCTION
One of the considerable consequences of all phases of petroleum exploration, exploitation and utilization is the introduction of harmful substances into the environment. When such substances are introduced in considerable quantity, they challenge the natural resilience of the environment, and the result is that the delicate equilibrium of the environmental ecosystem is broken, and the environment becomes polluted (Agbaire and Esiefarienrhe, 2009). The components of ecosystem affected respond in different ways. While man and animals may respond by moving away from abused environment, plants, which lack locomotive ability do not, instead they respond by adjustment of some physiological, anatomical and biochemical structure and functions. The nature and magnitude of the plants response depend on the nature and magnitude of the environmental abuse. When the abuse overwhelms the normal resilient property of the plants to accommodate extraneous circumstances, the plants begin to elicit biochemical and physiological changes as a way to ameliorate the impacts associated with the environmental pollution. The plants, so subjected are said to be under stress, resulting in the production of certain biochemical agents as oxidative
- - stress factors (O 3, H2O2, O 2, 0H, N0 2). Oxidative stress is a central factor in a number of biotic and abiotic stress phenomena that occur when there is a serious imbalance in any plant cell compartment between the production of reactive oxygen species (ROS) and antioxidant defence, leading to physiological and metabolic changes. The presence of high concentration of ROS in
2 cells causes oxidative damage to photosynthetic functions and vital bio-molecules, and disrupt cellular metabolism
Crude oil spillage is pollutant that affects the environment and the ecosystem. It also introduces compounds and elements that caused oxidative stress to plants within the polluted environment. Also oil pollution exerts adverse effects on plants indirectly by making toxic minerals in the soil available to plants. A primary cause of death in oiled mangroves is disruption of gas exchange when aerial root are coated with oil and can no longer supply oxygen to underground roots (cable roots) in hypoxic soils.
Hydrocarbons also can enter mangroves through the root system and be trans-located to and accumulate in the leaves. They can destroy membranes and interrupt transpiration or poison other biochemical pathways.
Toxicity varies with oil composition, relative amounts of oil and dispersants and developmental stage of plants. It was reported that dispersed oil accumulated rapidly in seedlings than in larger trees, but that reverse was true for un-dispersed oil. Some physiological effects in plants may occur years after contamination by oil due to elevated mutation rates. The blockage of gaseous exchange will definitely lead to
- accumulation of gaseous elements and compounds such as CO 2, CO, O 2, and couple with the absorption of hydrocarbons through the roots will have damaging effects on the plants (National Academies Press, 2003). Apart from the hydrocarbon absorbed through the roots other metal elements are also introduced into the plants through the root from the crude oil polluted soil environments. All these will interfere with the normal plants
3 biochemical and physiological activities, because they will create oxidative stress in the plants
However, cells are protected partly, from reactive oxygen species and other injurious products by enzymatic and non-enzymatic defence mechanisms (Bowler, et al 1992,
Elstner and Oswald, 1994). Defence against reactive oxygen species is provided by the scavenging properties of molecules. But in spite of this the different chemical activities, and elements and compounds introduced into the plants as a result of these crude oil pollution will definitely lead to change in the total ash values of the plants when compared to those from non-crude oil polluted environments. Also the oxidative stress brought to these plants as result of this pollution will affect the stomata and epidermal cells of the leaves of these plants when compared with those from non-crude oil polluted environments. Most importantly, the level of the antioxidants like ascorbic acid and phenolic compounds will increase in a situation when there is oxidative stress. Also the level of lipid peroxide will rise in situation of oxidative stress. These were the five indicators used to evaluate the oxidative stress cause by crude oil polluted environments on the five chosen medicinal plants, namely, Chromolaena odorata (L) king and H.E Robins, Psidium guajava (L), Aspilia africana (Pers.) C.D.Adams, Manihot esculenta Crantz and Gongronema latifolium Benth.
4
Aims and Objectives;
The principal aim of this study was to determine if there was oxidative stress in medicinal plants located at the site of crude oil polluted environments located in Afam at oyigbo local government area of River state.
Significance of the study
The efficacy of medicinal plants in their therapeutic usage depends on the stability of their biochemical constituents. Alteration of the active ingredients by interference with polluting agents would not only affect their potency for medicinal uses, but would also be another source of introduction of harmful substances into the user system.
Evaluation of oxidative stress indicators would be a pointer to the extent the medicinal plants in the crude oil polluted environments are negatively affected. This will serve as guide to herbal users on where safe and potent medicinal plants should be collected in the wide or area where medicinal plants can safely be cultivated in these crude oil bearing communities.
5
CHAPTER TWO
Literature review of crude oil po llution of environment in Nigeria
It is a known fact that Nigeria is an oil producing nation. This crude oil deposit is mostly located in the Niger delta region of the country. The exploration of crude oil in this region has lead to problem of pollution. This has caused serious environmental problem, affecting both land and water resources in the region. Water, which is one of the essential compounds for living organisms, is affected badly by this pollution. This directly or indirectly affects both plant and animal lives and their activities.
There are different categories of oil pollutions. According to the U.S. Fish and Wildlife
Service, oil spills can be classified into five categories: very light oils, light oils, medium oils, heavy oils and very heavy oils.
Very light oils, such as gasoline and jet fuel, are extremely toxic to marine organisms, but evaporate rapidly in water so cleaning spills of this type is unnecessary. Light oils, such as diesel, leave a residue in water and have long–term consequences on ocean life. Although light oils have fewer toxins than very light oils, they are still damaging. Nevertheless, light oil spills can be effectively cleaned.
Medium oils, including crude oils like petroleum, do not evaporate quickly. As such, these oils can devastate marine communities residing in intertidal areas, or areas between high and low waters. Medium oils are especially threatening to birds
6 and mammals as they can adhere to their feathers, hair, or fur. Cleaning up medium oils is most successful if done immediately following the spill.
Heavy oils, on the other hand, are less likely to evaporate in water and can be exceptionally detrimental to aquatic life. Heavy oils are known to injure birds and mammals that come in contact with the contaminated site. Decontaminating areas in which heavy oils have been spilt is also very challenging.
Very heavy oils, also known as Group V oils, are capable of hovering and diffusing into water, affecting animals like lobster, which subsist on ocean floors. While Group V oils are not as toxic as the lighter oils, finding and pinpointing these oils is a difficult task
(oil spills, 2012)
In Nigeria, oil exploration which has been on for over fifty years has caused a lot of damages to the environment. This happened through oil spillage from oil pipes, thereby leading to oil pollution of the environment. Many terrestrial and aquatic lives have been affected by these oil spillages which pollutes their habitats. Human activities have also been affected by the pollution of the environment. Occupations such as fishing and
farming have also been affected. Many aquatic lives have been lost to these oil
spillages, because the normal conditions such as water salinity, PH, oxygen tension,
which sustained their lives have been badly changed. Some of these aquatic organisms
have either died or migrated out of the region. This has affected the productivity of
fishermen in these places. Farmers are not left out. Many farmlands have been
destroyed by oil spillage leading to poor yield and low productivity. One report put it in
this way; Nigeria has intense environmental degradation due to various discharges
7
being in the various state of matter that is solid, liquid and even gaseous states. Each of
these various states have contributed to degrading land, water bodies and the
atmosphere at large. In 1995 serious cases of pollution of well water and streams with
petroleum effluent have been reverted around the Kaduna refinery. River Ibeno serves
as the commonest sink for all water borne waste produced by the industry on its banks.
It is pertinent to observed that strong pressures of Mobil producing Nigeria Unlimited in
Alesa-Eleme and its crude oil exploration activities such as drilling, pipe laying and sea
transportation within the marine environment are bound to create an impact which will
inevitable extend to surrounding land and air environment, there by producing a new
dimension into this delicate ecosystem.
According to the Department of Petroleum Resources (DPR) of the Federal Ministry of
Petroleum in Nigeria, as reported by Nasiru (2012), the Niger Delta is one of the world’s largest Wetlands’ and the largest in Africa. It encompasses four main flourish and rich ecological zones. They are coastal barrier Islands, mangroves, fresh water swamp forest and lowland rainforests. This area in particular has been greatly impacted by oil spillage.
The NNPC in its annual report, places the quantity of oil jettisoned into the Niger Delta environment yearly at two thousand, three hundred (2,300) cubic meters with an average of three hundred (300) individual spills annually. In another report, it states that the total amount of oil in barrels spilled between 1960 and 1997 is upwards of one hundred million barrels (Green Peach Oil Briefing, 1993). Oil Spillage has a major impact on the ecosystem into which it is released. It destroys the mangrove forests which are especially susceptible to oil; this is mainly because
8 it stores up oil in the soil and re-releases it annually with inundation. In onshore areas, most pipelines and flow lines are laid above the ground and are more than twenty-four years old.
Even one of the oil company in the region, Shell, admits that most of the facilities has fifteen years estimated life span. Consequently, the Niger Delta has faced one calamity to another due to poor management, thus the continuous criticisms over multinational oil companies
’management in the Niger Delta Region. The revenues and incomes generated by the petroleum industry have contributed very significantly to the economic well being of Nigeria as a whole, petroleum exploration and production has however posed several environmental economic and social problems.
The Nigerian coastal areas within the offshore regions are zones of functional activities.
Since oil exploration is not properly regulated, hazardous developments are bound to occur and this extends through normal wave force to other regions. For instance, the major effects of major hazards such as NNPC spill in 1992, Oshaka oil spill in 1993, and Ibeno spill in
1998 has resulted to serious environment problems in the Niger Delta. The magnitude of crude oil pollution and damage of environmental value occasioned by multi-national oil companies operations in the Niger Delta of Nigeria is incredible. It is noteworthy that, the devastating consequences of the spill of this crude in Eleme Local Government Area Ogoni with its eventual hazards on both aerial, and human safety. The oil spill related problems in
Eleme include the defoliation of terrestrial environs tantamount to an irreversible chain effect on the bio-diversity death of mangrove vegetation, contamination of rivers, stream and groundwater supply, destroying aquatic and terrestrial lives leading to the extinction of plants and animals species (Nasiru, 2012).
9
The medicinal plants in this region are not left out. One should expect the production of secondary metabolites by these plants to be affected by the changes in their environment cause by these oil pollutions. Drugs from plants are products of secondary metabolites.
Plants produce both primary and secondary metabolites. Metabolites are organic compounds synthesized by organisms using enzyme-mediated chemical reactions called metabolic pathways. Primary metabolites have functions that are essential to growth and development and are therefore present in all plants. In contrast, secondary metabolites are variously distributed in the plant kingdom, and their functions are specific to the plants in which they are found. Secondary metabolites are often coloured, fragrant, or flavour compounds and they typically mediate the interaction of plants with other organisms. Such interactions include those of plant-pollinator, plant-pathogen, and plant-herbivore.
Secondary metabolites can also be said to be chemicals produced by plant which are not required by the plant for its survival. They may be regarded as wastes, but they may be produced in time of distress, danger or adverse condition for defensive purpose. For example, drugs such as quinine from Cinchona bark, atropine from Atropa belladonna and a host of others are products of secondary metabolites. Changes in the environmental conditions caused by this pollution by crude oil may indeed affects the production of secondary metabolites by medicinal plants used in this investigation.
10
Fig 1: A section of Imo River polluted by oil at Afam in Oyigbo local government area of River state, Nigeria, with oil floating on the water.
Oil exploration activities do not only affect the land and water only, it also affects the
atmosphere and the air compositions, as much type of gases are released into the air.
Flare of gases from these oil walls in this delta region also affects the atmospheric air. The Niger Delta region of Nigeria has suffered all forms of pollution and degradation arising from oil and natural gas exploitation. These include a decrease in agricultural yield, depression in flowering and fruiting in Okro and palm trees, deformities in children, liver damage and skin problems, increasing concentrations of airborne pollutants, acidification of soils and rainwater, corrosion of metal roofs and significant increases in concentrations of sulphates, nitrates and dissolved solids, with associated socio-economic problems (Temi, 2001). All these have serious implications
11 to the lives and health of all living organism in every type of ecosystems in this delta region and beyond.
What are Medicinal Plants?
According to Smith (2012), Medicinal plants are plants which have a recognized medical use. They range from plants which are used in the production of mainstream pharmaceutical products to plants used in herbal medicine preparations. Herbal medicine is one of the oldest forms of medical treatment in human history, and could be considered one of the forerunners of the modern pharmaceutical trade. Medicinal plants can be found growing in numerous settings all over the world.
Some medicinal plants are wild crafted, meaning that they are harvested in the wild by people who are skilled at plant identification. Sometimes plants cannot be cultivated, making wild crafting the only way to obtain them, and some people believe that wild plants have more medicinal properties. Wild crafting can also be done to gather medicinal plants for home use, with people seeking out plants to use in their own medicinal preparations.
Other medicinal plants may be cultivated. One of the advantages of cultivation is that it allows for greater control over growing conditions, which can result in a more predictable and consistent crop. Cultivation also allows for mass production, which makes plants more commercially viable, as they can be processed in large numbers and priced low enough that people will be able to afford them. (Smith, 2012)
Plants form the main ingredients of medicines in traditional systems of healing and have
12 been the source of inspiration for several major pharmaceutical drugs. Roughly 50,000 species of higher plants (about 1 in 6 of all species) have been used medicinally. This represents by far the biggest use of the natural world in terms of number of species.
Most species are used only in folk medicine, traditional systems of formal medicine using relatively few (e.g. 500-600 commonly in Traditionally Chinese Medicine). Around
100 plant species have contributed significantly to modern drugs. The use of medicinal plants is increasing worldwide, related to the persistence and sometimes expansion of traditional medicine and a growing interest in herbal treatments.
The medicinal uses of plants grade into their uses for other purposes, as for food, cleaning, personal care and perfumery. Plants are used in medicine to maintain and augment health - physically, mentally and spiritually - as well as to treat specific conditions and ailments. (Idu, 2012) It is clear from the above explanation that medicinal plants have been with man right from time, and have served to meet his medical needs. These medicinal plants are grown both by cultivation as well as grown as wild plants. Therefore, any changes in the environment and the ecosystem of these plants may affect their medicinal activities.
. Literature review of medicinal plants use for this investigation
Five plants are used for this work. These are Psidium guajava Linn (Myrtaceae) , Aspilia
africana (Pers.) C.D.Adams, (Asteraceae), Chromolaena odorata. (L) king and H.E
Robins (Asteraceae), Gongronema latifolium Benth (Asclepiadaceae) and Manihot
esculenta.Crantz (Euphorbiaceae).
13
Psidium guajava : This plant is use both for nutritional and medicinal purposes. A lot of work has been done to establish these facts. (Shah et al., 2011), has demonstrated pharmacologically that the plant can be use to manage hyperactive gut disorders. It has been shown that Guava is rich in tannins, phenols, triterpenes, flavonoids, essential oils, saponins, carotenoids, lectins, vitamins, fiber and fatty acids.
Guava fruit is higher in vitamin C than citrus (80 mg of vitamin C in 100 g of fruit and contains appreciable amounts of vitamin A as well. Guava fruits are also a good source of pectin - a dietary fiber. The leaves of guava are rich in flavonoids, in particular, quercetin. Much of guava's therapeutic activity is attributed to these flavonoids. The flavonoids have demonstrated antibacterial activity. Quercetin is thought to contribute to the anti-diarrhea effect of guava; it is able to relax intestinal smooth muscle and inhibit bowel contractions. In addition, other flavonoids and triterpenes in guava leaves show antispasmodic activity. Guava also has antioxidant properties which are attributed to the polyphenols found in the leaves (Taylor, 2004). Also, in a study with guinea pigs (in 2003) Brazilian researchers reported that guava leaf extracts have numerous effects on the cardiovascular system which might be beneficial in treating irregular heart beat (arrhythmia). Previous research indicated guava leaf provided antioxidant effects beneficial to the heart, heart protective properties, and improved myocardial function. In two randomized human studies, the consumption of guava fruit for 12 weeks was shown to reduce blood pressure by an average 8 points, decrease total cholesterol levels by 9%, decrease triglycerides by almost 8%, and increase "good"
HDL cholesterol by 8%. The effects were attributed to the high potassium and soluble
14 fiber content of the fruit (however 1-2 pounds of fruit was consumed daily by the study subjects to obtain these results!). In other animal studies guava leaf extracts have evidenced analgesic, sedative, and central nervous system (CNS) depressant activity, as well as a cough suppressant action. The fruit or fruit juice has been documented to lower blood sugar levels in normal and diabetic animals and humans. Most of these studies confirm the plant's many uses in tropical herbal medicine systems.
Chromolaena odorata : It is a medicinal plants used in traditional health care
(Erinoso and Aworinde, 2012). It has been established that extracts from the plant can be use in treating skin infection (Egharevba et al., 2008). The extract can also be use in
arresting bleeding, and as such promote wound healing (Adetutu et al., 2011). The
aqueous extract and the decoction from the leaves of this plant have been used throughout Vietnam for the treatment of soft tissue wounds and burn wounds (Phan., et al 1996)
Aspilia africana : Like Chromolaena, it is widely used in ethno medical practice in
Africa for its ability to stop bleeding, even from a severed artery, as well as promote rapid healing of wounds and sores and for the management of problems related to cardiovascular diseases (Eweka, 2009). Infusion of a liquid made from the leaves is taken by children and can also be mixed with clay as a medicine for stomach trouble It has been reported that the plant is effective against malaria ( Plasmodium falciparum ) infection. It has been classified among substances with a low potential for toxicity, with an LD 50 averaging 6.6g/Kg body weight (Eweka, 2009). The methanolic and aqueous
15 extracts of the leaves of A. Africana have exhibited differential anti-infective activities on both Gram-positive and Gram-negative bacteria species (Eweka, 2009). A. Africana has many other additional uses such as palliative properties as documented by (Okoli and
Akah, 2007) that the leaves of A. africana possess constituents capable of arresting wound bleeding, inhibiting the growth of microbial wound contaminants and accelerating wound healing which suggest good potentials for use in wound care , alleviating menstrual cramps and dysmenorrhoeal (which are not documented) probably because empirical studies had not been carried out on them to authenticate their efficacy.
Chemical constituents of the plant had also been studied for instance, the chemical constituents of the leave includes the following; alkaloids (6.350±0.841%), tannins (0.188±0.035%), saponins (2.260±0.15%) flavonoids (2.006±-0.11%) and phenols
(0.109±0.15%), respectively. The concentration per 100 g of vitamins in the leaf of the plant are as follows ascorbic acid (11.00±0.15 mg) niacin (3.045±0.081 mg) thiamine
(1.940±0.025 mg), riboflavin (0.135±0.100 mg). The mineral elements determined are per 100 g Calcium (246±4.345 mg), Nitrogen (213.00±6.600) and Phosphorus
(25.00±0.500), zinc (34.675±0.54), nickel (10.975±1.330), Selenium (Se)
(10.875±1.880), Boron (B) (9.675±2.045) and lead (Pb) (7.425±1.050), these results show that the list may possess the medicinal potentials as claimed by tradomedical practitioners ( Abii and Onuoha, 2011)
16
Gongronema latifolium : this plant has been studied and investigated in many ways.
Thus over the years, several reports have appeared on Gongronema latifolium a leafy vegetable locally known in the South-Eastern part of Nigeria as Utasi or Utazi.
The proximate analyses and chemical compositions of the herb have been reported as well as antimicrobial activities and hypoglycaemic effects (Etta et al., 2012). A study of the nutritional composition of this plant with that of bitter leaf put the result as follows; the lipid extract, ash, crude fibre and nitrogen free extractives, oxalate, phytate and tannin of the plants are within expected ranges. They however had unexpectedly high crude protein content: 27.20 and 21.69 per cent. Potassium, phosphorus, calcium and cobalt were the most abundant mineral elements. G. latifolium and V. amygdalina leaf oils are 50.22 and 24.54 per cent saturated; 39.38 and 65.45 per cent polyunsaturated, respectively. Palmitic and oleic acids were the major monounsaturated fatty acids.
Degrees of un-saturation are 0.46 and 0.41, respectively. Major essential amino acids are leucine, valine and phenylalanine. Proportions of essential to non-essential amino acid are 43.37 and 49.84 per cent, respectively (Eleyinmi., et al 2008).
Manihot esculenta ; it was regarded to have some medicinal uses but not much work
has been done scientifically to prove most of the claims on its traditional medical uses.
There are claims that botanical products currently made from the cassava have
anticancer properties, but available scientific evidence does not support these claims. A
British researcher identified the cassava genes involved in making hydrogen cyanide in
the early 1990s. In collaboration with cancer specialists in Spain, she has conducted
17
studies of the linamarase gene. They added this gene to a virus, which was then
injected into rat brain tumors. These tumors were killed when the rats were infused with
linamarin (American cancer society, 2008). Further research is needed to determine if
this technique would work in human. However, the leaves of some species of the plant
are used for nutritive and medical purposes apart from the facts that plant tubers are
very good source of carbohydrates.
Effects of crude oil pollution on the environment Environment here referred to both the terrestrial and aquatic environments. In both the terrestrial and aquatic environments, pollution by crude oil causes a lot of destruction to the ecosystems.
General composition of crude oil: Crude oil and petroleum are complex mixtures of several polycyclic aromatic compounds and other hydrocarbons. The major hydrocarbon classes found in diesel fuel are the normal alkanes (rapidly degraded), branched alkanes and cycloalkanes (difficult to identify), the isoprenoids (very resistant to biodegradation), the aromatics, (fairly identified and much more soluble than other hydrocarbons), and finally the polar ones containing mainly sulphur, oxygen and / or nitrogen compounds. Typical composition of crude oil based on distillation properties is illustrated in Table1a while Table 1b shows some examples and aqueous solubility of hydrocarbon compounds commonly analyzed in crude oil. Non-hydrocarbon compounds may also be found in crude oil and they include porphyrins and their derivatives. Metals that could be found in crude oil via their association with porphyrins include nickel,
18 vanadium, iron, zinc, cobalt, titanium and copper. Some priority contaminant of petroleum hydrocarbons and crude oil include benzene, heptanes, hexane, isobutene, isopentane, PAHs such as benzo[a] anthracene, benzo[b] pyrene etc. (Onwurah et al.,
2007)
Distillation fraction Composition (%) Chain length Gasoline 11.2 C4-C10
Naphtha 18.1 C4-C10
Kerosene 16.9 C10 -C20
Gas oil 15.7 C15 -C40
Heavy gas 25.8 C40 and above
Table 1(a). Some distillation components of Nigerian Brass crude oil
19
0 Components Examples Solubility (mg/L) 28+ _2 C
Alkanes n-butane 101.00
n-decane 0.05
Branched alkanes 2-methylpentane 78
Cycloalkanes Cyclohexane 55 Olefins 1-pentene 148
Monoaromatic Benzene 1760
Toluene 470
Ethyl benzene 140
Polyaromatic Naphthalene 30
Phenols Phenol 82,000
2,6-dimethyl phenol 4,600
2,4,6-trimethyl phenol 14,000
Table (b). Solubility (in water) of some common components easily analysed in crude oil.
Spills in populated areas often spread out over a wide area, destroying crops and aquacultures through contamination of the groundwater and soils. The consumption of dissolved oxygen by bacteria feeding on the spilled hydrocarbons also contributes to the
20
death of fish. In agricultural communities, often a year’s supply of food can be destroyed. Because of the careless nature of oil operations in the Delta, the environment is growing increasingly uninhabitable. People in the affected areas complain about health issues including breathing problems and skin lesions; many have lost basic human rights such as health, access to food, clean water and their ability to work. In some places a crust of ash and tar caused by fires after oil spills had been left for decades. Even in the mangroves and creeks, one can find water polluted by crude oil at different levels of pollutions. Also, note that Pollution from over 50 years of oil operations in the region has penetrated further and deeper than many may have supposed, the report says. Some areas that seemed unaffected on the surface are severely contaminated underground and need urgent action to protect the health of fishing and farming communities (Robyn, 2011). There are some bodies of water in these areas which may not look like oil polluted water but closer study will show that they are contaminated. Since these pollutions had taken long time, it will surely have serious effect on plants and animals in these environments.
In the British Petroleum (BP) oil spill of 2010, 4.9 million barrels of crude oil were spilt in the Gulf of Mexico. According to Time , thousands of dead invertebrates like starfish and coral were found. Unfortunately, these species play an essential role in the ecosystems to which they belong, thereby impacting many other marine populations.
Similarly, many dolphin offspring were found dead along the Gulf Coast. Oyster beds were also devastated by the oil spill; in fact, it could take ten years for the population to reach its former size.
21
The 1989 Exxon Valdez oil spill was equally catastrophic. According to BBC News, the oil killed over 250,000 seabirds, 2,800 sea otters, 250 bald eagles, 300 harbor seals, and 22 killer whales, as well as countless herring and salmon.
In addition to killing many sea dwellers, oil spills can also impact the health of those that survive. Oil can modify invertebrate feeding habitats, disrupt their shell development, and cause slow suffocation. Bottom-dwelling invertebrates are especially at risk when oil accumulates at the shoreline. Many bottom-dwellers can survive oil contamination; however, they transmit these toxins to their predators, leading to increased concentration of the toxins in higher species. From oil spills, fish can experience impeded growth, respiratory and cardiac malfunction, and stunted larval development. As a result, survival rates for offspring are low.
Oil spills can similarly thwart plant development. They can also spur growth of certain algae populations. When oil directly contacts birds, it can get in their feathers, which impedes their abilities to fly. As a result, many birds drown while others die of hypothermia. If oil is ingested, kidney, liver and lung damage often results, usually followed by death. Other side effects include an inability to reproduce, abnormal behaviors, a debilitated immune system, and skin irritability.
Humans can also be affected by oil spills. In Ogoniland, Nigeria, for example, the people have dealt with nearly 50 years of oil production and water contamination. Many communities are faced with dangerous levels of carcinogens, cancer causing agents.
22
In one such community, families are drinking water polluted with benzene, a type of carcinogen, at a concentration 900 times that considered to be safe. In other areas of
Ogoni land, nearly eight centimeters of oil were found on top of the water. This horrific spill has so far killed tens of thousands of people, as well as livestock, and is predicted to take up to 30 years to reach its former clean state. Altogether, it will cost approximately $1 billion to rebuild the area. The Shell Oil Company, which was responsible for the spill, has neglected the impact this spill has had on the Nigerians.
They have, however, taken responsibility for the recent 2008 and 2009 oil spills.
The physicochemical properties of the water are greatly adjusted when the water is polluted by petroleum. The PH which determines the acidity and alkalinity of the water is also affected by the pollution. The salinity which describes or determines the salt content of the water is also affected by the pollution. Other parametric properties of the water, like the total solid (TS) which is the sum total suspended solid (TSS) and total dissolved solid (TDS) of the water are also affected by the pollution. Turbidity, floc formation organoleptic properties are all adjusted by the crude oil pollution. Oil is persistent--both in its desirability in our modern age and its damaging effects in the environment when it spills. It sticks around. When crude oil ends up in water, it forms a buoyant layer on the water. That layer spreads into a thin slick but in weeks can weather into thick, tarry globs--a tough “skin” trapping fresher oil inside. Storms and weather also break up oil into small drops that can disperse as small oil particles. As these particles collide in the water with suspended sediment, they form “tarballs” of oil,
23 sand, algae and other debris. And while slicks and tarballs can be broken down by light and by micro-organisms and plants, it takes a long time. Floating oil can become stuck onto shorelines, or oil particles can collect enough sediment to sink to the water bottom and remain there (Slick Science, 2011). When oil is spilled in the ocean, it initially spreads in the water (primarily on the surface), depending on its relative density and composition. The oil slick formed may remain cohesive, or may break up in the case of rough seas. Waves, water currents, and wind force the oil slick to drift over large areas, impacting the open ocean, coastal areas, and marine and terrestrial habitats in the path of the drift.
Oil that contains volatile organic compounds partially evaporates, losing between 20 and 40 percent of its mass and becoming denser and more viscous (i.e., more resistant to flow). A small percentage of oil may dissolve in the water. The oil residue also can disperse almost invisibly in the water or form a thick mousse with the water. Part of the oil waste may sink with suspended particulate matter, and the remainder eventually congeals into sticky tar balls. Over time, oil waste weathers (deteriorates) and disintegrates by means of photolysis (decomposition by sunlight) and biodegradation
(decomposition due to microorganisms). The rate of biodegradation depends on the availability of nutrients, oxygen, and microorganisms, as well as temperature. If oil waste reaches the shoreline or coast, it interacts with sediments such as beach sand and gravel, rocks and boulders, vegetation, and terrestrial habitats of both wildlife and humans, causing erosion as well as contamination. Waves, water currents, and wind
24 move the oil onto shore with the surf and tide. Although marine transportation accidents can result in such oil spills, they account for only about 5 percent of the waste oil that enters the ocean annually. Beach sand and gravel saturated with oil may be unable to protect and nurture normal vegetation and populations of the substrate biomass. Rocks and boulders coated with sticky residue interfere with recreational uses of the shoreline and can be toxic to coastal wildlife (Ogallala, 2013). Thus there are a lot of changes in the water when polluted with crude oil. These changes have strong disturbances in the ecosystem and the animals and plants in their natural habitats are not spared.
Crude oil spills; impacts on plants
Crude oil spills has a variety of negative impacts to all forms of life in the ecosystem of where it occurs, because it pollutes the environment. Pollution in this context means an alteration or distortion of ecosystem by introducing foreign bodies or increasing the concentration of the pre-existing natural components beyond the limit of environmental self restoration. Depending on the magnitude and length of time of its existence, the delicate ecological stability may break down, resulting in the destruction of lives in the environment. While man and other animals may relocate from the impacted site, plants will not owing to their immobility. Thus the most affected biotic component of the ecosystem in the crude oil spill is plant
Heavy metals available to the soil during crude oil spills have been implicated as causative agent in the ethiology of oxidative stress in plants (Dietz et al., 1999). Excess
25 heavy metal pollution of plant may stimulate the formation of free radicals and reactive oxygen species, resulting in the formation of oxidative stress (Hall, 2002, Dietz et al.,
1999). Lead and mercury are reputed to cause an increase in ascorbic acid levels in two
Oryza sativars (Mishra and Choudhuri, 1999), this indicate that the plants were experiencing oxidative stress. (Del vos et al., 1992) reported that copper and cadmium induced oxidative stress in plants. Oxidative stress, in this context, is defined as all of the effects such as cellular damage, caused by the active forms of oxygen (superoxide
- - O 2), hydrogen peroxide (H 2O2), hydroxyl radical (OH ) and singlet oxygen (O 2). The
displacement of certain bioactive elements in plants enzyme and pigments by heavy
elements or metals resulting from crude oil spills hinders some biochemical process at
the cellular levels, thus enhancing the accumulation of reactive species and dangerous
radicals, injurious to plants. This produces oxidative stress, which may lead to
physiological changes in plants before exhibiting visible damage to leaves (Dohmen et
al., 1990)
Crude oil spills may cover the leaves and thereby inhibiting stomata movement, photosynthesis and growth. Apart from screening out sunlight, oil on leaves blocks stomata and lowers their conductance to carbon dioxide, simultaneously interfering with photosystem ii.
26
Oxidative stress; phytopathology, phytopathogenesis
Phytopathology
Different disease presentations resulting from oxidative stress in plants have been reported (Hussain., et al 2013) cited the presentation on plants resulting from cadmium induced stress, It inhibits the seed germination disturbs the photosynthetic metabolism and transpiration rate, reduces enzymatic and non enzymatic activities disturbs water homeostasis and ionic relations mineral nutrition induces synthesis of reactive oxygen species, and strongly reduced the biomass production. The cadmium stress causes chlorosis and leaf and root necrosis resulting in stunted growth in the majority of the plants. Physiological and anatomical presentations of oxidative stressed plants include; stunted growth, reduction in leaf area, vascular atrophy,Interveinal necrosis, stomata diameter reduction, foliar distortion and cell wall elasticity (Baszynski et al., 1980)
The interference of agents of oxidative stress with plants enzymes especially kinctin results in the increasing of the relative content of the plant`s leaf because of the reduction in stomata aperture. This in turn leads to plant leaf turgidity. Leaf turgidity results in the creation of osmotic pressure on the cellular integrity, often culminating in the folialysis, veinal rupture, cuticular and waxy distortion and general disorganisation of the foliar physio-chemical functions.
Bacterial and viral attacks and exacerbation occur when vegetations are weakened by oxidative stress. Blights, notches patchy derma necrosis chlorosis are the resultant effects of oxidative stress in synergistic interaction with microbial agents.
27
An oxidative stress factor-ozone binds to plasma membranes and it alters metabolism.
As a result, stomata aperture are poorly regulated, chloroplast thylakoid membranes are damaged, rubisco is degraded and photosynthesis is inhibited, and the plants begin to manifest symptoms of diseased condition.
Phytopathogenesis
Oxidative stress is a situation occasion by an imbalance in rate of oxidation of cellular substrates and the antioxidant activities of certain enzymes and vitamins, caused by a number of factors, namely environmental factors and Biochemical factor
Environmental stress factors:
Most environmental stresses are affecting the production of active oxygen species in plants causing oxidative stress (Smirnoff, 1993, Hendry, 1994). The balance between the production of activated oxygen species and the quenching activity of antioxidant is upset which often results in oxidative stress
Ultraviolet disrupts metabolic processes in plants, including photosynthesis, respiration, glucose assimilation and phosphorylation There have been many reports on the deleterious physiological effects on plants exposed to high level of ultraviolet radiation, which may increase if the stratophoric ozone concentration decreases. The destructive action results from both direct and indirect mechanisms involving endogenous sensitizers and the generation of active oxygen species.
28
Physiological and biochemical effects include effects on enzyme, stomata resistance, concentration of chlorophyll, protein and lipids reduction in leaf area, and tissue damage Metal induced stress resulting from phytotoxic metals form stable complex that generate active oxygen whose destructive effects on cellular status causes phyto-oxidative stress. Binding of metals to the important sulflydryl group of enzymes exacerbates the phytotoxic action of the metals leading to phyto-oxidative stress.
Biochemical factor in oxidative stress formation:
Atmospheric pollutants such as ozone (O 3) and sulphur iv oxide (SO 2) have been implicated in free radical formation and hence in the formation of oxidative stress.
Ozone, which originates from a natural photochemical degradation of nitrous oxide
(NO 2), seems to have greater effect to plant than other forms of pollution. In oil exploration activities such as gas flaring, results in the formation nitrous oxide and sulphur oxide, among others.
Exposure of plants to sulphur oxide results in tissue damage and release of stress ethylene from both photosynthetic and non-photosynthetic tissues. The sulphur oxide causes a shift in cytoplasmic PH. The product concentration in cytoplasm is one of the most important factors resulting in cellular activity. When cells are exposed to sulphur oxide an appreciable acidification of the cytoplasm occur because this gas react with water to form sulphurous acid which may be converted to sulphuric acid. These result in
29 loss of photosynthetic function caused by inhibition of SH-conting light activated enzymes of the chloroplast. The oxidation of sulphite to sulphate in the chloroplast also
- gives rise to the formation of active oxygen (O 2 ). The oxidation of sulphite is initiated by light and is mediated by photosynthetic electron transport.
In oil exploration activities involving gas flaring and oil spills gaseous and particulate pollutants results in the decrease of atmospheric oxygen depending on the degree of pollution, the plants exposed to the polluted environment may suffer hypoxia or anoxia.
A decrease in adenylate energy change, cytoplasmic acidification, anaerobic formation, elevation in cytosokic Ca 2+ concentration, changes in the redox state and a decrease in the membrane barrier function, are the main features caused by lack of oxygen.
Hypoxia may also lead to the formation of reactive oxygen species, and lipid peroxidation.
30 The medicinal plants under study
Five plants were used in this investigation. These are Chromolaena odorata (L) king and H.E Robins, Psidium guajava (L), Aspilia africana (Pers.) C.D.Adams, Gongronema latifolium Benth and Manihot esculenta Crantz.
Chromolaena odorata :
Scientific classification
Kingdom: plantae
(Unranked): Angiosperms
(Unranked): Eudicots (Unranked): Asterids Order: Asterales Family: Asteraceae Genus: Chromolaena Species : C. Odorata Binomial name: Chromolaena odorata (L) king and H.E Robins
Fig. 2: Matured Chromolaena odorata with flowers
31 Chromolaena odorata is a species of flowering shrub in the sunflower family,
Asteraceae . It is native to North America, from Florida and Texas to Mexico and the
Caribbean, and has been introduced to tropical Asia, west Africa, and parts of Australia.
Common names include Siam Weed, Christmas Bush, and Common Floss Flower.
Local names are Awolowo, Akintola. It is sometimes grown as a medicinal and ornamental plant. It is used as a traditional medicine in Indonesia. The young leaves are crushed, and the resulting liquid can be used to treat skin wounds It was earlier taxonomically classified under the genus Eupatorium, but is now considered more closely related to other genera in the tribe Eupatorieae. “Siam Weed is a big bushy herb or sub-shrub with long rambling, but not twining branches. In open areas it spreads into tangled, dense thickets up to 2 m tall, and higher when climbing up vegetation. Many paired branches grow off the main stem. The base of the plant becomes hard and woody while the branch tips are soft and green. The leaves are arrowhead-shaped, 5–
12 cm long and 3–7 cm wide, with three characteristic veins in a ‘pitchfork’ pattern. They grow in opposite pairs along the stems and branches. As the species name ‘odorata’ suggests, the leaves emit a pungent odour when crushed. Clusters of 10–35 pale pink– mauve or white tubular flowers, 10 mm long, are found at the ends of branches. The seeds are dark coloured, 4–5 mm long, narrow and oblong, with a parachute of white hairs which turn brown as the seed dries. Siam weed is native to Tropical America, but is now naturalized throughout the tropics ( Basu and Eby, 2007)
Chromolaena odorata is considered invasive weed of field crops in its introduced range, and has been reported to be the most problematic invasive species within
32 protected rainforests in Africa (Wikipedia, 2014). Some of the attributes that contribute to Chromolaena’s success as an invader are its rapid growth rate, its allelopathic
properties and, in the dry season, its huge seed production (up to about 1 million seeds per plant per flowering season) and, due to high levels of essential oils, its high flammability. Although it readily invades disturbed areas, it can also invade natural habitats that are subject to minimal disturbance. It has a very plastic morphology in open areas; it forms a compact shrub up to 3m tall, whereas under trees or on forest margins it can scramble up to 10m to emerge over the canopy. However, it does not tolerate complete shade. Chromolaena spreads rapidly due to its small, light seeds attached to a pappus, which are dispersed both by air currents and on animals and vehicles.
Chromolaena displays considerable morphological variation within its native range.
Flower colour varies from white through pale lilac to blue, and other aspects of flower morphology can also vary (e.g. shape of bracts, diameter of capitula). Leaves and stems range from glabrous to hairy and as a consequence, their colour varies too. Plant architecture is variable (lax to upright), as is the odour of crushed leaves and stems. In some areas of the neotropics (e.g. the Caribbean and Central America), high local variability is evident, while in others (parts of the South American mainland), it is locally uniform in morphology.
There appear to be two primary centres of invasion by Chromolaena in the Old
World. Each centre of invasion is characterised by plants which are morphologically
33 homogeneous within that centre of invasion, but morphologically distinct between the centres of invasion. In the first, more widespread centre of invasion, Chromolaena probably spread throughout Asia and Oceania from the Calcutta Botanic Garden, where it had been planted in the mid-19 th century. This form of Chromolaena appears to have been taken from Sri Lanka to West Africa in the first half of the 20 th century. From here, it spread and is now present from the Gambia to northern Angola and Tanzania.
Because this constitutes a secondary introduction it is considered as part of the first centre of invasion. A second form of Chromolaena appeared in south-eastern South
Africa in the 1940s, from where it spread throughout climatically suitable areas of the subcontinent. Since the biology of plants in these two centres of invasion is apparently also distinct in some important characteristics, both these forms have been characterised as ‘biotypes’ and functionally, although not strictly, can be considered to be separate species. They are referred to from here on as the Asian/West African biotype and the southern African biotype .” “It is a highly successful plant that has
colonized diverse ecological areas of tropical lands. Its ability to survive long spells of
drought as occasioned in many tropical areas and its propensity to resume active
growth at the commencement of the rains is unparalleled. Its soil-enriching prowess and
its tendency to infiltrate new areas have recommended it as a plant of choice in fighting
desert encroachment especially in the Sudano-sahelian ecological zones of Africa.
The medicinal use of Chromolaena odorata has also not gone unnoticed. The
astringent properties of the leaf extracts of Chromolaena odorata on the blood vessel
(Iwu 1993) has made it a popular plant in the prevention of blood loss from wounds,
34 also its anti-microbial properties has made it a popular choice in disinfecting and treating open wounds (Odugbemi, 2006). The antihelmintic properties of the aqueous extracts of Chromolaena odorata have also been widely known among the peasant population of Asia and Africa. Its popularity as an effective therapy against diarrhoea, malaria fever, tooth ache, diabetes, skin diseases, dysentery and colitis has been severally documented (Odugbemi, 2006, Akinmoladun and Akinloye, 2007) ( Aro et al.,
2009)
Psidium guajava (guava)
Scientific classification
Kingdom: Plantae
(Unranked): Angiosperms
(Unranked): Eudicots
(Unranked): Rosids
Order: Myrtales
Family: Myrtaceae
Sub-family: Myrtoideae
Tribe: Myrteae
Genus: Psidium Species: P. guajava
Binomial name: Psidium guajava (L) 35
Tropical Guavas are known scientifically as Psidium guajava they are the best tasting with the largest fruit with the most juice. These are the most frost tender Guavas. Tropical Guavas grow up to 10 to 15 feet high & wide. Strawberry
Guavas, Psidium lucidum are shrubby trees with tart but very flavourful fruit that is smaller than a Tropical Guava. Strawberry Guavas are very productive & grown 12 feet high & wide. Pineapple Guavas, is a South American plants are related to other
Guavas. Their fruit is tangy with a citrus flavour. This is the most frost tolerant variety.
Pineapple guavas grow to 15 feet high & wide & have wonderful gnarled trunks & make good substitutes for olive trees. Owing to its hardy nature, guava is grown successfully in tropical and subtropical regions up to 1, 500 m above mean sea-level. Best quality guavas are obtained where low night temperatures (10 oC) prevail during winter. It tolerates high temperatures and drought conditions in North India during summers but it is susceptible to severe frost as it can kill the young plants. An annual rainfall of about
100 cm is sufficient during the rainy season (July- September). The rains during
harvesting period, however, deteriorate the quality of fruits.
Guava is cultivated on varied types of soils, heavy clay to very light sandy soils.
Nevertheless, very good quality guavas are produced in river-basins. It tolerates a soil
pH of 4.5- 8.2. Maximum concentration of its feeding roots is available up to 25cm soil
depth. Thus the top soil should be quite rich to provide enough nutrients for accelerating
new growth which bears fruits. (Wikipedia, 2014)
Guava fruit is edible and very nutritious. It has a lot of Phytochemicals which are very
beneficial to mankind. The leaves are use in the treatment of fever and malaria.
36 Psidium guajava is an important food crop and medicinal plant in tropical and subtropical countries it is widely used like food and in folk medicine around the world.
Fig. 3: A section of Psidium guajava (guava) tree
Different pharmacological experiments in a number of in vitro and in vivo models have been carried out. Also have been identified the medicinally important phyto- constituents. A number of metabolites in good yield and some have been shown to possess useful biological activities belonging mainly to phenolic, flavonoid, carotenoid, terpenoid and triterpene. Extracts and metabolites of this plant, particularly those from leaves and fruits possess useful pharmacological activities. A survey of the literature shows P. guajava is mainly known for its antispasmodic and antimicrobial properties in the treatment of diarrhoea and dysentery. It has also been
37 used extensively as a hypoglycaemic agent. Many pharmacological studies have demonstrated the ability of this plant to exhibit antioxidant, hepatoprotection, anti- allergy, antimicrobial, anti-genotoxic, anti-plasmodial, cytotoxic, anti-spasmodic, cardio active, ant cough, ant diabetic and anti-inflammatory activities, supporting its traditional uses (Gutierrez et al., 2008)
The plant has very good food or nutrient value. This can be summarised as follows:
Guavas are low in calories and fats but contain several vital vitamins, minerals,
and antioxidant poly-phenolic and flavonoid compounds that play a pivotal role in
prevention of cancers, anti-aging, immune-booster, etc.
The fruit is very rich source of soluble dietary fibre (5.4 g per 100 g of fruit, about
14% of DRA), which makes it a good bulk laxative. The fibre content helps
protect the colon mucous membrane by decreasing exposure time to toxins as
well as binding to cancer-causing chemicals in the colon.
Guava-fruit is an excellent source of antioxidant vitamin-C. 100 g fresh fruit
provides 228 mg of this vitamin, more than three times the DRI (daily-
recommended intake). Outer thick rind contains exceptionally higher levels of
vitamin C than central pulp.
Scientific studies shown that regular consumption of fruits rich in vitamin C helps
the body develop resistance against infectious agents and scavenge cancer
causing harmful free radicals from the body. Further, the vitamin is required for
collagen synthesis within the body. Collagen is the main structural protein in the
38 human body required for maintaining the integrity of blood vessels, skin, organs,
and bones.
The fruit is a very good source of Vitamin-A, and flavonoids like beta-carotene,
lycopene, lutein and cryptoxanthin. The compounds are known to have
antioxidant properties and are essential for optimum health. Further, vitamin-A is
also required for maintaining healthy mucus membranes and skin. Consumption
of natural fruits rich in carotene is known to protect from lung and oral cavity
cancers.
• 100 g of pink guava fruit provides 5204 µg of lycopene, nearly twice the amount
that in tomatoes. (100 g tomato contains 2573 µg of lycopene). Studies suggest
that lycopene in pink guavas prevents skin damage from UV rays and offers
protection from prostate cancer.
• Fresh fruit is a very rich source of potassium. It contains more potassium than
other fruits like banana weight per weight. Potassium is an important component
of cell and body fluids that helps controlling heart rate and blood pressure.
• Further, the fruit is also a moderate source of B-complex vitamins such as
pantothenic acid, niacin, vitamin-B6 (pyridoxine), vitamin E and K, as well as
minerals like magnesium, copper, and manganese. Manganese is used by the
body as a co-factor for the antioxidant enzyme, superoxide dismutase . Copper is
required for the production of red blood cells. (nutrition and you, 2013)
It is very clear from the above summary that the health benefits of Psidium guajava
are enormous 39
Aspilia Africana
Scientific classification:
Kingdom: plantae
(Unranked): Angiosperms
(Unranked): Eudicots
(Unranked): Asterids
Order: Asterales
Family: Asteraceae
Genus: Aspilia
Species : A. Africana
Binomial name: Aspilia africana , (Pers.) C.D.Adams
Aspilia is a genus of flowering plants in the daisy family. Historically, Aspilia africana has been used in Mbaise and most Igbo speaking parts of Nigeria to prevent conception suggesting potential contraceptive and anti-fertility properties. Leaf extract and fractions of A. africana effectively arrested bleeding from fresh wounds, inhibited microbial growth of known wound contaminants and accelerated wound healing process
(Oluyemi et al., 2007) . Aspilia africana is a common weed of field crops in West Africa found in fallow land, especially in the forest zone. It is a scrambling perennial herb varying in height from 60 cm to about 1.5 m depending On rainfall (Agyakwa and
Akobundu, 1987). The flowers are showy yellow florets and the fruits are bristly and minutely hairy with 4 angled schemes about 5 mm long. There are several indigenous 40 uses of the leaves and flowers of this plant. The most notable being the use of it to stop bleeding and fast healing of wounds. They are used in the treatment of rheumatic pain as well as bee and scorpion stings. The plant is used to treat different diseases in different ecological zones due to varying chemical composition as a result of various ecological conditions of different places. In Kenya, they are used to kill intestinal worms in Uganda, it is used to treat gonorrhoea. The methanol extract of the leaves are reported to cure malaria and respiratory problems. A concussion of the leaves are used to cure eye problem and as a lotion for the face to relieve headache. They are also used to cure ringworm and dysentery (Abii and Onuoha, 2011).
Investigations has been done on the chemical constituents of the leaves of Aspilia africana, in an attempt to provide a profile that gives a scientific backing to various trado-medical claims and uses of the leaves of Aspilia africana
Fig. 4: Aspilia africana plant with flowers 41
Semi-woody herb is a perennial woody root-stock to 2 m high, very polymorphic with at least four varieties recognized in the Region, occurring throughout the Region on waste land of the savannah and forested zones, and widely distributed across tropical
Africa. The plant is a weed of cultivated land and fallows. It is of very rapid growth. It has a somewhat aromatic carroty smell.
In Nigeria it is grazed by cattle and sheep, and is much used in the Western State as a food for rabbits and hares. It has high crude protein content. The plant has a wide reputation and use as a haemostatic, and in Liberia it is even credited with the capacity of arresting bleeding of a severed artery. The fresh bruised leaves and flowers are used.
Their application is said to draw up exudations (hence the Hausa vernacular name meaning ’draw up mucus’) and to promote rapid healing. Besides using the fresh leaf on cuts, the Ijaw of Southern Nigeria will also apply leaf-ash onto wounds and sores. A decoction has been recommended for use in treating pulmonary haemorrhage, and haemostasis is thought to be due to vaso-constriction. In Tanganyika a root-decoction is taken for tuberculosis and in Ghana the leaves are made into a cough-medicine for children. In Uganda a leaf-concoction is taken for gonorrhea. Water in which leaves have been squeezed with a little salt and lime-juice added is much used in Ghana as an eye- medicine to remove corneal opacities. In Nigeria such water with only salt added is used as an eye-lotion for sun-blindness. The leaf-sap is also an eye-medicine in Tanganyika, dripped into the eyes for eye-pains of no apparent origin. The Igbo instill a leaf-decoction into the eyes for headache, and a decoction is used to wash the face and eyes to relieve feverish headache in Ghana and in Nigeria. A leaf-infusion is taken in Ghana to assist in 42 childbirth and also in Nigeria. The leaf sap and a leaf-decoction are rubbed onto the breasts and made into an inhalation in Tanganyika to promote milk-flow. An infusion mixed with white clay is taken by the Akan people of Ghana for stomach-troubles. Ijaw of
Southern Nigeria squeeze out the leaf-sap which is given with salt to revive a fainting person or to someone suffering from an attack of nerves. The Yoruba use the plant to cure
Craw-craw and it features in an incantation to this end: the Yoruba name yun yun means
‘scratch’. In francophone territories it is used in fumigation to remove guinea-worm. The plant is said to be used in decoction for washing horses, and the leaves to be admixed with other materials for plastering mud floors. Amongst the Hausa, superstitious uses are prominent as a love-philtre, etc. A charm prepared from the plant and tied around the forehead attracts the ‘glad eye’ (kalankuwa means a headband); or a youth hides the plant in a maiden’s house. To the Ijaw this species is male, its female counterpart being
Melanthera scandens (Compositae) (Burkill, 1985)
Manihot esculenta
Scientific Classification
Kingdom: Plantae
(unranked): Angiosperms
(unranked): Eudicots
(unranded) Rosids
Order: Malpighiales
Family: Euphorbiaceae
Subfamily: Crotonoideae
Tribe: Manihoteae 43
Genus Manihot
Species: M. esculenta
Binomial name Manihot esculenta. Crantz
The cassava root is long and tapered, with a firm homogeneous flesh encased in a detachable rind, about 1mm thick, rough and brown on the outside. Commercial varieties can be 5 to 10 cm in diameter at the top, and around 15 cm to 30 cm long. A woody cordon runs along the root's axis. The flesh can be chalk-white or yellowish.
Cassava roots are very rich in starch and contain significant amounts of calcium (50 mg/100g), phosphorus (40 mg/100g) and vitamin C (25 mg/100g). However, they are poor in protein and other nutrients. In contrast, cassava leaves are a good source of protein (rich in lysine) but deficient in the amino acid methionine and possibly tryptophan
Fig. 5: Picture o fManihot esculenta (cassava) plant
44
Nutritional profile of cassava
Cassava root is essentially a carbohydrate source. Its composition shows 60–65 percent moisture, 20–31 percent carbohydrate, 1–2 percent crude protein and a comparatively low content of vitamins and minerals. However, the roots are rich in calcium and vitamin C and contain a nutritionally significant quantity of thiamine, riboflavin and nicotinic acid. Cassava starch contains 70 percent amylopectin and 20 percent amylose. Cooked cassava starch has a digestibility of over 75 percent. Cassava root is a poor source of protein. Despite the very low quantity, the quality of cassava root protein is fairly good in terms of essential amino acids. Methionine, cysteine and cystine are, however, limiting amino acids in cassava root.
Cassava is attractive as nutrition source in certain ecosystems because cassava is one of the most drought-tolerant crops, can be successfully grown on marginal soils, and gives reasonable yields where many other crops do not grow well. Cassava is well adapted within latitudes 30° north and south of the equator, at elevations between sea level and 2000 meters above sea level, in equatorial temperatures, with rainfalls of 50 millimeters to five meters annually, and to poor soils with a pH ranging from acidic to alkaline. These conditions are common in certain parts of Africa and South America.
Cassava is a highly productive crop in terms of food calories produced per unit land area per unit of time, significantly higher than other staple crops. Cassava can produce food calories at rates exceeding 250,000cal/hectare/day compared with 176,000 for rice, 110,000 for wheat, and 200,000 for maize (corn).Cassava, like other 45 foods, also has anti-nutritional and toxic factors. Of particular concern are the cyanogenic glycosides of cassava (linamarin and lotaustralin). These, on hydrolysis, release hydrocyanic acid (HCN). The presence of cyanide in cassava is of concern for human and for animal consumption. The concentration of these anti-nutritional and unsafe glycosides varies considerably between varieties and also with climatic and cultural conditions. Selection of cassava species to be grown, therefore, is quite important. Once harvested, bitter cassava must be treated and prepared properly prior to human or animal consumption, while sweet cassava can be used after simple boiling.
Ethnomedicine
The bitter variety leaves are used to treat hypertension, headache, and pain. As cassava is a gluten-free, natural starch, its use in Western cuisine as a wheat alternative for sufferers of celiac disease is becoming common. (Wikipedia, 2014)
Gongronema latifolium Scientific Classification Kingdom: Plantae Subkingdom: Viridaeplantae Phylum: Magnoliophyta Subphylum: Spermatophytina Infraphylum: Angiospermae Class: Magnoliopsida Subclass: Lamiidae
46 Superorder: Gentiananae
Order: Gentianales Family: Asclepiadaceae Genus: G. latifolium Botanical name: Gongronema latifolium Benth .
Fig. 6: Gongronema latifolium (Utazi) plant
Gongronema Latifolium belongs to the family of asclepiadaceae family. The plant common name is amaranth globe. The parts commonly used are leaves, stem and root.
The origin of the plant is traced to Nigeria in West Africa. Gongronema Latifolium is called madumaro by Yoruba ethnic group in Nigeria. It is a rainforest plant which has been traditionally used in the South Eastern part of Nigeria over the ages for the management of diseases such as diabetes, high blood pressure etc. 47
The result of phytochemical screening of ethanolic root extract of Gongronema
Latifolium by Antai et al., (2009) reported that the root contains polyphenols in
abundance. Alkaloids, glycosides and reducing sugars were also present in moderate
amounts. Atawodi, (2005) reported that Gongronema Latifolium has antioxidant
potential. Sonibar. and Gbile, (2008) reported that Gongronema Latifolium has anti-
ashmatic potential. Akuodor et al., (2010) reported that Gongronema Latifolium has anti- plasmodia activity; this supports the traditional use of the leaf extract of the plant for local treatment of malaria. Akuodor, (2010) and his team in their review state that
Gongronema Latifolium is used in South Eastern Nigeria to treat various ailments such as cough, loss of appetite, malaria and stomach disorders.
The liquor usually obtained after the plant is sliced and boiled with lime juice or infused with water over three days is usually taken as a purge for colic and stomach pains. Various parts of the plant, particularly the stems and leaves are used as chewing sticks or liquor in Sierra Leone. It is also used to treat symptoms related to worm infections. Gongronema Latifolium is good for maintaining healthy blood glucose level, and has antibacterial activity.
The earlier studies of the researchers have shown that ethanolic leaf extract of
Gongronema Latifolium possess analgesic effects. The stem bark extract of
Gongronema Latifolium has equally been reported to have anti-ulcerative property.
Akuodo et al., (2012) in a collaborative study reported that ethanol extract of
Gongronema Latifolium leaves when evaluated were found to possess anti-ulcer,
48 analgesic and antipyretic activities. The plant enjoys reputation as a remedy for inflammation, bacteria, ulcer, malaria, diabetes and analgesic.
Antioxidant properties of the medicinal plants under study
The medicinal plants in this study are known plants with antioxidant properties. The antioxidant properties of these plants have been investigated for instance the work on the `In vitro Antioxidant Activity of Extracts from Chromolaena Odorata by Amatya and
Tuladhar, (2011), Showed that The antioxidant activities of ethanolic extracts of leaf, stem, root and defatted flower parts were evaluated by β-carotene bleaching and 1, 1- diphenyl-2-picryl-hydrazyl (DPPH) free radical scavenging assays. Leaf and flower revealed the good antioxidant activities in both methods. In β-carotene bleaching assay, ethanol extracts of leaf and flower exhibited the relative antioxidant activity values as
0.95± 0.029 and 0.86±0.05, respectively. The Antioxidant properties of chlorophyll- enriched and chlorophyll-depleted polyphenolic fractions from leaves of Vernonia amygdalina and Gongronema latifolium , had also been reported (Fasakin et al.,2011).
Faleye and Ogundaini, (2012) reported that compound with strong antioxidant properties were isolated from Aspilia africana . For Psidium guajava, it had been shown that guava leaf extracts comprise effective potential source of natural antioxidants (Qian and Nihorimbere, 2004). The antioxidant activities of Manihot esculenta had also been evaluated. For instance it had been shown that phenolic compounds contributed to the antioxidant activity of cassava (Yi, et al., 2011). It is very clear from previous studies that these medicinal plants have antioxidant activities. 49
CHAPTER THREE
Experimental
3.1. Materials and apparatus : These include laboratory and research microscope with photomicrograph, spectrophotometer (spectro 21D of Pec medicals, USA,) electric heated oven, Gallenkamp for ash value, etc.
3.2. Collection of test and control samples of medicinal plants.
The samples used in this experiment were collected from the appropriate sites. The
sites chosen satisfy the condition of been oil polluted environment, for test samples
and non oil polluted environment for control samples. Thus two sites were chosen
for the collection of the samples. These were oil polluted area of Imo River in Afam
of Oyigbo local government area of River state of Nigeria. This area has long history
of oil pollution. One can see oil floating on top of the river. The oil because of having
been released on the water for long time has undergone some changes and thus
having more effects on the water and its environments. The other site is where the
control samples were collected. The site is in Oguta of Oguta local government area
and Mgbidi in Oru west local government both in Imo state of Nigeria. The samples
were collected during the day, and were authenticated by Mr. A.O Ozioko (Botanist)
of International Centre for Ethinomedicine and Drug development, Nsukka, Enugu
State.
50
3.3. Quantitative determination of ascorbic acid contents of the samples
The ascorbic acid contents of the medicinal plants used were determined in
accordance with the method in official method of analysis of the Association of
Official Analytical Chemist (AOAC), 2005.
0.5g of each of the samples was macerated with 20mls of 4.0 % oxalic acid for
10mins, centrifuged for 5min. Then 1ml of the supernatant was transferred into duplicate tubes. Then 9mls of 2, 6, dichlorophenol indophenols (12mg/L) was added, the absorbance was taken at 520nm against water. The amount or the ascorbic acid contents was calculated in mg/100g of samples and recorded for both the test and the control samples.
3.4. Quantitative determination of the total phenolic compounds in the samples
This was determined by Folin Ciocalteu method; 0.1g was weighed and macerated with
50mls of 80% ethanol for 20minutes, and centrifuged for 5minutes. 1ml was transferred into triplicate tubes. 4ml of water was added, and mixed 0.5ml of phenol reagent was added mixed and allowed to stand for 5minutes. 2mls of 20% sodium carbonate was added mixed and allowed to stand for 30minutes. The absorbance was taken at 650nm the blank. The result was calculated in mg/100ml and recorded for both the test and control samples.
51
3.5 Quantitative determination of lipid peroxide content of the samples
This was determined by thiobarbituric acid (TBA) method; 0.1g of sample was weighed and added to 2mls of water and 3mls, 20% tri-chloro-acetic acid. It was macerated for
20minutes, centrifuged for 10minutes at 3000rpm. 1ml of the supernatant was transferred into triplicate test tubes. 2mls of 1% thiobarbituric acid was added in 20%
T.C.A. It was boiled in the water bath for 30minutes and cooled. The absorbance was taken at 600nm. The lipid peroxide content was calculated in mg/100g and recorded.
3.6. Determination of the ash values of the leaves of the samples
The ash values of the medicinal plants were determined by the parametric method
in accordance with the method in official method of analysis of the Association of
Official Analytical Chemist (AOAC), 2005
a. Total ash ; Ten crucibles were weighed separately and the weight of each of them
were noted and recorded. Weight of the leaves to be used for each of the samples
for both the tests samples and the control samples, were taken and recorded. The
leaves were transferred into the crucible, the weight of the crucible and the leave for
each of the ten samples were taken and recorded. The crucibles and the samples
were put in the hot electric furnace and heated at about 500 0c until the samples
were well ashed. After this the ash and the crucible for each of the samples were
weighed after cooling in the desiccators and 52 their weight were noted and recorded. The percentage total ash for each of the samples both for the test and control samples were calculated and recorded
b. Water soluble ash : Half of the amount of total ash was weighed and put into the
test beaker; 10ml of distilled water was added and then boiled. This was filtered with
a filter paper which had been weighed previously. The filter paper with residue was
dried. It was re-weighed and the weight of the residue determined. This was
subtracted from the original weight to determine the value of the water soluble ash.
The percentage water soluble ash was calculated and recorded for all the test and
control samples of the medicinal plants.
c. Acid insoluble ash: Half of the total ash was weighed and put in a small beaker
10ml of dilute HCl was added to the beaker boiled and shaken very well. It was then
filtered with a previously weighed filter paper, and dried. The residue was weighed
with the filter paper. The weight of the residue was then determined by subtracting
the weight of the filter paper from the total weight of the residue and filter paper. The
percentage acid insoluble ash was calculated and recorded for both the test and
control samples.
d. The sulphated ash; 0.5g of each of the samples were weighed and put in the
crucibles and heated to ash, then 2ml of concentrated sulphuric acid was added and
then heated for the sulphated ash to occur. The sulphated ash was weighed and the
percentage sulphated ash was calculated and recorded.
53
3.7. The epidermal cell and stomata
The epidermal cells and stomata of the leaves were examined with microscope and the photographs taken using photomicrograph. The stomata, epidermal cells, as well as the presence or absence of stress were well identified, by examining the photomicrograph and noting the difference and similarities between the test samples and the control samples. These were done for both the upper and the lower epidermis of the leaves.
3.8. Phytochemical screenings of the medicinal plants
The phytochemical screenings of the medicinal plants were carried out for each group of plant secondary metabolites as follows;
a. Flavonoids:
Total flavonoid content was evaluated according to a colorimetric assay with Aluminum
Chloride. ( Zhishen et al., 1999). 1 ml aliquot of plant extract was added to a 10 ml volumetric flask containing 4 ml of distilled water, followed by the addition of 0.3 ml of solution of NaNO 2 (0.5 g/L). After 5 min, 0.3 ml of a 1 g/L solution of AlCl 3 was added
and 6 min later, 2 ml of NaOH (1 mol/L) was added to the mixture. The total volume was
made up to 10 ml with distilled water. The solution was mixed and the absorbance was
measured at 510 nm against water blank. Catechin was used as the standard for the
construction of a calibration curve and the concentrations were
54 expressed as catechin equivalents (mg/100g). The flavonoid contents were calculated and recorded.
b. Tannins
Tannin content was determined by (Khan et al., 2011) method: 1g of the sample was macerated with 50ml methanol, filtered; 5mls of it was put into 3 test tubes. 0.3ml of ferric chloride in 0.1M HCl was added, then 0.3ml of 0.0008M K(CN) (1:10 dilution
0.008M) , mixed and the absorbance was read at 720nm. The amount of tannin was calculated and recorded.
c. Steroids The steroid content was determined by the method of (Alexander and Griffiths, 1993).
1g of the sample was macerated with 20mls of ethanol, 2mls of it was taken into 3 test tubes, then 2mls of cholesterol colour reagent was added and left for 30mins. The absorbance was taken at 550nm. The amount of the steroid was calculated and recorded.
d. Terpenoids The terpenoid content was determined according to method of (Snell and Snell,
1962). 1g of the sample was macerated with 50mls of ethanol and filtered, then 2.5mls of the filtrate was added to 2.5mls of 5% aqueous phosphomolybdic acid solution and
2.5ml concentrated sulphuric acid was gradually added and mixed. It was allowed to stand for 30min to cool and then made up to 12.5ml with ethanol. The absorbance was taken against the reagent blank at 700nm. The amount of the terpenoids was calculated and recorded.
55
e. Saponins
The saponin content was determined by method of (Uematsu, et al., 2000). 1g of the
sample was macerated with 10mls petroleum ether. It was decanted into a beaker and to the residue was added 10ml of petroleum ether, macerated and decanted into the same beaker. The combined filtrate was evaporated to dryness. The residue was dissolve with 6mls of ethanol and 2mls was transferred into a test-tube and chromogen was added and left for 30min (chromogen = 2:23ml of Iron stock: conc H 2SO 4). The
absorbance was taken at 550nm. The saponin content was then calculated and
recorded.
f. Alkaloids
This was done using (Harborne, 1923) method. 1g of the sample was macerated with
20mls of ethanol and 20% H 2SO 4 (1:1) for 5mins and filtered. To 1ml of the filtrate was
added 5mls of 60% H 2SO 4. Then after 5mins, 5mls of 0.5% formaldehyde in 60%
H2SO 4 was added and mixed and allowed to stand for 3h. Then the absorbance was read at 565nm. The amount of the alkaloids was then calculated and recorded.
g. Protein
This was determined by the (kjeldahl, 1883) method: 2ml of sample was added to 3mls of water, 0.5ml of Nessler`s reagent was added and allowed to stand for 15min. The absorbance was taken immediately at 490nm.The amount of protein was then calculated and recorded.
h. Resin:
This was done by (Michael, et al., 2009) method; 1ml of acetone extract 56 was transferred into a centrifuge tube and centrifuge for 5min, 0.2ml of the supernatant was transferred into another tube containing 1.8ml acetone nitride and the absorbance was read at 272nm against the blank. The resin content was then calculated and recorded
i. Sugar:
This was determined by (Alexander and Griffiths, 1993) method ; to 1ml of the test
sample was added1ml of alkaline copper reagent. It was boiled for 5mins.1ml of
phosphomolybdic acid reagent was added. 7ml of water was added to make up to 10ml.
then the absorbance was taken at 420nm. The amount of sugar was then calculated
and recorded.
3.9: statistical analysis
Data obtained were analyzed using on-way analysis of variance (ANOVA).
Differences between Means were accepted at P< 0.05. Results were presented as
Mean ± SEM.
57
CHAPTER FOUR
RESULTS
4.1. Quantitative determination of ascorbic acid contents of the samples
From the table1below, P. guajava was noted to have ascorbic content value of
105.5±0.6 and 88.7±0.4 in the test and the control sample respectively. C. odorata have the ascorbic acid content value of 99.1±0.4 and 59.1±0.7 in the test and the control sample respectively. The ascorbic acid content value of A. africana is 104.9±0.8 and
85.5±0.7 in the test and the control samples respectively. These plants showed significant difference in the ascorbic acid content values for the test and the control samples. However, there was no significant difference in the ascorbic acid content values of the test and the control samples of G. Latifolium (test=157.7±1.1, control=156.5±1) and M. esculenta (test=21.5±.5, control=18.6±0.55) as the values obtained were relatively close.
Table1: Result for the ascorbic acid contents of the samples
Medicinal plant Test (mg/100g) Control (mg/100g) Psidium guajava 105.5 ±0.6 88.7±0.4
Gongronema latifolium 157.7±1.1 156.5±1
Chromolaena odorata 99.1±0.4 59.1±0.7
Manihot esculenta 21.5± 0.5 18.6±0.55
Aspilia africana 140.9±0.8 85.5±0.7
Values are mean ± SEM, n = 3, p <0.05 58
4.2 . Quantitative determination of the total phenolic compounds in the samples
On the table 2 below, it was noted that the total phenolic compounds in P. guajava is
673.9±10.9 and 573.5±12.2 for the test and the control samples respectively. That of G.
latifolium is 13.9 ± 0.45 and 196.3±1.9 for the test and the control samples respectively
C. odorata have values of 391.3±5.4 and 300.8±5.1 for the test and the control samples respectively. M. esculenta have total phenolic compounds of 194±1 and 236.4±5.1 for
the test and the control samples respectively, while that of A. africana is 93.3±0.65 and
689±5.5 for the test and the control samples respectively. Though there were significant
differences in the amount of the total phenolic compounds of the test and the control
samples of all the plants, that of G. latifolium and A. africana very much when compare
to their test and the control samples.
Table 2: Result for the total phenolic compounds in the samples Medicinal plant Test (mg/100ml) Control (mg/100ml)
Psidium guajava 673.9±10.9 573.5±12.2
Gongronema latifolium 13.9 ± 0.45 196.3±1.9
Chromolaena odorata 391.3±5.4 300.8±5.1
Manihot esculenta 194±1 236.4±5.1
Aspilia africana 93.3±0.65 689±5.5
Values are mean ± SEM, n = 3, p <0.05
59
4.3. Quantitative determination of lipid peroxide content of the samples
The result for the lipid peroxide contents of the samples showed that there were significant difference between the values for the test and the control samples for all the plants. The values for P. guajava (188.4 ± 0.93 and 252.8± 0.7) for the test and the control samples respectively, G. latifolium (120.1± 1.1and 361.2 ± 1.6) for the test and the control samples respectively and A. africana (91.2 ± 0.3 and 124.3± 1.1) for the test and the control samples respectively, showed that the lipid peroxide content for the control samples were higher than those of the test samples. The values for C. odorata
(350.6± 5.2 and 195.1± 1.1) for the test and control samples respectively and M. esculenta (239.7± 1.3 and 117.8 ± 0.85) for the test and the control samples respectively showed that the lipid peroxide contents for the test samples were higher than those of the control samples.
Table 3: Result for lipid peroxide content of the samples
Medicinal plant Test (mg/100g) Control (mg/100g)
Psidium guajava 188.4 ± 0.93 252.8± 0.7
Gongronema latifolium 120.1± 1.1 361.2 ± 1.6
Chromolaena odorata 350.6± 5.2 195.1± 1.1
Manihot esculenta 239.7± 1.3 117.8 ± 0.85
Aspilia africana 91.2 ± 0.3 124.3± 1.1
Values are mean ± SEM, n = 3, p <0.05
60
4.4. The ash values of the leaves of the samples
a. Total ash
The difference in the total ash values for the test and the control samples are not much, except in the case of A. africana . In the overall the test samples are in the higher side than the control samples
Table 4: Result for Total ash of the samples
Medicinal plant Percentage (%) Total Percentage (%) Total ash Test samples ash Control samples Psidium guajava 6 5.5
Chromolaena odorata 11 10
Manihot esculenta 8 7
Aspilia africana 22 13
Gongronema latifolium 16 14
61
b. water soluble ash
The result for water soluble ash is also similar to that of the total ash value there are
no much variation except in the case of A. africana and G. latifolium
Table 5: Result for the water soluble ash
Percentage (%) water Percentage (%) water soluble Medicinal plants soluble ash, test samples ash, control samples 3 3.5 Psidium guajava
6 5 Chromolaena odorata
5 4 Manihot esculenta
12 5 Aspilia africana
8 5 Gongronema latifolium
.
62
c. Acid insoluble ash
The result for the acid insoluble ash showed that only A. Africana and G. Latifolium
have significant difference between the values for the test and the control samples. On
the overall the test samples have higher values than the control samples.
Table 6: Result for the acid insoluble ash
Percentage ( %) acid insoluble Percentage (%) acid insoluble Medicinal plants ash, test samples ash, control samples 0.4 0.5 Psidium guajava
2 1.5 Chromolaena odorata
0.6 0.41 Manihot esculenta
4 2 Aspilia africana
8 2 Gongronema latifolium
63
d. Sulphated ash
There was significant difference in test and control sample of the sulphated ash when we consider the result of C. odorata , (4 and 1.62) for test and control sample respectively though the value for the test sample is on the higher side. On the result for
M. esculenta (4 and 6) and G. latifolium (7 and 14.5) for the test and the control samples respectively there was significant difference, but with the values for the control samples on the higher side. There was no significant difference for the values of the test and the control samples for P. guajava and A. africana.
Table 7: Result for the sulphated ash
Percentage (%) sulphated Percentage (%) sulphated Medicinal plant ash, test samples ash, control samples 5 5.5 Psidium guajava
4 1.62 Chromolaena odorata
4 6 Manihot esculenta
6 6 Aspilia africana
7 14.5 Gongronema latifolium
64
4.5. Phytochemical screening of the medicinal plants
Table 8: Result for Quantitative screening of the phytochemical contents of the medicinal plants, in mg/100g, but proteins are in percentage . Alkaloids Terpenoids Saponin Tannins Flavonoids Steroids Resin Sugar Proteins Medicinal plants
P. guajava (t) 12012.5±100 7686.1±57 230.3±4.9 2304.7±7.2 858.6±3.7 3519.0±7 53.4±0.8 893.3±3.4 25.7±0.4
P. guajava (c) 14166.7±16.7 9306.2±96.9 219.1±4 3178.7±10.7 171.7±2.2 8420.7±10 34.6±1.2 969.6±15 22.5±0.7
C. odorata (t) 5416.7±16.7 5453.5±23.3 240.2±2.4 984.4±9.8 255.7±2.2 1512.1±12 48.9±1.1 794.5±5.3 15.7±0.5
C. odorata (c) 9652.1±24 3302.3±28.9 240.7±2.2 710.9±9.6 226.0±2 1008.6±19 39.7±1.7 469.2±4.5 13.0±0.8
M. esculenta (t) 4929.2±10 9136.6±31.7 227.3±4.9 578.7±5.7 152.3±1.9 1720.7±8 26.5±0.8 869.3±9.9 29.2±0.4
M. esculenta (c) 1943.8±23.6 3639.5±20.3 221.5±2.3 687.5±5.8 175.5±1.8 1260.4±10 21.4±0.8 98.4±2.3 32.7±0.4
A. africana (t) 22429.2±85.4 2104.7±7.7 175.2±2.4 812.5±3.8 150.0±1.5 636.2±6.9 69.4±1.3 426.5±4.8 17.6±0.6
A africana (c) 13875.0±62.5 3124.0±23 158.3±2.9 3085.9±7.1 190.2±3.4 1082.8±6 57.7±1.2 589.2±5.3 25.0±1
G. latifolium (t) 2012.5±23.8 2337.2±11.4 237.4±2.8 617.2±4.9 39.7±1.2 1148.3±3 41.9±1.1 594.6±2.7 16.6±0.7
G. latifolium (c) 3193.8±28.1 3728.6±25.6 226.2±1.9 867.2±4.9 171.7±1.2 1260.4±6 36.4±0.8 665.2±7.4 20.2±0.9
Values are ± SEM, n =3, p <0.05
65
4.6. The epidermal examination
The epidermal characteristics of the leaves of the test and control samples
of the medicinal plants are as shown in Figs. 7 – 16.
Fig.7a: photomicrograph of upper epidermis fig. 7b: photomicrograph of upper of test sample Aspilia africana, showing epidermis of control sample of A. some damaged stomata and cuticular africana , with normal cells and stomata abrasion MAG X400 MAG X400
Fig.8a: photomicrograph of lower Fig.8b: photomicrograph of lower epidermis of test sample of A. africana epidermis of control sample of A.africana showing normal stomata and showing normal epidarmal cell and stomata cuticular abrasion MAG X400 MAG X400 66
Fig. 9a: photomicrograph of upper Fig.9b: photomicrograph of upper epidermis of test sample of C. odorata epidermis of control sample of C. odorata showing damaged stomata and cuticular showing normal cell and no stomata abrasion MAGX400
Fig.10: photomicrograph of lower epidermis of test sample of C. odorata showing some damaged stomata most of the epidermal cells are normal with few cuticular abrasion MAG X400
67
Fig.11a photomicrograph of lower Fig.11b photomicrograph of lower epidermis of test sample of P. guajava epidermis of control sample of P. guajava with some stomata opened and closed with opened stomata MAG X400 MAG X400
Fig.12a: Photomicrograph of upper fig.12b: Photomicrograph of upper epidermis epidermis of test sample of P. guajava of control sample of Psidium guajava showing some closed and damaged with some opened stomata and leaf vein stomata, but with normal epidermal cells
MAG X400 MAG X400
68
Fig. 13a: Photomicrograph of test sample fig. 13b Photomicrograph of control sample of Manihot esculenta lower epidermis of Manihot esculenta lower epidermis with normal epidermal cell and stomata with normal epidermal cell and no stomata MAGX400 MAG. X400
Fig. 14a: Photomicrograph of test sample fig. 14b: Photomicrograph of control sample of Manihot esculenta upper epidermis of Manihot esculenta upper epidermis with normal stomata and epidermal cell with normal epidermal cell and stomata MAG. X400 MAG. X400
69
Fig. 15a: Photomicrograph of test sample fig.15b: Photomicrograph of control sample of Gongronema latifolium lower epidermis of Gongronema latifolium lower epidermis with normal epidermal cell and stomata with normal epidermal cell and stomata MAG. X400 MAG. X400
Fig.16a: Photomicrograph of test sample fig.16b: Photomicrograph of control sample of Gongronema latifolium upper epidermis of Gongronema latifolium upper epidermis with normal epidermal cell and stomata with normal epidermal cell and stomata MAG. X400 MAG. X400
70
CHAPTER FIVE
DISCUSSION AND CONCLUSION
Psidium guajava : This plant which is an excellent source of antioxidant vitamin-C
(nutrition and you, 2013), had ascorbic acid content in which there a significant differences between the value of that of the test and the control samples. The ascorbic acid content of the test is on the higher side. Also the same observation was made with regard to the total phenolic compounds and the lipid peroxide, were significant differences occur between their content for the test and the control samples. The contents of the test samples are more than those of the control samples. Moreover
Psidium guajava has antioxidant properties which are attributed to the polyphenols found in the leaves (Taylor, 2004). The production of these phenolic compounds is expected to increase in condition of oxidative stress.
However, there were no significant difference between the values for the test samples and the control samples with regard to the result on the ash values. On the stomata and epidermal cells, it is evidence from the photomicrograph that the stomata of the plant are well distributed on both the upper and lower epidermis, except that the stomata on the upper epidermis on the test sample are closed and some are damaged. This is possible since it has been shown by (McAinsh, 1994)) that oxidative stress affects the stomata response. The cells of both the control and the test samples were not damaged and there are little differences in the sizes of the stomata openings for those that are opened.
71
The result of the ascorbic contents, phenolic compounds and lipid peroxides as oxidative stress indicators showed that the different environments had different effects on the medicinal plants, thereby giving positive indication of oxidative stress. This is because most environmental stresses affect the production of reactive oxygen species in plants causing oxidative stress (Smirnoff, 1993, Hendry, 1994). However with regard to the ash values, stomata and epidermal cells there were no much difference and thus did not give positive indication to oxidative stress.
The result of the phytochemical screening showed that while there were some which had significant difference between their values for the test and the control samples, others had no much difference. The values of alkaloid, tannins terpenoids, steroids and sugar showed significant difference in which the control sample had more of these chemicals than the test sample, while only flavonoid showed significant difference in which the value for the test sample is more than that of the control sample. The remaining phytochemical namely resins saponins and proteins had no significant difference in their values for the test and the control samples.
The environment might have affected the production of some of these phytochemical.
This might have lead to the significant difference in the quantity of these phytochemical between the test and control samples. Since they are more in the control sample, it showed that the productions of these phytochemical are negatively affected in the test sample from the crude oil polluted environment. Flavonoids which have higher content in the test sample may be link to the environmental factor. P. guajava on its own is a good source of flavonoids and some flavonoids are antioxidants. It could be that the 72 crude oil polluted environment might have induced stress in the plants leading to increase production of the flavonoid to serve as antioxidant.
Gongronema latifolium : The values of ascorbic acid content of the test and control samples did not show any significant difference. However, since this plant have antioxidant potential (Atawodi, 2005), these two oxidative stress indicators namely total phenolic compounds and lipid peroxide were affected. Thus there were significant difference between the values for the test and the control samples when we consider the results of the total phenolic compounds and the lipid peroxide. The control sample had more of these than the test sample.
The value of the ash values showed that the test sample higher values for total ash, water soluble ash and acid insoluble ash with significant differences. However the control sample had higher value of the sulphated ash than the test sample.
On the stomata and the epidermal cells, it was observed that there were not much difference between the test and the control samples. The stomata and the epidermal cell did not show any evident of stress as a result of the environmental differences
The phytochemical screening showed that there were significant differences in amount of alkaloids, terpenoids, tannins, flavonoids and sugar found in the test and the control samples. The control sample had more of these phytochemical than the test sample. There were no significant differences in the amount of resins, proteins, steroid and saponins found in the test and the control samples. Though the environment affected the amount of some of these phytochemical produced by these plants there were no sufficient evidence to show that any of these sample is under oxidative stress. 73
Chromolaena odorata: The result of the ascorbic acid content showed that there were no significant difference between the quantity in the test and the control. However, there were significant difference in the amount of the phenolic compound and lipid peroxide found in the samples, were the amount found in the test sample were more than those of the control samples
There were no significant differences in the values of the ash values between the test and the control samples of this plant, except for the value of the sulphated ash in which that of the test is more than that of the control. Thus the environment did not affect the ash values for the two samples from different environments.
Examination of the stomata and the epidermal cells of the plant samples showed that some of the stomata of the test sample were destroyed and the epidermal cells were also affected. This is possible because it had been shown by (Samuilov et al ., 2008) that the action of reactive oxygen species can lead to the death of guard cells. Whether this was as the result of stress from environmental factor is not very clear as there were varied results from other oxidative stress indicators.
On the phytochemical screening, it was observed that terpenoid, tannins, steroids and sugar had significant differences between the test and the control samples. They were more in the test sample than the control sample. Only alkaloids were more in the control sample than the test sample. There were no significant differences in the amount of resin, proteins, saponins and flavonoid found in the test and control samples of the plant.
74
The fact that the amounts of phytochemicals produced by the test sample were not affected by the crude oil polluted environment showed that the plants were not under serious stress. Though the result on total phenolic compound and lipid peroxide suggest presence of oxidative stress, they are at this level not sufficient for us to conclude that the test sample was under oxidative stress, as other oxidative stress indicators did not give positive results.
Manihot esculenta: The amount of the ascorbic acid content and the total phenolic compounds, which is responsible for the antioxidant activity of the plant (Yi et al ., 2011), found in the test and the control samples had not much significant differences. The values for the lipid peroxide was higher in the test sample when compare with that of the control sample, and this is not sufficient to prove the presence of oxidative stress in the plant.
The result of the ash values showed that there were no any significant difference in all of them between the test and the control samples.
A very close look at the epidermal cells and the stomata of the two samples showed that there was no significant difference between the test and the control samples. The stomata and the epidermal cells were within normal appearance and cannot be said to be under stress. The environment did not affect the test sample from crude oil polluted environment.
The quantitative screening phytochemical contents of the test and control samples showed that there were significant differences and more of alkaloids, terpenoids, steroids and sugars in the test sample than the control samples. Only tannins were 75 more in the control sample than the test sample. There were no significant difference in amount of resins, saponin, flavonoids and proteins found in the test and the control samples of the plant. The crude oil polluted environment did not have any adverse effect on the production of these phytochemicals, and the test sample cannot be said to be under stress.
Aspilia africana: The ascorbic acid content was higher in the test sample than the control sample, but there were more of the phenolic compounds in the control sample where there was wide difference between the amount in the test sample and the control sample also, the quantity of the lipid peroxide was more in favour of the control sample than the test sample though the difference was not much.
The result of the ash values showed that there were much significant differences between the test and control sample. The values of the total ash, acid insoluble ash and water soluble were higher for the test sample than those of the control sample. Only the sulphated ash gave equal values for both the test samples and control samples.
Evaluation and careful observation of the stomata and the epidermal cell showed interveinal necrosis and cuticular abrasions on the test sample than the control sample.
This is because it has been shown by (Baszynski et al., 1980) that plants can present
interveinal necrosis in time of oxidative stress. It also affect stomata resistance Thus
from the photomicrograph and the picture of the samples, the test samples showed
some element of stress.
On the phytochemical screening, the values of alkaloid, terpenoids, and tannins are
higher in the test sample than the control sample with significant differences. On the 76 other hand the values of steroids and sugar are more in the control sample than the control sample with significant differences. There were no much difference in the values of saponins, flavonoids, resins and proteins in the test and control samples.
Looking at the result of the phytochemical contents, it was observed that the phytochemical contents were not affected by the environment when we compared the test result with that of the control. In fact, some of the phytochemical like alkaloids, terpenoids and tannins were more in the test sample than the control sample. When consider the fact that some of other oxidative stress indicator were positive for oxidative stress, we may not rule out of the possibility of other changes manifesting later.
Conclusion
In conclusion, from the discussion above, we say that the pollution of the environment by crude oil spills has a lot of effects on some medicinal plants found in these environments of which one is oxidative stress, as shown by the oxidative stress indicators. However, the extent of the oxidative stress in these medicinal plants differ from one plant to another some show strong evidence of oxidative stress, while others show little or no oxidative stress at all. From this investigation we conclude that only
Psidium guajava showed sufficient evidence of oxidative stress.
77
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86
Appendix
Ascorbic acid standard curve
0.4
0.35 y = 0.0693x R² = 0.9968 0.3
0.25
Absorbance 0.2
0.15 0 0 1 0.07 0.1 2 0.14 3 0.2 0.05 4 0.29 0 5 0.34 0 2 4 6
Concentration (mg/100ml )
Flavonoid std curve 0.9
0.8 y = 0.0162x 0.7 R² = 0.9986 0.6 0.5 0.4
Absorbance 0.3 0 0 0.2 10 0.162 20 0.341 0.1 30 0.498 40 0.649 0 50 0.796 0 20 40 60 Concentration (mg/100ml) 87
Crude Nitrogen std curve (current ) 0.7 y = 0.1885x 0 0 0.6 R² = 0.9725 0.303 0.055 0.5 0.606 0.094 0.909 0.166 0.4 1.212 0.265 0.3 1.515 0.331 Absorbance 1.818 0.318 0.2 2.121 0.406 2.424 0.391 0.1 2.727 0.51 0 3.03 0.607 0 1 2 3 4
Concentration (ug/ml.)
Lipid peroxidation standard curve (curr) 1.4
1.2 y = 0.0666x R² = 0.9193 1
0.8 Absorbance
0 0 0.6 1.67 0.006 3.33 0.013 5 0.084 0.4 6.67 0.415 8.33 0.62 10 0.732 11.67 0.66 0.2 13.33 0.985 15 0.978 16.67 1.201 0 0 5 10 15 20 Concentration (mg/100ml.) 88
Saponin standard curve
2.5
y = 3.0225x 2 R² = 0.9303
1.5 0 0 0.09 0.374 0.18 0.702 1 Absorbance 0.27 1.108 0.36 1.287 0.45 1.454 0.5 0.54 1.632 0.63 1.85 0.72 1.904 0 0 0.2 0.4 0.6 0.8 Concentration (mg )
0.7 Tannin std curve 0.6 y = 0.0328x 0.5 R² = 0.9945
0.4
0.3 Absorbance
0 0 0.2 2 0.077 4 0.118 0.1 6 0.202 8 0.253 0 10 0.308 12 0.409 0 5 10 15 20 25 14 0.47 Concentration (mg/100ml.) 16 0.547 18 0.56 20 0.663
89
Terpenoid std curve 0.6
y = 0.0433x 0.5 R² = 0.9432
0.4
0.3 Absorbance 0.2
0 0 2.48 0.068 0.1 3.1 0.19 4.14 0.218 6.2 0.305 0 12.4 0.499 0 5 10 15
Concentration (mg/ml.)
ALKALOID STANDARD CURVE
1.4 1.2 y = 0.0123x 1 0.8 0.6 0.4 ABS ABS 0.2 0 0 50 100 150 CONC(mg/dl)
90
1.8 Resin standard curve 1.6 y = 0.068x 1.4 R² = 0.9999 1.2
1
0.8 Absorbance 0.6
0.4
0.2 0 0 5 0.34 0 10 0.676 15 1.021 0 10 20 30 20 1.352 Concentration (mg/100ml.) 25 1.706
Total phenol std curve 0.45 y = 0.1759x + 0.0634 0.4 R² = 0.9562 0.35 0.3
0.25 0 0 0.2 0.2 0.128 0.4 0.146 Absorbance 0.15 0.6 0.188 0.8 0.213 0.1 1 0.244 1.2 0.29 0.05 1.4 0.317 1.6 0.339 0 1.8 0.368 0 0.5 1 1.5 2 2.5 2 0.399 Concventration (mg/100ml,)
91
160
140
120
100 test 80 control 60
40
20
0 P. guajava G. latifolium C. odorata M. esculenta A. africana
Histogram chart 1, showing the ascorbic acid content of the of the leaves of the test and control samples of the medicinal plants in mg/100g
25
20
15 test
10 control
5
0 P. guajava C. odorata M. esculenta A. africana G. latifolium
Histogram chart 2, showing the percentage total ash of the leaves of the test and control sample of the medicinal plants
92
12
10
8 test 6 control 4
2
0 P. guajava C. odorata M. esculenta A. africana G. latifolium
Histogram chart 3, showing the percentage water soluble ash of the leaves of the test and control samples of the medicinal plants
8 7 6 5 4 3 2 test 1 control 0 P. guajava C. odorata M. esculenta A. africana G. latifolium
Histogram chart 4, showing the percentage acid insoluble ash of the leaves of the medicinal plants samples
93
16 14 12 10 test 8 control 6 4 2 0 P. guajava C. odorata M. esculenta A. africana G. latifolium
Histogram chart 5, showing the percentage sulphated ash of the leaves of the test and control samples of the medicinal plants
700
600
500
400 test
300 control
200
100
0 P. guajava G. latifolium C. odorata M. esculenta A. africana
Histogram chart 6, showing result for the total phenolic compound in the leave of test and control samples of the medicinal plants, in mg/100g.
94
400
350
300
250 test 200 control 150
100
50
0 P. guajava G. latifolium C. odorata M. esculenta A. africana
Histogram chart 7 showing the result for the lipid peroxide of the test and control samples of the medicinal plants in mg/100g.
15000
10000
5000 test
0 control
Chart 8, showing the phytochemical content of P. guajava in mg/100g
95
10000 8000 6000 4000 test 2000 control 0 Series 3
Chart 9, showing the phytochemical content of C. odorata in mg/100mg
10000
8000
6000
4000 test 2000 control 0
Chart 10, showing the phytochemical content of M. esculenta in mg/100g
96
25000
20000
15000
10000 test 5000 control 0
Chart 11, showing the phytochemical content of A. africana in mg/100mg
4000 3500 3000 2500 2000 1500 1000 test 500 control 0
Chart 12, showing the phytochemical content of G. latifolium in mg/100g