SELECTION, PRODUCT CHARACTERISTICS AND METABOLITE SPECTRUM OF A COMMON STARTER CULTURE FOR FUFU AND USI PRODUCTION BY Kubrat Abiola, OYINLOLA B.Sc., M.Sc. Microbiology (Ibadan) (Matriculation Number: 99994) A Thesis in the Department of Microbiology, submitted to the Faculty of Science in partial fulfilment of the requirements for the degree of DOCTOR OF PHILOSOPHY of the UNIVERSITY OF IBADAN July 2015 ABSTRACT Fermented cassava products like fufu and usi (edible starch) are important staple foods in many African homes. Natural fermentation time is usually long resulting in slower acidification process and inconsistent nutritional composition of products which could be overcome with the use of starter culture. However, most available starter cultures are used for single food fermentation and are uneconomical. This necessitates the development of a starter culture for multiple related food products to reduce cost. Hence this study was designed to produce a common starter culture for the production of fufu and usi. Cassava varieties TME 30572, TME 4(2)1425 and TME 50395 were obtained from the International Institute of Tropical Agriculture, Ibadan and landraces from Bodija market. Fresh, peeled, chipped and grated cassava tubers were spontaneously fermented in the laboratory. Lactic Acid Bacteria (LAB) were isolated from the fermenting mash and identified phenotypically. Genotypically identified starters were selected based on screening for starch hydrolysis, linamarase and pectinase enzyme production, antimicrobial compound production and rate of acidification using standard methods. The starters were utilised singly and randomly combined to initiate fermentation for production of fufu and usi. Un-inoculated fermentation mash served as control. Rate of production of organic acids, various sugars, metabolic enzyme assays, nutritional and anti-nutritional content of the resulting mashes were monitored using standard procedures. Best starter was applied in the final production of fufu and usi. Shelf-life of the products were evaluated and compared with the control. Data were subjected to descriptive statistics and ANOVA technique at p=0.05. Ninety-eight LABs were identified as Lactobacillus plantarum (50.0%), L. acidilactici (12.2%), L. brevis (11.3%), L. fermentum (10.3%), L. delbrueckii (8.2%), L. mesenteroides (6.0%), and L. lactis (2.0%). Screened isolates did not hydrolyse starch but produced pectinase, linamarase alongside hydrogen peroxide, diacetyl and lactate with a rapid decrease in medium pH (6.5 - 3.6). Selected potential starters were genotypically identified as L. pentosus F2A (A), L. plantarum subsp. argentolarensis F2B (B), L. plantarum F2C (C), L. plantarum U2A (G) and L. paraplantarum U2C (I). The best starter combination CGI gave significant reduction in fermentation pH (7.1 - 3.7) and lactic acid ranged between 0.04mg/ mL and 6.9mg/mL. Sugars produced include xylose ii (3.2µg/mL), arabinose (1.4µg/mL), fructose (26.2µg/mL), glucose (30.3µg/mL) and sucrose (99.7g/mL). Enzyme assay revealed peak amylase (10.1U/mL) and pectinase (4.4U/mL) activities at 24 hours as well as linamarase (0.8U/mL) at 48 hours in fufu, whereas, in usi, highest linamarase (0.7U/mL) and pectinase (1.0U/mL) activities were recorded at 72hours with no amylase activity. The CGI-produced fufu and usi had significant reduction in phytate (0.3-0.1mg/g and 0.3-0.27mg/g), tannin (35.4-34.0mg/g and 35.4-32.3mg/g), cyanide (0.1-0.05mg/g and 0.1-0.0mg/g), and moisture (7.3%-5.1% and 7.3%-5.4%) content while total protein content increased (1.0-1.3% and 1.0-1.8%) respectively. Starter fermented fufu and usi had shelf-life of five days while control had three days. The selected starter was able to ferment both fufu and usi to yield products with improved nutritional content, better shelf-life and reduced anti-nutritional composition. This could be employed in the production of indigenous fermented foods. Keywords: Lactic acid bacteria, Starter culture, Fufu and Usi, Shelf-life, Fermented food Word count: 495 ii i DEDICATION This thesis is dedicated to the memory of my late father, Murtala Olayinka OGUNMOLA. iv ACKNOWLEDGEMENTS I am grateful to Almighty Allah, without whom this thesis would not have manifested, and for His continued guidance. I am especially thankful for the gift of life. To him alone be the glory and adoration forever. I owe a lot of gratitude to my Supervisor, Prof. Anthony Abiodun ONILUDE. I appreciate your constant guidance, thorough, conscientious and constructive criticisms and suggestions from the inception of work to date. The totality of your contribution is immeasurable and highly influential to the success of this thesis. Thank you for being my teacher and mentor. My thanks also go to the academic staff of the Department of Microbiology, under the headship of Prof. O. E. Fagade, for their dedication and valuable suggestions at every stage of the study. The enabling environment provided and interest in my progress deserve commendation. Furthermore, the support and assistance from the non-academic staff cannot be overlooked. Thanks to you all. I am greatly indebted to my immediate family members. I want to thank my husband Dr M.A. Oyinlola for his supports, understanding, perseverance and tolerance; the children, Oyinkansola, Olaoluwa and Moyosola, I thank you for the affection offered. It is with you by my side that I am able to record success. A special thank you also goes to my mother, Omowumi Ogunmola for your love and prayers. I could not have had a better mum. I render appreciation to my siblings, Folake, Olutayo, Olanrewaju and Babatunde, for your prayers and support. I thank my parents- in-law, Alhaji Ganiyu and Bolaji Oyinlola for those words of encouragement and prayers. A special thank you goes to Prof. Labode Popoola and Dr S.M. Wakil, through whom I had the opportunity of having a great scholar, as a Supervisor. I also thank my fellow colleagues: Emmanuel Garuba, Ukponobong Antia, Muyiwa Adeyemo, Damilola Seyi-Amole and other friends too numerous to mention. Your names will remain evergreen in my mind. Kubrat Abiola, OYINLOLA July 2015. v CERTIFICATION I certify that this work was carried out by Kubrat Abiola, OYINLOLA under my supervision in the Department of Microbiology, University of Ibadan, Ibadan. Nigeria. ANTHONY ABIODUN ONILUDE B.Sc., M.Sc. (Ife) Ph.D. (Ibadan) Professor of Microbiology, University of Ibadan, Nigeria. v i TABLE OF CONTENTS Page ABSTRACT ii DEDICATION iv ACKNOWLEDGEMENT v CERTIFICATION vi TABLE OF CONTENTS vii LIST OF TABLES xiv LIST OF PLATES xvi LIST OF FIGURES xvii LIST OF ABBREVIATIONS xix CHAPTER ONE INTRODUCTION 1.1 Background 1 1.2 Statement of Problem 3 1.3 Justification 3 1.4 Scope of the research 3 1.5 Aim and Objectives 4 CHAPTER TWO LITERATURE REVIEW 2.1 Cassava 5 2.1.1 Origin of cassava 5 2.1.2 Importance and consumption 6 2.1.3 Shelf life 7 2.1.4 Cassava varieties 7 2.1.5 Nutritional composition 8 vi i Content Page 2.1.6 Anti-nutritional factors of cassava 11 2.2 Fermentation 13 2.2.1 Definition 13 2.2.2 Advantages of fermentation 13 2.2.3 Types of fermentation 15 2.2.3.1 Homolactic acid fermentation 16 2.2.3.2 Heterolactic acid fermentation 17 2.2.4 Fermented foods 17 2.3 Cassava fermentation 20 2.3.1 Submerged fermentation 20 2.3.2 Microbiology of cassava fermentation 21 2.3.3 Cassava products 22 2.3.3.1 Fufu 23 2.3.3.2 Usi 24 2.4 Starter culture 24 2.4.1 Definition 24 2.4.2 Properties of starter culture 28 2.4.3 Effect of starter on cassava fermentation 28 2.4.4 Metabolite spectrum of cassava fermentation 30 CHAPTER THREE MATERIALS AND METHODS 3.1 Sample collection 33 3.1.1 Cassava samples 33 3.1.2 Organic acids and sugar standards 33 3.1.3 Test pathogenic isolates 33 3.2 Media and sample preparation 33 3.2.1 Media preparation 33 3.2.2 Sample Preparation and fermentation 34 vi ii Content Page 3.3 Total titratable acidity of fermenting mash 34 3.4 pH of fermenting mash 34 3.5 Isolation and identification of organisms from fermenting cassava tubers 34 3.5.1 Isolation technique 3.5.2 Characterization of selected isolates 34 3.5.2.1 Morphological Characterization 35 i Macroscopic 35 ii Microscopic 35 3.5.2.2 Biochemical Characterization 35 i Catalase test 35 ii Motility test 36 iii Voges-Proskauer test 36 iv Oxidase test 36 v Indole test 37 vi Methyl red test 37 vii Starch Hydrolysis 37 viii Production of ammonium from arginine 37 ix Sugar fermentation test 37 3.5.3 Identification of organisms 38 3.6 Molecular characterization 38 3.6.1 DNA extraction 38 3.6.2 DNA amplification 38 3.6.3 Profile sequencing 39 3.7 Screening for potential starters 39 3.7.1 Starch hydrolysis 39 3.7.2 Production of linamarase 39 3.7.3 Production of pectinase 40 3.7.4 Acidification of growth medium by selected isolates 40 3.7.5 Production of Lactic acid 40 3.7.6 Production of Hydrogen peroxide 40 3.7.7 Production of Diacetyl 41 3.7.8 Antimicrobial activity against pathogenic organisms 41 ix Content Page 3.7.8.1 Preparation of Cell-Free Filtrate 41 3.7.8.2 Agar Well Diffusion Test 41 3.8 Fermentation studies with identified isolates 42 3.8.1 Preparation of cassava for controlled fermentation 42 3.8.2 Determination of inoculum size 42 3.9 Performance studies on fermenting mash 43 3.9.1 pH of fermenting mash 43 3.9.2 Total titratable acidity of fermenting mash 43 3.9.3 Nutritional Analysis of fermenting mash 43 3.9.3.1 Determination of moisture content 43 3.9.3.2 Determination of crude protein 44 3.9.3.3 Determination of crude fat 44 3.9.3.4 Determination of ash content 45 3.9.3.5 Determination of fibre content 45 3.9.3.6 Determination of total carbohydrate 45 3.9.4 Anti-nutritional factors of the fermenting cassava mash 45 3.9.4.1Determination of cyanide content 45 3.9.4.2 Estimation of phytic acid 46 3.9.4.3 Estimation of tannin content 46 3.10 Estimation of enzyme activities during fermentation 47 3.10.1 Enzyme crude extracts 47 3.10.2 Amylase activity 47 3.10.3 Linamarase assay 47 3.10.4 Pectinase assay 48 3.11 Analysis of metabolites in fermenting mash 48 3.12 Selection of starter 49 3.13 Optimization studies on selected starter 49 3.13.1 Determination of inoculum size 49 3.13.2 Determination of optimal pH 49 3.13.3 Determination of optimal incubation temperature 49 3.13.4 Determination of optimal salt concentration 50 x Content Page 3.13.5 Effect of agitation on growth 50 3.13.6 Effect of carbon sources on growth 50 3.13.7 Effect of nitrogen sources on growth 50 3.13.8 Effect of different incubation time on growth 51 3.14 Application of starter in traditional processing of fufu and usi under optimized growth conditions and performance study on the products 51 3.14.1 Inoculum preparation 51 3.14.2 Preparation of Fufu and Usi 51 3.14.3 Nutritional analysis of mash fermented with starter grown under optimised condition 52 3.14.4 Sensory evaluation of products 52 3.14.5 Microbiological assessment and physical evaluation of products during storage 53 3.15 Statistical Analysis 53 CHAPTER FOUR RESULTS 4.1 Physico-chemical analysis of spontaneously-fermenting cassava for fufu and usi 54 4.2 Microbial load during spontaneous fufu and usi fermentation 54 4.3 Isolation of Lactic Acid Bacteria from fermenting cassava mash 58 4.4 Screening for potential starters among selected Lactic Acid Bacteria 61 4.5 Molecular identification of screened potential starters 68 4.6 Fermentation of cassava with identified potential starters 71 4.6.1 pH of starter-fermented cassava mash 71 4.6.2 Total titratable acidity (%) of starter-fermented cassava mash 71 x i Content Page 4.6.3 Proximate analysis of starter-fermented cassava for fufu and usi production 77 4.6.4 Anti-nutritional factors of starter-fermented cassava for fufu and usi production 80 4.6.5 Selection of a common starter for both products 82 4.6.6 Enzyme activities during starter fermentation for fufu and usi production 85 4.6.7 Analysis of organic acid and sugars during fufu and usi fermentation 89 4.7 Optimization of growth conditions for starter CGI 95 4.7.1 Effect of different pH values on the growth of selected starter 95 4.7.2 Effect of different incubation temperature on the growth of selected starter 95 4.7.3 Effect of different salt concentrations on the growth of selected starter 98 4.7.4 Effect of different agitation speed on the growth of selected starter 98 4.7.5 Effect of different carbon sources on the growth of selected starter 98 4.7.6 Effect of different nitrogen sources on the growth of selected starter 98 4.7.7 Effect of different incubation time on the growth of selected starter 103 4.8 Effect of optimization of starter growth conditions on proximate and anti-nutritional factors (%) of fermented fufu and usi mashes 103 4.9 Sensory evaluation of starter-fermented fufu and usi 107 4.10 Microbial load during fufu and usi storage at room temperature 107 xi i CHAPTER FIVE 5.0 DISCUSSION, SUMMARY AND CONCLUSION 115 CONTRIBUTION OF THE STUDY TO KNOWLEDGE 136 REFERENCES 137 APPENDIX 185 xi ii LIST OF TABLES Table Content Page 2.1 Nutrient Composition of fresh cassava roots per 100g of edible portion 10 2.2 Different classification of fermented foods 19 8 4.1 Microbial count (10 CFU/mL) of spontaneously-fermenting cassava for fufu and usi 57 4.2 Colonial morphology of randomly selected Lactic Acid Bacteria 59 4.3 Biochemical characteristics of selected LAB. 60 4.4 Frequency of occurrence (%) of selected Lactic Acid Bacteria during spontaneous cassava fermentation. 62 4.5 Starch hydrolysis, linamarase and pectinase enzyme production by selected isolates 63 4.6 Acidification (pH) of growth medium by selected isolates 64 4.7 Quantity of lactic acid produced in g/L by selected strains of Lactic Acid Bacteria 65 4.8 Quantity of hydrogen peroxide produced in µg/L by selected strains of Lactic Acid Bacteria 66 4.9 Quantity of diacetyl produced in g/L by selected strains of Lactic Acid Bacteria 67 4.10 Antagonistic effect of selected isolates against test pathogenic organisms. 69 4.11 Molecular identification of selected potential starters 70 4.12 Single and randomly combined LAB isolates used as potential starters for fufu and usi production. 73 4.13 pH during starter fermentation of cassava for fufu and usi production 75 4.14 Total titratable acidity of mash during starter fermentation of cassava for fufu and usi production 76 4.15 Proximate composition (%) of starter fermented cassava for fufu production after 72-hour fermentation 78 xi v Table Content Page 4.16 Proximate composition (%) of starter fermented cassava for usi production after 72hours fermentation 79 4.17 Anti-nutritional factors of fresh and starter-fermented cassava for fufu and usi production 81 4.18 Selection of a common starter using pH and anti- nutritional factors 83 4.19 Selection of a common starter using proximate composition 84 4.20 Effect of optimization of starter growth conditions on nutritional contents (%) of fermented fufu and usi mashes 105 4.21 Effect of optimization of starter growth conditions on anti- nutritional contents (mg/g) of fermented fufu and usi mashes 106 4.22a Sensory evaluation of starter-fermented and spontaneously fermented usi 109 4.22b Sensory evaluation of starter-fermented and spontaneously fermented fufu 110 4.23a Microbial load during usi storage at room temperature 111 4.23b Microbial load during fufu storage at room temperature 112 x v LIST OF PLATES Plate Page 4.1 Cassava fermentation with single and combined starter cultures using plastic bioreactors 74 4.2 Laboratory prepared starter-fermented usi and fufu 108 th 4.3 Spoilage symptoms on usi and fufu on the 7 storage day 114 xv i LIST OF FIGURES Figure Content Page 2.1 Flow chart for traditional fufu and usi production 26 4.1 pH of spontaneously-fermenting cassava for fufu and usi 55 4.2 Titratable acidity (%) of spontaneously-fermenting cassava for fufu and usi. 56 4.3 Phylogenetic relationship between identified starters and closely related organisms in the GenBank. 72 4.4 Amylase activities in starter fermented and un-inoculated cassava mash during fufu and usi fermentation 86 4.5 Pectinase activities in starter fermented and un-inoculated cassava mash during fufu and usi fermentation 87 4.6 Linamarase activities in starter fermented and un-inoculated cassava mash during fufu and usi fermentation 88 4.7 Lactic acid quantities at different time intervals during spontaneous and starter (CGI) fermentation for fufu 90 4.8 Lactic acid quantities at different time intervals during spontaneous and starter (CGI) fermentation for usi. 91 4.9a Sugar quantities at different time intervals during starter (CGI) fermentation for usi. 93 4.9b Sugar quantities at different time intervals during starter (CGI) fermentation for usi. 93 4.10a Sugar quantities at different time intervals during starter (CGI) fermentation for fufu. 94 4.10b Sugar quantities at different time intervals during starter (CGI) fermentation for fufu. 94 4.11 Effect of different pH values on the growth of selected starter (CGI) 96 4.12 Effect of different incubation temperature on the growth of selected starter (CGI). 97 4.13 Effect of different NaCl concentrations on the growth of selected starter (CGI) 99 xv ii Figure Content Page 4.14 Effect of different agitation speed on growth of selected starter (CGI). 100 4.15 Effect of different carbon sources on the growth of selected starter (CGI). 101 4.16 Effect of different nitrogen sources on the growth of selected starter (CGI) 102 4.17 Effect of different incubation time on the growth of selected starter (CGI) 104 xv iii LIST OF ABBREVIATIONS % Percentage ANOVA Analysis of Variance AOAC Association of Official Analytical Chemists Bp Boiling point cal Calorie CDM Chemically defined medium CFU Colony forming unit cm Centimetre DM Dry matter DNA Deoxyribonucleic acid DNSA Dinitrosalysilic acid FAO Food and Agricultural Organization of United Nations FW Fresh weight g Gram g/L Gram per litre H2O2 Hydrogen peroxide H2SO4 Sulphuric acid HCl Hydrogen chloride HCN Hydrogen cyanide kcal Kilocalorie kg Kilogramme KH2PO4 Potassium dihydrogen phosphate KMnO4 Potassium permanganate L Litre LAB Lactic acid bacteria m Meter M Molar MEA Malt Extract Agar MEGA Molecular Evolution Genetics Analysis mg Milligramme mg/g Milligram per gram mL Millilitre xi x mm Millimetre MRS Mann Rogosa Sharpe NAD+ Nicotinamide Adenine dinucleotide NADH Nicotinamide Adenine dinucleotide hydrogenase NaOH Sodium hydroxide nm Nanometer O C Degree Celcius OD Optical density PCA Plate Count Agar pKa Acid dissociation constant PNPG Para-nitrophenyl-B-D-glucopyranoside ppm Parts per million rpm Revolution per minute rRNA Ribosomal ribonucleic acid SD Standard deviation SON Standards Organization of Nigeria Sp. Species (singular) spp. Species (plural) SSF Solid state fermentation TRIP Tuber and Root Improvement Programme USA United States of America UV Ultraviolet v/v Volume per volume VRBGA Violet Red Bile Glucose Agar w/v Weight per volume β Beta μg/mL Microgram per millilitre μL Microlitre μM Micromolar x x CHAPTER ONE INTRODUCTION 1.1 Background to the study In Africa, cassava is very important to the people because fermented cassava products are known to constitute a major part of the daily diets in many homes. It is cultivated widely as a food crop, ranked as the world′s sixth most important (Soccol, 1996) and fourth on the list of major food crops in developing countries after rice, wheat and maize (Mingli et al., 1992). De Bruijin and Fresco (1989) reported a progressive increase in demand of fermented cassava products yearly as a result of the high energy content due to the fact that it provides averagely more than 50% of daily energy intake. However, cyanogenic glucosides inherent in cassava usually restrict its use as a food crop (Koch et al., 1992; Peifan et al., 2004) even though there is an endogenous linamarase (β-glucosidase), an enzyme which can easily hydrolyse linamarin, situated in the cell wall (Mkpong et al., 1990). It was reported that the endogenous linamarase could not completely breakdown the linamarin (Ikediobi and Onyike, 1982; Mkpong et al., 1989) thus, bringing about the addition of an exogenous linamarase during fermentation, which is by far the most important and widely used means of processing cassava (Oyewole, 1992; Nweke et al., 2002) to reduce cyanogenic toxicity (Ikediobi and Onyike, 1982). Fermentation, the oldest method of food processing, started over 6000 years ago (Holzapfel, 2002) in which the traditional methods and outdated techniques of producing fermented foods were based on spontaneous fermentation due to naturally occurring microorganisms in the environment and on the raw materials. However, fermentation durations were long due to the lag phase of the organisms, thus, yielding a longer acidification process and making it difficult to produce an end product of consistent quality. Developing countries cannot continue to be dependent on the historic methods for food processing because of factors such as increasing populations, 1 drought and other natural disasters, inadequate food production as well as other associated problems such as long fermentation time, inconsistencies in final products and the presence of pathogenic organisms, all because it depended on chance inoculation from the environment. Common research approaches have included isolation and characterization of microorganisms that could be used as starter culture with modifications to fermentation regimes. To date, little of this research has been put to use. Part of technology considerations suggested by Baseline Consultancy Report for Cassava in 2010 included the use of isolated starter cultures in maintaining product quality. Therefore, an improved fermentation method that will not compromise the quality and safety of the product would be through the use of starter cultures which are preparations or materials containing large number of viable microorganisms which may be added to facilitate improved and controlled fermentation process (Holzapfel, 1997, 2002). Fufu and Usi (edible starch) are among the products of cassava fermentation in Africa (Etejere and Bhat, 1985). Fufu is an important basic commodity, ranked next to gari as a native food of most Nigerians (Sanni et al., 1998) and widely eaten in many parts of West Africa and the Tropics (Sanni, 1989). It is a sticky cassava mash which is cooked in boiling water and consumed with soup. It is eaten mostly in the Eastern and Western parts of Southern Nigeria as well as some other areas of West and Central Africa; and unlike other fermented cassava products, it has very intense odour (Lancaster et al., 1982). Usi is an indigenous food of the Itsekiri and Urhobo in Southern Nigeria, who also refer to it as edible starch. It is a very pasty, light yellow food eaten with any oil or pepper soup. The starch is precipitated out of the solution pressed out of the grated cassava during the preparation of gari and sometimes, obtained from grated cassava, soaked directly in water (Etejere and Bhat, 1985). Both cassava products undergo lactic acid fermentation by several microorganisms, thus yielding various metabolites which confer positive effects such as preservation, flavour development, cyanide reduction and changes in functional properties on the final product (Akindahunsi et al., 1999). 2 The use of starters will provide a means of standardising the production process resulting in products of uniform quality and contributes to reduction in processing time. Furthermore, such starter will have the ability to detoxify, while retaining the desirable organoleptic qualities of the product, grow rapidly to significantly shorten fermentation time, rapidly drop the pH and increase the acidity, as acidic conditions inhibit the growth of and toxin production by pathogens (Mugula et al., 2002). 1.2 Statement of Problem Constantly, there is an increase in demand for fermented cassava products because they are high energy yielding foods but indigenous spontaneous fermentation have been characterized with longer acidification time, inconsistencies in the nutritional composition and quality of the final products. Use of starter culture has brought some improvement on fermented cassava products, but most available starter cultures are used for single food fermentation and are uneconomical. 1.3 Justification The ability to isolate strains of microorganisms with desirable physiological and metabolic characteristics for use as starter culture will result in a high degree of control over the fermentation process, thus, maintaining consistency. Furthermore, the possibility of developing a common starter culture for multiple related food products will reduce cost and be of economic importance. 1.4 Scope of the research In order to establish the selection process, this study was approached in four phases, namely; isolation, characterization, identification and screening for potential starters; utilization of potential starters both singly and in combination for controlled fermentation; physiological studies and optimisation of growth conditions of selected starter(s); utilisation of the starter(s) in fufu and usi fermentation and final product assessment. 3 1.5 Aim and objectives The overall aim of this study is to select a common starter culture for the production of fufu and usi. Specifically, this study was designed to:  Isolate, characterize and identify lactic acid bacteria involved in the fermentation of cassava to produce fufu and usi.  Screen for, and genotypically identify potential starters.  Utilise selected isolates solely and in combination to ferment cassava in fufu and usi production, monitoring microbiological, nutritional and technological properties as well as derived metabolites during the fermentation processes, thus, select isolate(s) of best fit.  Carry out optimization of growth studies on the potential starter(s) selected  Apply the selected starter(s) in the production of the two products and analyse the end products. 4 CHAPTER TWO LITERATURE REVIEW 2.1 Cassava Cassava (Manihot esculenta Crantz), a 1-2 m high woody shrub with an edible root is a perennial crop. It is made up of an ariel part (2-4 m) and an underground part (edible root) clustered around the lowermost part of the plant and extends about 60 cm on all sides (Pandey et al., 2000). A single root may weigh as much as four kilogram under convinient conditions. The number of roots per plant at harvest varies from 2 to 7, each averaging 27.7 to 43.3 cm long and from 4.5 to 7.4 cm in diameter. A central vascular core, the cortex (flesh) and the phelloderm (peels) makes up a mature root. Cassava peels account for 10–12% of the total dry matter of the root and are 1–4 mm thick (Nartey, 1979). Cassava, one of the most useful tropical crops widely exploited as a cheap energy source in Africa, Asia, South America and India (de Bruijin and Fresco, 1989) was however castigated as an inferior food crop (Kwatia, 1986), poor people′s crop (Hahn and Keyser, 1985) and a dangerous crop (Cheok, 1978) because of major limitations like low protein content, short postharvest shelf life and the presence of toxic cyanogenic glucosides. 2.1.1 Origin of cassava In tropical Africa cassava has assumed the status of a security and industrial crop, alleviating the food crisis in many war-torn and drought ravaged parts of Africa because it can be cultivated throughout the year without regard to the seasons. It produces high yield and grows with limited water (Hahn and Keysar, 1985; Oyewole, 2002). Although, it was reported to have originated in Venezuela, South America during 2700B.C. (Soccol, 1996), Philips (1982) established its being introduced to Nigeria and other parts of West Africa by the Portuguese. It was estimated by 5 FAO that 37% of the 13.4 million ton world production was produced by African countries. The main cassava producing countries of Africa include Nigeria, Benin, Kenya, Zambia, Tanzania, Uganda, Ghana, Zimbabwe, Democratic Republic of Congo and Mozambique (Okigbo, 1980; Nweke et al., 2002). In 1996, Soccol reported Africa as the largest producer with about 53% of the world′s production. Although, it was cultivated in about 88 countries, only 5 countries accounted for 67% of the production and these include Nigeria, Brazil, Thailand, Zaire and Indonesia. Purseglove (1968) reported that cassava was known around the North of the River Niger in 1914 but it has now become one of the most important staple food crops of the inhabitants in Nigeria and known by different names among the ethnic groups of the country. The Yoruba call it gbaguda or ege, the Hausa, rogo, karaza or doyar kudu, the Ibo, akpu, abacha or jigbo, the Benin, igari, the Efik, iwa unene while the Urhobo refer to it as imidaka (Etejere and Bhat, 1985) 2.1.2 Importance and Consumption of cassava Cassava plays a major role in efforts to alleviate the African food crisis because of its efficient production of food energy, year-round availability, tolerance to extreme stress conditions, and suitability to present farming and food systems in Africa (Hahn and Keyser 1985, Hahn et al., 1987). Much recognition has been dwelled on the importance of cassava and among reported ones are its use as source of income and raw material in the manufacturing of processed food, animal feed as well as industrial products (Beléia et al., 2004). Wider utilization of cassava products can be a catalyst for rural industrial development and raise the income for producers, processors and traders, contribute to the food security status of its producing and consuming households as well as its high efficiency in converting solar energy to starch (Dufour et al., 1996). Total world cassava use was projected to increase from 172.7 million ton to 275 million ton over a period of 27 years (1993-2020), using the International Food Policy Research Institute baseline data whereas, a higher prediction of demand and production growth puts the 2020 production at 291 million tons (Scott et al., 2000). In both projections, cassava use in Africa is equivalent to 62% of total world production with an average of about 102 kg/person/year or 220 kcal/person/day (Giraud, 1993) and Nigeria is the current leading producing country (FAO, 2008). Almost all the 6 cassava produced globally is used for human consumption, either in natural form as flour or in fermented forms and other products with only 5% being used industrially (Bokanga and Otoo, 1991; FAO, 2002; Ajao & Adegun, 2009). There is an increase in demand yearly as it was reported by Seigler and Pereira in 1981 to have been consumed by 300 million people. However, Cock (1982, 1985), FAO (2000) and Mroso (2003) reported it to be consumed by more than 500 million people worldwide, including 80 million from Nigeria alone (Okafor and Ejiofor, 1990). 2.1.3 Shelf life of cassava Other tuber crops such as yam and sweet potato are not as perishable as cassava roots (Poulter, 1995). Physiological deterioration (primary or secondary) occurs 2-3 days after harvesting, followed by microbial deterioration 3-5 days thereafter (Rickard and Coursey, 1981; Rickard, 1985; Akingbala et al., 2005). The primary deterioration is an endogenous physiological process called vascular streaking brought about by damage to the roots during postharvest handling which results in a fine blue-black or brown discoloration. This usually occurs when phenolic compounds present in the tubers are converted to coloured compounds called quinines and the process is catalyzed by the enzyme, polyphenol oxidase which acts on the phenolic compounds in the presence of oxygen (Rickard and Coursey, 1981; Rickard, 1985; Sakai et al., 1986). Furthermore, dehydration caused by the physical damage to the tubers worsens the conditions thus, making the tissues become portals of entry for pathogens leading to the secondary deterioration by microorganisms. The microbial spoilage involves rotting, softening or fermentation of the tissue by microorganisms (Rickard and Coursey, 1981; Uritani et al., 1984). 2.1.4 Cassava varieties There are many natural cassava varieties (cultivars) and are classified according to morphological traits as well as taste, cyanide content, average yield, disease performance and pubescence (MIC, 2007; Gbadegesin et al., 2013). However, in recent times, a number of regional programs have been initiated to breed improved varieties of cassava to increase yield and resistance to diseases. Various studies have shown that the physicochemical, functional and other quality characteristics of cassava 7 products are significantly affected by varietal chemical composition differences such as dry matter, starch content and quality (Safo-Kantanka and Owusu-Nipah, 1992). Even though more than 5,000 cassava cultivars are recognized globally (Best, 1993; Gade, 2003; IFAD/FAO, 2005), several improved cassava varieties have been recommended and released in Nigeria (IITA, 2004). Among the most commonly grown are TME 30572, 4(2)1425, 92/0326 and NR8082. The major genetic factor that determines quality of roots is dry matter content and more recently, 42 new improved genotypes have been made available to farmers (IITA, 2004) with qualities like multiple resistance/tolerance to cassava mosaic disease and other major problems of cassava, bacterial blight disease, anthracnose, green mite, and mealybug. They are high yielding varieties, suitable for use as food and livestock feed as well as a raw material in industry. 2.1.5 Nutritional composition of cassava Cassava roots, though deficient in protein (less than 1.5% of fresh weight), are rich in carbohydrates (31% of fresh weight) with most of it being present as starch hence, its utilization by most people in tropics as sources of carbohydrate (Blagbrough et al., 2010). The edible starchy flesh comprises some 80- 90% total weight of the root as water, forming the major components (Wheatley et al., 1993; Harris and Koomson, 2011) and it is between the range of 60.3 - 87.1% (Padonou et al., 2005; Zvinavashe et al., 2011). Water is an important parameter in the storage of cassava with low levels giving desirable and relatively longer shelf life (Padonou et al., 2010; Harris and Koomson, 2011). Cassava contains about 1-2% protein which makes it a predominantly starchy food (Charles et al., 2005). The protein content is low (1% - 3%) on a dry matter basis (Buitrago, 1990) and between 0.4 and 1.5% per 100 g fresh weight (Bradbury and Holloway, 1988). However, about 50% of the crude protein in the roots consists of whole protein and the other 50% in the form of free amino acids (primarily glutamic and aspartic acids) and non-protein components such as nitrite, nitrate and cyanogenic compounds (Zvinavashe et al., 2011). The root also has high content of dietary fibre, magnesium, sodium, riboflavin, nicotinic acid, thiamine and citrate (Bradbury and Holloway, 1988) but was reported to be low in iron and vitamin A (McDowell and Onduro, 1983). 8 Even though, cassava tuber has been criticized for its low and poor protein content, it produces more weight of carbohydrate per unit area than other staple food crops under comparable agro-climatic conditions thus, being an energy-dense food and therefore 3 3 ranked high for its energy value of 250 x 10 cal/ha/day as compared to 176 x 10 for 3 3 3 rice, 110 x 10 for wheat, 200 x 10 for maize, and 114 x 10 for sorghum (Okigbo, 1980; Jisha et al., 2010). Montagnac (2009) and Zvinavashe et al. (2011) reported that the root is a physiological energy reserve with high carbohydrate content, which ranged from 32% to 35% on a fresh weight (FW) basis, and from 80% to 90% on a dry matter (DM) basis. Raw cassava root however, has more carbohydrate than potatoes and less carbohydrate than wheat, rice, yellow corn, and sorghum on a 100 g basis (Montagnac, 2009). The lipid content in cassava roots ranges from 0.1% to 0.3% on a fresh weight basis with values ranging from 0.1% to 0.4% (Charles et al., 2005) and 0.65% (Padonou et al., 2005) on a dry weight basis. This content is relatively low compared to maize and sorghum, but higher than potato and comparable to rice. The lipids are either non-polar (45%) or glycolipids (52%) according to Hudson and Ogunsua (1974), mainly galactose-diglyceride (Gil and Buitrago, 2002). Predominant fatty acids include palmitate and oleate (Hudson and Ogunsua, 1974). 9 Table 2.1: Nutrient Composition of fresh cassava root per 100g of edible portion Nutrients Unit Cassava root Food energy Kjoules 0.61 Water G 62.5 Carbohydrate G 34.7 Protein G 1.20 Fat G 0.3 Calcium Mg 33 Iron Mg 0.7 Vitamin A I.U Trace Thiamine Mg 0.06 Riboflavin Mg 0.03 Niacin Mg 0.06 Vitamin C Mg 36 Source: FAO Food Composition (Nweke et al., 2002) 1 0 2.1.6 Anti-nutritional factors of cassava Cassava has significant deficiencies that restrict its usefulness as a food source, one of which is the precence of cyanogenic glucosides; linamarin (2-β- Dglucopyranosyloxy-2-methylpropanenitrile) and lotaustralin [(2R)-2- β - Dglucopyranosyloxy- 2-methylbutyronitrile] derived from valine and isoleucine, respectively (Peifan et al., 2004; Koch et al., 1992). The ratio of linamarin to lotuastralin in leaves and roots was reported to be about 93:7 by Nartey (1978) and Liangcheng et al. (1995) while less than 83% linamarin was recounted by other researchers (Cereda and Mattos, 1996; Kimaryo et al., 2000). Linamarin is a β-glucoside of acetone cyanohydrin and ethyl-methylketone- cyanohydrins which is stored in the vacuoles of the cassava cells (McMahon et al., 1995) and whose β-linkage can only be broken down under high pressure, temperature and use of mineral acids. Linamarin is bitter thereby making high-cyanide cassava (>100ppm) bitter, thus called bitter cassava. Cyanogens are present in variable concentrations ranging from 300 to 500 ppm (El Tinay et al., 1984) depending on the type of cassava. Bitter varieties, which contain higher amounts of cyanogenic glucosides therefore needs processing to remove the toxic compounds before consumption, whereas, sweet varieties (˂100 ppm) have low levels and can be eaten fresh (Rosling, 1990). Despite this, populations which use cassava as main staple food, mainly grow the bitter varieties due to their higher yields (Mozambique Ministry of Health, 1984), resistance to insects, pests and therefore rely on processing methods for detoxification. Cardoso et al. (2005) puts the range of total cyanide content to be from 1 to 1550 ppm. Although hydrocyanic concentrations of 15 to 400 mg/kg of fresh weight in cassava varieties are reported, more frequent values fall within the interval of 15 – 150 mg/kg (Cereda and Mattos, 1996) even though the minimum tolerant level recommended by Standard Organisation of Nigeria is 50 mg/kg (SON, 1985). There are, however, cassava varieties which contain concentrations above 1000 mg/kg (Cereda and Mattos, 1996). These variations were reported to be due to different rate of biosynthesis, degradation or transport (Elias et al., 1997). Environmental factors, the cultivar and growth conditions have also been documented (Cooke, 1978; Bradbury et al., 1991). Cassava plant has endogenous linamarase (β-glucosidase), an enzyme which can easily hydrolyse linamarin, situated in the cell wall (Mkpong et al., 1990). The 1 1 grating or mincing of the roots permits, through the cell structure damage, the release of endogenous linamarase able to hydrolyse linamarin into glucose and acetone O cyanohydrin (Conn, 1969), under optimum conditions at 25 C and pH range 5.5 – 6.0 (Cereda and Mattos, 1996). As reported by Mkpong et al. (1989) as well as Ikediobi and Onyike (1982), the endogenous linamarase content could not permit the complete breakdown of linamarin. However, it was demonstrated (Ikediobi and Onyike 1982) that it is possible to lower the cyanogen toxicity by the introduction of an exogenous linamarase during fermentation. Many authors (Ikediobi and Onyike, 1982; Padmaja and Balagopal, 1985; Okafor and Ejiofor, 1990) have suggested the inoculation of fermenting cassava with a linamarase-producing microorganism. This reduces the cyanogen content because the microorganisms produce linamarase which break down the linamarin (Guyot et al., 1998). Fermentation of cassava with water is the simplest method to reduce the cyanide content (Cumbana et al., 2007; Bradbury and Denton, 2010) as the water will facilitate swelling of the cells and allow linamarase to come into contact with linamarin leading to hydrolysis (Bradbury, 2006). Uyoh et al. (2009) also observed that using unchanged water during fermentation will reduce cyanide content significantly. Reduction in the cyanide content of fermenting cassava as reported by Westby and Choo (1994) ranged from 65 to 110 mgHCN/kg for a period of 12 to 96 hours while Onyesom et al. (2008) puts the range between 7.02 and 2.70 mgHCN/100g cassava wet weight for a period of 24-96 hours. Although, much of these toxic components were removed during processing of cassava, a quantity still remains, depending on the process used according to Nartey (1981) as well as Nambisan and Sundaresan (1985). Consumption of cassava which still contains residual levels of cyanogenic compound can result in chronic diseases such as goitre, cretinism, topical ataxic neuropathy, iodine deficiency, destructions of cells and tropical diabetes (Cock 1982; Tylleskar et al., 1992). Since cassava is completely devoid of protein and highly deficient in vitamins and minerals, it causes malnutrition as well, when consumed solely. 1 2 2.2 Fermentation 2.2.1 Definition Adams (1990) describes fermentation as a form of energy-yielding microbial metabolism in which an organic substrate, usually a carbohydrate, is partially oxidised, and an organic carbohydrate acts as the electron acceptor. But according to Campbell- Platt (1987), Fermentation is the subjecting of food to the action of microorganisms or enzymes so that desirable biochemical changes cause significant modification to the food. It was also defined as the process by which alcohol or lactic acid was produced by living cells in solutions that contain sugars (Ribéreau-Gayon et al., 2000). William and Dennis (2011) as well as Wikipedia (2012) described fermentation as the conversion of carbohydrates to alcohol and carbondioxide or organic acids using yeast and/or bacteria, under anaerobic conditions. Fermentation of food has however been termed as one of the oldest method of food preparation and preservation (Pederson 1971; Steinkraus et al., 1983; Campbell-Platt, 1994) used as far back as 6000 BC in the Middle East, although then, it was without any knowledge of the roles of the microorganisms involved (Caplice and Fitzgerald, 1999). 2.2.2 Advantages of fermentation Fermentation processes are used for production of a vast number of valuable products using various fermentation media (substrates) and microorganisms (Mshandete, 2011). It is widely used to transform and preserve food because of its low technology, energy requirements and the unequaled organoleptic qualities it confer on the final product (Daeschel et al., 1987). Some of the importance of fermentation includes flavour enhancement, improved nutritional quality, preservation, detoxification, inhibitory metabolite production, food edibility and income generation. Fermentation tend to make food more palatable by enhancing its aroma and flavour thereby, making the organoleptic properties of fermented food more popular than the unfermented one in terms of consumer acceptance (Blandino et al., 2003; Osungbaro, 2009). The microorganisms and metabolites responsible for these changes have been described (Ramaite and Cloete, 2006). However, the specific mechanisms by which flavour is generated are still subject to investigation. Fermentation is unique in that it modifies the unfermented food in diverse ways, resulting in new sensory properties in the fermented product (Leroy and De Vuyst, 2004). 1 3 A number of foods especially cereals are poor in nutritional value, and they constitute the main staple diet of the low income populations. Fermentation has however, been shown to improve the nutritional value and digestibility of these foods (Obiri-Danso et al., 1997; Nout, 2009). It was reported that the activities of microbial enzymes which include amylases, proteases, phytases and lipases were enhanced by the acidic nature of the fermentation products at a temperature range of 22-25ºC (Mokoena et al., 2005), thus, modifying the primary food products through hydrolysis of polysaccharides, proteins, phytates and lipids respectively. Anti-nutrients such as phytic acid and tannins in foods were also reduced through lactic fermentation leading to increased bioavailability of minerals such as iron, protein and simple sugars (Sripriya et al., 1997; Chelule et al., 2010) By lowering pH below 4.0 through acid production, lactic fermentation hinders the growth of pathogenic microorganisms which cause food spoilage, food poisoning and diseases (Ananou et al., 2007). The production of lactic and acetic acids resulting in pH decrease and increase in titratable acidity as reported by Abdel and Dardir (2009) and Olukoya et al. (2011) accounted for the overgrowth of desirable bacteria in food than other non-desirable food spoilage bacteria, thus, prolonging the shelf life of fermented food. Detoxification of toxins (mycotoxin) in food through fermentation has also been documented over the years (Mokoena et al., 2005, 2006; Schnurer and Magnusson, 2005; Chelule et al., 2010; Dalie et al., 2010). Fermentation has been reported to be more advantageous than using alkaline ammonia treatment because of its mild nature which preserves the nutritive value and flavour of decontaminated food (Bata and Lasztity, 1999). Furthermore, the mycotoxins are irreversibly degraded without leaving any toxic residues and this was believed to be through the toxin-binding effect or an enzymatic interaction (Zinedine et al., 2005). Endogenous linamarase production or the use of a linamarase-producing microorganism as starter culture during fermentation has also been reported to detoxify cassava (Ravi and Padjama, 1997; Sweeney and Dobson, 1998). Some of the inhibitory compounds produced against unwanted bacteria include hydrogen peroxide, carbon dioxide, diacetyl, broad spectrum antimicrobials such as reuterin, organic acids and bacteriocins (De Vuyst and Vandamme, 1994; Oyewole, 1997), which act as deterrents for pathogenic enteric bacteria and non-acid tolerant bacteria (Olukoya, 2011). This antagonistic effect by the organic acids was believed to have resulted from their action on the bacterial cytoplasmic membrane which 1 4 interferes with the maintenance of membrane potentials and inhibits active transport (Sheu et al., 1972; Eklund, 1989; De Vuyst and Vandamme, 1994a). Hydrogen peroxide inhibitory mechanism was said to be mediated through the strong oxidizing effect on membrane lipids and cell proteins (Lindgren and Dobrogosz, 1990). Carbon dioxide can directly confer an anaerobic environment which is toxic to aerobic food microorganisms through its action on cell membrane and its ability to lower internal and external pH (De Vuyst and Vandamme, 1994a). (De Vuyst and Vandamme, (1994a) and Motlagh et al. (1991) ascertained that gram negative bacteria, yeasts and moulds are more sensitive to diacetyl and its mode of action was believed to be due to its interference with utilization of arginine. Bacteriocins, regarded as extracellularly released primary or modified products of bacterial ribosomal synthesis by Jack et al. (1995) have a relatively narrow spectrum of bactericidal activity. African locust bean, oil bean and cassava, for example are inedible in their unfermented state but according to Odunfa (1983), during fermentation, they become edible as a result of hydrolysis of the indigestible components which are removed by the action of microorganisms. Fermented food production has also been found to provide a source of income to a lot of people around the world. Anon (1995) reported in the FAO of nation that there was value added through processing and marketing of raw products. About 60% of workforces in sub-Sahara Africa are employed in small scale food processing sector and between ⅓ to ⅔ in manufacturing of agricultural raw materials (Anon, 1989; Conroy et al., 1995). It was also observed that rural-urban migration and the associated social problems were reduced by the generation of employment opportunities in the rural areas and small scale food industries (Aworh, 2008). 2.2.3 Types of fermentation Soni and Sandhu (1990) described four main fermentation processes namely, alcoholic, lactic acid, acetic acid and alkali fermentations. Alcoholic fermentation brings about the production of ethanol, and yeasts are the predominant organisms. Products of such fermentation include wine and beer. Lactic acid fermentation produces foods such as fermented milks and cereals, mainly carried out by Lactic Acid Bacteria. A second group of bacteria of importance in food fermentations are the acetic acid producers from the Acetobacter species which converts alcohol to acetic acid in the presence of excess oxygen. Alkali fermentation often takes place during the 1 5 fermentation of fish and seeds, popularly known as condiment (McKay and Baldwin, 1990). Chisti (1999) however, classified most commercially useful fermentations as solid state or submerged, each possessing particular advantages over the other and have been reviewed (Pandey et al., 2000; Pérez-Guerra et al., 2003; Cauto and Sanromán, 2006). Although different, the processes are influenced by numerous factors, including temperature, pH, nature and composition of the medium, dissolved oxygen, dissolved CO2, operational system (batch, fed batch, continuous), mixing and most especially, fermentation microorganisms. Variations in these factors could affect the rate of fermentation, product spectrum and yield, organoleptic properties of the products (taste, texture, smell and appearance), nutritional qualities and other physico chemical properties (Chisti, 1999). During lactic fermentation, all lactic acid bacteria produce lactic acid from hexoses and since they lack functional heme-linked electron transport chains and a functional Kreb‟s cycle, they obtain energy via substrate level phosphorylation. The pathways by which hexoses are metabolised thereby, divide Lactic Acid Bacteria into two groups: homofermentative and heterofermentative (Kockova et al., 2011) 2.2.3.1 Homofermentative lactic fermentation Homofermentative bacteria transform nearly all the sugars they utilise, especially glucose into lactic acid thus, the major or sole end product. Homofermenters use the Embden-Meyerhof-Parnas pathway to generate two moles of lactate per mole of glucose and produce approximately, twice the energy per mole of glucose as heterofermenters (Kockova et al., 2011). The pathway includes a first phase involving the conversion of hexose to pyruvate (glycolysis). The terminal electron acceptor is pyruvate which is further reduced to lactic acid (Khalid, 2011). Lactate dehydrogenase catalyzes reduction of the keto group in pyruvate to a hydroxyl, converting both molecules of pyruvate to lactate as NADH is oxidized to NAD+ (Wikipedia, 2012). Examples of homolactic acid fermentation involve the production of gari, fufu, lafun etc. C6H12O6 → 2 CH3CHOHCOOH Glucose Lactate 1 6 2.2.3.2 Heterofermentative lactic fermentation Heterofermentative bacteria use the pentose phosphate pathway in which NADH is converted to NAD+ in the reaction catalyzed by pyruvate and alcohol dehydrogenase. One molecule of pyruvate is converted to lactate while the other is converted to ethanol and carbon dioxide or acetate (acetic acid) through the enzyme acetate kinase (Wikipedia, 2012). The final products of this pathway are equimolar amount of lactic acid and ethanol in a slightly aerated environment as well as lactate and acetate in an aerated environment (Khalid, 2011). An example of heterolactic acid fermentation is production of burukutu. C6H12O6 → CH3CHOHCOOH + C2H5OH + CO2 Glucose Lactic acid Ethylacohol Carbodioxide 2.2.4 Fermented foods A great majority of the foods we eat are fermented; from bread and cheese to yogurt, beer, wine, coffee, vinegar, pickled vegetables, sausages, to mention only a few. Harlender (1992) defined fermented foods as those products which have been subjected to the effect of microorganisms or enzymes to cause desirable biochemical changes. Holzapfel (2000) described them as palatable and wholesome foods prepared from raw or heated raw materials. However, Eisenbrand (2005), in the DFG Commission on Food Safety referred to fermented foods as consumable products, generated from either thermally treated or untreated raw materials of plant/animal origin which have characteristic sensory and nutritional value as well as properties determining shelf life and hygiene, conferred on them by microorganisms and/or enzymes from the raw materials. The history of fermented foods has early records in Southeast Asia, where China was regarded as the cradle of mold-fermented foods, and in Africa where the Egyptians developed the concept of the combined brewery bakery. The early Egyptian beers were probably quite similar to some of the traditional opaque sorghum, maize, or millet beers found in various African countries today (Hesseltine, 1981). When consumed by humans, it was reported that fermented foods often introduce microflora that inhabit the human body (Ross et al., 2002; Reid et al., 2003; Picard et al., 2005). This probiotic effect and the reduced level of pathogenic bacteria observed in them are especially important when it comes to developing countries 1 7 where fermented foods have been reported to reduce the severity, duration and morbidity of diarrhoea (Mensah, 1997; Kimmons et al., 1999). Nutritionally, they also have reduced antinutritional composition (Paredes-López and Harry, 1988), increased concentrations of vitamins, minerals and protein when measured on a dry weight basis (Adams, 1990). They are generally safe and wholesome because the dominant microorganisms involved in their fermentations do not appear to be associated with any health risks. These beneficial microorganisms serve to some extent in safeguarding against pathogens and spoilage organisms. Acidification to pH values of less than 4.2 constituted a major safety concern because a number of metabolites, such as acetic acid, hydrogen peroxide and bacteriocins, produced during the fermentation process, exhibit antimicrobial properties which may contribute to the safety of lactic fermented foods in particular (Holzapfel, 2002). Fermented foods have been classified in many different ways according to the views of different authors. It could be according to the kind of microorganism involved (Yokotsuka, 1982) or based on commodity (Campbell-Platt, 1987; Odunfa, 1988; Kuboye, 1985). Nigeria has a variety of people and culture, thereby, difficulty in choosing one national dish. Abdel and Dardir (2009) and Adebayo et al. (2010) stated that each area has its own regional favourite food and such is dependent on custom, tradition and religion. The fermentation processes for such native food constitute a vital body of indigenous knowledge used for food preservation, acquired by observations or experience, and passed on from generation to generation (Aworh, 2008; Chelule et al., 2010). 1 8 Table 2.2: Different classification of fermented foods Yokotsuka (1982) Campell-Platt Odunfa Sudanese (Dirar (1987) (1988) 1993) 1. Alcoholic 1. Beverages 1. Starchy roots 1. Kisser – staples beverages (yeast) 2. Cereal products 2. Cereals 2. Millet – sauces 2. Vinegars 3. Dairy products 3.Alcoholic Beverages and relishes for the (Acetobacter) 4. Fish products 4. Vegetable protein staples. 3. Milk products 5. Fruit and 5 Animal protein 3. Marayiss – beers (Lactobacilli) Vegetable products and other alcoholic 4. Pickles 6. Legumes drinks (Lactobacilli) 7. Meat products 4. Akil-munasabat – 5. Fish or meat 8. Starch crop food for special (Enzymes and products occassions Lactobacilli) 9. Miscellenous 6. Plant protein products. (moulds with or without Lactobacilli and yeasts) Source: (Dirar, 1993) 1 9 2.3 Cassava fermentation Fermentation of cassava was reported to be the most important and widely used means of processing cassava (Oyewole, 1992; Nweke et al., 2002). As of date, the ancient traditional processing of cassava is still being used and this practice is however, afflicted with so many problems because it depended on chance inoculation from the environment (Oyewole, 1990, 1995). Thus, the fermentation period is rather slower, with inconsistent quality of the products from one processor to the other as well as from one production batch to the other even by the same processor, and from one season to the other (Oyewole and Sanni, 1995). Improvements in cassava processing, which have been employed over the years helped to reduce the duration of processing to economically viable limits, maximise the detoxification process and improve the physical and nutritional qualities of the products. While the methods of fermentation vary from one locality to another (grated root fermentation, underwater/soaking fermentation, and mold fermentation), cassava fermentation in sub-Saharan Africa has been categorized either as solid state or submerged (Oyewole, 1992). 2.3.1 Submerged Fermentation Submerged fermentation is the cultivation of microorganisms in liquid nutrient medium (Renge et al., 2012) in which the bioactive compounds are secreted into the fermentation medium (Subramaniyam and Vimala, 2012). Submerged fermentation of cassava involves the soaking of cassava roots under water for 3 -5 days, causing the root to soften and swell, thus having a combined effect of enabling linamarase and linamarin to mix as well as leaching of cyanogens (Westby and Choo, 1994). The softened roots can be easily broken into pieces by hand, passed through a sieve to remove the fibre, leaving a smooth paste. Initially, a mixed microflora was reported to be involved, but it was later dominated by lactic acid bacteria (Oyewole and Odunfa, 1988; Achi and Akubor, 2000; Obilie et al., 2004; Kostinek et al, 2005). The size to which the roots were cut prior to soaking was also found to affect the rate of fermentation and the quality of product (Oyewole and Odunfa, 1992). Different groups of lactic acid bacteria isolated from submerged fermenting cassava includes Lactobacillus cellobiosus, E. bulgaricus, L. brevis, L. coprophilus, L. plantarum and Leuconostoc mesenteroides with Lactobacillus plantarum being predominant during the last 36 hours. 2 0 2.3.2 Microbiology of cassava fermentation Starch is a complex carbohydrate that can be degraded either by microorganisms that produce α-amylase or an inducible/constitutive amylase to produce simple sugars which can then be readily metabolised by many microorganisms. The indigenous natural fermentation has been reported to involve mixed colony of microorganisms such as molds, bacteria and yeasts (Anthony and Chandra, 1997; Kobawila et al., 2005; Ekundayo and Okoroafor, 2012). Even though, microbial size in food is usually small, their influence on the nature of the food, especially in terms of flavour, and other organoleptic properties, is profound (Okafor, 2009). Thus, fermentation products are based on the microorganisms involved in the fermentation and the type of bacterial flora developed in each fermented food varies based on water activity, pH, salt concentration, temperature and substrate composition (Blandino et al., 2003). These microorganisms are harmless to the consumer and produces enzymes such as proteases, amylases and lipases that hydrolyze food complexes into simple non-toxic products with desirable texture and aroma that makes them palatable for consumption (Steinkraus, 1997). Numerous authors have linked a wide spectrum of microorganisms to cassava fermentation and these includes Bacillus, Leuconostoc, Klebsiella, Corynebacterium, Lactobacillus, Aspergillus, Candida, Geotrichum, Streptococcus, Enterococcus, Aerococcus and Pediococcus species (Oyewole and Odunfa, 1988; Anthony and Chandra, 1997; Hirayama and Rafter, 1999; Holzapfel, 2002; Blandino et al., 2003). Furthermore, yeasts and molds such as Saccharomyces, Candida, Kluyeromyces, Aspergillus, Rhizopus, Mucor, Penicillium and Debaryomyces were also reported (Wouters et al., 2002; Omemu et al., 2007). However, rapid growing Lactic Acid Bacteria (LAB) are the most common prominent microorganisms for fermentation and preservation of foods. Their importance was known to be associated mainly with their safe metabolic activity while growing in foods, utilising available sugar for the production of organic acids and other metabolites. Their common occurrence in foods, coupled with their long-lived use contributed to their natural acceptance as GRAS (Generally Recognised as Safe) for human consumption (Aguirre & Collins, 1993). Lactobacillus plantarum has been shown to be the predominant LAB specie in sour cassava starch (Ngaba and Lee, 1979; Amoa-Awua et al., 1996; Ben Omar et al., 2000; Lacerda et al., 2005; Kostinek et al., 2007), even though, most species were found not to produce α-amylase and this 2 1 was quite surprising because cassava has about 84% of the carbohydrates in the form of starch (Ketiku and Oyenuga, 1972). Sanni et al. (2002) stated that only a few amylolytic LABs have been isolated from starchy fermented foods in Africa but more studies over time, had led to the discovery of more (Diaz-Ruiz et al., 2003; Putri et al., 2011a, 2011b; Mukisa et al., 2012) . In other cases, there are many kinds of fermented foods in which the dominating processes and end products are dependent on a mixture of endogenous enzymes and mixed microbial cultures which were earlier reported to have originated from the native microflora of the raw materials utilised in most of the traditional food fermentation processes according to Anthony and Chandra (1997). Some of the inhibitory compounds formed during fermentation include organic acids (e.g. palmitic, pyruvic, lactic, acetic, propionic and butyric acids), alcohols (mainly ethanol) aldehydes and ketones (acetaldehyde, acetoin, 2-methyl butanol) (Campbell- Platt, 1994). These varieties of metabolites are antagonistic in action to competing bacteria (Breidt and Fleming, 1997). The inhibition has been attributed to the protonated form of the acids, which are uncharged and may therefore cross biological membranes, thus inhibiting growth due to lowered pH of the cytoplasm and/or accumulation of anions inside the cell (Adams, 1990; Russel, 1992; Breidt & Fleming, 1997). In other words, they interfere with the maintenance of cell membrane potential, inhibiting active transport and a variety of metabolic functions as well as reducing intracellular pH (Ross et al., 2002). 2.3.3 Cassava products As earlier stated, cassava roots are bulky with about 70% moisture content, and therefore transportation of the tubers to urban markets is difficult and expensive. Moreso, the raw roots and uncooked leaves are not palatable and they contain varying amounts of cyanide which is toxic to humans and animals. Therefore, cassava must be processed into various forms in order to increase the shelf life, facilitate transportation and marketing, reduce cyanide content and improve palatability. Traditionally, cassava roots are processed by various methods into numerous products and utilised in various ways according to local customs and preferences. In some countries, the leaves are consumed as vegetables, and many traditional foods are processed from cassava roots and leaves. The nutritional status of cassava can also be improved through fortification with other protein-rich crops. Processing reduces food losses and stabilizes seasonal fluctuations in the supply of the crop. Various traditional processing methods are 2 2 known which include boiling, smoking, drying and fermentation while some of the products of these processes in Nigeria include gari, abacha, lafun, usi, fufu, tapioca cakes etc. 2.3.3.1 Fufu Fufu is traditionally produced and marketed as a fermented wet, pasty food product, which is also made into porridge in boiling water before consumption. Mostly consumed in the Eastern and Western parts of Nigeria as well as some other parts of West and Central Africa, it is known as chikwuangue or chikwange in Zaire; fufu or foo-foo in Southern Nigeria and akpu in some parts of Eastern Nigeria (Okafor et al., 1984; Longe, 1990). Traditional fufu fermentation involves peeling and washing cassava roots that are manually cut into different sizes by using a hand knife and soaking in earthen pots or drums of water for 3 to 5 days to undergo lactic acid fermentation. Reports indicated that during soaking, the pH value decreases, the root softens and this facilitates the reduction in potentially toxic cyanogenic compounds (Oyewole and Odunfa, 1992; Westby and Choo, 1994; Oyewole et al., 2001; Aworh, 2008; Uyoh, et al., 2009). The soft roots were broken with clean hands and the fibres removed by sieving which is done by adding water to the retted mass on a sieve. The starch suspension is allowed to sediment for about 24 hours after which the water is decanted. Fine, clean starch is further dewatered by putting in raffia or cotton bags and pressed with heavy stones overnight (Oyewole and Odunfa, 1989). To prepare for consumption, a quantity of the slurry containing about 25% of fufu paste in water was boiled in an open pan. After continuous stirring using a wooden rod, strong dough was formed (Kwatia, 1986; Ayankunbi et al., 1991; Anon, 1994). This method is commonly reported among the Yoruba tribe. The Igbo and Efik tribes of Eastern Nigeria who refer to it as nni akpu and udep utim respectively, rolls the starch into large balls, wrap it with wilted plantain leaves and partially steam cook. The balls are removed and pounded in a wooden mortar to give a fine, smooth and soft mash. It is further rolled into small balls, wrapped in leaves and thoroughly steamed, after which the balls are finally pound together (Etejere and Bhat, 1985). The cooked fufu is usually eaten warm with fish, meat, vegetable stew or soup. However, unlike other fermented cassava products, it has a very strong odour (Lancaster et al., 1982; Okafor et al., 1998) and considered by consumers to be of good quality when it has a smooth 2 3 texture, characteristic aroma and creamy white, grey or yellow colour (Akingbala et al., 1991; Oyewole and Odunfa, 1992; Blanshard, 1994). Variability in the quality of fufu could be as a result of chance inoculation involving varieties of microorganisms (Oyewole and Sanni, 1995), little or no control over the process (Oyewole, 1997), roots cut size (Okafor et al., 1984), difference in dry matter content (Hahn, 1989), root quality and fermentation water. 2.3.3.2 Usi Usi is one of the major native foods of the Itsekiri and Urhobo in Southern Nigeria, who also refer to it as edible starch (Etejere and Bhat, 1985). At household level, different techniques are used to obtain the fermented starch. It may be precipitated from the solution pressed out of the grated cassava roots or from grated cassava that is soaked directly in water. The starch is produced either in a wet form or more commonly, dried. The cassava roots were peeled, washed and grated. The grated pulp is steeped for 2-3 days in a large quantity of water. The mixture is stirred and filtered through a piece of cloth sieve. The filtrate stands overnight and the supernatant is then decanted. The fine starch paste is collected and put in a wide metal pan that is already smeared with red palm oil. Water is added and then stirred with the hand to dissolve completely. The pot is put on fire and the solution constantly stirred with a wooden rod until it is converted to a very sticky, light yellow mass. This is eaten with any oil or soup (Etejere and Bhat, 1985). 2.4 Starter culture 2.4.1 Definition Spontaneous fermentation has been used for the production of verieties of fermented foods based on the microflora present in the fermentation environment as well as the raw material (Vogel et al., 2002) and it has been reported that the initiation of a natural fermentation takes long with high risk for failure (Holzapfel et al., 2000). During the long lag phase which was characterised with microorganisms physicochemically equilibrating with their environment, contaminating organisms from the environment slowly increase in number and compete for nutrients in order to produce metabolites. In addition, spontaneous fermentation has been reported by Oyewole and Sanni (1995) to lead to product inconsistencies since the end-product was dependent on the types and number of microorganism in the raw material. Thus, 2 4 the use of a preparation containing a large number of viable microorganisms was recommended, as this would lead to rapid acidification of fermentation process, the inhibition of spoilage and pathogenic organisms (Holzapfel, 1997, 2002), as well as a product with consistent quality. A starter culture may therefore be defined as a preparation or material containing large numbers of viable microorganisms, which may be added to accelerate a fermentation process. Being adapted to the substrate, a typical starter facilitates improved control of a fermentation process and predictability of its products (Holzapfel, 1997). Leroy and De Vuyst (2004) further define it as a microbial preparation of large numbers of cells containing at least one microorganism which is added to a raw material to produce a fermented food by accelerating its fermentation process. However, according to the DFG Senate Commission on Food Safety, “starter cultures are preparations of live microorganisms or their resting forms, whose metabolic activity has desired effects in the fermentation substrate, the food”. The early technologies in starter usage include the transfer of an old batch of fermented products to a new batch (back-slopping) and the indigenously-derived cultures (Westby et al., 1997). It was customary in the beginning when cereals were fermented by their natural flora, to put aside pieces called „sours‟ or „starters‟ for fermenting subsequent batches. 2 5 Cassava tuber Cassava Tuber Peel and cut into small sizes Peel and Wash Wash & put in pots (earthen) or drums and add water Ferment (3-4 days) Grate and add water Sieve out the starchy solution Steep for 3-4 days Allow to sediment Filter out starch Sediment packed into cotton & squeezed Solution Rolled into balls & wrapped Allow to sediment Partial steam cooking Decant Pound in mortar to a fine, smooth mash Sediment mixed with Wrapped into small balls & cook thoroughly palm oil and cook Usi Fufu Figure 2.1: Flow chart for traditional fufu (Oyewole and Odunfa, 1989) and usi (Etejere and Bhat, 1985) production 2 6 This back-slopping process often resulted in irregularities and unpredictability that led to the development and use of defined starter cultures (Holzapfel, 2002). Commercial starter cultures generally originated from food substrates or from the processes in which they were applied (Holzapfel, 2002), even though there has been a trend to isolate wild-type strains from traditional products (Beukes et al., 2001; De Vuyst et al., 2002; Leroy and De Vuyst, 2004). Edward et al. (2012) envisaged that the microorganisms isolated from the samples taken would be ideal for the development of a starter culture, as they would be typical of the fermentation and well adapted to the ecological conditions. Since the majority of starter cultures are natural isolates of the desirable microorganisms found normally in the substrates (Holzapfel, 2002; De Vuyst and Vancanneyt, 2007), a wide variety of species of organisms have been used in the food industry and many have been investigated for their potential use as starter cultures (De Vuyst and Neysens, 2005; Gaggiano et al., 2007). As at 2009, it was reported that relatively few LABs, out of millions of specie, have been isolated from starchy fermented foods in Africa and used as starter cultures (Yao et al., 2009), even though some reports indicated that Lactobacillus brevis, L. fermentum, L. plantarum, L. reuteri, Pediococcus pentosaceus and P. acidilactici exhibited superior performance in lactic fermented cereal, root crops and vegetable products (Steinkraus, 1996, 1997; Holzapfel, 1997; Lee, 1997; Oyewole, 1997). Some of the earlier studies that reported the use of defined starter cultures include Guiraud et al. (1998), Guiraud and Raimbault (1993) and Kimaryo et al. (2000) whom mentioned the use of the amylolytic L. plantarum as a starter culture during cassava fermentation for gari and kivunde respectively. Okafor et al. (1998) inoculated L. coryneformis and Saccharomyces sp. as a starter culture in cassava mash for gari production. Oyewole (1990) and Okolie et al., (1992) have both reported successfully producing acceptable fufu by the use of isolated pure starter cultures and the survival of these microorganisms was studied when placed alone or mixed in different carriers (Okafor et al., 1999). Various other reports have shown the utilization of starters during cassava fermentation (Kimaryo et al., 2000; Egounlety et al., 2007; Asmahan and Muna, 2009; Yao et al., 2009; Padonou et al., 2010). Starter cultures could be acquired in a ready-to-use, highly concentrated frozen /freeze dried form, or being propagated (Høier et al., 1999; Buckenhuskes, 1993; Hansen, 2002), ensuring the principles of hazard analysis critical control point (HACCP). 2 7 However, continuous propagation of starters from the original mixed microflora of traditional foods over time has led to the loss of certain beneficial strains as a result of natural ecosystem alterations. Moreso, certain metabolic characteristics of some starters, encoded in the plasmid are sometimes lost during propagation and these factors have been reported to reduce the biodiversity of starter cultures (Asmahan, 2010). This has prompted more studies towards the isolation of more strains even though typical commercial starters remain highly expensive which make people to still use small batches from a previous fermentation. 2.4.2 Properties of a starter culture Spontaneous food fermentations are neither predictable nor controllable. Pure cultures isolated from mixed populations of traditional fermented foods exhibit a diversity of metabolic activities, which vary even among strains. These include variability in growth rate, adaptation to a particular substrate, ability to degrade antinutritive compounds, antimicrobial properties, flavour and quality attributes and competitive growth behaviour in mixed cultures (Holzapfel, 1997). Therefore, the selection of suitable starter must be based on its ability to compete for survival, antagonise pathogens and spoilage microorganisms, produce acid or alcohol rapidly, confer desirable organoleptic changes, produce primary metabolites, degrade antinutritive and improve nutritive factors, detoxify and have probiotic features (Holzapfel, 1997). 2.4.3 Effects of starter on cassava fermentation Even though Hansen (2002) stated that the primary activity of the culture in a food fermentation is to convert carbohydrates to desired metabolites as alcohol, acetic acid, lactic acid or CO2, the introduction of starter culture to initiate a fermentation process is expected to bring about certain physical, chemical, organoleptic and nutritional changes to the products and such have been reported by numerous authors (Hernandez-Jover et al., 1997; Johanningsmier et al., 2007; Ogunbanwo et al., 2008; Ojokoh, 2009; Henshaw and Ikpor, 2010; Adetunde et al., 2011; Ahaotu et al., 2011; Hati et al., 2013). .The potential starters for cassava fermentation are pre-selected on the basis of suitable technological characteristics such as enzyme (amylase, pectinase, glucosidase, phytatse etc) and antimicrobial compound (bacteriocin, lactic acid, diacetyl, hydrogen 2 8 peroxide) production, survival rate, hydrolysis of complex carbohydrates, and rapid acidification of the fermentation process (Kostinek et al., 2007). Reduction of anti-nutritional compounds has been an important factor to be considered during the analysis of fermented products. Although lactic fermentation has been shown in certain studies to have reduced the phytate content, which is an antinutritive component of some cereals (Lopez et al., 1983; Mahajan and Chauhan, 1987; Svanberg and Sandberg, 1988; Khetarpaul and Chauhan, 1989; Svanberg et al., 1993), phytic acid-degrading ability is relatively rare among pure LAB cultures. Some L. plantarum strains have however, been reported to be capable of degrading phytic o acid on incubation at 37 C for 120 hours (Holzapfel, 1997). Phytase activity was not detectable in Bacillus sp. associated with the fermentation of the African locust bean Parkia biglobosa which was used in the preparation of iru or dawadawa (Aderibigbe and Odunfa, 1990). Detoxification during the fermentation of cassava was reported by Westby and Choo (1994) to be dependent primarily on microbial activity, even though, endogenous linamarinase present in cassava play a significant role in the process. Different starter cultures such as Lactobacillus sp, Bacillus sp, yeasts and molds play important roles during cassava processing (Ejiofor and Okafor, 1981; Hahn, 1989; Essers et al., 1995; Amoa-Awua and Jakobsen, 1995; Amoa-Awua et al., 1996; Olasupo et al., 1997) but an experiment comparing the effects of spontaneous fermentation, back-slopping and the use of starter cultures for the reduction of cyanogenic glucosides in cassava (Kimaryo et al., 2000) revealed that all three types of fermentations contributed significantly to the detoxification of cassava, with the starter-fermented (L. plantarum strains) cassava, having the best results. Rapid acidification rate is an important factor in cassava fermentation as this creates an unsuitable environment for spoilage and pathogenic microorganisms as well as hastening the entire process. It was reported by Henshaw and Ikpoh (2010) that after 96-hour controlled fermentation of cassava for fufu production, L. plantarum, Bacillus subtilis, Klebsilla sp and L. mesenteroides reduced the initial pH of fermenting mash from 6.2 to 3.68, 4.90, 4.88 and 4.68 respectively, with L. plantarum showing the highest acid-producing ability and this had earlier been reported by Oyewole (1990) who associated L. plantarum strain with high acid production during cassava fermentation for the production of fufu. Also, initial pH reduction from 5.62 to 3.05, 2 9 3.37 and 3.65 by L. plantarum, L. mesenteroides and S cerevisiae respectively, was reported during starter selection (Edema and Sanni, 2008). The production of acid, diacetyl, hydrogen peroxide and some other inhibitory compounds, in the fermenting meals were significantly higher in substrates fermented with starter cultures than in the spontaneously fermented ones as shown in the studies conducted by Edema and Sanni (2008) as well as Ogueke (2008). Nutritional, organoleptic and sensory qualities of starter fermented foods are expected to improve after the fermentation process. Cook (1994) and Steinkraus (1996) reported that the use of selected starter cultures for fermentation during processing of foods has been found to improve shelf life, nutritional status and to promote safety. Furthermore, Opere et al. (2012) observed increased nutritive value, acceptable organoleptic characteristics, and acceptable flavour properties in the fermentation of cereal gruel with starter culture. Ogunbanwo et al. (2013) also reported improved organoleptic property in burukutu while using mixed starter culture. Other numerous studies have shown the effect of starters on the nutritive and general sensory qualities of fermented foods (Kristek et al., 2004; Singh et al., 2012; Kabuo et al., 2013; Hasan et al., 2014). In general, the prospect of applying starter cultures will be achieved only if benefits, such as reduction of costs (e.g. energy), reduced fermentation duration, reduced risk of spoilage (increased shelf-life), improved process control, sensory quality (taste, aroma, visual appearance, texture, consistency), safety attributes (e.g. lower risk of diarrhoea, detoxification of cassava) and reduced preparation procedures for the final product, were perceived (Holzapfel, 2002). 2.4.4 Metabolite spectrum of cassava fermentation Through fermentation, microorganisms growing on inexpensive carbon sources can produce valuable products such as simple sugars, amino acids, nucleotides, vitamins and organic acids which had been proven to enhance flavour or increase nutritive values (Demain 1980). Similarly, microorganisms also produce a range of metabolites during food fermentation processes that can suppress the growth and survival of undesirable microflora in foods (Ross et al., 2002). Caplice and Fitzgerald (1999) reported that these metabolites not only increase the shelf life and microbiological safety of a food, but also make foods more digestible and reduce toxicity of the substrate. 3 0 Organic acids have been reported to be naturally present in foods or they are synthesized either during biochemical metabolic processes or bacteria metabolism (Akalin et al., 2002; Soyer et al., 2003; Karadeniz, 2004). They have important roles in food because they affect the organoleptic properties, stability, nutrition, acceptability and maintaining quality (Santalad et al., 2007). These acids have, not only antimicrobial effects but the organic acid-producing organisms are also considered as antimicrobial agents due to acid and antibodies production which lowers the pH of food/substrate to suppress the growth of other microorganisms. It has been reported that acetic and propionic acids produced by LAB strains may interact with cell membranes, and cause intracellular acidification and protein denaturation (Huang et al., 1986). Earnshaw (1992) stated that both are usually more antimicrobially effective than lactic acid due to their higher pKa values (lactic acid 3.08, acetic acid 4.75, and propionic acid 4.87), and higher percentage of undissociated acids. Acetic acid has been shown to have more inhibitory property than lactic acid bacteria towards Listeria monocytogenes (Ahmad and Marth, 1989; Richards et al., 1995), and toward the growth and germination of Bacillus cereus (Wong and Chen, 1988). It also acted synergistically with lactic acid, decreasing the pH of the medium, thereby increasing the toxicity (Adams and Hall, 1988). These organic acids are used as flavouring agents and the acceptability of food products depends largely on the flavour profiles, which are complex and are type- specific. Akalin et al. (2002) reported that those flavour profiles are influenced by many substances, e.g organic acids, sulphur compounds, lactones, methyl ketones and alcohols, as well as phenolic substances (Seitz 1990; Urbach, 1993). The flavour substances are formed as a result of the hydrolysis of fatty acids, normal bovine metabolic processes, and bacterial growth during food processing (Adda et al., 1982). Quantitative determination of organic acids is important to flavour studies for nutritional reasons and as an indicator of bacterial activity (Pham and Nakai 1983). With the use of High Performance Liquid Chromatography, number of authors have attempted to use organic acid content as an indicator of microbial metabolism and classification parameter of various food products (Marsili, 1985; Panari, 1986; Bevilacqua and Califano, 1989; Lombardi et al., 1994; Lues and Botha, 1998; Califano and Bevilacqua, 1999; Aka et al., 2008; Nour et al., 2010; Yuwono et al., 2011; Miguel et al., 2014). Mugula et al. (2003) reported the production of lactate during togwa fermentation whereas lactate, butyrate, acetate and formate were reported during 3 1 fermentation of ogi and maize dough by Banigo and Muller (1972) as well as Mensah et al. (1991), respectively. Sanni et al. (1999), during the analysis of cereal based beverages in Nigeria, detected the presence on lactic, acetic, malic, succinic and formic acids. In general, the organic acids are not only recommended as food additives by FAO for taste, aroma and texture but are also food preservatives. Among the organic acid metabolites are citrate, acetate, propionate, fumarate and most importantly, lactate which was the first organic acid to be produced on an industrial scale by fermentation in 1880 (Zadow, 1992) through controlled fermentation from the hexose sugars, molasses, corn, or milk. 3 2 CHAPTER THREE MATERIALS AND METHODS 3.1 Sample collection 3.1.1 Cassava samples Healthy cassava varieties TME 30572, TME 4(2)1425 and TME 50395 were obtained from the Tuber and Root Improvement Programme (TRIP), International Institute of Tropical Agriculture, Ibadan as well as local variety from a retailer at Bodija market, Ibadan and transported to the Microbial Physiology and Biotechnology laboratory, Department of Microbiology, University of Ibadan. 3.1.2 Organic acid and sugar standards The linamarin analogue (4-nitrophenyl-B-D-glucopyranoside), acetonitrile, KH2PO4 (HPLC grade), organic acids and sugar standards were obtained from Sigma (Sigma- Aldrich, Germany) 3.1.3 Test Isolates Test pathogenic organisms isolated from food samples were obtained from Food Microbiology and Biotechnology Unit of the Department of Microbiology, University of Ibadan. 3.2 Media and Sample preparation 3.2.1 Media preparation De Mann Rogosa and Sharpe (MRS) Agar, Violet Red Bile Glucose Agar (VRBGA), Malt Extract Agar and Plate Count Agar were used for Lactic Acid Bacteria enumeration, coliforms, yeast and total bacterial count respectively. They were prepared according to manufacturer‟s instruction (Appendix I, II, III and IV), o sterilized at 121 C and 15 psi for 15 minutes. 3 3 3.2.2 Sample Preparation and fermentation The method of Oyewole and Odunfa (1989) was used in which the cassava tubers were sorted by visual assessment, peeled, washed with clean tap water, and cut into small sizes (3-5 cm). Two hundred gram (200 g) of the cassava was submerged in 2 L tap water in plastic fermenters of 10 L capacity for the 72-hour fermentation at room temperature. The method of Etejere and Bhat (1985) was used for the fermentation of usi. Two hundred gram (200 g) cassava roots were grated and the pulp steeped in 2 litres tap water in a 10 L capacity plastic fermenter for 3 days. Samples of fermenting mash were taken every 24 hours and analyzed for microbiological and physicochemical parameters. 3.3 Total titratable acidity of fermenting cassava mash Titratable acidity was determined using the standard titration procedure for total titratable acidity (TTA) described by Lonner et al. (1986). Ten gram (10 g) of the fermenting mash was mixed with 90 mL sterile distilled water and homogenized. The mixture was filtered through Whatman Filter paper (No.1) and the filtrate titrated against 1M NaOH using 1% phenolphthalein as indicator. Acid equivalent is the amount of NaOH consumed in mL and each mL of 1M NaOH is equivalent to 90.08 mg of lactic acid. 3.4 pH of the fermenting cassava mash Ten gram (10 g) of fermenting cassava was weighed, mixed with 90 mL of sterile distilled water, homogenized and filtered through Whatman Filter paper (No.1). The pH of the resulting filtrate was then measured using a pH meter (H19107, Hanna), after standardizing with phosphate buffer solution at pH 4.0 and 7.0. 3.5 Isolation and identification of organisms from fermenting cassava tubers 3.5.1 Isolation technique Ten gram (10 g) of each sample was aseptically added to 90 mL of sterile peptone water and homogenized for 2 minutes. Samples were further diluted in a -9 -5 -7 tenfold serial dilution up to 10 . One millilitre (1 mL) from dilutions 10 , 10 and 3 4 -9 10 was introduced into sterile petri dishes and sterile PCA, VRBGA, MEA and MRS agar were added and allowed to solidify (Harrigan and MacCance, 1976). MRS agar o o culture plates were incubated at 30 C for 48 hours and the others at 30 C for 24 hours after which microbial counts were carried out. Representative LAB colonies were picked randomly from the plates and purified by repeated sub-culturing on fresh agar o plates. Pure cultures were grown on MRS agar slants and kept at 4 C for further use. o The stock cultures were stored at -4 C in glycerol broth for subsequent use. 3.5.2 Characterization of selected isolates Isolates were characterized and identified on the basis of their morphological (microscopic, macroscopic) and biochemical properties. 3.5.2.1 Morphological Characterization i Macroscopic: They were observed based on shape, colour, elevation, size, edge and surface on respective agar plate. ii Microscopic: This was carried out using the simple staining technique described by Norris and Ribbond (1971). A thin smear of the isolate was made on a clean glass slide and heat-fixed. Two drops of Crystal violet were applied onto the smear for 60 seconds. It was washed with water and stained with Grams iodine solution for 1 minute. The stain was decolourized by flooding the slides with ethanol until no more violet coloration was observed. Two drops of counter stain Safranin reagent were added for a minute, rinsed with water and blotted dry using filter paper. Microscopic observation was carried out under the oil immersion objective (Fisher Scientific, USA). Gram positive organisms were characterized by purple colouration after counter staining while Gram negative cells were pink in colour. Their shapes were also observed. 3.5.2.2 Biochemical Characterization i Catalase test Twenty four (24) hour old culture was obtained by sub-culturing on MRS agar o plates and incubated at 30 C for 24 hours. A drop of freshly prepared 3% hydrogen peroxide solution was added to a clean glass slide. With the aid of a sterile wire loop, the culture was aseptically picked onto the slide (Seeley and Van Demark, 1972). 3 5 Evolution of gas as white froth indicated catalase positive reaction. Catalase negative isolates were selected for further tests. ii Motility test In semi-solid agar medium, motile organisms swarm and gave a diffuse spreading growth that was easily visually recognized. Agar agar (0.2% w/v) was dissolved in 100 mL nutrient broth and 10 mL of the medium was dispensed into test o tubes. The medium was sterilized at 121 C, 15 psi for 15 minutes and allowed to set in a vertical position. Inoculation was done with a sterile needle by making a single stab down the centre of the tube to about half the depth of the medium and incubated at o 37 C for 48 hours. Non-motile organisms gave growth that were confined to the stab line, had sharply defined margins and left the surrounding medium clearly transparent. Motile organisms gave diffuse hazy growth that spread throughout the medium (Olutiola et al., 2000) iii Voges-Proskauer test The selected isolates were each inoculated into sterile glucose phosphate broth (Glucose 0.5 g, KH2PO4 0.5 g, Peptone 0.5 g and distilled water 100 mL) and o incubated at 37 C for 2-5days. Aliquot (1 mL) of 66% alpha-naphtol solution and 1 mL of 10% NaOH was added and the culture thoroughly shaken. Appearance of a pale pink colouration within 5 minutes was noted as positive reaction. The solution was left for a while to check for slow reaction in case of false negative result (Reuter, 1970). iv Oxidase test Whatman No. 1 filter paper was soaked in oxidase reagent (1% aqueous tetramethyl-p-phenylenediamine dihydrochloride). Sterile wire loop was used to touch colony of the test isolates, then transferred onto the reagent spot on the filter paper. Formation of a very deep purple coloration within 10 seconds indicated a positive reaction, while absence of a deep coloration indicated negative reaction (Seeley and Van Demark, 1972). 3 6 v Indole test Each isolate was inoculated into sterile peptone water medium (Peptone 9 g, o distilled water 100 mL) and incubated at 37 C for 48 hours. Five to six drops of ‟ Kovac s reagent were then added to the culture. Development of rose pink coloration on the surface of the culture indicated positive reaction of indole production. No colour change was recorded as negative (Kovac, 1956). vi Methyl red test Glucose phosphate peptone broth (Glucose 0.5 g, KH2PO4 0.5 g, Peptone 0.5 g and distilled water 100 mL) was prepared as described by Harrigan and McCance (1966). Ten millilitres (10 mL) of the broth was dispensed into screw cap tubes and o sterilized at 121 C and 15 psi for 15 minutes. Inoculation with 24 hour old culture was o aseptically done and incubated at 37 C for 2-5 days. After incubation, a few drops of methyl red indicator were added to the culture and a resultant definite red colouration was considered positive result. vii Production of ammonium from arginine This test was done to detect the ability of the organisms to produce deaminase enzyme which enables them to break down proteins to release ammonia. Modified MRS broth (10 mL) containing 0.3% arginine (Appendix V) was dispensed into screw o cap tubes and sterilized at 121 C and 15 psi for 15 minutes. Loopful of cultures were o inoculated aseptically and incubated at 37 C for 5-7 days. Uninoculated tube served as control. A loopful of Nesslers reagent was added to a loopful of the culture on a clean glass slide and observed. Presence of ammonia is indicated by the formation of orange to brown colour while pale yellow or no colour change indicated no reaction (Olutiola et al., 2000) viii Sugar fermentation test This test was for investigating the ability of the bacteria isolates to utilize different sugars. 10 g of different sugars (Glucose, maltose, mannitol, sucrose, melibiose, galactose, fructose, sorbitol, raffinose, lactose, xylose, arabinose, inositol and sorbose) were dissolved in 100 mL distilled water and each was sterilized at o 121 C, 15 psi for 10 minutes. 1 mL of each sugar solution was added to 9 mL of sterile 3 7 modified MRS broth in test tubes, as the sole carbon source with methyl red indicator to give a final concentration of 1% (v/v). Each tube contained an inverted Durham tube. The tubes were inoculated with 24 hour old LAB cultures and incubated at 30°C for 5 days. Un-inoculated tubes served as control. Positive reaction with acid production was indicated by a colour change from red to yellow and/or gas production which was collected in the Durham tubes (Olutiola et al., 2000) 3.5.3 Identification of organisms Selected isolates were identified based on comparison of the results obtained ‟ from the macroscopic, microscopic and biochemical characterization with the Bergey s Manual of Determinative Bacteriology (Holt et al., 1994). 3.6 Molecular Identification 3.6.1 DNA extraction The DNA was extracted according to the procedure of Pitcher et al. (1989) with some modifications. Bacteria culture grown overnight was centrifuged at 13000 g for 2 minutes. Cell pellet was suspended in lysis buffer (pH 8.0) containing 25 mM Tris-HCl (Sigma), 10 mM EDTA, 50 mM sucrose, 10 mg/mL lysozyme and incubated o at 37 C for 30 minutes. 0.5 mL aliquots of the mixture of 5M guanidine thiocyanate, 0.1N EDTA and 0.5% N- lauroyl sarcosine sodium salt (Sigma, England) was added o and incubated at 30 C for 15 minutes. Precipitation was done using chloroform: isoamyalcohol (24:1) and centrifugation at 13000 g for 10 minutes. Upper protein precipitate was removed, added to isopropanol and centrifuged for 5 minutes. Resultant pellets were then washed in 70% ethanol. Purification was done by dissolving in 1X TE buffer (10 mM Tris-HCl and 1 mM EDTA, pH=8.0) containing 10 mg/mL of RNase and incubated at 37 °C for 30 minutes. 3.6.2 DNA Amplification The bacterial universal primers 27F (5'-AGAGTTTGATCCTGG CTCAG-3') and 1492R (5'-GGTTACCTTGTTACGACTT-3') previously shown to be useful for identification of Lactic Acid Bacteria (Lane, 1991) were used to amplify approximately 600 base pairs of the genomic 16S conserved region. Amplification method of Tajabadi et al. (2013) was used. The PCR reaction was carried out in a total volume of 25 μL with a reaction mixture with the following: 1× 3 8 Taq Master Mix (Promega, UK), deoxynucleoside triphosphates, 1.5 mM MgCl2, 0.25 mM forward primer, 0.25 mM reverse primer and 0.4 mg of genomic DNA. The reaction mixture in micro-centrifuge tube was amplified in a thermocycler PCR o system (Techne-Progene, UK) in which an initial heating of 95 C for 3 minutes was o o followed by 40 cycles of denaturation at 95 C for 30 seconds, annealing at 55 C for 55 o seconds, extension at 72 C for 1 minute, and terminating with a 10-minute final o incubation at 72 C. 3.6.3 Profile sequencing The successful PCR amplification products were sequenced using an ABI Bigdye 3.1 sequencing kit (Applied Biosystems, California USA) on ABI 3730XL automated sequencing analyzer at Laragen Inc., Culver City, California. Sequences (Appendix VI) obtained were compared with sequences listed in GenBank database using the NCBI Basic Local Alignment Search Tools (BLAST) and identified based on the closest relative (Altschul et al., 1997). The 16S rRNA gene clone sequences that had 99-100% similarity to sequences deposited in GenBank were designated as belonging to the corresponding species and strains. Phylogenetic analysis was conducted utilising computer software plan and neighbour-joining methods with MEGA (6) package. All bacterial 16S rRNA gene sequences were entrusted in GenBank using their accession numbers. 3.7 Screening for potential starters 3.7.1 Starch hydrolysis Selected LAB isolates were streaked on sterile modified MRS agar containing 0.4% (w/v) soluble starch as the sole carbon source. The cultures were incubated at 37°C for 24 hours, after which they were flooded with Gram‟s iodine. Degradation of starch was evident by zone of inhibition around the streak, indicating starch utilisation (Edward et al., 2012). 3.7.2 Production of linamarase A medium for testing linamarase (in the form of β –glucosidase) production was prepared by adding 0.1 g of 4-nitrophenyl-B-D-glucopyranoside (Sigma Aldrich, 3 9 Germany) to 100 mL 0.666 M NaH2PO4 (pH 6). The mixture was dissolved and filter- sterilized. The test culture was grown on MRS agar for 24 hours at 30°C. Colonies were picked from the plates using a sterile loop and were emulsified in physiologic saline to McFarland Turbidity Standard No. 3. Thereafter, 0.75 mL of culture was added to 0.25 mL of the test medium. It was incubated at 30°C overnight. Positive isolates that produced β -glucosidase degraded the linamarin analogue and changed the colour of the mixture from colourless to distinct yellow (Edward et al., 2012) 3.7.3 Production of pectinase Chemically defined medium (CDM) containing pectin (Appendix VII) for the screening for pectinase enzyme was used. The test culture was grown on MRS agar for 18 hours at 30°C. Colonies were picked from the plates using a sterile loop and streaked on the sterile CDM plate, then incubated for 24 hours. Colonies showing zone of inhibition upon flooding with 1% cetyltrimethyl ammonium bromide were confirmed as pectinase producers (Altan, 2004). 3.7.4 Acidification of growth medium by selected isolates MRS broth was prepared from a single batch with initial pH 6.50. Aliqouts (10 o mL) were dispensed into screw cap tubes and autoclaved at 121 C and 15 psi for 15 minutes. The selected isolates were inoculated into each tube and incubated at 30°C. Acid production was determined by measuring the pH of the culture medium at 24, 48 and 72 hours respectively (Kostinek et al., 2005). 3.7.5 Production of Lactic acid The amount of lactic acid produced in the fermented cassava was determined by the standard titration procedure for total titratable acidity (Lonner et al., 1986). Twenty five millilitres (25 mL) of broth culture was titrated with 1M NaOH using phenolphthalein as indicator. Acid equivalent is the amount of NaOH (mL) consumed. Each mL of 1M NaOH is equivalent to 90.08 mg of lactic acid. 3.7.6 Production of Hydrogen peroxide Hydrogen peroxide production was determined by measuring 25mL of broth culture into a 100mL flask. To this was added 25 mL of dilute H2SO4 (10%). The preparation was then titrated with 0.1M potassium permanganate (KMnO4). The end 4 0 point was the point at which the pale pink colour persisted for 15 seconds before de- colourization. Each mL of 0.1M KMnO4 is equivalent to 1.701 mg of H2O2 (Sanni et al., 1995). The volume of H2O2 produced was then calculated as follows: KMnO (mL) KMnO (M )M .E.100 H 2O2 concentration = 4 4 H 2SO4 (mL) sample(mL) 3.7.7 Production of Diacetyl To 25 mL each of the broth culture, 7.5 mL of hydroxylamine solution (1M) was added in two flasks (one flask was for residual titration). Both flasks were titrated with 0.1M HCl to a greenish yellow end point using bromophenol blue as indicator (Sanni et al., 1995). The equivalence factor of HCl to diacetyl is 21.52 mg. The concentration of diacetyl produced was calculated as follows: R  S 100E Ak  W where Ak (mg) is the percentage of diacetyl, R the volume of 0.1M HCl used in residual titration, S the volume of 0.1M HCl used in titration of sample, E the equivalence factor and W is the volume of sample. 3.7.8 Antimicrobial activity against pathogenic organisms 3.7.8.1 Preparation of Cell-Free Filtrate MRS broth (10 mL) was inoculated with selected isolates and incubated at 37°C for 48 hours. A cell-free solution was obtained by centrifuging the bacterial culture at 6000 rpm for 15 minutes followed by filtration of the supernatant through 0.2 mm pore size filter, thus, obtaining cell free filtrate (Kalalou et al., 2004). 3.7.8.2 Agar Well Diffusion Test The antimicrobial activity of the isolated LAB (cell free filtrate) against Escherichia coli, Bacillus cereus, Proteus species, Salmonella sp., Corynebacterium sp. and Shigella sp. was performed by the agar well diffusion assay. Loopful of the test o pathogenic bacteria were incubated in brain heart Infusion (BHI) broth at 30 C for 24 hours. Petri dishes containing 20 mL of sterile Muller Hinton agar were streaked with 0.1 mL of 24 hour-old broth culture of each pathogenic bacterium. Wells were cut on 4 1 the agar using sterile 5 mm corkborer and filled with 100 μL of cell-free filtrate. The plates were stored for 2 hours in the middle compartment of the refrigerator and then incubated at 37°C for 24 hours. The antimicrobial activity was determined by measuring the diameter (mm) of inhibition zone around the wells (Schillinger and Lucke, 1989). 3.8 Fermentation studies with identified isolates This was carried out by inoculating the isolates singly and in-combination into a 10 L capacity bioreactor containing cassava in cut pieces and grates, soaked in 2 L sterile distilled water (pH 7.1) for fufu and usi production respectively. 3.8.1 Preparation of cassava for controlled fermentation Cassava tubers for fufu were cut into small pieces of about 3 – 5 cm long while those for usi were grated. Cut/grated cassava tubers (200 g) were sterilized using 0.1% HgCl in 70% ethanol followed by rinses with sterile distilled water (Adetunde and Onilude, 2010). 3.8.2 Determination of Inoculum size The selected lactic acid bacteria were inoculated into sterile MRS broth and o incubated at 30 C for 48 hours. Aliquot (1 mL) of broth culture was introduced into a fresh sterile broth and incubated for 24 hours. At the end of incubation period, the broth was centrifuged (Himac CR21GII, Japan) at 5000 rpm for 10 minutes. The supernatant was decanted while the pellet was washed with sterile distilled water and re-centrifuged before being suspended in sterile normal saline. The inoculum size was determined using McFarland standard. Dilutions were made with sterile normal saline to McFarland standard (No 4) using a spectrophotometer (Cecil CE 1011, Cambridge, England) to give 0.669 optical density at 600 nm, 9 resulting in an approximate cell density of 1.2×10 CFU/mL (McFarland, 1907). Aliquots (5 mL) of each resultant diluent was used as inoculum in all cases either singly or in-combination to inoculate 200 g of the cassava soaked in 2 litres sterile distilled water and fermentation was allowed for 72 hours at room temperature. 4 2 3.9 Performance studies on fermenting mash 3.9.1 pH of fermenting mash Ten gram (10 g) of fermenting cassava samples was mixed with 90 mL of sterile distilled water, homogenized and filtered through a Whatman‟s Filter paper (No. 1). The pH was then measured at 24 hours interval using a pH meter (H19107 Hanna), after standardizing with phosphate buffer solution of pH 4.0 and 7.0. 3.9.2 Total titratable acidity of fermenting mash Titratable acidity was determined using the standard titration procedure for total titratable acidity (TTA) as described by Lonner et al. (1986). Ten gram of the fermenting mash was mixed with 90 mL sterile distilled water and homogenized. The mixture was filtered through Whatman filter paper (No. 1) and the filtrate titrated against 1M NaOH using 1% phenolphthalein as indicator. Acid equivalent is the amount of NaOH consumed in mL. Each mL of 1M NaOH is equivalent to 90.08 mg of lactic acid. 3.9.3 Nutritional Analysis of fermenting mash Methods described by AOAC (2005) were used to estimate moisture content, crude protein, fat, fibre, ash and total carbohydrate. 3.9.3.1 Determination of moisture content A metallic dish was dried in an oven (Fisher scientific, model 655F, USA) at o 110 C for 10 minutes to a constant weight. It was cooled in a desiccator for 30 minutes and weighed. Five gram of the sample was placed in the dish and weighed. The dish o with the sample was then dried in an oven at 105 C for 4 hours to achieve a constant weight and was quickly transferred to the desiccator to cool. It was re-weighed immediately after cooling with minimum exposure to the atmosphere. The loss in weight of the sample during drying was the moisture content. 4 3 3.9.3.2 Determination of crude protein The crude protein content of fermenting cassava mash was determined by the micro Kjeldahl method using a protein conversion factor of 5.80. Sample (0.5 g) was weighed and placed on filter paper, then folded and dropped into a Kjeldahl digestion tube. Three gram (3 g) digesting mixed catalyst (CuSO4 + Na2SO4) and 25 mL of concentrated Na2SO4 were added to the sample in the digestion tube. The mixture was transferred to the Kjeldahl digestion apparatus with the heater regulated at a temperature below the boiling point of the acid until frothing ceased. The mixture was allowed to boil vigorously as the temperature increased, until a clear (light) green colour was obtained. The digest was allowed to cool, then transferred into 100 mL volumetric flask and diluted with distilled water to make up 100 mL. Ten millilitres (10 mL) aliquot of the digest was introduced into the distillation jacket of the micro steam distillation apparatus that was connected to the main, as the water in the distiller flask boils. 20 mL of 40% (w/v) NaOH was added to the digest in the distillation jacket. 50 mL of 40% boric acid was measured into 250 mL conical flasks and four (4) drops of methyl red indicator were added each. The conical flask containing the mixture was placed onto the distillation apparatus with the outlet tube inserted into the conical flask and NH3 was collected through the condenser. The distillation continued until 25 mL of the distillate was trapped into the boric acid solution and colour changes from red to yellow. The distillate was then titrated against 0.02 M HCl and the titre values recorded. Percentage crude protein was calculated (AOAC, 2005). 3.9.3.3 Determination of crude fat The fat content was determined by direct Sohxlet extraction using petroleum o ether (bp 40–60 C) as solvent (AOAC, 2005). Sample (0.5 g) contained in a filter paper was placed in the sohxlet extraction apparatus (Osk, Japan) and the set-up was placed on a heating mantle. The heat source was adjusted so that the solvent boiled gently and refluxed several times for 6 hours until the ether had siphoned over and the barrel of the extractor was empty. On removal, the filter paper containing the sample o was dried in an oven at 50 C to a constant weight and percentage lipid (fat) was then calculated (AOAC, 2005). 4 4 3.7.3.4 Determination of ash content The ash content of the fermented cassava was determined by placing a pre- weighed crucible with 5 g of the sample on a burning flame for 15 minutes until smoke ceased. It was then transferred into a muffle furnace (Gallenkamp, England), and o heated at 550 C for 4 hours, leaving a white ash. The crucible was removed, immediately covered and placed in a desiccator to cool. The weight was measured and the percentage ash was calculated (AOAC, 2005). 3.9.3.5 Determination of fibre content Cassava sample (5 g) was weighed into a round bottom flask. Sulphuric acid 100 mL, 0.25 M) was added and the mixture boiled for 30 minutes. The hot solution was quickly filtered under suction. The residue was thoroughly washed with hot water until acid free. This was further transferred into labeled flask and 100 mL of hot 0.3 M sodium hydroxide solution was added. The mixture was allowed to boil again under reflux for 30 minutes and filtered quickly under suction. Insoluble residue was washed with hot water until it was base free, dried to a constant weight in an oven at 100ºC for 2 hours, cooled in desiccator and weighed. The weighed samples were then incinerated, and reweighed. Percentage crube fibre content was calculated (AOAC, 2005). 3.9.3.6 Determination of total carbohydrate The total carbohydrate content was calculated by difference in which the sum of the percentage moisture, ash, crude lipid, crude protein and crude fibre was subtracted from 100%. 3.9.4 Anti-nutritional factors of the fermenting cassava mash 3.9.4.1 Determination of cyanide content The cyanide content of the fresh and fermenting cassava was determined using the method of Ojimelukwe (1997). Twenty gram (20 g) of the crushed cassava was homogenized in 200 mL distilled water for 10 minutes. The homogenate was incubated for 18 hours at room temperature after which 100 mL of 5% NaHCO3 was added to it before distillation. After distillation, the filtrate was collected and titrated 4 5 against 0.2% iodine solution using 1% starch as indicator. Cyanide content was calculated using the titre value. 3.7.3.7 Estimation of phytic acid The phytic acid was determined using the procedure described by Markkar et al. (1993). Two gram (2g) of sample was weighed into 250 mL conical flask. A hundred millilitre concentrated HCl acid (2% v/v) was used to soak the sample in the conical flask for 3 hours and then filtered through a double layer of hardened filter papers. The filtrate (50 mL) was placed in 250 mL beaker and 100 mL of distilled water was added to give proper acidity. Ten millilitres (10 mL) of 0.3% (w/v) ammonium thiocyanate solution was added to the solution as indicator. The solution was titrated with standard iron chloride solution, which contained 0.00195 g iron per millilitre. The end point colour was slightly brownish-yellow which persisted for 5 minutes. The percentage phytic acid was calculated using the titre value. 3.7.3.8 Estimation of tannin content The method described by Markkar et al. (1993) was adopted. Four hundred milligram (400 mg) of sample was placed in 500 mL conical flasks and 40 mL diethyl ether containing 1% acetic acid (v/v) was added, then the mixture was properly mixed to remove the pigment materials. The supernatant was carefully discarded after 5 minutes and 20 mL of 70% aqueous acetone was added and the flasks were sealed with cotton plug covered with aluminum foil, then kept in shaker for 2 hours for extraction. The content of each flask was filtered through Whatman filter paper (No. 1) and samples (filtrates) were used for analysis. Aliquot (50 mL) of tannin extract from the sample was introduced into test tubes and the volume was made up to 100 mL with distilled water. Folin-Ciocalteua reagent (0.5mL) was added to each and mixed properly after which 2.5 mL of 20% (w/v) sodium carbonate solution was added and further mixed. The mixture was kept for 40 minutes at room temperature, and absorbance was read at 725 nm using spectrophotometer (Cecil CE 1011, Cambridge, England). Tannin concentration was estimated from the tannic acid standard curve. 4 6 3.10 Estimation of enzyme activities during fermentation 3.10.1 Enzyme crude extract An 80 mL sample of 0.1 M citrate buffer, pH 6.5, was added to 40 g of cassava o mash, homogenized and kept overnight at 4 C. The mixture was centrifuged (Himac CR21GII Hitachi, Japan) at 12,000 rpm for 30 minutes. The resulting filtrate was used for enzyme assay (Ampe and Brauman, 1995). 3.10.2 Amylase activity This was determined by DNS method (Miller, 1959). One millilitre (1 mL) of sample was added to 1 mL of standard starch solution (containing 1% soluble starch in 0.1 M phosphate buffer pH 7) and incubated at room temperature for 30 minutes. Reducing sugars were estimated by stopping the reaction with the addition of 1 mL Di- nitrosalicylic acid (DNS) reagent, boiled for 5 minutes in a water bath (Clifton, England) and cooled under running tap water. Thereafter, 2 mL of sterile distilled water was added to dilute the solution. The absorbance of the resulting solution was determined at 540 nm with spectrophotometer (Cecil CE 1011, Cambridge, England) against a reagent blank. A standard graph was generated using standard glucose solution (Appendix VIII). One unit of amylase activity was taken as the amount of enzyme in 1 mL of crude amylase that produced 1 mg of reducing sugars under the standard assay conditions (Panduranga et al., 2010). 3.10.3 Linamarase assay Quantification of linamarase was based on the degradation of linamarin analogue, para-nitrophenolgalactosidase (PNPG) and determination of the released p- nitrophenol (Ikediobi et al., 1980). The assay medium included 0.5 mL of enzyme extract and 1 mL of PNPG (5 mM) in 0.01 M phosphate buffer (pH 6.8). The mixture was incubated for 15 minutes at 65°C, and the reaction was terminated by the addition of 2 mL of 0.2 M borate buffer (pH 9.8). Colour of the released p-nitrophenol was measured at 425 nm against enzyme blank using spectrophotometer (Cecil CE 1011, Cambridge, England). A standard graph was generated using standard PNPG solution (Appendix IX). One unit of linamarase activity was expressed as the amount that caused a change in absorbance of 0.01 unit against an enzyme blank under the defined assay conditions (Ugwuanyi et al., 2007). 4 7 3.10.4 Pectinase assay Pectinase enzyme was assayed using the colorimetric method as outlined by Miller (1959). Five milliliters (5 mL) of cell free supernatant was incubated with 2% pectin in 0.1 M acetate buffer (pH 6.0) and the reaction mixture was incubated at 40°C for 10 minutes. After adding 1.0 mL of DNSA reagent (without sodium potassium tartarate), the mixture was boiled for 5 minutes at 90°C. The reaction was stopped by adding 1 mL of Rochelle‟s salt (sodium potassium tartarate - Sigma, USA). The mixture was further diluted by adding 2 mL of distilled water after which the absorbance was read at 595 nm using spectrophotometer (Cecil CE 1011, Cambridge, England) to estimate the reducing sugar released. A standard graph was generated using standard glucose solution. One unit of pectinase activity was defined as the amount of enzyme which liberated 1 μm glucose/minute (Karthik et al., 2011). 3.11 Analysis of metabolites in the fermenting mash Metabolites were analyzed by the high performance liquid chromatography (HPLC) according to the method of Andersson et al. (2007), with slight modification to the flow rate. The organic acid concentration was measured using HPLC (CECIL CE4200, Cambridge England) equipped with a C18 column (Ultra Aqueous, 5µm, 150mm x 4.6 mm, Restek). The mobile phase was 50 mM KH2PO4 buffer with 5% acetonitrile (pH 2.5 adjusted by 37% HCl) at a flow rate of 1.0 mL/minute. Samples (20µL) were injected manually for analysis with ultra violet detection at 210 nm. Samples for acid analysis were taken at 24 hour intervals and centrifuged (Himac CR21GII Hitachi, Japan) at 10 000 rpm for 10 minutes at 4°C. The supernatant was diluted with the mobile phase and 37% hydrochloric acid in the ratio 0.1:0.8:0.1 mL respectively and further filtered through a 0.2 µm syringe filter. Sugar concentration was determined using the same HPLC system as above but equipped with a Series 200 refractive index (RI) detector (Perkin-Elmer), guard column, and an ion exchange column (Aminex HPX87-P, BioRad). The column was kept at 85°C in a column oven for optimal performance. Prior to analysis, the samples were centrifuged at 10 000 rpm for 10 minutes at 4°C. The supernatant was diluted with water and filtered through a 0.2 µm syringe filter. Samples (20 µL) were injected -1 manually and water, at a flow rate of 1.0 mL min was used as the mobile phase. The analysis was externally calibrated using standard solutions as prepared for the samples. Peak area from the chromatograms were evaluated by comparison to standard 4 8 curves generated (Sigma Aldrich, Germany) with known concentrations of organic acids (lactic acid, acetic acid, propionic acid and butyric acid) as well as sugar standards (sucrose, glucose, fructose, xylose, maltose, ribose, arabinose, lactose and rhamnose). 3.12 Selection of starter The selection of a common starter for the two products was based on improved nutritional contents, reduced anti-nutritional contents and rapid acidification. Results were statistically analysed using SPSS 20.0 software package and the most common to both food products was selected for further use. 3.13 Optimization studies on selected starter 3.13.1 Determination of inoculum size The inoculum size was determined using McFarland standard. Dilutions were made using 18-hour-old culture of the selected starter with sterile normal saline to McFarland standard (No 4) using a spectrophotometer (Cecil CE 1011, Cambridge, England) to give 0.669 optical density at 600nm, resulting in an approximate cell 9 density of 1.2×10 CFU/mL (McFarland, 1907). 3.13.2 Determination of optimal pH Effect of different pH values on starter growth was determined by inoculating 7 0.1 mL of 18-hour-old culture (10 ) onto sterile MRS broth that was adjusted to different pH values (3.5, 4.5, 5.5, 6.5, and 7.5). The broth culture was incubated for 24 hours and the optical density (OD) determined at 600 nm using spectrophotometer (Cecil CE 1011, Cambridge, England) (Tofan et al., 2002). 3.13.3 Determination of optimal incubation temperature Effect of different incubation temperatures on starter growth was determined 7 by inoculating 0.1 mL of 18-hour-old culture (10 ) onto sterile MRS broth and o o o o o incubated at different temperatures (25 C, 30 C, 37 C, 40 C and 45 C) for 24 hours. The OD at 600 nm was determined (Tofan et al., 2002). 4 9 3.13.4 Determination of optimal salt concentration Optimal salt concentration for starter growth was determined by inoculating 0.1 7 mL of 18-hour-old culture (10 ) onto modified MRS broth containing different NaCl o concentrations (2%, 4%, 6%, 8% and 10%). They were incubated at 30 C for 24 hours and OD at 600 nm was determined (Tofan et al., 2002) 3.13.5 Effect of agitation on growth Agitation intensity had effect on the mixing and oxygen transfer rates in a culture medium and thus influences cell growth and product formation (Wijbenga et al., 1991). Broth cultures were subjected to different agitation speed using an incubator 7 shaker (Thermoscientific, USA). Aliquot (0.1 mL) of 18-hour old culture (10 ) was inoculated into sterile MRS broth and subjected to agitation at 50 rpm, 100 rpm, 150 rpm, 200 rpm and 250 rpm respectively. OD at 600 nm was determined after 24 hours using a spectrophotometer (Cecil CE 1011, Cambridge, England). 3.13.6 Effect of carbon sources on growth The effect of different carbon sources (starch, glucose, galactose, maltose and lactose) on growth of selected starter was determined according to the method of Enitan et al. (2011) with modification to the optical density used. Overnight culture (0.1mL) was inoculated into modified MRS broth containing 0.2 g MgSO4.7H2O, 0.05 g MnSO4.4H2O, 5 g sodium acetate, 1.5 g of K2HPO4, 1.5 g of KH2PO4, 10 g peptone, 5 g yeast extract, 1 mL of Tween 80 per litre of distilled water and each sugar at 2% o w/v was introduced as the sole carbon source. Incubation was done at 30 C and optical density at 500 nm determined using a spectrophotometer (Cecil CE 1011, Cambridge, England) after 24-hour incubation. 3.13.7 Effect of nitrogen sources on growth The effect of different nitrogen sources (peptone, yeast extract, casein, urea and [NH4]2SO4) on growth of the selected starter was determined by inoculating 0.1mL of overnight culture into modified MRS broth containing 0.2 g MgSO4.7H2O, 0.05 g MnSO4. 4H2O, 5 g sodium acetate, 1.5 g KH2PO4, 6% w/v glucose, 1.0 mL vitamin solution (0.2 g vitamins B6, 0.1 g riboflavin and 0.1 g folic acid in 100 mL of 20% ethanol) per litre of distilled water and each listed substrate at 2% w/v as its sole o nitrogen source. Incubation was done at 30 C for 24 hours and the optical density was 5 0 determined at 500 nm using a spectrophotometer (Cecil CE 1011, Cambridge, England) (Enitan et al., 2011). 3.13.8 Effect of different incubation time on growth The growth rate was determined by inoculating 0.1 mL of 18-hour old culture o into sterile MRS broth. It was incubated at 30 C and samples withdrawn at 24-hour intervals for three days. The optical density at 620 nm was determined using a spectrophotometer (Cecil CE 1011, Cambridge, England) (Manca De Nadra et al., 2003). 3.14 Application of starter in traditional processing of fufu/usi under optimized conditions and performance studies on the products 3.14.1 Inoculum preparation The selected starter(s) were cultivated on sterile MRS agar. The culture plates o were incubated at 30 C for 48 hours. Pure colonies were introduced onto a fresh broth o (30 mL) using the determined optimal growth conditions and incubated at 30 C for 24 hours. At the end of the incubation period, the broth was centrifuged (Himac CR21GII o Hitachi, Japan) at 5000 rpm for 10 minutes at 4 C. The supernatants were decanted while the pellets were washed with sterile distilled water and re-centrifuged before being re-suspended in sterile distilled water. Dilutions were made with sterile normal saline to McFarland standard (No 4) using a spectrophotometer (Cecil CE 1011, Cambridge, England) to give 0.669 optical density at 600 nm, resulting in an 9 approximate cell density of 1.2×10 CFU/mL. Aliqout (5 mL) of each diluent was used singly and in-combination as inoculum for the fermentation process. An un-inoculated experiment served as the control (McFarland, 1907). 3.14.2 Preparation of Fufu and Usi Fufu: The fermented roots were taken out after 72-hour fermentation, broken with clean hands and the fibres removed by sieving through a muslin cloth after the addition of water. The starch suspension was allowed to sediment in a large container for about 24 hours after which the water was decanted. The fine, clean starch was thereafter dewatered by putting in cotton bags and pressed with heavy stones overnight (Oyewole and Odunfa, 1989). A quantity of the slurry containing about 25% of fufu 5 1 paste in water was boiled in an open pan. After continuous stirring using a wooden rod, strong dough (ready-to-eat fufu) was formed (Kwatia, 1986; Ayankunbi et al., 1991; Anon, 1994). Usi: The fermented grated root mixture was stirred and filtered through a piece of cloth (sieve). The filtrate stands overnight and the supernatant was decanted. The fine starch paste was collected and put in a wide metal pan already smeared with red palm oil. Water was added and then stirred with the hand to dissolve completely. The pot was put on fire and the solution constantly stirred with a wooden rod until being converted to a very sticky, light yellow mass (Etejere and Bhat, 1985). 3.12.3 Nutritional analysis of cassava fermented with starter cultivated under optimized growth condition Methods described by AOAC (2005) were used to estimate crude protein, fat, carbohydrate, fibre, ash and moisture contents as earlier described. Anti-nutritional content (tannin, phytic acid and cyanide) were also estimated as earlier described. The results were compared to un-inoculated samples (control) as well as the samples obtained from the starter that was not subjected to optimized growth conditions. 3.12.4 Sensory evaluation of products The acceptability of the cassava products was determined through sensory evaluation using Hedonic scale test. The products were presented in small plates and coded, to avoid bias from panelists. Modified method of (Iwe, 2002) was used in which a 20-man trained panel each that was familiar with each food product was set up to determine their general acceptability. Panellists were provided with drinking water to rinse their mouth after tasting each sample. A 7-point Hedonic rating on the degrees of acceptability was conducted with score 7 having like extremely, 6- like very much, 5- like moderately, 4- indifferent, 3- dislike moderately, 2- dislike very much and 1 having dislike extremely for the following attributes (texture, colour, taste, odour and overall acceptability). Each product was considered against a control (spontaneously fermented) product. 5 2 3.12.5 Microbiological assessment and physical evaluation of products during storage Samples of stored fufu and usi were taken at 24 hours interval for microbial enumeration over a 7-day period. One gram (1 g) each of starter-fermented and the spontaneously-fermented fufu and usi were homogenized in 9 mL of sterile peptone -9 water. Ten-fold serial dilution was done aseptically and 0.l mL of 10 dilution was introduced into sterile petri dishes. Sterile Plate Count Agar, MRS agar, Violet Red Bile Glucose Agar, Malt Extract Agar, Potato Dextrose Agar and Manitol Salt Agar for the enumeration of total bacteria, LAB, coliforms, yeast, mold and Staphylococcus respectively, were poured into the plates, allowed to set and incubated. Microbial count was done thereafter on each agar medium. Furthermore, spoilage symptoms (appearance of mold and slime, change in colour, texture odour and firmness) were also observed for a period of 7 days (Aderiye et al., 2006). Each product was considered against a control (spontaneously-fermented) product. 3.15 Statistical Analysis Data obtained were subjected to statistical analysis. Duncan‟s Multiple Range Test was used to separate mean values and Analysis of variance (ANOVA) at 5% level of significance was used to determine differences. Data in tables were presented as mean ± standard deviation while figures were presented as mean ± error bars. 5 3 CHAPTER FOUR RESULTS 4.1 Physico-chemical analysis of spontaneously-fermenting cassava for fufu and usi production A rapid decrease in pH was first observed within the first 24 hours, after which the reduction was gradual till the end of the fermentation during the spontenous fermentations for both fufu and usi (Figure 4.1). The mean pH of the spontaneously fermented cassava showed decrease in pH values with increase in fermentation time for both products. The initial pH for fufu and usi mashes at zero hour was 7.1 ± 0.31. The values decreased, reaching its minimum 4.43 ± 0.31 and 4.27 ± 0.21, respectively at 72 hours. The pH values for fermenting usi mashes were slightly reduced than those for fufu. A progressive increase in the percentage titratable acidity was observed during fufu th fermentation while a decrease from the beginning of fermentation up till the 48 hour occurred during usi fermentation with a slight increase at 72 hours. The mean titratable acidity (%) of fermenting fufu ranged from 0.027 ± 0.01 to 0.393 ± 0.12 while those for usi was between 0.039 ± 0.01 and 0.084 ± 0.03 as illustrated in Figure 4.2. 4.2 Microbial load during spontaneous fufu and usi fermentation The mean microbial count on Plate Count Agar, De Mann Rogosa Sharpe agar, Violet Red Bile Glucose Agar and Malt Extract Agar for total bacteria, Lactic Acid Bacteria, coliforms and yeasts, respectively, were shown in Table 4.1. During usi 8 fermentation, the highest total count (7.50×10 CFU/mL) was observed at 24 hours 8 8 while a sharp decrease (1.76×10 CFU/mL and 0.13×10 CFU/mL) was at 48 and 72 hours, respectively. There was an increase in LAB count at the beginning of 8 8 fermentation up till 24 hours from 2.9×10 CFU/mL to 5.5.7×10 CFU/mL, but 8 8 decreased at 48 hours to 0.91×10 CFU/mL and further 0.07×10 CFU/mL at 72hours. 5 4 8 7 6 5 4 fufu usi 3 2 1 0 0 24 48 72 Time (Hours) Figure 4.1: pH of spontaneously-fermenting cassava for fufu and usi 5 5 pH 0.5 0.45 0.4 0.35 0.3 0.25 Fufu 0.2 Usi 0.15 0.1 0.05 0 0 24 48 72 Fermentation time (Hours) Figure 4.2: Titratable acidity (%) of spontaneously-fermenting cassava for fufu and usi 5 6 Titratable acidity (%) 8 Table 4.1: Microbial count (10 CFU/mL) during spontaneous fermentation of cassava for fufu and usi production 8 Time (Hours)/Products/ Microbial counts (10 CFU/mL) Fresh cassava 24 48 72 Sample F U F U F U F U TBC 6.17±5.28 4.10±1.57 4.43±2.47 7.50±1.04 5.29±4.69 1.76±2.81 3.44±4.37 0.13±0.04 LC 2.03±1.86 2.90±0.85 2.67±1.36 5.70±0.3 3.43±1.50 0.91±1.38 4.03±1.69 0.07±0.04 CC 1.56±1.29 1.22±0.33 1.37±0.48 1.19±0.53 1.09±0.98 0.39±0.61 0.16±1.18 0.06±0.02 YC 2.10±0.54 1.90±1.02 2.12±0.71 1.50±1.21 0.91±0.88 0.72±1.10 0.82±1.21 0.56±0.09 Values are mean ± SD, n=3; F (Fufu), U (Usi), TBC (Total bateria count), LC (LAB count), CC (Coliform count), YC (Yeast count). 5 7 8 Coliform count also decreased from the peak value (1.22×10 CFU/mL) at the beginning of 8 8 8 fermentation to 1.19×10 CFU/mL, 0.39×10 CFU/mL and 0.06×10 CFU/mL at 24, 48 and 72 hours respectively. Total bacteria count was maximal at the beginning of fermentation 8 8 (6.17×10 CFU/mL) and the least (3.44×10 CFU/mL) was at 72 hours during fufu fermentation. There was gradual increase in LAB count as fermentation progressed with 8 8 count ranging between 2.03×10 CFU/mL and 4.03×10 CFU/mL. Decrease in coliform count 8 8 (1.56×10 CFU/mL to 0.16×10 CFU/mL) was however observed with increase in fermentation time. In both fermentations, yeast count decreased till the end of the 8 8 fermentation ranging between 1.9×10 CFU/mL and 0.56×10 CFU/mL in usi while it was 8 8 between 2.10×10 CFU/mL and 0.82×10 CFU/mL in fufu. In general, the total bacterial count in usi increased within the first 24 hours, but decreased at 48 hours and further at 72 hours, whereas, a lower value at the beginning of th fermentation up till 24 hours which later increased at the 48 hour was observed in fufu. LAB count increased with increase in fermentation time in fufu while it increased up till 24 hours nd and then decreased gradually in usi. Coliform and yeast counts decreased till the 72 hour in both fermentations. 4.3 Isolation of Lactic Acid Bacteria from fermenting cassava mash Ninety-eight (98) Lactic Acid Bacteria isolates were obtained from the fermenting cassava mashes for both fufu and usi. The isolates were characterized and identified based their cultural, morphological, physiological and biochemical properties. The colonial morphology of the randomly selected isolates varied from small, medium to big colonies, shiny, creamy and whitish in colour. Most were concave while some were either flat or embedded on the agar mdium (Table 4.2). Biochemical characterization showed the isolates to be Gram positive colonies of medium, short and long rods while some are cocci in shape, catalase negative, non motile and do not hydrolyze starch and gelatin. Some produced ammonia from arginine and grew at 6.5% NaCl. Hydrogen sulphide was produced by all isolates, but they were negative to methyl red test. Varied sugar utilization pattern was observed by the organisms where glucose, sucrose, fructose, lactose and maltose were fermented by all isolates. Sorbose, inositol and starch were not utilized as shown in Table 4.3. 5 8 Table 4.2: Colonial morphology of randomly selected Lactic Acid Bacteria Isolate Elevation Colour Surface Edge Opacity Shape Size Number of code randomly selected isolates I Convex White Shiny Entire Opaque Circular Small 17 II Convex White Shiny Entire Opaque Circular Big 8 III Flat Cream Shiny Irregular Opaque Star like Medium 10 IV Embedded Cream Shiny Irregular Opaque Spindle Small 6 V Convex Cream Shiny Irregular Opaque Spindle Small 6 VI Embedded Cream Shiny Entire Opaque Circular Big 8 VII Convex Cream Shiny Entire Opaque Circular Medium 27 VIII Flat Cream Shiny Irregular Opaque Irregular Medium 10 IX Flat Cream Shiny Entire Opaque Circular Small 6 Key I - F1E, F1F, FIG, F1J, F1K, LFA, F2A, F2B, F2C, F2E, F2F, F3I, F3J, F3N, U1A, U1B, U1D II - LFF, U1C, U2I, LFK, U3A, U3G, U3J, U3P III - F3M, U1N, LFJ, U3C, U3H, LFB, F2J, F3B, F3L, U1M IV - U3P, LFE, F2K, U1P, U2J, U2P V - U2C, U2D, U2F, F1I, U1K, U2L VI – F1H, F1L, F2N, F3D, LFH, LFG, LFI, U1F VII - U2H, U2K, U2L, U3D, U3E, U3F, U3I, U3K, F1A, F1B, F1D, F2G, F2I, F3A, F3C, F3E, F3F, F3G, F3H, U1E, U1G, U1H, U1I, U1J, U1L, U2A, U2B VIII - LFC, F2D, F3K, U3O, U3B, U2E, U1O, U2M, U3N, U2L IX - F1C, F1J, LFD, F2H, F2L, F2M 59 Table 4.3: Biochemical characteristics of selected LAB isolates I + rod - - + + - - - + - - + + + + + + + + + + + + - - - - 49 L. plantarum II + cocci - - + - - - - + - - + + - - - - + - + + - - - - - 6 Leuconostoc. mesenteroides III + rod - - + + - - - + - - + + + + + - - + + - + + - - - - 12 L. acidilactici IV + rod - + + + - - - + - - + + + - + + + + + + + + - + - - 10 L fermentum V + rod - - + + - - - + - - + + + - + - - + - - - - - - - - 8 L. delbruekii VI + rod - - + + - - - + - - + + + + + + + + + - + + - + - - 11 L. brevis VII + cocci - + + - - - - + - - + + + - + + + + - + + - - + - - 2 L. lactis Key: + positive, - negative I - F1A, F1B, F1D,F1E, F1F, FIG,F1K, LFA, F2A, F2B, F2C, F2E, F2F, F2G, F2I, F3A, F3C, F3E, F3F, F3G, F3H,F3I, F3J, F3N, U1A, U1B, U1D, U1E, U1G, U1H, U1I, U1J, U1L, U2A, U2B, U2C, U2D, U2F, U2G, U2H, U2K, U2L,U2Q U3D, U3E, U3F, U3I, U3K, U3M. II – F1C, F1J, LFD, F2H, F2L, F2M III – F1H, F1L, F2N, F3D, LFH, F3M, U1N, LFJ, U3C, U3H, U3N,U3L IV – F1I, LFC, F2D, F3K, U3O, U3B, U2E, U1O, U2M, U1K V – LFB, F2J, F3B, F3L, U1M, U1P, U2J, U2P VI – LFE, F2K, LFF, U1C, U2I, LFK, U3A, U3G, U3J, U3P, U1F VII – LFG, LFI 60 Isolate code Grams rxn Shape Catalase NH3 from Arginine 4% NaCl 6.5%NaCl 8% NaCl Motility Starch hydr MR VP Gelatin H2S pdtn Glucose Maltose Mannitol Sucrose Melibiose Galactose Fructose Sorbitol Raffinose Lactose Xylose Starch Arabinose Inositol Sorbose No. of occurence Probable id The selected Lactic Acid Bacteria were identified as Lactobacillus plantarum (50.0%, 50.0%), Lactobacillus fermentum (8.0%, 12.5%), Lactobacillus brevis (6.0%, 16.7%), Leuconostoc mesenteroides (12.0%, 0.0%), Lactobacillus delbruekii (8.0%, 8.3%), Lactobacillus acidilactici (12.0%, 12.5%) and Lactobacillus lactis (4.0%, 0.0%) for fufu and usi, respectively (Table 4.4). L. plantarum had the highest percentage of occurrence in both fermentations hence selected for further screening. 4.4 Screening for potential starters among selected Lactic Acid Bacteria All the forty-eight (48) selected Lactobacillus plantarum strains did not hydrolyze starch, when grown on modified MRS agar but produced pectinase as well as linamarase in the form of β –glucosidase, a linamarin analogue when grown in a medium containing 4-nitrophenyl-B-D-glucopyranoside (Table 4.5). Rate of acid production in growth medium as monitored showed decrease in values with increasing incubation time, ranging from a starting pH of 6.50 to 3.58 after 72- hour incubation. The least pH at 24 hours was 4.62 by isolate F2B, 4.05 at 48 hours by U2C and 3.58 at 72 hours by isolate F2B (Table 4.6). Twenty (20) isolates that produced the lowest pH values after 72 hours incubation were selected for further screening (production of antimicrobial compounds). Production of lactic acid, hydrogen peroxide and diacetyl by the selected isolates was shown in Tables 4.7- 4.9 respectively. Lactic acid concentration produced ranged from 1.10 g/L by isolate U3K to 1.78 g/L by isolate U2C at 24 hours, 1.22 g/L by isolate F1B to 2.45 g/L by isolate F2A at 48 hours and 0.57 g/L and 2.55g/L by isolates F1F, U3K, respectively. The highest hydrogen peroxide concentration produced was 0.629 µg/L by isolate F2A at 24 hours while the least was 0.136 µg/L by isolate U1D at 72 hours. Least concentration (1.08 g/L) of diacetyl was the produced by both F2A and U1H at 24 hours while the highest was 2.86 g/L by F1B at 48 hours. 6 1 Table 4.4: Frequency of occurrence (%) of selected lactic acid bacteria during spontaneous cassava fermentation Probable identity Fufu Usi Freq. of occurrence % occurence Freq. of occurrence % occurence L. plantarum 25 50.0 24 50.0 L. fermentum 4 8.0 6 12.5 L. brevis 3 6.0 8 16.7 Leuc. 6 12.0 - - mesenteroides L. delbruekii 4 8.0 4 8.3 L. acidilactici 6 12.0 6 12.5 L. lactis 2 4.0 - - Total 50 100 48 100 6 2 Table 4.5: Starch hydrolysis, linamarase and pectinase enzyme production by selected isolates Isolates Starch hrdrolysis Linamarase Pectinase F1A - + + F1B - + + F1D - + + F1E - + + F1F - + + F1G - + + F1K - + + LFA - + + U1A - + + U1B - + + U1D - + + U1E - + + U1G - + + U1H - + + U1I - + + U1J - + + U1L - + + F2A - + + F2B - + + F2C - + + F2E - + + F2F - + + F2G - + + F2I - + + F2J - + + U2A - + + U2B - + + U2C - + + U2D - + + U2F - + + U2G - + + U2H - + + U2K - + + U2L - + + F3A - + + F3C - + + F3E - + + F3F - + + F3G - + + F3H - + + F3I - + + F3J - + + F3N - + + U3D - + + U3E - + + U3F - + + U3I - + + U3K - + + U3M - + + Key: - negative, + positive 6 3 Table 4.6: Acidification (pH) of growth medium by selected isolates Isolates Time (Hours)/ pH 24 48 72 k n F1A f 5.33 * 4.86 4.52 m y q F1B 5.26 4.53 4.43 l x p F1D 5.27 4.58 4.50 q zb w F1E 5.06 4.33 4.25 s s o F1F 5.04 4.68 4.51 e f i F1G 5.35 5.06 4.64 g g g F1K 5.32 4.98 4.71 j o r LFA 5.29 4.76 4.41 l n f U1A 5.27 4.79 4.73 h k n U1B 5.31 4.86 4.52 n zf za U1D 5.15 4.22 4.17 i d c U1E 5.30 5.10 4.82 d i k U1G 5.36 4.90 4.62 x zc w U1H 4.94 4.29 4.25 g g g U1I 5.32 4.98 4.71 k s l U1J 5.28 4.68 4.61 c b j U1L 5.41 5.18 4.63 zc zh zd F2A 4.75 4.06 4.01 zd zj ze F2B 4.62 3.66 3.58 z zc z F2C 4.89 4.29 4.18 y zb v F2E 4.93 4.33 4.27 d e F2F 5.36 5.09e 4.74 i d c F2G 5.30 5.1 4.82 h j h F2I 5.31 4.87 4.68 k s l F2J 5.28 4.68 4.61 za zg zc U2A 4.82 4.13 4.05 i d c U2B 5.30 5.10 4.82 zb zi zd U2C 4.76 4.05 4.01 l q l U2D 5.27 4.73 4.61 u ze v U2F 4.99 4.26 4.26 h l d U2G 5.31 4.85 4.76 k s l U2H 5.28 4.68 4.61 g h l U2K 5.32 4.92 4.61 j v s U2L 5.29 4.60 4.39 w zb y F3A 4.95 4.33 4.22 v zf z F3C 4.97 4.22 4.18 q u n F3E 5.06 4.64 4.52 i r h F3F 5.30 4.72 4.68 n zd x F3G 5.15 4.28 4.23 l p t F3H 5.27 4.74 4.31 m m F3I 5.32 4.82 4.59 a c b F3J 5.44 5.17 4.93 t w t F3N 5.00 4.59 4.31 o ze zb U3D 5.12 4.26 4.09 j t m U3E 5.29 4.65 4.59 r z v U3F 5.05 4.42 4.26 b a a U3I 5.42 5.31 5.10 p za x U3K 5.07 4.35 4.23 m g U3M 5.30i 4.82 4.71 *The means reported with the same superscript in each column indicated no significant difference (p≤0.05). Bold values: Least pH values selected for further screening 6 4 Table 4.7: Quantity of lactic acid produced in g/L by selected strains of Lactic Acid Bacteria Time (Hours) Isolates Lactic acid (g/L) 24 48 72 a m F2A m1.24 * 2.45 0.74 hij i g F2B 1.48 1.69 1.13 b j h F2C 1.71 1.62 1.03 e h i F2E 1.57 1.81 0.98 ghi b f U2A 1.50 2.34 1.23 a c l U2C 1.78 2.18 0.78 c j k U2F 1.64 1.64 0.83 e n j F1B 1.57 1.22 0.94 efg k m F1D 1.54 1.47 0.74 l d l F1E 1.33 2.10 0.79 l f o F1F 1.30 1.93 0.57 d k m U1D 1.59 1.47 0.72 jk k g U1H 1.45 1.48 1.15 ij g e F3C 1.46 1.86 2.11 n m n F3A 1.18 1.26 0.69 j l c F3E 1.41 1.37 2.39 de g b U3D 1.58 1.88 2.46 o e d U3F 1.10 1.99 2.25 fgh g U3J 1.51 1.86 2.55a def d g F3G 1.55 2.07 1.14 *The means reported with the same superscript in each column indicated no significant difference (p≤0.05) 6 5 Table 4.8: Quantity of hydrogen peroxide produced in µg/L by selected strains of Lactic Acid Bacteria Time (Hours) Isolates Hydrogen peroxide (µg/L) 24 48 72 F2A a a c 0.629 * 0.340 0.204 b e b F2B 0.442 0.272 0.221 j b b F2C 0.238 0.323 0.221 l i e F2E 0.187 0.204 0.170 i f d U2A 0.272 0.255 0.187 g c d U2C 0.323 0.306 0.187 h h c U2F 0.306 0.221 0.204 f d c F1B 0.340 0.289 0.204 e g b F1D 0.357 0.238 0.221 c i c F1E 0.391 0.204 0.204 h i a F1F 0.306 0.204 0.374 g g g U1D 0.323 0.238 0.136 d i d U1H 0.374 0.204 0.187 k b f F3C 0.221 0.323 0.153 c f c F3A 0.391 0.255 0.204 j i c F3E 0.238 0.204 0.204 d i c U3D 0.374 0.204 0.204 e j b U3F 0.357 0.187 0.221 d i d U3J 0.374 0.204 0.187 g d c F3G 0.323 0.289 0.204 *The means reported with the same superscript in each column indicated no significant difference (p≤0.05) 6 6 Table 4.9: Quantity of diacetyl produced in g/L by selected strains of Lactic Acid Bacteria Time (Hours) Isolates Diacetyl (g/L) 24 48 72 i i F2A j1.08 * 1.91 1.72 f d cd F2B 1.72 2.35 2.18 je c b F2C 1.91 2.51 2.31 d a a F2E 2.00 2.82 2.51 c ef e U2A 2.11 2.25 2.10 i g fgh U2C 1.51 2.10 1.98 j hi i U2F 1.09 1.97 1.77 a a a F1B 2.33 2.86 2.52 d g h F1D 1.98 2.10 1.90 h h i F1E 1.56 2.00 1.72 e de de F1F 1.91 2.32 2.13 c b c U1D 2.10 2.61 2.23 j f fg U1H 1.08 2.22 1.96 g hi i F3C 1.62 1.98 1.72 ki hi i F3A 1.52 1.98 1.77 b c b F3E 2.21 2.51 2.32 d g f U3D 1.99 2.13 2.01 a b b U3F 2.33 2.64 2.38 h h i U3J 1.56 1.99 1.70 f g gh F3G 1.73 2.13 1.93 *The means reported with the same superscript in each column indicated no significant difference (p≤0.05) 6 7 Most of the isolates inhibited the growth of test pathogens by showing zones of inhibition around the colonies. Only a few showed no inhibition zones at all, most especially against Corynebaterium specie (Table 4.10). All the isolates inhibited the growth of E. coli with zones ranging from 5 to 7 mm. All except F3E and U3K inhibited the growth of Salmonella with inhibition zones between 4 and 9 mm. The growth of Bacillus cereus was not inhibited by F1D and while only F1F showed no inhibition against Shigella sp. The best five (5) overall producers after being analyzed statistically, using Duncan Multiple Range Test at 0.05% level of probability, across the three compounds were isolates F2A, F2B, F2C, U2A and U2C. They were thus selected as the potential starters and identified genotypically. 4.5 Molecular identification of screened potential starters. The result of the amplified nucleotide sequences (Appendix IX) of the five (5) selected isolates was shown in Table 4.11. On the basis of the database information available on National Centre for Biotechnology Information (NCBI) site using the Basic Local Alignment Search Tool (BLAST), the isolates were classified and identified using the highest percentage similarity with organism of the nearest homology. All the isolates belong to the family Lactobacillaceae and genus Lactobacillus. Isolate F2A was identified as Lactobacillus pentosus F2A with accerssion number KJ778115, having showed 99% homology alignment with Lactobacillus pentosus strain 405 in the GenBank. F2B was L. plantarum subsp. argentolarensis F2B (KJ778116) with 99% nucleotide homology with L. plantarum subsp. argentolarensis strain Ni1031 while F2C was L. plantarum F2C (KJ77117) showing 99% homology with L. plantarum strain 097. 100% similarity was observed between isolate U2A, which was L. plantarum U2A (KJ78118) and L. plantarum P2 whereas U2C was 99% homologous with L. paraplantarum DSM 10667 nucleotide sequence in the NCBI Genbank. 6 8 Table 4.10: Antagonistic effect of selected isolates against test pathogenic organisms. Test pathogenic organisms Zone of inhibition (mm) Isolates E. coli Proteus Salmonella B. cereus Shigella Corynebacterium sp. sp. sp. sp. F2A 6 2 6 4 12 NI F2B 6 2 8 4 9 NI F2C 6 3 7 3 11 NI F2E 7 5 9 3 10 NI U2A 9 NI 7 6 10 NI U2C 6 2 7 2 9 7 U2F 6 NI 8 6 8 3 F1B 6 NI 7 3 4 6 F1D 6 2 4 NI 9 4 F1E 7 2 9 4 10 8 F1F 5 2 5 4 NI NI U1D 6 3 9 3 10 3 U1H 6 2 8 5 11 3 F3C 5 6 8 3 10 NI F3A 7 2 7 6 7 NI F3E 6 NI NI 5 9 NI U3D 5 8 8 6 10 3 U3F 5 5 5 8 9 NI U3J 6 NI NI 3 10 3 F3G 7 3 5 2 9 2 NI - No Inhibition. 6 9 Table 4.11: Molecular identification of selected potential starters Isolate Closely related Percentage Base pair Identification Accerssion code species/GenBank similarity analyzed number Accerssion number (%) F2A Lactobacillus 99 517 Lactobacillus KJ778115 pentosus 405 pentosus (AB775188.1) F2B L. plantarum subsp. 99 496 L. plantarum KJ778116 argentolarensis subsp. Ni1031(AB598953.1) argentolarensis F2C Lactobacillus 99 521 Lactobacillus KJ778117 plantarum 097 plantarum (JN560914.1) U2A Lactobacillus 100 500 Lactobacillus KJ778118 plantarum P2 plantarum (EU167523.1) U2C Lactobacillus 99 520 Lactobacillus KJ778119 paraplantarum paraplantarum DSM10667 (NR117813.1) 7 0 Phylogenetic tree constructed using the Molecular Evolution Genetics Analysis (MEGA) version 6 established the relationship among the organisms and their nearest homologies (Figure 4.3). The neighbour joining method revealed the clustering of the identified isolates into four clusters of closely related strains. 4.6 Fermentation of cassava with identified potential starters The genotypically identified potential starters were utilised singly and in- combination (Table 4.12) to ferment cassava for fufu and usi production (Plate 4.1). 9 Inoculum size of approximately 1.2×10 CFU/mL as earlier described was used for each organism and 5 mL of single and randomly combined starters were used as the final inoculum. 4.6.1 pH of starter-fermented cassava mash Decrease in pH values with increasing fermentation time was observed in both fufu and usi fermentations with values ranging from initial pH of 7.10 to 3.68 in fufu and 3.53 in usi. The least pH value after 72-hour fermentation in fufu was 3.68 by the combined starter CGI (Lactobacillus plantarum F2C/L. plantarum U2A/L. plantarum U2C) while usi had 3.53 by the same starter combination. The un-inoculated control however had the highest pH values throughout the fermentation processes with a minimum of 4.12 (fufu) and 4.05 (usi) at 72 hours (Table 4.13). 4.6.2 Total titratable acidity (%) of starter-fermented cassava mash The highest percentage total titratable acidity in the form of lactic acid produced at 24 hours during fufu fermentation was 0.77% by the combined starter AB (L. pentosus F2A/L. plantarum subsp. argentolarensis F2B). Reduction (0.41%) in the maximal quantity was observed at 48 hours while a slight increase (0.45%) in value was produced at 72 hours by L. plantarum F2C. During usi fermentation, the maximal total titratable acidity reduced with increase in fermentation time. Combined starter CG (L. plantarum F2C/ L. plantarum U2A) produced the highest titratable acidity (0.279%) at 24 hours whereas, the value decreased to 0.025% at 48 hours with a much lower quantity (0.002%) at 72 hours by the same starter. It was however noted that the observed highest values in both fermentations were produced by different starter cultures (Table 4.14). 7 1 11 Lactobacillus pentosus 405 0.0000 5 0.0000 Lactobacillus plantarum E7304 0.0000 10 0.0000 Lactobacillus plantarum P2 0.0031 0.00006 Lactobacillus pentosus124-2 0.0000 0.0000 66 Lactobacillus paraplantarum DSM 10667 76 0.0000 0.0434 Lactobacillus plantarum TW57-4 90 0.0932 0.0136 Lactobacillus plantarum KLDS 1.0725 0.4341 0.0902 46 0.0000 Lactobacillus plantarum subsp argentolarensis F2B (KJ778116) 0.1927 Lactobacillus plantarum F2C (KJ778117) 97 0.0000 0.4544 16 0.0000 Lactobacillus paraplantarum U2C (KJ778119) 0.0000 24 Lactobacillus pentosus F2A (KJ778115) 0.0537 0.0000 0.0000 0.0000 Lactobacillus plantarum U2A (KJ778118) Lactobacillus plantarum 097 95 0.0000 0.3266 18 Lactobacillus plantarum subsp. argentoratensis Ni1031 0.0000 0.0000 20 Lactobacillus plantarum PM411 0.0000 0.0000 Lactobacillus plantarum C21-41 0.0000 Lactobacillus plantarum AF1 0.8638 Figure 4.3: Phylogenetic relationship between identified starters and closely related organisms in the GenBank. 7 2 Table 4.12: Single and randomly combined LAB isolates used as potential starters for fufu and usi production. Codes Isolates combinations A Lactobacillus pentosus F2A B Lactobacillus plantarum susp. argentolarensis F2B C Lactobacillus plantarum F2C G Lactobacillus plantarum U2A I Lactobacilluc paraplantarum U2C AB L. pentosusF2A + L. plantarum subsp.argentolarensis F2B ABC L. pentosus F2A + L. plantarum subsp.argentolarensis F2B +L. plantarum F2C ABCG L. pentosus F2A + L. plantarum subsp.argentolarensis F2B +L. plantarum F2C +L. plantarum U2A ABCGI L. pentosus F2A + L. plantarum subsp.argentolarensis F2B +L. plantarum F2C +L. plantarum U2A +L. paraplantarum U2C BC Lactobacillus plantarum susp. argentolarensis F2B + L. plantarum F2C BCG L. plantarum subsp.argentolarensis F2B +L. plantarum F2C +L. plantarum U2A BCGI L. plantarum subsp.argentolarensis F2B +L. plantarum F2C +L. plantarum U2A+ L. paraplantarum U2C CG L. plantarum F2C +L. plantarum U2A CGI L. plantarum F2C +L. plantarum U2A +L. paraplantarum U2C GI L. plantarum U2A+L. paraplantarum U2C Control Un-inoculated 7 3 X Y Z Plate 4.1: Cassava fermentation with single and combined starter cultures using plastic bioreactors X – Main improvised bioreactor Y – Cassava samples Z – Fermenting liquor collection outlet 7 4 Table 4.13: pH during starter fermentation of cassava for fufu and usi production Time (Hour)/Cassava product/ pH Potential starter 24 48 72 F U F U F U c* d m i k j A 4.23 4.22 3.78 3.76 3.68 3.75 g e e e d e B 4.17 4.14 3.95 3.9 3.81 3.88 c k f i g j C 4.18 3.95 3.93 3.76 3.77 3.75 d c e c e c G 4.21 4.28 3.95 4.04 3.79 3.99 k k d j b k I 4.11 3.95 3.96 3.74 3.95 3.73 e h f h f AB 4.20 4.05 3.93 3.77 3.78 3.76i b g b l c n ABC 4.28 4.08 4.05 3.66 3.82 3.64 g a l a d b ABCG 4.17 5.16 3.82 4.33 3.81 4.03 g j j g f h ABCGI 4.17 3.99 3.87 3.78 3.78 3.78 m i g j g l BC 4.01 4.02 3.91 3.74 3.77 3.72 j e h e i f BCG 4.12 4.14 3.89 3.9 3.74 3.86 h g d m g o BCGI 4.15 4.08 3.96 3.53 3.77 3.53 l f k d i d CG 4.10 4.1 3.83 3.94 3.74 3.91 k e i k j m CGI 4.11 4.14 3.88 3.72 3.71 3.7 i f c f h g GI 4.13 4.1 3.98 3.87 3.75 3.81 a b a b a a Un-inoculated 4.75 4.93 4.25 4.2 4.12 4.05 A - Lactobacillus pentosus F2A, B – L. plantarum subsp. argentolarensis F2B, C - L. plantarum F2C, G – L. plantarum U2A, I – L. paraplantarum F- Fufu, U- Usi.*The means reported with the same superscript in each column indicated no significant difference (p≤0.05). 75 Table 4.14: Total titratable acidity (%) during starter fermentation of cassava for fufu and usi production Potential starter Time (Hour)/Cassava product/Total titratable acidity 24 48 72 F U F U F U A c* d b d f d 0.549 0.252 0.360 0.023 0.252 0.0020 g f c f f f B 0.342 0.198 0.198 0.018 0.252 0.0016 b g i g a g C 0.594 0.189 0.108 0.017 0.459 0.0015 j h g h j h G 0.270 0.180 0.135 0.016 0.144 0.00145 i e h e l e I 0.306 0.225 0.117 0.020 0.117 0.00182 a i n j j i AB 0.774 0.153 0.054 0.014 0.144 0.00123 d j k g j ABC 0.522 0.144 0.099j 0.013 0.243 0.00117 e b g b k b ABCG 0.504 0.270 0.135 0.024 0.126 0.00219 h i k j i i ABCGI 0.315 0.153 0.090 0.014 0.180 0.00124 g f e f c f BC 0.342 0.198 0.153 0.018 0.324 0.0016 l h l i f h BCG 0.216 0.180 0.072 0.016 0.252 0.00145 m k l l e k BCGI 0.189 0.135 0.072 0.012 0.261 0.0011 n a m a b a CG 0.153 0.279 0.063 0.025 0.342 0.0023 k g f g d g CGI 0.252 0.189 0.144 0.017 0.315 0.0015 c g a g l g GI 0.549 0.189 0.414 0.017 0.108 0.0015 f c d c h c Un-inoculated 0.387 0.261 0.162 0.023 0.189 0.0021 A- Lactobacillus pentosus F2A, B – L. plantarum subsp. argentolarensis F2B, C - L. plantarum F2C, G – L. plantarum U2A, I – L. paraplantarum F- Fufu, U- Usi. *The means reported with the same superscript in each column indicated no significant difference (p≤0.05). 76 4.6.3 Proximate analysis of starter-fermented cassava for fufu and usi production The proximate analysis of the fresh cassava indicated moisture content to be 7.28%, crude protein 1.02%, crude fat 0.48%, crude fibre 1.75%, ash 1.57% and the total carbohydrate 89.65%. Table 4.15 showed the proximate analysis of fufu after 72-hour fermentation. The moisture content ranged between 5.10% by the combined starter CGI (L. plantarum F2C/L. plantarum U2A/L. paraplantarum U2C) and 8.61% by starter C (L. plantarum F2C). Only two starters (L. plantarum F2C and L. paraplantarum U2C) showed higher moisture content (8.61% and 7.72%) than the un-inoculated experiment (control) which had 7.28%. Protein content ranged between 0.73% and 1.34%. The combined starter CGI had the highest protein content of 1.34% even though combined starter ABC and ABCGI also had higher protein content (1.24% and 1.16%) than the fresh cassava (1.02%). L. plantarun U2A had the least crude fat content (0.24%) from an initial 0.48%, although, some starters showed higher fat content (0.52 - 0.91%) than the fresh cassava after fermentation. L. paraplantarum U2C recorded the highest crude fibre (3.28%) and ash content (2.85%) while the total carbohydrate was maximal at 91.33% by the combined starter GI (L. plantarum U2A/L. paraplantarum U2C). As shown in Table 4.16, the proximate composition of usi after fermentation indicated that moisture content was between 5.34% and 9.38% by combined starter GI (L. plantarum U2A/ L. paraplantarum U2C) and CG (L. plantarum F2C/L. plantarum U2A) respectively, even though, starters Lactobacillus plantarum F2A, combined starter CGI (Lactobacillus plantarum F2C/L. plantarum U2A/L. paraplantarum U2C) and the un-inoculated batch had values that were not significantly different from the lowest moisture content (5.34%). Increased protein contents ranging from 1.14% to 1.82% was observed in most samples after fermentation. The highest (1.82%) however, was by the combined starter CGI (Lactobacillus plantarum F2C/L. plantarum U2A/L. paraplantarum U2C). Meanwhile, reduction in protein content was also observed in the un-inoculated sample as well as when combined starter GI (L. plantarum U2A/L. paraplantarum U2C) and ABCGI were used. All the starter- fermented samples had reduced fat content except L. paraplantarum U2C which had 2.06% and the least (0.04%) observed when combined starter BC was used. 7 7 Table 4.15: Proximate composition (%) of starter fermented cassava for fufu production after 72 hour fermentation Proximate composition (%) Potential Moisture Crude Crude fat Crude fibre Ash Total starters content protein carbohydrate† i f b i j c A 5.51±0.06 * 0.91±0.05 0.83±0.04 1.73±0.04 1.19±0.01 89.83±0.06 c g e j c l B 7.23±0.04 0.89±0.02 0.52±0.02 1.58±0.03 1.68±0.03 88.10±0.08 a d h fgh f m C 8.61±0.05 1.02±0.12 0.45±0.02 1.83±0.03 1.57±0.04 86.52±0.07 g e k d h b G 5.63±0.06 0.98±0.11 0.24±0.02 2.00±0.02 1.29±0.01 89.86±0.14 b g j a a n I 7.72±0.04 0.89±0.02 0.29±0.02 3.28±0.03 2.85±0.07 84.97±0.09 k d gh c j c AB 5.38±0.04 1.02±0.12 0.42±0.02 2.15±0.01 1.19±0.02 89.84±0.10 f b d efg h g ABC 6.18±0.03 1.24±0.13 0.39±0.01 1.86±0.06 1.29±0.02 89.04±0.14 d f e c g l ABCG 6.70±0.04 0.91±0.05 0.58±0.02 2.18±0.03 1.48±0.02 88.15±0.06 l c c b b i ABCGI 5.16±0.02 1.16±0.13 0.69±0.02 2.41±0.03 1.79±0.03 88.79±0.13 e d c i g j BC 6.58±0.06 1.02±0.12 0.69±0.03 1.69±0.03 1.48±0.03 88.54±0.15 d d ef hi i h BCG 6.70±0.03 1.02±0.12 0.54±0.05 1.70±0.02 1.24±0.06 88.80±0.21 g g ef k k f BCGI 5.62±0.05 0.89±0.02 0.55±0.05 1.40±0.03 1.14±0.03 89.10±0.07 h h a hi e e CG 5.6±0.044 0.81±0.13 0.91±0.04 1.70±0.02 1.61±0.03 89.37±0.11 n a i def d d CGI 5.10±0.09 1.34±0.05 0.33±0.06 1.91±0.04 1.67±0.02 89.65±0.15 m i jk ghi l a GI 5.11±0.01 0.73±0.13 0.28±0.01 1.77±0.03 0.78±0.06 91.33±0.14 j f a de b f Uninoculated 5.48±0.04 0.91±0.05 0.90±0.04 1.90±0.02 1.78±0.02 89.03±0.12 Fresh cassava 7.28±0.03 1.02±0.12 0.48±0.01 1.75±0.03 1.57±0.04 87.90±0.11 Values are means ±SD, n=3 A - Lactobacillus pentosus F2A, B – L. plantarum subsp.argentolarensis F2B, C - L. plantarum F2C, G – L. plantarum U2A, I – L. paraplantarum U2C. * The means reported with the same superscript in each column indicated no significant difference (p≤0.05). † By difference 7 8 Table 4.16: Proximate composition (%) of starter fermented cassava for usi production after 72-hour fermentation Proximate composition (%) Potential Moisture Crude Crude fat Crude fibre Ash Total starters content protein carbohydrate† c h c bcdef bc d A 6.18±2.25 * 1.02±0.12 0.31±0.02 1.93±0.05 0.93±0.02 89.63±0.18 abc h d bcdef c g B 7.32±0.06 1.02±0.12 0.20±0.01 1.94±0.05 0.89±0.02 88.63±0.16 abc f d bcdef def i C 7.35±0.02 1.24±0.13 0.29±0.02 1.96±0.02 0.78±0.03 88.37±0.13 abc b b ef ef j G 7.36±0.10 1.68±0.13 0.42±0.02 1.78±0.02 0.77±0.03 87.99±0.13 bc c a def de k I 6.56±0.06 1.38±0.13 2.06±0.03 1.81±0.02 0.81±0.02 87.38±0.09 bc c de f de f AB 6.29±0.02 1.38±0.13 0.16±0.01 1.58±0.06 0.79±0.03 88.80±0.16 bc f bc abc a i ABC 6.79±0.03 1.24±0.13 0.39±0.02 2.24±0.02 1.03±0.01 88.31±1.28 bc d d a bc h ABCG 6.64±0.06 1.34±0.05 0.29±0.02 2.35±0.04 0.93±0.03 88.45±0.06 bc i f abcde f c ABCGI 6.45±0.03 0.95±0.12 0.06±0.01 2.14±0.02 0.74±0.05 89.66±0.1 bc b f ab f f BC 6.34±0.55 1.68±0.13 0.04±0.01 2.31±0.03 0.74±0.05 88.89±0.08 d e e abcde b e BCG 5.39±0.03 1.28±0.05 0.14±0.01 2.14±0.02 0.90±0.04 89.55±0.04 ab f bc cdef d l BCGI 8.41±0.06 1.24±0.13 0.38±0.03 1.89±0.02 0.82±0.02 87.26±0.15 a g d abc a m CG 9.38±0.03 1.14±0.08 0.27±0.03 2.21±0.02 1.01±0.01 85.99±0.05 c a de a g d CGI 5.36±0.04 1.82±0.13 0.16±0.03 2.35±0.05 0.69±0.02 89.62±0.08 c j d abcd bc b GI 5.37±0.07 0.85±0.05 0.28±0.03 2.17±0.03 0.93±0.06 90.40±0.15 c i d cdef h a Uninoculated 5.68±0.42 0.95±0.12 0.29±0.02 1.91±0.03 0.57±0.03 90.60±0.52 Fresh cassava 7.28±0.03 1.02±0.12 0.48±0.01 1.75±0.03 1.57±0.04 87.90±0.11 Values are means ±SD, n=3 A - Lactobacillus pentosus F2A, B – L. plantarum subsp.argentolarensis F2B, C - L. plantarum F2C, G - L. plantarum U2A, I - L. paraplantarum U2C. * The means reported with the same superscript in each column indicated no significant difference (p≤0.05). † By difference 7 9 Fibre content increased after fermentation ranging between 1.78% and 2.35% from an initial 1.75%, except for the combined starter AB (L. pentosus F2A/L.plantarum subsp. argentolarensis F2B) which had reduced fibre content (1.58%). All the starters yielded samples with reduced ash content after fermentation when compared to the fresh cassava with values ranging between 0.51% and 1.03%. Nonetheless, the highest ash content (1.03%) was observed with the combined starter ABC (L. pentosus F2A/L. plantarum subsp. argentolarensis F2B/L.plantarum F2C). Total carbohydrate ranged from 85.99% to 90.6%. The highest value (90.6%) after fermentation was observed in the spontaneously fermented sample while the least (85.99%) was by the combined starter CG (L. plantarum F2C/L. plantarum U2A). 4.6.4 Anti-nutritional factors of starter-fermented cassava for fufu and usi The anti-nutritional factors of the fresh and starter-fermented cassava for fufu and usi were shown in Table 4.17. The fresh cassava tuber had 0.30 mg of phytic acid, 35.40 mg tannin and 0.10 mg cyanide per gram of the tuber. After 72-hour fufu fermentation, the phytic acid ranged between 0.1 mg/g to 0.5 mg/g with a minimum value of 0.1 mg/g by numerous starter combinations (A, AB, ABCG, ABCGI, CGI and CG). It was however, observed that starter combination ABC (L. pentosus F2A/L. plantarum subsp. argentolarensis F2B/L.plantarum F2C) and G (L. plantarum U2A) had higher phytic acid values (0.4 mg/g and 0.5 mg/g) than the fresh cassava. Tannin ranged between 34 mg/g and 62.7 mg/g with the least value of 34 mg/g by the combined starter CGI (L. plantarum F2C/L. plantarum U2A/L. paraplantarum U2C) while all other starter-fermented samples yielded tannin content that was higher than that of the fresh cassava. Cyanide content was not detected in samples fermented with starters B (L. plantarum subsp. argentolarensis F2B), BC (L. plantarum subsp. argentolarensis F2B/L.plantarum F2C), CG (L. plantarum F2C/L. plantarum U2A) and BCGI. 8 0 Table 4.17: Anti-nutritional factors (mg/g) of fresh and starter (CGI)-fermented cassava for fufu and usi production Potential starters Cassava Product/ Anti-nutritional factors (mg/g) Phytate Tannin Cyanide Fufu Usi Fufu Usi Fufu Usi A defg0.1±0.00 * a g b b c0.32±0.003 45.7±0.35 46.4±0.04 0.08±0.004 0.2±0.01 cd a f e c d B 0.2±0.01 0.32±0.003 45.9±0.19 35.5±0.03 0.00 0.1±0.00 defg bc i a a b C 0.2±0.01 0.28±0.00 42.1±0.02 64.0±0.02 0.1±0.00 0.6±0.03 a bc k c b a G 0.5±0.01 0.28±0.00 37.5±0.13 40.6±0.02 0.08±0.004 0.8±0.02 defg c l i bc d I 0.2±0.00 0.277±0.0004 37.3±0.13 33.8±0.02 0.05±0.00 0.1±0.01 fg bc h k a e AB 0.1±0.00 0.28±0.001 43.2±0.13 33.0±0.01 0.1±0.00 0.00 ab c b g bc e ABC 0.4±0.02 0.277±0.0004 61.8±0.06 34.6±0.02 0.05±0.00 0.00 efg bc e l bc d ABCG 0.1±0.00 0.28±0.00 46.3±0.16 32.3±0.01 0.05±0.00 0.1±0.01 efg bc c e b e ABCGI 0.1±0.00 0.28±0.00 55.5±0.06 35.5±0.03 0.08±0.004 0.00 def c a i c e BC 0.2±0.01 0.275±0.00 62.7±0.07 33.7±0.01 0.00 0.00 de c d e b e BCG 0.2±0.01 0.275±0.00 43.7±0.39 37.5±0.02 0.08±0.004 0.00 cd c m f c e BCGI 0.2±0.01 0.275±0.00 36.9±0.09 35.3±0.02 0.00 0.00 fg b n d c e CG 0.1±0.00 0.289±0.0002 35.9±0.26 39.0±0.02 0.00 0.00 g c o l bc e CGI 0.1±0.00 0.277±0.0004 34.0±0.28 32.3±0.01 0.05±0.00 0.00 defg c j h bc e GI 0.2±0.01 0.277±0.0004 39.9±0.17 34.4±0.03 0.05±0.00 0.00 bc b g j bc e Un-inoculated 0.3±0.01 0.29±0.001 45.7±0.01 33.4±0.01 0.05±0.00 0.00 Fresh cassava 0.30±0.004 35.40±0.03 0.10±0.01 A - Lactobacillus pentosus F2A, B – L. plantarum subsp. argentolarensis F2B, C- L. plantarum F2C, G – L. plantarum U2A, I – L. paraplantarum U2C. Values are means ±SD, n=3; * Means reported with the same superscript in each column indicated no significant difference (p≤0.05). 81 In usi, cyanide was not detected in most of the samples fermented with starters where as, phytic acid value was within the range of 0.275 mg/g and 0.32 mg/g with starters BC (L. plantarum subsp. argentolarensis F2B/L.plantarum F2C), BCG (L. plantarum subsp. argentolarensis F2B/L.plantarum F2C/L. plantarum U2A) and BCGI (L. plantarum subsp. argentolarensis F2B/L.plantarum F2C/L. plantarum U2A/L. paraplantarum U2C), having the least phytic acid values. The tannin content after fermentation ranged between 32.3 mg/g and 64 mg/g. The combined starter CGI had the least (32.3 mg/g) tannin. 4.6.5 Selection of a common starter culture for both products The starter combination that had the most reduced anti-nutritional factor, improved proximate composition and a faster acidification was considered for further study. Five starters with the best results for the above mentioned parameters were considered, after which the most common was selected. For usi, starters BC, BCG and BCGI had the least phytate (0.275 mg/g), followed closely by I, ABC, CGI and GI having 0.277 mg/g. It was however observed that the values were not significantly different at 5% level of probability. The CGI and ABCG starter fermented samples had the least tannin (32.30 mg/g). Absence of cyanide compound was observed in majority of the starter fermented samples. For fufu, the least phytate (0.10 mg/g) and tannin (34.0 mg/g) was produced by the combined starter CGI while cyanide compound was not detected in starters C, BC, BCGI and CG. The combined starter CGI (Lactobacillus plantarum F2C/Lactobacillus plantarum U2A/Lactobacillus paraplantarum U2C) was the most common starter to reduce phytate, tannin and cyanide in both fufu and usi even though there are some other starters that reduced the anti-nutritional factors, but were not common among the three factors (Table 4.18). Lactobacillus petosus F2A and the combined starter BCGI had the least pH values of 3.68 and 3.53 for fufu and usi, respectively after the fermentation process. Since the least pH values were produced by different starters, the next reduced values (3.71 and 3.7) were considered and these were from samples fermented with starter CGI. Increased protein content and lower moisture content were crucial factors in fermented foods. The least moisture content (5.10% and 5.36%) and highest protein (1.34% and 1.88%) recorded during starter fermentation of fufu and usi respectively were observed when the combined starter CGI was used (Table 4.19). 8 2 Table 4.18: Selection a common starter among the most reduced pH values and anti-nutritional factors after the 72-hour cassava fermentation for fufu and usi production pH Phytate Tannin Cyanide Fufu Usi Fufu Usi Fufu Usi Fufu Usi A BCGI CGI BC CGI CGI B BCG CGI* ABC CG BCG - ABCG BC BCGI CG CGI AB BCGI - AB BCGI CG BCG BC ABCG CGI - Control CG CGI GI I A CG - BC CGI GI *Starters in bold are the most common among the parameters analysed and thus, selected for further work. A - Lactobacillus pentosus F2A, B – L. plantarum subsp.argentolarensis F2B, C - L. plantarum F2C, G - L. plantarum U2A, I - L. paraplantarum U2C. 8 3 Table 4.19: Selection of a common starter among the most improved proximate composition after the 72-hour cassava fermentation for fufu and usi production Moisture content Crude protein Crude fat Ash content Carbohydrate Fufu Usi Fufu Usi Fufu Usi Fufu Usi Fufu Usi CGI* CGI CGI CGI G ABCGI I CG GI Control GI GI ABC G GI BC ABCGI BCG G GI ABCGI Control G BC I BCG B GI A ABCGI AB A ABCGI I CGI CGI CGI A AB CGI Control BCG A AB ABC AB CG B CGI A *Starters in bold are the most common among the parameters analysed and thus, selected for further work. A - Lactobacillus pentosus F2A, B – L. plantarum subsp.argentolarensis F2B, C - L. plantarum F2C, G - L. plantarum U2A, I - L. paraplantarum U2C. 84 In general, starter combination of CGI, comprising of Lactobacillus plantarum F2C, Lactobacillus plantarum U2A and Lactobacillus paraplantarum U2C was the most frequent and common to both product and thus, selected for further experiment. 4.6.6 Enzyme activities during starter fermentation for fufu and usi production The assay for amylase, pectinase and linamarase during the fermentation indicated their production and activities. The selected starter combination (Lactobacillus plantarum F2C/Lactobacillus plantarum U2A/Lactobacillus paraplantarum U2C) showed decrease in amylase activity with increase in fermentation time in both fermentations. Starter-fermented fufu mash had highest activity (10.1 U/mL) at 24 hours, 5.18 U/mL and 3.93 U/mL at 48 and 72 respectively, whereas, activity in usi ranged between 4.37 U/mL and 2.17 U/mL. In fufu, the un- inoculated experiment (control) had reduced activities than the starter-fermented batch, except at 72 hours, while amylase activities (1.04 U/mL, 0.76U/mL) were detected at 48 and 72 hours, respectively in usi (Figure 4.4). Pectinase activity during fufu fermentation decreased with increase in fermentation time in both the starter and un-inoculated fermentations. The activities ranged from 4.44 U/mL at 24 hours, to 3.81 U/mL at 48 hours and 1.74 U/mL at 72 hours. The control experiment had a higher activity (6.32 U/mL) than the starter-fermented batch at 24 hours after which the activities were lower than the latter throughout the fermentation. In usi, activity was detected at 24 hours (0.88 U/mL) and 72 hours (1.01 U/mL), but not at 48 hours (Figure 4.5). Starter-fermented experiment had higher linamarase activities than the control in both products. Fufu fermentation had 0.17 U/mL linamarase activity at 24 hours, increased gradually to 0.83 U/mL at 48 hours and had a sharp decrease to 0.39 U/mL at 72 hours while usi had a slight decrease from 0.38 U/mL at 24 hours to 0.34 U/mL at 48 hours and subsequently increased to 0.71 U/mL at 72 hours (Figure 4.6). However, it was generally observed that the un-inoculated (control) fermentation for th usi had none of the enzyme activities at the 24 hour and most of the enzyme activities in the starter fermented experiments were higher than the in the un-inoculated experiments. 8 5 12 10 8 6 24hours 48hours 4 72hours 2 0 SFF CF SFU CU Fufu Usi Figure 4.4: Amylase activities in starter (CGI)-fermented and un-inoculated cassava mash during fufu and usi fermentation Key: SFF- Starter-fermented fufu; CF- Spontaneously-fermented fufu (control); SFU- Starter- fermented usi; CU- Spontaneously-fermented usi (control) 8 6 Amylase avtivity (U/mL) 7 6 5 4 24hours 3 48hours 72hours 2 1 0 SFF CF SFU CU Fufu Usi Figure 4.5: Pectinase activities in starter (CGI)-fermented and un-inoculated cassava mash during fufu and usi fermentation Key: SFF- Starter-fermented fufu; CF- Spontaneously-fermented fufu (control); SFU- Starter- fermented usi; CU- Spontaneously-fermented usi (control) 8 7 Pectinase activity (U/mL) 1.6 1.4 1.2 1 0.8 24hours 48hours 0.6 72hours 0.4 0.2 0 SFF CF SFU CU Fufu Usi Figure 4.6: Linamarase activities in starter (CGI)-fermented and un-inoculated cassava mash during fufu and usi fermentation Key: SFF- Starter-fermented fufu; CF- Spontaneously-fermented fufu (control); SFU- Starter- fermented usi; CU- Spontaneously-fermented usi (control) 8 8 Linamarase activity (U/mL) 4.6.7 Analysis of organic acids and sugars during fufu and usi fermentation Since flavour enhancement, increase in nutritive values, suppression of growth of undesirable microflora, thus, increasing shelf life and general acceptability of fermented products have been attributed to the production of fermentation metabolites, the organic acids analyzed during fermentation process involving the utilization of the selected starter combination comprising L. plantarum F2C, L. plantarum U2A and L. paraplantarum U2C indicated that lactic acid was the major organic acid produced throughout the fermentation process with little traces of acetic acid. It has a retention time of 2 minutes 6 seconds (02mins:06secs). Production of other acids (butyric and propionic) was not detected. During fufu fermentation, lactic acid was not detected at zero hour, 0.51 mg/mL was produced at 24 hours after which a subsequent decrease (0.04 mg/mL) was observed at 48 hours and a much higher increased value of 1.23 mg/mL at 72 hours whereas, in the un-inoculated control, the acid was not produced at the zero hour but after which its production was decreasing with increase in fermentation time. It was also observed that higher quantities were produced in the starter fermented experiment than in the uninoculated batch throughout the fermentation process (Figure 4.7). Production of lactic acid was detected at zero hour, during usi fermentation. A slight decrease was observed from zero to 24 hours, after which there was an increase in quantity till the end of fermentation, with the highest value (6.91 mg/mL) at 72 hours. 2.71 mg/mL, 2.68 mg/mL and 4.38 mg/mL were produced at 0, 24 and 48 hours, respectively. In the un-inoculated experiment, there was increase in quantity produced with increasing time. Gradual increase (0.22 mg/mL, 0.97 mg/mL and 1.38 mg/mL) was observed from zero to 48 hours, after which production was more than doubled (3.98 mg/mL) at 72 hours. It was also noted that higher lactic acid was produced in the starter fermented batch than the un-inoculated batch (Figure 4.8). However, acetic acid was detected at the beginning of the fermentation (0 hour) both in the starter-fermented fufu (0.14 mg/mL) and usi (0.15 mg/mL). 8 9 1.4 1.2 1 0.8 Starter-fermented 0.6 Un-inoculated 0.4 0.2 0 0 24 48 72 Fermentation time (Hours) Figure 4.7: Lactic acid quantities at different time intervals during the spontaneous and starter (CGI) fermentation for fufu. 9 0 Lactic acid (mg/mL) 8 7 6 5 4 Starter fermented 3 Uninoculated 2 1 0 0 24 48 72 Fermentation time (Hours) Figure 4.8: Lactic acid quantities at different time intervals during the spontaneous and starter (CGI) fermentation for usi. 9 1 Lactic acid (mg/mL) Xylose, arabinose, fructose, glucose and sucrose were all detected during the analysis of the cassava mashes using HPLC. Other sugars such as ribose, rhamnose, maltose and lactose, even though present, had quantities below the detectable level in both fufu and usi. Figures 4.9 and 4.10 showed the results of the obtained sugar quantities. Xylose quantity increased from 1.95µg/mL at the beginning of fermentation to 3.09 µg/mL at 24 hours after which it further decreased to 1.28 µg/mL and 0.43 µg/mL at 48 and 72 hours respectively during starter (CGI) fermentation for fufu. The quantities produced in usi were lower except at 72 hours, with values ranging between 0.47 µg/mL and 3.2 µg/mL. However, in both fermentations, the un-inoculated (control) samples recorded increase in xylose quantity with increasing fermentation time. There was gradual increase from 0.38 µg/mL at the beginning of fermentation to 0.42 µg/mL at 24 hours and further to 1.77 µg/mL and 2.14 µg/mL at 48 and 72 hours respectively during spontaneous fufu fermentation. The values for usi also increased from 0.46 µg/mL to 0.64 µg/mL, 2.34 µg/mL and 2.48 µg/mL with increase in fermentation time. It was however observed that overall highest xylose (3.20 µg/mL) was produced at 72 hours in starter (CGI)-fermented usi samples. Quantities of arabinose produced in the starter (CGI)- fermented samples of both fufu and usi had slight increase from 0-24 hours with values ranging between 0.39 µg/mL and 0.61 µg/mL, however, a decrease, from 0.35µg/mL to 0.15µg/mL at 48 and 72 hours were observed in fufu sample while an increased peak value (1.37 µg/mL) was recorded in usi sample. Gradual increase in quantity with an increase in fermentation time characterized the un-inoculated usi fermentation with values ranging between 0.41 µg/mL and 1.13 µg/mL. Much higher quantities of fructose, glucose and sucrose were observed in all experimental batches. Fructose quantity decreased with increasing fermentation time in the starter (CGI)-induced fermentation for both products. Fufu had values ranging between 21.9 µg/mL and 25.4 µg/mL, while usi had between 24.1 µg/mL and 26.1 µg/mL respectively. There was an increase in quantities produced, with increasing fermentation time in the un-inoculated usi sample, with the highest value (26.2 µg/mL) at 72 hours. 9 2 120 100 80 0hr 60 24hrs 40 48hrs 72hrs 20 0 Xylose Arabinose Fructose Glucose Sucrose -20 Sugar Figure 4.9a: Sugar quantities at different time intervals during starter (CGI) fermentation for usi. 120 100 80 0hr 60 24hrs 40 48hrs 72hrs 20 0 Xylose Arabinose Fructose Glucose Sucrose -20 Sugar Figure 4.9b: Sugar quantities at different time intervals during spontaneous fermentation for usi production. 9 3 Sugar concentration (µg/mL) Sugar concentration (µg/mL) 90 80 70 60 50 0hr 40 24hrs 30 48hrs 20 72hrs 10 0 -10 Xylose Arabinose Fructose Glucose Sucrose Sugar Figure 4.10a: Sugar quantities at different time intervals during starter (CGI) fermentation for fufu production. 100 90 80 70 60 0hr 50 24hrs 40 30 48hrs 20 72hrs 10 0 -10 Xylose Arabinose Fructose Glucose Sucrose Sugar Figure 4.10b: Sugar quantities at different time intervals during spontaneous fermentation for fufu production. 9 4 Sugar concentration (µg/mL) Sugar concentration (µg/mL) Furthermore, the starter (CGI)-fermented samples had higher fructose quantities than the un-inoculated batches except at 48 and 72 hours during un-inoculated usi fermentation where values higher than those of the starter-fermented samples were observed. During usi fermentation, highest glucose quantity (30.3 µg/mL) was recorded at the beginning of the fermentation while the least (24.4 µg/mL) was at 24 hours. The least glucose (25.9 µg/mL) quantity was produced at 48 hours while the highest produced (28.8 µg/mL) was observed at 72 hours. However, a gradual increase in quantity was observed with increase in fermentation time in the un-inoculated usi samples with values ranging between 24.3 µg/mL and 26.8 µg/mL. The starter (CGI)-fermented samples had higher glucose quantities than the un-inoculated during fufu fermentation. Sucrose quantities were the highest in both fermentations, with peak values (82.9 µg/mL and 86 µg/mL) at 48 hours during starter (CGI)-fermented and un-inoculated fufu samples respectively. The highest value observed during usi fermentation was 99.7 µg/mL at 72 hours, in the un-inoculated sample. 4.7 Optimization of growth conditions for selected starter 4.7.1 Effect of different pH values on the growth of selected starter (CGI) Different pH ranges (3.5 – 7.5) supported the growth of the starter culture except at pH 3.5 when observed at 600 nm. There was a gradual increase in growth with increasing pH values, ranging between 0.22±0.13 and 1.73±0.15. Optimal starter growth (1.73±0.15) was observed at pH 7.5, closely followed by 1.24±0.17, 1.14±0.21 and 0.22±0.13 at pH 6.5, 5.5 and 4.5 respectively (Figure 4.11). 4.7.2 Effect of incubation temperature on the growth of selected starter (CGI) o The starter culture (CGI) grew within incubation temperature range of 25 C o and 45 C with growth at 600 nm ranging from 0.07±0.04 to 1.68±0.05. Optimal o temperature was at 30 C because it recorded the highest growth (1.75±0.05). This was o o closely followed by 1.68±0.05 at 37 C. Lower growth values were observed at 40 C o (0.24±0.35) and 45 C (0.07±0.04) (Figure 4.12). 9 5 2 1.8 1.6 1.4 1.2 1 Starter 0.8 0.6 0.4 0.2 0 3.5 4.5 5.5 6.5 7.5 pH Figure 4.11: Effect of different pH values on the growth of selected starter CGI. 9 6 OD (600 nm) 2 1.8 1.6 1.4 1.2 1 Starter 0.8 0.6 0.4 0.2 0 25 30 37 40 45 Incubation temperature (oC) Figure 4.12: Effect of different incubation temperature on the growth of selected starter CGI. 9 7 OD (600 nm) 4.7.3 Effect of salt concentrations on the growth of selected starter (CGI) Figure 4.13 showed the effect of various NaCl concentrations on starter growth. The optimal salt concentration for the growth of the starter was 2%, with the highest growth value 0.88±0.45 at 600 nm. Next to this was 10% salt concentration with growth measurement of 0.51±0.01 after which much less growth values were obtained at 4% (0.18±0.02), 6% (0.21±0.01) and 8% (0.17±0.01), respectively. 4.7.4 Effect of agitation on the growth of selected starter (CGI) The effect of agitation on microbial growth was monitored within the range of 50 to 250 revolutions per minute. Growth ranged between 0.89±0.02 and 1.86±0.04 at 600 nm. The optimum agitation speed, resulting in highest growth (1.86±0.04) was 100 rpm. This was closely followed by speed at 50 rpm. Speed at 200, 150 and 250rpm produced starter growth of 1.03±0.13, 0.07±0.04 and 0.89±0.02 respectively (Figure 4.14) 4.7.5 Effect of different carbon sources on the growth of selected starter (CGI) Figure 4.15 showed the effect of the different carbon sources (glucose, maltose, starch, lactose and galactose) that could support the growth of the starter optimally at 500 nm optical density. Lactose gave the least growth value (0.06±0.1), followed by galactose (0.14±0.02), starch (0.18±0.05), glucose (0.22±0.07) and then maltose, with the highest growth value (0.34±0.22), thus, making maltose the optimal carbon source for the growth of the starter culture (Figure 4.16). 4.7.6 Effect of different nitrogen sources on the growth of selected starter (CGI) Figure 4.16 showed the effect of different nitrogen sources on the growth of the starter at 500 nm optical density. The highest growth (0.59±0.07) was detected when yeast extract was used as the sole nitrogen source. This was closely followed by casein, having 0.45±0.04. No significant difference was observed in starter growth (0.31) when urea and (NH4)2SO4 were used. 9 8 1.2 1 0.8 0.6 Starter 0.4 0.2 0 2 4 6 8 10 NaCl concentration (%) Figure 4.13: Effect of NaCl concentrations on the growth of selected starter CGI 9 9 OD (600 nm) 2 1.8 1.6 1.4 1.2 1 Starter 0.8 0.6 0.4 0.2 0 50 100 150 200 250 Agitation speed (rpm) Figure 4.14: Effect of different agitation speed on growth of selected starter CGI. 10 0 OD (600 nm) 0.45 0.4 0.35 0.3 0.25 0.2 Starter 0.15 0.1 0.05 0 Glucose Galactose Lactose Maltose Starch Carbon sources Figure 4.15: Effect of different carbon sources on the growth of selected starter CGI. 10 1 OD (600 nm) 0.7 0.6 0.5 0.4 0.3 Starter 0.2 0.1 0 peptone Y.E Casein Urea (NH4)2SO4 Nitrogen sources Figure 4.16: Effect of different nitrogen sources on the growth of selected starter CGI. 10 2 OD (600 nm) 4.7.7 Effect of different incubation time on growth of starter Gradual increase in growth at 620nm with increasing incubation time was observed. Growth increased from 1.46±0.24 to 1.67±0.08 and eventually 1.83±0.08 at 24, 48 and 72 hours, respectively, thus, making 72 hours the optimal incubation time for the cultivation of the starter (Figure 4.17) 4.8 Effect of optimization of starter growth conditions on the proximate and anti-nutritional factors (%) of fermented fufu and usi mashes Proximate analysis of the cassava mashes fermented with starter that was cultivated using the optimal growth conditions was evaluated. Significantly improved nutitional content was generally observed when compared with the starter that was not subjected to optimal growth conditions during fufu fermentation. When the starter that was subjected to optimal growth conditions was used for fufu fermentation, moisture content of the mash was reduced (5.32% to 5.00%), protein content increased (1.28% to 1.52%), crude fat was reduced (0.73% to 0.60%), crude fibre and ash content increased (1.89% to 1.90% and 1.19% to 1.21%) respectively, while reduction in carbohydrate content (92.1% to 91.0%) was also observed. Meanwhile, the results obtained for usi also indicated an increase in protein content (1.82% to 1.98%) and slight decrease in crude fibre (2.35% to 2.32%). However, moisture, fat and ash contents increased in samples that were fermented using the starters cultivated under the optimized growth conditions. Results were presented in Table 4.20. The effect of optimization of on the anti-nutritional factors was also analyzed. As shown in Table 4.21, there was no significant difference at 5% level of probability in the phytate content of both optimized and non-optimized usi fermentations, whereas, the fufu mash had an increased value of 0.30mg/g in sample fermented with starter grown using optimal growth conditions. Significant reductions were observed in the tannin content of usi (33.2 mg/g to 33.0 mg/g) and fufu (35.8mg/g to 34.9 mg/g) while there was no significant difference in the cyanide content (0.05mg/g) of both optimized and non-optimized fufu fermentations. Cyanide content was not detected in usi samples. However, samples fermented with starter grown using the optimal growth conditions as well as those of the non optimised growth conditions had reduced values of anti-nutrients than in the fresh cassava. 10 3 2.5 2 1.5 Starter 1 0.5 0 0 24 48 72 Incubation time (Hours) Figure 4.17: Effect of different incubation time on the growth of selected starter CGI 10 4 OD (600 nm) Table 4.20: Effect of optimization of starter growth conditions on nutritional content (%) of fermented fufu and usi mashes Proximate Fresh Fermentation with Fermentation using cassava starter (CGI) optimal growth conditions Fufu Usi Fufu Usi a d c e b Moisture 6.82±0.04 * 5.32±0.07 5.36±0.04 5.00±0.1 5.38±0.06 e d b a c Protein content 1.12±0.15 1.28±0.06 1.82±0.13 1.52±0.07 1.98±0.15 c a e b d Crude fat 0.50±0.03 0.73±0.08 0.36±0.03 0.60±0.05 0.41±0.05 e d a c b Crude fibre 1.72±0.02 1.89±0.02 2.35±0.05 1.90±0.12 2.32±0.1 a c e b d Ash 1.39±0.07 1.19±0.05 0.69±0.02 1.21±0.02 0.81±0.04 c a d b e Carbohydrate 89.10±0.19 92.10±0.11 87.77±0.08 91.00±0.11 87.2±0.1 Data are means±SD, n=3. *The means reported with the same superscript in each row indicated no significant difference (p≤0.05) 10 5 Table 4.21: Effect of optimization of startergrowth conditions on anti-nutritional factor (mg/g) of fermented fufu and usi mashes Parameter Unfermented Fermentation with Fermentation using cassava starter (CGI) optimal growth conditions Fufu Usi Fufu Usi a d c b c Phytic acid 0.34±0.00 * 0.20±0.002 0.29±0.00 0.30±0.00 0.29±0.004 a b d c e Tannin 36.1±0.05 35.8±0.19 33.2±0.03 34.9±0.22 33.0±0.01 a b c b c Cyanide 0.10±0.02 0.05±0.00 0.00 0.05±0.003 0.00 content Data are means±SD, n=2. *The means reported with the same superscript in each row indicated no significant difference (p≤0.05) 10 6 4.9 Sensory evaluation of fufu and usi Using a 7-point Hedonic scale with a 20-man panel for the analysis of product quality, both products (Plate 2) were generally accepted by the panellists. Results shown in Table 4.22a indicated that the colour, odour and texture of the starter- fermented usi were well accepted than that of the spontaneously-fermented product (control). However, there was no significant difference in the taste and elastic quality of both products. Furthermore, the overall acceptability of the starter-fermented usi was higher than the control product with 5.29 points as against 5.21. Starter-fermented fufu had more acceptable colour and odour while no significant difference was observed in taste. The control product had a stronger elastic quality and texture even though the overall acceptability was higher in the starter- fermented fufu (Table 4.22b). 4.10 Microbial load during fufu and usi storage at room temperature. The estimation of microbial load during storage of fufu and usi at room temperature for a period of seven days was presented in Tables 4.23a and 4.23b, respectively. Total bacterial count was found to increase with increasing storage period in both starter-fermented and control products with values ranging between 9 9 5.82x10 CFU/g on day 1 to 8.51x10 CFU/g on day 7 in the starter-fermented fufu 9 9 (SFF) and 5.24x10 CFU/g (day 1) to 7.51x10 CFU/g (Day 7) in the starter-fermented usi (SFU). The total count obtained for the control products was higher, ranging from 9 9 9 9 6.19x10 CFU/g to 9.28x10 CFU/g and 6.52x10 CFU/g to 8.78x10 CFU/g in the fufu (CF) and usi (CU), respectively. Lactic acid bacteria count increased in all the products as storage period progressed 9 9 with values within the range of 0.93x10 CFU/g and 2.91x10 CFU/g from the first, to the last storage day. Coliform count was higher within the first and second days, but rd th slightly reduced on the 3 and 4 day, beyond which an increase was observed, mostly with higher values in the control products, till the last storage day. The count ranged 9 9 9 9 between 0.87x10 CFU/g to 3.44x10 CFU/g (SFF), 2.22x10 CFU/g to 3.21x10 9 9 9 CFU/g (CF), 1.01x10 CFU/g to 3.73x10 CFU/g (SFU) and1.93x10 CFU/g to 9 4.09x10 CFU/g (CU). 10 7 Usi Fufu Plate 4.2: Laboratory prepared starter-fermented usi and fufu. 10 8 Table 4.22a: Sensory evaluation of the starter-fermented and spontaneously- fermented usi Sensory parameters Starter-fermented Spontaneously- usi fermented usi a Taste a4.25±0.5 * 4.26±0.5 a b Colour 6.43±0.5 5.86±0.9 a b Odour 5.07±0.5 5.00±0.8 a b Texture 5.21±1.1 5.14±0.6 a a Elastic quality 5.63±0.8 5.64±0.5 a b Overall acceptance 5.29±0.5 5.21±0.4 Data are means±SD, n=20. *The means reported with the same superscript in each row indicated no significant difference (p≤0.05) 10 9 Table 4.22b: Sensory evaluation of starter-fermented and spontaneously fermented fufu Sensory parameters Starter-fermented Spontaneously- fufu fermented fufu b a Taste 4.57±0.5 5.07±0.4 a b Colour 5.71±0.5 4.79±0.7 a b Odour 5.14±0.7 4.29±0.8 b a Texture 4.43±0.5 5.00±0.8 b a Elastic quality 5.29±0.5 5.71±0.5 a b Overall acceptance 5.24±0.7 5.21±0.6 Data are means±SD, n=20. *The means reported with the same superscript in each row indicated no significant difference (p≤0.05) 11 0 Table 4.23a: Microbial load during usi storage at room temperature over a 7-day period 9 Samples Storage period (Days)/Microbial count (x10 CFU/g) 1 2 3 4 5 6 7 Starter- TBC 5.24 5.97 5.90 6.32 6.91 7.28 7.51 fermented usi LC 1.72 1.51 1.82 2.31 2.90 2.78 2.93 CC 1.01 2.0 1.10 1.02 2.35 2.93 3.73 YC 0.91 1.28 1.54 2.02 1.97 2.0 2.23 MC ND ND ND ND 0.11 0.31 0.64 SC 0.31 0.72 1.05 1.39 1.52 1.71 1.79 Spontaneously- TBC 6.52 8.11 8.36 9.35 9.91 8.41 8.78 fermented usi LC 1.31 1.68 2.0 2.91 2.33 2.9 3.1 CC 1.93 1.99 2.38 3.81 3.89 3.8 4.09 YC 0.87 1.51 1.73 1.31 2.51 2.72 1.93 MC ND ND ND 0.72 0.96 1.23 1.82 SC 0.58 1.31 2.84 1.77 2.93 1.95 2.32 TBC-Total bacteria count; LC-LAB count; CC-Coliform count; YC- Yeast count; MC-Mold count; SC- Staphylococcus count, ND-Not detected 11 1 Table 4.23b: Microbial load during fufu storage at room temperature over a 7-day period 9 Samples Storage period (Days)/Microbial count (x10 CFU/g) 1 2 3 4 5 6 7 Starter- TBC 5.82 6.38 6.79 7.13 7.01 7.62 8.51 fermented fufu LC 1.51 1.98 2.36 2.77 2.86 2.88 2.61 CC 0.87 2.91 2.11 1.53 1.84 2.65 3.44 YC 0.34 1.53 1.73 1.91 2.01 1.89 2.02 MC ND ND ND 0.13 1.51 1.71 1.53 SC 1.72 1.90 0.8 1.31 1.64 1.82 1.84 Spontaneously- TBC 6.19 7.34 7.91 8.28 8.53 9.01 9.28 fermented fufu LC 0.93 0.98 1.23 1.38 2.51 2.35 2.91 CC 2.22 2.73 2.21 1.62 1.71 2.53 3.21 YC 0.73 1.81 2.0 2.53 2.48 2.45 2.99 MC ND ND 0.75 1.12 2.31 2.7 2.91 SC 2.15 2.62 1.98 1.89 2.51 2.93 2.81 TBC-Total bacteria count; LC-LAB count; CC-Coliform count; YC- Yeast count; MC-Mold count; SC- Staphylococcus count, ND-Not detected 11 2 Staphylococci count was generally not as high as the coliform count. Increasing rd th growth was observed, with slight decrease on the 3 and 4 day in SFF, CF and CU, after which the count increased until the last storage day th th Mold growth was not detected until the 4 and 5 day in SF and SU, but was observed rd as early as the 3 day in the spontaneously-fermented products. Spoilage symptoms were not observed in all products on the first day, till the third storage day. On the fourth day, visible spoilage symptoms were observed in the form of whitish moldy growth, watery film and off colour in the spontaneously- fermented fufu while usi had off-yellow colour with soft and loosened texture. The symptoms were more prominent as storage period progressed, with slight visible microbial growth on starter-femented fufu and usi as well towards the end of day 5. th By the end of the 7 day, profuse spoilage was evident as off odour, slime and moldy growth in all products, brown coloured patches in fufu and off-yellow in usi as well as loosened texture (Plate 3). 11 3 Spoilt usi Spoilt fufu K Plate 4.3: Usi and Fufu showing spoilage symptoms such as whitish moldy growth, watery film, loose texture and off colour on the 7th storage day. 114 CHAPTER FIVE 5.0 DISCUSSION Cassava still remains an important root crop especially in developing countries, where it had been utilized by processing into various food products through different procedures. Fermentation of cassava has been reported to be the most important and widely used means of processing the roots (Oyewole, 1992; Nweke et al., 2002) into different fermented cassava products. Numerous authors have fermented cassava into various products like gari (Okafor, 1977; Abe and Lindsay, 1978; Ngaba and Lee, 1979; Oyewole and Odunfa, 1988; Osho et al., 2010; Edward et al., 2012), fufu (Oyewole and Odunfa, 1989, 1992; Oyewole, 1990; Oyewole and Sanni, 1995; Brauman et al., 1996; Oyewole et al., 2001; Obadina et al., 2006; Henshaw and Ikpoh, 2010), lafun (Oyewole and Odunfa, 1988; Oyewole, 1991; Nwabueze and Odunsi, 2007; Padonou et al., 2009) and so on. The submerged fermentation of cassava roots have been reported to be associated with reduction in pH of the fermenting medium, thus, increasing the acidity and acidification as a result of production of organic acids (Olukoya et al., 2011). Rapid acidification is advantageous for the process because it creates unsuitable environment for spoilage and pathogenic organisms, furthermore, hastening the fermentation process. During the spontaneous fermentation of cassava for fufu and usi production in this study, the reduction in pH values with increasing fermentation time and increasing acidity observed has previously been reported in numerous literatures. Obadina (2006) observed a decrease from an initial pH of 6.95 to 3.78 after 96 hours of cassava fermentation for fufu. During cassava fermentation to lafun and fufu, initial pH of 6.50 and 6.90 were reduced to 3.80 and 4.10, respectively after the fermentation process (Oyewole, 1995). Kobawila et al. (2005) also reported a decrease in initial pH of 7.20 to 3.80. Similar observations were reported with other substrates as well. Edema and Sanni (2008), observed a reduction in initial pH from 5.62 to 3.05, 3.37 115 and 3.65 after fermentation of maize sour dough by different microorganisms during starter selection experiment. They earlier reported a final pH of 3.71 during the spontaneous fermentation of maize meal (Edema and Sanni, 2006). The reduced pH at the end of the fermentation could be as a result of the metabolic breakdown of starch into different organic acid metabolites. Mathew et al. (1995) had earlier linked the increasing acidity which results in lowering of pH values to microbial activities which converts carbohydrates to organic acids. This was also in accordance with Oyewole (1990) and Malonga et al. (1993, 1996) who attributed it to the production of mixed acids by lactic acid bacteria. Lactic acid fermentation resulting in pH levels lower than 4.2 has been reported to constitute a major food safety factor (Holzapfel, 1997). Lower pH values observed in usi could be due to the effect of grating prior to fermentation since Oyewole and Odunfa (1992) and Okafor (1984) earlier linked rate of acidification during fermentation to the root cut size. The indigenous fermentation microflora is a complex and mixed colony of microorganisms (Kobawila et al., 2005; Ekundayo and Okoroafor, 2012). The growth and succession pattern of these organisms were reported to be dependent on factors such as water activity, pH, substrate, temperature and so on (Blandino et al., 2003), thus, the microbial as well as resulting physicochemical interactions eventually regulates the number and types of microorganisms that survives to the end of the fermentation process (Brauman et al., 1996; Padonou et al., 2009). Gradual increase in LAB count with increasing fermentation time in fufu up till the end of fermentation as observed was similar to the reports of Wakil and Osamwonyi (2012), Tetchi et al. (2012) as well as Ekundayo and Okoroafor (2012). This could either be due to increase in acidity or low oxygen tension which favours growth of facultative anaerobes in submerged fermentation. It was, however, not surprising that subsequent drop in pH could lead to increase in LAB count since it has been reported that LAB predominate at low pH of fermenting medium (Soomro et al., 2002). Increase in total bacteria count reported by Ekundayo and Okoroafor (2012) during fufu fermentation contrasted with the result obtained in this study where highest total count was at thebeginning of the fermentation while the least was observed at 72 hours. An increasing total counts as observed by Tetchi et al. (2012) also contradicted those observed in this study. Similar to Ekundayo and Okoroafor (2012), a decrease in coliform count was observed during fermentation. Furthermore, while studying the 116 survival of enteropathogens during sausage and cassava fermentations by Maftar et al. (1993) and Ogunbanwo et al. (2004) respectively, decrease in non-lactic acid bacteria with time was also reported. Olsen et al. (1995) however, linked this to the interaction among the microbial flora of the indigenous fermented foods that usually caused an apparent reduction in the total count of non lactics when reduction after 24 hours was observed in steeped maize. It could therefore be said that increase in the number of more acid-tolerant organisms which in turn creates an unfavourable environment for non acid-tolerant ones might be the reason for the decrease in number of non-lactics, whereas, their high counts at the onset of fermentation could be linked to the fermentation water, materials and handling. Yeast counts decreased throughout the fermentation process. This was contrary to Brauman et al. (1996), who reported that yeasts appeared at the end of retting during cassava lactic acid fermentation as well as Kobawila et al. (2005) and Tetchi et al. (2012), who observed yeasts to have increased and becomes important at the end of retting. The observed decrease after 48 hours of fermentation could be due to a more acidic environment, brought about by increasing number of Lactic Acid Bacteria. Lactic Acid Bacteria are described as an important group of organisms during fermentation and numerous species have consistently been isolated (Okafor, 1978; Abe and Lindsay, 1978; Ngaba and Lee, 1979; Oyewole and Odunfa, 1988; Oyewole, 1995; Kobawila et al., 2005). Oyewole and Odunfa, (1990) studied their spectrum since it was established that more than one specie was involved. Apart from other bacterial genera associated with cassava fermentation, the different lactic acid bacteria species isolated in this study (Lactobacillus plantarum, L. fermentum, L. brevis, L. delbrueckii, L. acidilactici, L. lactis and Leuconostoc mesenteroides) had earlier been reported to be involved in fermentation of cassava into numerous products. It has also been established that it is important to isolate predominant strains from fermentation batches for starter development. The observation of 50% each of randomly selected organisms from each fermentation being identified as Lactobacillus plantarum which was also the most prevalent was in line with reports of numerous authors that confirmed the predominance of lactic acid bacteria especially Lactobacillus plantarum during cassava fermentation (Ngaba and Lee, 1979; Oyewole 117 and Odunfa, 1988, 1990; Amoa-Awua et al., 1996; Ben Omar et al., 2000; Lacerda et al., 2005; Kostinek et al., 2007; Adetunde et al., 2011; Oyedeji et al., 2013). Kobawila et al. (2005) reported the predominance of Lactobacillus sp. to be 73.3% during the fermentation of cassava roots and leaves to produce bikedi and ntoba mbodi, cassava products from Congo. 99% of the total microflora during traditional lactic acid fermentation for fufu was observed to be LAB, as reported by Brauman et al. (1996). Also, Olukoya, (1995) detected the prevalence of L. plantarum (28.6%) while isolating from nine (9) different African fermented foods with similar report by Ishola and Adebayo-Tayo (2012) during the isolation of LAB from different foods. Apart from fermented foods, Ogunjobi et al. (2010) observed 72.5% predominance of L. plantarum from the fermenting core of silage. All this could be attributed to the fact that lactic acid bacteria can withstand the acidic environment created due to the gradual decrease in pH through the production of various organic acids during the fermentation process. Furthermore, L. plantarum has been shown to have less complex nutritional requirements when compared to other Lactobacillus spp. (Hammes et al., 1992) Screening of organisms for starter development during cassava fermentation involves the ability of the microorganisms to produce microbial enzymes (amylase, linamarase, pectinase) which are essential for starch hydrolysis, cyanide detoxification and tissue disintegration, the ability to rapidly acidify the fermentation process as well as production of antimicrobial compounds which antagonize unwanted pathogens (Kostinek et al., 2007; Edward et al., 2012). Screening of strains in this study for starter development showed all isolates to be negative to starch hydrolysis. This result was similar to earlier reports of Ketiku and Oyenuga, (1972) which confirmed most LAB as non-amylolytic even though, 84% of the cassava carbohydrate is in the form of starch. Amylase activity during fermentation could then be said to induced or constitutive as well as from other micro flora. Sanni et al. (2002) also reported the isolation of only a few amylolytic LAB. However, contrary to earlier reports and that of this study, Oyewole, (1995) reported over 80% of the total screened lactobacilli from fermented cassava as being amylolytic and Giraud et al. (1993) showed that L. plantarum strain A6 isolated from cassava produced an extracellular amylase. All the screened strains produced linamarase, an enzyme which is responsible for detoxification. Hydrolysis of cyanogenic glucosides was earlier reported (Okafor et 118 al., 1986; Vasconcelos et al., 1990; Louembe et al., 1997; Kobawila et al., 2003) during lactic fermentation of cassava. Several microorganisms including Bacillus sp. (Amoa- Awua and Jakobsen, 1995), lactic acid bacteria (Cohen, 1994), Lactobacilli, Leuconostoc, Streptococci and yeasts (Obilie et al., 2004), Aspergillus, Fusarium, Penicillium and Trichoderma strains (Yeoh et al., 1995) are known for their detoxification activities during cassava fermentation. It could however be ascertained that the selected screened isolates can hydrolyse cassava cyanogens through their linamarase production even though, an endogenous linamarase could be inherent in the cassava. Good pH reduction (a fast lowering of the pH) is important to accelerate fermentation process as well as reduce the levels of contaminating microorganisms which can compete with the starters for nutrients (Holzapfel, 2002) and also a critical factor in developing flavour and aroma of foods (Montet et al., 2006; Panda et al., 2007). The decrease in pH values during the fermentation of cassava roots resulted from the production of organic acids by lactic acid bacteria, which constitute the dominant micro flora (Malonga et al., 1993; Malonga et al., 1996). Similar pH reduction as observed in the screened isolates was reported (Kobawila et al., 2005; Coulin et al., 2006) with a decrease in values with increasing fermentation time. This trend could result from the fact that LAB dominated till the end of the process, thus the production of acid metabolites most especially lactic acid. A faster acidification rate, than the previous spontaneous fermentation was observed in which the final pH value after 72 hours fermentation was between 4.43 and 4.97, whereas, the starter fermented experiment had between 3.53 and 3.80 even though both initial pH was within the neutral level. Same trend was also observed in the un-inoculated batches. It has been established that the preservative action of starter culture in food is being attributed to a wide range of metabolites produced during fermentation (Caplice and Fitzgerald, 1999). The potential of LAB to produce antimicrobial compounds has attracted much attention as such compounds can be used to prevent food spoilage and inhibit the growth of food pathogens (Foulquie et al., 2006). LAB has been reported to release toxic substances such as hydrogen-peroxide, diacetyl, organic acids and bacteriocins into the fermenting medium during food fermentation (Mataragas et al., 2002). However, varied antimicrobial compound (lactic acid, diacetyl and hydrogen peroxide) concentrations as observed in this study might be as a result of being 119 produced by different strains since Tannock, (2004) linked production level and proportion to be dependent on strains, medium compounds and physical parameters. The inhibition zone of the selected potential starters in this study against pathogens ranged between 2 mm and 12 mm. Similar antagonistic effect has been reported from LAB isolated from different sources; dairy products (Saranya and Hemashenpagam, 2011), fermented maize products (Omemu and Faniran, 2011), fermented fish (Liasi et al., 2009), poultry meat (Adesokan et al., 2008) and cow milk (Olanrewaju, 2007). As observed in this work, similar antagonistic effect was reported by Adesokan et al. (2008) where LAB species were tested against Staphylococcus aureus, Pseudomonas aeruginosa, Candida albicans, Escherichia coli and Proteus vulgaris. Obadina et al. (2006) also showed the antagonistic effects of Lactobacillus plantarum on Salmonella thyphii, S. aureus, E.coli and Bacillus substilis. The ability of LAB to inhibit the pathogenic organisms however, could to be due to the fact that they produce certain antagonistic substance which interferes with the metabolic activities of such pathogens, thus, inhibiting their growth. The molecular identification of the five (5) identified potential starters further confirmed their identification to strain level. The genus Lactobacillus consists of a genetically and physiologically diverse group of Gram positive, rod shaped, catalase negative, non-spore forming bacteria (MacFaddin, 2000). Nucleotide base-sequences of 16S ribosomal DNA (rDNA) of Lactobacillus spp. has been reported to provide accurate basis for phylogenetic identification and analysis (Tabatabace et al., 2005). The use of starter cultures has been described as an appropriate control and optimization of fermentation problems of variations in organoleptic quality and microbiological stability observed in African indigenous fermented food (Sanni, 1993; Kimaryo et al., 2000). Inoculation of starters (singly and in-combination) to initiate both fermentations, had similar trend of decreasing pH values with increase in fermentation period. The least pH values (3.68 and 3.53) for fufu and usi respectively, after 72 hours fermentation, was in accordance with the report of Henshaw and Ikpoh (2010) who observed decrease in pH values from 6.20 to 3.67 after 72 hours fermentation of cassava with L plantarum starter. Kristek et al. (2004) however reported a higher range during the inoculation of L. mesenteroides and L. lactis ssp. lactis starters on sauerkraut fermentation. They obtained a final pH within the range 3.86-3.90. This is still based on the fact that fermentation pH reduces (due to increasing hydrogen ion content) as acidity increases, with increase in fermentation 120 time. Meanwhile, variation in the pH of fufu and usi could be as a result of the cassava cut size since Oyewole (1990) associated the rate of acidification to be affected by the size which the tubers were cut. It was however noted that the un-inoculated fermentations had higher pH values than the starter-fermented experiments. This confirms one of the properties of starter cultures which involves rapid rate of acidification. Furthermore, starter cultures have been shown to have significant effect on fermented foods (Adegbehingbe, 2014). At the end of 72 hours fufu fermentation, some of the starters used increased the protein content significantly while most of the starters did same during usi fermentation as well. This, according to Erukainure et al. (2010), could be attributed to the ability of the organisms to secrete extracellular protease during the fermentation process. Similar increase was reported by Oboh and Elusiyan (2007) and Adeyemi et al. (2012) while fermenting cassava tubers with single starters of Sacharomyces cerevisiae and Rhizopus oligosporus as well as during the fermentation of Chrysophyllum albidum seeds with Aspergillus niger. Other authors that reported an increased protein content during fermentation using different substrate includes Egounlety (1994), Michodjehoun et al. (2005) and Ogunlakin et al. (2012). Tortora et al. (2002) linked it to be due to the structural proteins that are an integral part of the microbial cell, but Igbabul et al. (2014) attributed it to the increase in microbial mass during fermentation, causing extensive hydrolysis of protein molecules to amino acid and other simple peptides. However, usi had higher increased values than fufu even though, same cassava roots were used. This makes such effect to be attributed to the starters. Most of the starter fermented mashes had reduced moisture contents which could be an indicator for stable shelf life. Similar reduction, as observed in this study, had earlier been linked to activities of fermenting microorganisms, since water is essential for growth and cell metabolism as well as loss of moisture along with the leaching of nutrients (Tiruha et al., 2014). Igbabul et al. (2014) reported that the decrease in moisture after fermentation may probably be due to the soft and porous texture after fermentation resulting in maximum moisture loss. However, Wang et al. (1999) reported moisture reduction as a function of many factors such as temperature, time, humidity, etc. Contrary results were reported (Omafuvbe et al., 2004; Ogueke et al., 2010; Adegbehingbe, 2014) during the fermentation of African locust bean and melon seed where moisture content was confirmed to have increased and it was related 121 to hydrolytic activity of the fermenting organisms releasing moisture as part of their metabolic product. Oladunmoye (2007) and Fadahunsi et al. (2011) also corroborated that the increase could be as a result of decompositon of the substrates by the fermenting organisms which released moisture as one of their end products. In both fermentations, ash content was reduced when fermented with majority of the starters, but with higher values in fufu. Ash content is a factor of mineral availability in the fermented food, so such decrease could be ascribed to leaching of soluble mineral elements into the fermenting medium or as a result of enzymatic hydrolysis of food components into their absorbable forms. Similar decrease in ash content was reported by Atti (2000) during fermentation of millet. It was also consistent with the report of Michodjehoun et al. (2005) during fermentation of “Gowe” a traditional food made from sorghum, millet or maize, but contrary report by Sefa-Dedeh and Kluvitse (1995), observed an increase in ash content of fermented maize cowpea blends. Sanusi et al. (2013) and Adeyemi et al. (2012) also observed an increase in ash content when fermenting Jatropha curcas and Candida albidum seeds with fungi. A general increase in crude fibre was observed after both fermentations with very few decreases in fufu fermentation. This was similar to the findings of Oyewole and Ogundele (2001) who also reported an increase in crude fibre of fufu as fermentation progressed. However, Adeyemi et al. (2012) suggested that increasing crude fibre interferes with other nutrients thereby, making such nutrients unavailable for use. On the contrary, Igbabul et al. (2014), during the fermentation of cocoyam, observed a decrease in crude fibre which was reported as an indication of softening of the fibrous tissues during fermentation and the decrease was attributed to microbial bio-conversion of carbohydrates and lignocelluloses into protein. Similar findings were reported by Hwei-Ming et al. (1994) and Balagopalan (1996). General increase in total carbohydrate was observed in both fermentations even though, some samples had reduced values. The decrease observed in this study might be as a result of breakdown of carbohydrates into simple sugars and organic acids. Oboh (2006) reported a decrease in carbohydrate content during the fermentation of cassava peels for nutrient enrichment. Meanwhile, Igbabul et al. (2014) attributed increase in carbohydrate to the decrease in moisture content as fermentation time increased during the fermentation of cocoyam flour. 122 Cassava contains anti-nutritional compounds that affect the digestibility and absorption of nutrients. Since fermentation has been known to denature anti-nutritional factors and increase nutritional values (Montagnac et al., 2009b), decrease in contents would be expected and such was attributed to leaching and microbial activities (Nwosu, 2010). Tannin, phytic acid and cyanide were considered in this study. Tannin contents in fufu increased after fermentation with all the starters except one whereas, some decrease in values were observed for usi. A general reduction in cyanide content and phytic acid were observed in both fermentations. Since the quantity of anti- nutritional compounds has been linked to the varieties and root maturity (Nambisan and Sundareson, 1994; Montagnac et al., 2009a; Bandna, 2012/2013), inconsistent values obtained when compared with numerous works was justified. Meanwhile, variations among starters indicated varied enzyme activities. Significant reduction observed had been reported by many authors on other substrates as well (Fardiaz and Markakis , 1981; Sutardi and Buckle, 1985; Marfo et al., 1990; Chelule et al., 2010; Azeke et al., 2011; Efiong and Umoren, 2011; Adegbehingbe et al., 2014; Igbabul et al., 2014). Reduction in phytic acid below the initial 0.3 mg/g as observed in this study was lower than the value (7.05 mg/g and 97.07 mg/g) obtained by Oboh, (2006) and Igbabul et al. (2014) during the fermentation of cassava peels and cocoyam flour respectively. Phytate reduction has been attributed to the activity of the endogenous phytase enzyme from the raw ingredient and inherent microorganisms which are capable of hydrolyzing the phytic acid in the fermented food preparations into inositol and orthophosphate (Sandberg and Andlid, 2002). The few increase in cyanide content observed in this study was inherent in the fermentation with single starters, however, reduction could be said to be due to combined effect of the combined starters. Majority of the fermented mashes had reduced or no detectable cyanide at all after 72-hour fermentation. This trend had earlier been established, that cassava fermentation using water, is the simplest method to reduce cyanide content (Cumbana et al., 2007; Bradbury and Denton, 2010) as the water will swell the cells and allow linamarase to come into contact with linamarin, thus, initiating hydrolysis (Bradbury, 2006). Furthermore, Guyot et al. (1998) reported such effect to have emanated from the use of linamarase producing organisms during fermentation. Meryandini et al. (2011) reported a 50% decrease in hydrogen cyanide during the addition of cellulolytic bacteria to improve the quality of cassava flour 123 while (70-75%) and 85% decrease was observed by Kobawila et al. (2005) and Achinewhu et al. (1998) after 72 hours fermentation, respectively. This observation was also in line with that of Agbor-Egbe et al. (1995) confirming fermentation to be a very effective process for eradication of endogenous cyanic compounds in cassava roots. Lower values of cyanide 0.01 mg/g was observed by Igbabul et al. (2014) when compared to those obtained in this study (0.00, 0.05 and 0.08 mg/g). The reduction from an initial 0.1mg/g to (0-0.05 mg/g) by some starters satisfied the minimum tolerant level 0.05 mg/g (SON, 1985), even though, Tweyongyere and Katongole, (2002) puts the deleterious level at 0.03mg/g and FAO/WHO (1991) recommended 0.01 mg/g dry matter to avoid acute toxicity in humans. However, observed values deviated from the findings of Oboh et al. (2002) as well as Oboh and Akindahunsi, (2003) who put the usual cyanide content of cassava products in Nigeria as 0.019 mg/g for gari, 0.025 mg/g for fufu, and that of the cyanide content of some micro-fungi fermented cassava products was between 9.1 - 17.2 mg/kg. It was noted that cyanide was not detected in most of the usi mashes after fermentations unlike for fufu and this could be as a result of the effect of grating during usi production since Dixon et al. (1994) considered grating of cassava roots to be an important factor in linking the hydrolytic enzyme linamarase to intimate contact with linamarin. Since controlled fermentation enriches nutritive value of foods with respect to shelf-life, hygienic quality, nutrient improvement and anti-nutrient reduction (Adeyemi et al., 2012), the results of the anti-nutritional/nutritional composition as well as acidification of the fermentation processes were pooled together, analyzed statistically and a starter, common to both fermentations was selected. The selected starter (CGI) was made up of the combination of Lactobacillus plantarum F2C, Lactobacillus plantarum U2A and Lactobacillus paraplantarum U2C. The starter CGI showed a considerable reduction in pH values (3.70 and 3.71) for usi and fufu respectively. This could be termed safe, since Holzapfel (1997) reported that lactic acid fermentation resulting in pH levels lower than 4.2 constitutes a major food safety factor. This pH value after 72-hour fermentation was in agreement with Oyedeji et al. (2013) who obtained a pH value of 3.70 during cassava retting, thereby concluding that the low pH would make such food safe for consumption. It was also expressed that the acidic fermentation would be responsible for inactivation of toxin-producing and foodborne infectious pathogens (Lorri, 1993; Byaruhanga et al., 1999; Kunene et al., 1999; Ana et al., 2006). 124 The starter combination expressed a significant reduction in the anti-nutritional compounds inherent in the roots after fermentation. Phytic acid has been known to form insoluble salts with metals thus, making them unavailable for absorption in the body (Igbabul et al., 2014), so reduced level of phytic acid in foods improves the availability and absorption of required metal ions in the body. The starter reduced initial phytate of 0.30 mg/g to 0.1 mg/g during fufu fermentation and to 0.27 mg/g in usi. Adegbehingbe (2014) reported higher values (3.17, 3.18 and 3.23 mg/g) than those observed in this study when fermenting Lima beans with different starter cultures. The minimum values (32.3 mg/g and 34 mg/g) in tannin content for both fermentations were observed in the samples fermented with the selected potential starter. Reduced cyanide content of 0.05 mg/g was observed in fufu while no cyanide in usi characterized the selected starter-fermentation as well. Despite being a cheap source of food calories, cassava is nutritionally deficient in protein and its high consumption without adequate supplements exposes the vulnerable group to incidence of protein-energy malnutrition (FAO, 1984). Cassava contains about 1-2% protein which makes it a predominantly starchy food (Charles et al., 2005). Certain processing techniques have been employed to improve the protein content of cassava, among which was the utilization of starter for fermentation as it was expected that the organisms will secrete protease enzymes useful in the hydrolysis of protein molecules into simple absorbable amino acids (Erukainure et al., 2010; Igbabul et al., 2014). The selected starter combination had the most improved protein content after both fermentations from an initial 1.02% to 1.82% and 1.34% for usi and fufu, respectively. Furthermore, it also showed the least moisture content in both products and reduced moisture in foods was linked to a more stable and prolonged shelf life as this hinders the proliferation of spoilage microorganisms (Padonou et al., 2010; Harris and Koomson, 2011). Least moisture levels of 5.10% and 5.36% were observed. This was lower than the recommended standard 13% for edible cassava flour (Sanni et al., 2005) and it could be due to the fact that the samples in this study are in the wet form. Reduced carbohydrate content was observed when the starter combination was used. Even though cassava is a high caloric food crop, efforts has been made to improve it nutritionally, so carbohydrate reduction observed could be as a result of the starters hydrolyzing available carbohydrate in the form of starch into simpler absorbable nutrients. The mashes had reduced fibre content and this could be 125 due to the fact that the organisms utilized soluble fibre. Similar observation was reported by Srinorakutara et al. (2006). In view of this, the selected potential starter could be said to have satisfied the aspect of reduction of anti-nutritional compounds in both fermentations more than any other observed combination, thus, its selection. Monitoring the enzyme activities during fermentation with the selected starter gives a more insight into the process. The optimal activity with respect to fermentation time of amylase, linamarase and pectinase enzymes was analyzed. Interestingly, starch hydrolysis was negative during the screening for starters in growth medium, but amylase activity was observed during fermentation with starters. This could be ascribed to the fact that the amylase enzyme was not produced by the organisms as most LAB have been proven to be non amylolytic even as 84% of cassava carbohydrate was stored as starch (Ketiku and Oyenuga, 1972). Numerous authors listed Bacillus spp. (Olafimihan and Akinyanju, 1999; Pandey et al., 2000; Gupta et al., 2003), Escherichia spp, Pseudomonas, Proteus, Serratia and Rhizobium (Oliviera et al., 2007), Aspergillus, Rhizopus, Mucor, Neurospora, Penicillium and Candida (Pandey et al., 2000; Gupta et al., 2003) as prominent organisms used in the commercial production of amylase, and as expected, LAB was not that prominent. The amylase activity might be as a result of the enzyme production being induced in the presence of starch. During fufu fermentation, the highest amylase activity 10.1 U/mL observed was at 24 hours after which there was subsequent reduction till the end of fermentation. More carbohydrate in the form of starch would still be available at the beginning of fermentation which provides a more substrate for induction since the starters are non amylolytic, thus leading to maximal activity. As fermentation progresses, available starch reduces and this could account for the reduced activity at 48 and 72 hours respectively. Activity in the un-inoculated control increases with increasing fermentation time with more reduced values, except at 72 hours. Reduced activity at the beginning of fermentation could be linked to the fact that development and colonization of a spontaneous fermentation process will be slower than an inoculated experiment, thus, a slower metabolic activity. Meanwhile, a higher activity in the control at 72 hours might be as a result of the development of more amylolytic microflora. 126 Pectinases, during cassava fermentation are involved with root softening and it has been reported that cassava softening seems to be mediated by bacteria (Okafor et al., 1984; Oyewole, 1990; Brauman et al., 1995) since sterile roots soaked into sterile water did not soften. Pectinase activity reduced with increase in fermentation time in both starter fermented and un-inoculated fufu fermentation. Earlier reports of Ampe and Brauman (1995) showed similar decrease in pectinase activity with increase in fermentation time. High activity in both fermentations at 24 hours indicated that tissue disintegration was maximal at such time since pectinase enzyme was attributed to tissue disintegration, that is, breakdown of cassava texture (Amoa- Awua and Jakobsen, 1995). During usi fermentation, the detection of pectinase activity at 24 hours, unlike in the control experiment could be as a result of faster metabolic activities in the starter inoculated batch. Linamarase or β-D-glucosidase converts the cyanide containing compounds in cassava into acetone cyanohydrins, which spontaneously decomposes to hydrogen cyanide HCN (Rolle, 1998), then further dissolves readily in water or is released into the air. This detoxification process is by the endogenous linamarase released during fermentation (Ikediobi & Onyike, 1982) or the bacteria involved in retting (Okafor & Ejiofor, 1985; Giraud & Raimbault, 1992). A higher linamarase activity observed in both fermentations when compared with the uninoculated controls indicated that starter cultures produced more linamarase than the spontaneous microflora. Decrease in activity after 48 hours during starter inoculated fufu fermentation could be as a result of rapid detoxification of cyanogenic glucosides within the early hours of fermentation. Linamarase activity in usi was higher than in fufu fermentation at 24 hours and this might be linked to the fact that cassava for usi was grated, thus, enabling rapid contact of hydrolytic linamarase with linamarin since grating has been described as an important factor during cassava fermentation (Mkpong et al., 1990; Dixon et al., 1994). Brauman et al. (1996) also ascertain the slower rate of linamarin removal in fufu when compared to gari as a result of grating before processing. The observed increased activity in fufu at 48 hours was in agreement with the findings of Cooke et al. (1978) who reported elimination of linamarin after 48 hours and it was suggested that linamarase reached optimal pH at this stage. It might also be as a result of increased tissue disintegration in fufu. It was however observed that enzyme activities in the starter fermented batches were higher than in the un-inoculated cassava mashes. 127 Fermentation metabolites have a role to play in flavour enhancement, increased nutritive and organoleptic quality, suppression of undesirable organism which in turns leads to a stable shelf life and the general acceptability of fermented foods. Organic acid, which is among the numerous fermentation metabolites are naturally present in foods or they are synthesized during biochemical or bacteria metabolism (Akalin et al., 2002; Soyer et al., 2003; Karadeniz, 2004). They have important roles because they affect the organoleptic properties, stability, nutrition, acceptability and in maintaining quality (Santalad et al., 2007). They also improve the safety and preservation of foods (Soomro et al., 2002). Quantitative determination of lactic acid is important in fermented products for technical, nutritional, sensorial, and microbial reasons (Vodnar et al., 2010) During fermentation with the selected starter, lactic acid was the main organic acid produced throughout the fermentation, even though, mild traces of acetic and butyric acids were discovered with few other starters. This was in contrast to Aka et al. (2008) who observed numerous organic acids (lactate, fumarate, oxalate, propionate) during the production of a traditional sorghum beer. Miguel et al. (2014) reported the production of citric, succinic, acetic, lactic, malic, tartaric, propionic and oxalic acids, with lactic and acetic acid dominating, thus, linked to the presence of LAB throughout the fermentation. The range (0.04 mg/mL to 1.23 mg/mL) of the lactic acid produced in the starter fermented fufu was higher than the un-inoculated batch and this confirms the fact that a starter must have a rapid acidification rate through the production of acids. Bustos et al. (2004) predicted a maximum lactic acid concentration (58.9 g/L) after 96 hour fermentation while Narita et al. (2004) obtained 14.7 g/L lactic acid (73.5% recovery) from raw starch. Javanainen and Linko (1995) utilized mixed culture of L. amylovarus and L. casei to produce lactic acid (36 g/L) using barley flour as substrate. Decrease in quantity of lactic acid with increase in fermentation time in the un-inoculated control might be as a result of the proliferation of non-lactic microflora since spontaneous fermentation is invaded by numerous unwanted organisms. The non detection of the acid at zero hour in both starter fermented and control experiment indicated that metabolic activity at such stage is minimal. Increase in quantity produced with increasing fermentation time both in the starter fermented and un-inoculated experiments in usi could be due to the fact that LAB predominates and increases till fermentation lapse. This agrees with Aka et al. (2008) who reported an increase in lactate concentration (0.17 - 7.75 g/L) till the end 128 of beer fermentation as well as Farook et al. (2012) during the production of lactic acid using sugarcane molasses. Lactic acid fermentation has been studied since 1935 using different types of microorganism and fermentation operation conditions such as pH, carbon source, temperature, inoculum size, initial substrate conditions and nitrogen source (Hofvendal and Hagerdal, 1997) as these parameters and operating condition affect the optimal process. Research has therefore concerned the influence of carbon and nitrogen sources, other growth factors (amino acids and vitamins), and culture conditions such as temperature, pH, level of aeration etc., on microbial growth (Polak-Berecka et al., 2010). Even though the starter grew at different pH range (3.5-7.5), the observed optimal being 7.5 in this study was slightly higher than an optimal pH of 7.0, reported by Barbu (2008) for the growth of LAB. Hofvendhal et al. (1999) obtained a maximum growth rate for Lactococcus lactis at pH 5.75 while Kandler and Weiss (1986) however emphasized that except for some species of Lactobacillus and Leuconostoc, lactic acid bacteria are neutrophiles i.e., their optimal pH for growth lies between 5 and 9, attributing a much lower pH to be harmful to the group of bacteria as this causes acidification of cytoplasmic pH below a threshold value and subsequent inhibition of cellular functions (Kashket, 1987). Yusuf and Abdul-Hamid (2012) also reported an optimal pH of between 9 and 10 for the growth of Enterococcus faecium B3L3. Similar report was by Ishikawa et al. (2005) who observed an optimal pH of 9.5 for halophilic and alkaliphilic marine LAB, but a contrary report (Guyot et al., 2000) indicated that L. plantarum WSO and L. plantarum A6 grew at pH values as low as 4.5. This feature allows L. plantarum strains to participate in the last stage of natural lactic acid fermentations of plant material (Daeschel and Nes, 1995) because, even though highest growth was recorded at pH 7.5, the starter expresses a gradual increase in growth from pH value 4.5. Optimum temperature is the basic characteristic of all Lactic Acid Bacteria which helps to differentiate them from each other because temperature controls the growth of bacteria. It affects bacterial generation time according to the phase of growth as each species has a unique optimum growth temperature (Ahmed et al., 2006). The O optimum growth temperature for Lactobacilli was reported to be between 30 C and O O 40 C, but they can grow at temperatures ranging from as low as 5 C to an upper limit O of 53 C depending on species (http//www.positiveaction.co.uk). Unlike the optimum 129 O growth temperature (30 C) for L. plantarum strains observed in this study, Ahmed et al. (2006) discovered that different strains of L. lactis, S. cremoris and L. acidophilus O O isolated from camel milk showed maximum growth at 37 C and 40 C. The variations in optimum temperature even among strains have been reported to be due to their genetic makeup, as it was observed by Yu et al. (1983) that if the plasmid profile of two organisms is similar or even identical, there may be differences in their nucleotide composition, sequence or even both. Barbu (2008) also reported that even though Lactobacillus spp isolated from wheat epiphyte microbiota grew within temperature O o range of 28- 45 C, optimal growth was observed at 37 C. However, these optimum o temperatures are still within the 30-40 C range and this could be attributed to their mesophilic nature. Meanwhile, Leroi et al. (2012) reported that Lactococcus piscium CNCM I-4031, a new species described by Williams et al. (1990), grew well at low temperature (0°C) and reached maximum growth below 25°C, which is very uncommon among lactic acid bacteria. Garnier et al. (2010) had earlier explained such psychrotrophic behaviour to be as a result of over-expression of cold-adaptation proteins during growth at low temperature. Salt concentration is another important growth factor for Lactic Acid Bacteria. NaCl have been confirmed to affect the metabolism and growth of LAB which are involved in fermentation processes (Chikthimma et al., 2001; Johanningsmeier et al., 2012). The optimal salt concentration of the selected starter was 2%. This was in agreement with the report of Barbu (2008) who observed an optimal salt concentration for Lactobacillus strains to be between 2-4% after which a decline in growth occurred even though they grew up till 10% concentration. Lee (2010) however, discovered that LAB in doenjang, a fermented soybean paste was survivable between 0- 40% with optimal value at 30%. This value was way above those obtained in this study even though a range of 2- 10% was used. It has been reported that bacterial cells cultivated in a high salt concentration would experience a loss of turgor pressure, which would then affect the physiology, enzyme activity, water activity and metabolism of the cells (Liu et al., 1998) even though, some cells overcome this effect by regulating the osmotic pressure between the inside and outside of the cell (Kashket, 1987). Rao et al.(2004) concluded that a 4% salt concentration was optimal for Lactobacillus plantarum strains 541 and A6 to be used as starters. Similar to this, was the observation made by Abdulla et al. (2013) in which optimal salt concentration of 4% was discovered for Lactobacilli strains. Ishikawa et al. (2005) who worked with 130 halophilic and alkaliphilic marine LAB, reported an optimal concentration of between 2.5-3.0%. However, it should be noted that the selected starter was tolerable to concentration 2-10% and this was in accordance with the findings of Elizete and Carlos (2005) that Lactobacilli isolated from gastrointestinal tract of swine were tolerable to 4-8 % Nacl even though they are from different sources. It thus gave an indication of the osmo-tolerance level of the LAB strains. The effect of oxygen on cell growth is important because during the cell metabolism, oxygen has a significant effect on the physiological characteristics of the cells (Pearce et al., 2003) and agitation tends to transfer such oxygen within the growth medium rather than it being limited to the medium surface, it also helps in avoiding cell culture to sediment at the bottom of flasks and become rapidly oxygen depleted, thus, maintaining homogeneity and fluid-to-particle mass transfer (Khadijah et al., 2009; Gupta et al., 2010). However, stationary conditions for growth and agitation mode for fermentation have also been reported in some earlier studies (Roy et al., 1986) and differences in growth rates has been attributed to differences in metabolic pathways under aerobic and anaerobic conditions (Murphy and Condon, 1984). Similar to the results obtained in this study, was the findings of Yu et al. (2013) during the development of Fed-batch process for high cell density cultivation of Lactobacillus fermentum L1. They observed that cells grew stronger with agitation speeds of 50 and 100rpm, the optimal being 100rpm, but when agitation speed was increased to150 or 200 rpm, cell density declined and this was attributed to the fact that suitable agitation helped the mixing of cells and nutrients, meanwhile, strong agitation was proven to cause high DO (dissolved oxygen) or shear force which inhibited cell growth (Yu et al., 2013). Also in accordance were the reports of Ibrahim et al. (2010), who obtained same speed of 100rpm during their study on Lactococcus lactis and that of Gupta et al. 2010, who observed maximum growth of L plantarum at 100rpm during the fermentation of edible Irish brown seaweeds. However, contrary to these was the report of Chauhan et al. (2013) who varied agitation speed for growth of Acetobacter tropicalis between 150-600 rpm and discovered the optimal growth at 450rpm, during dextransucrase production. The evaluation of cell viability and dominance during fermentation is another important characteristic of a starter culture and dominance is usually exerted by fast and dominant growth that is, generation time (Soro-Yao et al., 2014). In contrast to the result of the effect of incubation time on growth of the starter in which increasing 131 incubation period led to increase in LAB growth, Ketut et al. (2012) observed that increasing incubation periods from 24 to 48 and 72 hours, respectively reduced the number of lactic acid bacteria. They ascribed the reductions to have appeared due to the drop in pH of kefir samples and longer incubation period affects the intracellular pH of the lactic acid bacteria, which inhibit the enzyme activity, ion transport and nutrient uptake, thus, retarding the growth and counts of the lactic acid bacteria. However, a similar report was that of Enitan et al. (2011) where LAB isolates showed an increase in growth as incubation period increases even though, a decline was observed after 72 hours. Another critical step in the development of economical fermentation process is selection and optimization of carbon and nitrogen sources (Naveena et al., 2005) since high growth activity of Lactobacilli has been reported to be affected by medium formulation (Polak-Berecka et al., 2010). LAB, according to Amrane, (2000) has complex nutritional requirements, due to their limited ability to synthesize their own growth factors such as B vitamins and amino acids. Therefore, they require some elements for growth, such as carbon and nitrogen sources, in the form of carbohydrates, amino acids, vitamins, and minerals. The cost, availability and stability of fermentation substrates are of interest, as they can represent up to 68% of the production costs of fermentation products (Kwon et al., 2000; Serna-Cock and Rodríguez-De Stouvenel, 2007). Yeast extract was found to be the most suitable nitrogen source in this study and this was corroborated by Tejayadi and Cheryan (1995), in an economic study for Lactobacilli growth, who discovered the largest contributor to be yeast extract accounting for about 38% of total production medium cost whereas, Monteagudo et al. (1993) had earlier related such to be as a result of having profound influence on the cell concentration. Furthermore, it was reported that buffering capacity of yeast extract could prevent excessive drops in pH, which would inhibit cell growth (Gaudreau et al., 1997). Enitan et al. (2011) also discovered highest L. casei growth when the medium was supplemented with yeast extract. In contrast, Fung et al. (2008) showed that meat extract, vegetable extract and peptone significantly influenced the growth of L. acidophilus. Maltose, closely followed by glucose was the best carbon source observed in this study. Other reports have shown the effect of glucose on cell numbers (Liew et al., 2005; Ibrahim et al., 2010). Bajpaj-Dikshit et al. (2003) and Polak-Berecka et al. (2010) however confirmed the combination of glucose and pyruvate to be more 132 suitable because it leads to higher biomass yield. Galactose was reported to be most suitable carbon source for L. bulgaricus growth during production of hydrogen peroxide (Enitan et al., 2011). Utilizing the optimal conditions to cultivate the starter and further utilization in cassava fermentation for fufu and usi, recorded an improved proximate composition as well as anti-nutritional factors, however, some insignificant effects were noticed. This might be as a result of the optimization effect (improved metabolism) on the starter. Evaluation of the starter-fermented fufu and usi in this study, alongside the spontaneously-fermented products by trained panellists, indicating overall acceptance for the starter-fermented products justified the effective utilization of Lactobacillus plantarum F2C, Lactobacillus plantarum U2A and Lactobacillus paraplantarum U2C for both food products. This resulted in drastic reduction in foul odour and improved colour. Although, texture and elastic quality of the products had been linked to high dry matter (Etudaye et al., 2012), they could still be dependent on preparation method. These observations thus, placed utilization of starter cultures in food fermentation above the traditional processes. Similar findings were reported (Ojokoh, 2009; Henshaw and Ikpoh, 2010; Adetunde et al., 2011; Singh et al., 2012; Opere et al., 2012; Ogunbanwo et al., 2013) in which utilization of starters have been linked to control of acid content of the fermenting medium, inhibit and discourage undesirable bacteria, control fermentation time, improve odour, flavour and nutritional value of fermented products. Ready-to-eat foods are increasingly becoming the main source of inexpensive, convenient and nutritious foods with immediate consumption or consumption at a later time without further processing or preparation (Mensah et al., 2002). The highly perishable nature of ready-to-eat fufu has emerged as a recurrent and cross cutting issue over time. The negative effect of this includes spoilage of food products and contamination by pathogenic microorganisms (Adeyemi, 2012). By increasing their numbers, utilizing nutrients, causing enzymatic changes, and contributing off-flavours through breakdown of a product or synthesis of new compounds, they can spoil a food, thus, leading to reduction in demand as well as inability to expand operations size. The microbial load of starter-fermented and spontaneously fermented fufu and usi, studied over a storage period of seven (7) days at room temperature, which indicated increase in total bacteria counts with increasing storage period was similar to the report of Odom et al. (2012). This was attributed to have been as a result of storage temperature, 133 pH and acidity which favour growth of microbes. Furthermore, in accordance with the works of Onilude et al. (2002, 2004) and Odom et al. (2012), LAB counts increased as storage period progressed. However, coliform count and other spoilage organism count were evident in the food products on the first and second storage days, after which rd th they slightly decrease in number on the 3 and 4 day. Beyond this, they grew profusely till the last day, leading to spoilage of the food samples. The acidity of stored foods was reported to increase with increasing number of days (Aderiye et al., 2006), but Odom et al. (2012) reported increase in the pH of stored foods with increasing storage days. This had earlier been attributed to the production of ammonia (Sarkar et al., 2006; Olawepo et al., 2001) which favours the proliferation of more spoilage organisms as well as fungal/yeast growth. The initial decrease could be due to inhibitory action of antimicrobial compounds by LAB. Spoilage symptoms were visible in starter-fermented fufu and usi towards the end of th the 5 day, unlike in the spontaneously-fermented products that had observable th spoilage properties on the 4 day of storage. Obadina et al. (2009) reported no visible changes on day 1 and 2, change in colour and odour on day 3 and more profuse th spoilage till the 7 day during their study on the spoilage of fufu, lafun and gari. The fact that the starter produced products stayed more than the control product could be ascribed to the effectiveness of the starter acting as preservative agent during storage. 134 Summary and Conclusion This study investigated the possibility of developing a common starter culture for fufu and usi fermentation, monitoring product characteristics. Ninety eight (98) Lactic Acid Bacteria were isolated from spontaneous cassava fermentation, with Lactobacillus plantarum having the highest frequency of occurrence. They were characterized and screened (starch hydrolysis, linamarase, pectinase and antimicrobial compounds production, rate of acidification and antagonistic effect on pathogens for possible use as starter cultures. The obtained screened isolates were genotypically identified as Lactobacillus pentosus F2A, L. plantarum F2B, L. plantarum F2C, L. plantarum U2A and L. paraplantarum U2C. They were further utilized as potential starters singly and in-combination to initiate the fermentation of fufu and usi. The best starter combination, comprising of L. plantarum F2C, L. plantarum U2A and L. paraplantarum U2C (CGI) gave rapid acidification rate from 7.10 to 3.7, improved moisture and protein contents and reduced anti-nutritional contents in both fufu and usi fermentations. Lactic acid was the major organic acid produced while sugars such as xylose, arabinose, glucose, sucrose and fructose were detected at different fermentation stages, with sucrose having the highest quantity. Amylase activity decreased with inceasing time in both fermentations and pectinase activity were higher during fufu fermentation. o The optimal growth conditions for the selected starter was at pH 7.5, 30 C incubation temperature, 2% NaCl concentration, 100rpm agitation speed, maltose as well as yeast extract as carbon and nitrogen sources. Optimization of starter growth conditions had a more improved effect on the nutritional and also reduced the anti- nutritional contents of both mashes. When compared with spontaneously-fermented products, the starter-fermented fufu and usi had increased overall acceptance and a more stable shelf-life. 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Biofuels, Bioproducts and Biorefining 5.2:151–164. 184 APPENDIX I MRS Agar g/L Peptone 10.0 Beef extract 10.0 Yeast extract 5.0 Glucose 20 Di-pottasium phosphate 2.0 Sodium acetate 5.0 Ammonium citrate 2.0 Magnesium sulphate 0.2 Manganese sulphate 0.05 Tween 80 1.08 Agar 15 II Violet Red Bile Glucose Agar g/L Yeast extract 3.0 Peptone 7.0 NaCl 5.0 Bile salt 1.5 Glucose 10 Crystal violet 0.002 Neutral red 0.03 Agar 12 III Malt Extract Agar g/L Malt extract 30.0 Peptone 5.0 Agar 15.0 IV Plate Count Agar g/L Tryptone 5.0 Yeast extract 2.5 Glucose 1.0 Agar 12 185 V Modified MRS with arginine g/L Peptone 10.0 Beef extract 10.0 Yeast extract 5.0 Glucose 20 Di-pottasium phosphate 2.0 Sodium acetate 5.0 Sodium citrate 2.0 Arginine 3.0 Magnesium sulphate 0.2 Manganese sulphate 0.05 Tween 80 1.08 VI Amplified nucleotide sequence Lactobacillus plantarum subsp. argentolarensis F2B AATTGTGCTTAACACATGCATGTCGAACGAACTTTGGTATTGATTGGTGCA TCATGATTTACATTTGAGTGAGTGGCGAACTGGTGAGTAACCGTGGGAAAC CTGCCCAGAAGCGGGGGATAACACCTGGAAACAGATGCTAATACCGCATA ACAACTTGGACCGCATGGTCCGAGTTTGAAAGATGGCTTGCTATCACTTTT GGATGGTCCCGCGGCGTATTAGCTAGATGGTGAGGTAACGGCTCACCATG GCAATGATACGTAGCCGACCTGAGAGGGTAATCGGCCACATTGGGACTGA GACACGGCCCAAACTCCTACGGGAGGCAGCAGTAGGGAATCTTCCACAAT GGACGAAAGTCTGATGGAGCAACGCCGCGTGAGTGAAGAAGGGTTTCGGC TCGTAAAACTCTGTTGTTAAAGAAGAACATATCTGAGAGTAACTGTTCAGG TATGACGGTATTTAACCAGAAGTACGTTATGCCCG Lactobacillus plantarum U2A CGTATTACCGCGGCTGCTGGCACGTAGTTAGCCGTGGCTTTCTGGTTAAAT ACCGTCAATACCTGAACAGTTACTCTCAGATATGTTCTTCTTTAACAACAG AGTTTTACGAGCCGAAACCCTTCTTCACTCACGCGGCGTTGCTCCATCAGA CTTTCGTCCATTGTGGAAGATTCCCTACTGCTGCCTCCCGTAGGAGTTTGGG CCGTGTCTCAGTCCCAATGTGGCCGATTACCCTCTCAGGTCGGCTACGTAT CATTGCCATGGTGAGCCGTTACCCCACCATCTAGCTAATACGCCGCGGGAC CATCCAAAAGTGATAGCCGAAGCCATCTTTCAAACTCGGACCATGCGGTCC AAGTTGTTATGCGGTATTAGCATCTGTTTCCAGGTGTTATCCCCCGCTTCTG GGCAGGTTTCCCACGTGTTACTCACCAGTTCGCCACTCACTCAAATGTAAA TCATGATGCAAGCACCAATCAATACCAGAGTTCGTTCGACTTGCATGTGTT AGGCA 186 Lactobacillus pentosus F2A GTATTACCGCGGCTGCTGGCACGTAGTTAGCCGTGGCTTTCTGGTTAAATA CCGTCAATACCTGAACAGTTACTCTCAGATATGTTCTTCTTTAACAACAGA GTTTTACGAGCCGAAACCCTTCTTCACTCACGCGGCGTTGCTCCATCAGAC TTTCGTCCATTGTGGAAGATTCCCTACTGCTGCCTCCCGTAGGAGTTTGGGC CGTGTCTCAGTCCCAATGTGGCCGATTACCCTCTCAGGTCGGCTACGTATC ATTGCCATGGTGAGCCGTTACCCCACCATCTAGCTAATACGCCGCGGGACC ATCCAAAAGTGATAGCCGAAGCCATCTTTCAAACTCGGACCATGCGGTCCA AGTTGTTATGCGGTATTAGCATCTGTTTCCAGGTGTTATCCCCCGCTTCTGG GCAGGTTTCCCACGTGTTACTCACCAGTTCGCCACTCACTCAAATGTAAAT CATGATGCAAGCACCAATCAATACCAGAGTTCGTTCGACTTGCATGTGTTA GGCAA Lactobacillus paraplantarum U2C TCTCGTATTACCGCGGCTGCTGGCACGTAGTTAGCCGTGGCTTTCTGGTTA AATACCGTCAATACCTGAACAGTTACTCTCAGATATGTTCTTCTTTAACAA CAGAGTTTTACGAGCCGAAACCCTTCTTCACTCACGCGGCGTTGCTCCATC AGACTTTCGTCCATTGTGGAAGATTCCCTACTGCTGCCTCCCGTAGGAGTTT GGGCCGTGTCTCAGTCCCAATGTGGCCGATTACCCTCTCAGGTCGGCTACG TATCATTGCCATGGTGAGCCGTTACCTCACCATCTAGCTAATACGCCGCGG GACCATCCAAAAGTGATAGCCGAAGCCATCTTTCAAACTCGGACCATGCG GTCCAAGTTGTTATGCGGTATTAGCATCTGTTTCCAGGTGTTATCCCCCGCT TCTGGGCAGGTTTCCCACGTGTTACTCACCAGTTCGCCACTCACTCAAATG TAAATCATGATGCAAGCACCAATCAATACCAGAGTTCGTTCGACTTGCATG TGTTAGGCA Lactobacillus plantarum F2C GTATTACCCGCGGCTGCTGGCACGTAGTTAGCCGTGGCTTTCTGGTTAAAT ACCGTCAATACCTGAACAGTTACTCTCAGATATGTTCTTCTTTAACAACAG AGTTTTACGAGCCGAAACCCTTCTTCACTCACGCGGCGTTGCTCCATCAGA CTTTCGTCCATTGTGGAAGATTCCCTACTGCTGCCTCCCGTAGGAGTTTGGG CCGTGTCTCAGTCCCAATGTGGCCGATTACCCTCTCAGGTCGGCTACGTAT CATTGCCATGGTGAGCCGTTACCCCACCATCTAGCTAATACGCCGCGGGAC CATCCAAAAGTGATAGCCGAAGCCATCTTTCAAACTCGGACCATGCGGTCC AAGTTGTTATGCGGTATTAGCATCTGTTTCCAGGTGTTATCCCCCGCTTCTG GGCAGGTTTCCCACGTGTTACTCACCAGTTCGCCACTCACTCAAATGTAAA TCATGATGCAAGCACCAATCAATACCAGAGTTCGTTCGACTTGCATGTGTT AGGCAAGAT 187 VII Chemically-defined medium for pectinase production (NH4)2SO4 2g KH2PO4 4g Na2HPO4 6g FeSO4.7H2O 1mg MgSO4 0.2g CaCl2 1mg H3BO3 10µg MnSO4 10µg ZnSO4 70µg CuSO4 50µg MoO3 10µg Pectin 5g Agar 15g Distilled water 1000mL VIII Glucose standard curve 0.35 y = 0.032x - 0.108 0.3 R² = 0.958 0.25 0.2 0.15 0.1 0.05 0 0 2 4 6 8 10 12 14 -0.05 -0.1 Glucose conc. (µmole) 188 Absosorbance at 540nm IX PNPG standard curve 1.4 y = 0.672x + 0.539 1.2 R² = 0.970 1 0.8 0.6 0.4 0.2 0 0 0.2 0.4 0.6 0.8 1 1.2 P-nitophenyl conc. (µmole) 189 Absorbance at 405nm X Lactic acid calibration curve (Retention time 02mins:06secs) 60 y = 0.119x R² = 0.982 50 40 30 Peak area Linear (Peak area) 20 10 0 0 100 200 300 400 500 Quantity (µg/mL) 190 Peak area (mAs) XI Acetic acid calibration curve (Retention time 02mins:11secs) 120 y = 0.257x 100 R² = 0.998 80 60 Area 40 Linear (Area) 20 0 0 100 200 300 400 500 Quantity (µg/mL) 191 XII: Representative chromatograms showing different lactic and organic acid peak areas HPLC chromatogram of starter-fermented fufu at 0 hour 192 HPLC chromatogram of starter-fermented fufu at 24 hours 193 HPLC chromatogram of starter-fermented fufu at 48 hours 194 HPLC chromatogram of spontaneously-fermented fufu at 24 hours 195 HPLC chromatogram of spontaneously-fermented fufu at 72 hours 196 HPLC chromatogram of starter-fermented usi at 0 hour 197 HPLC chromatogram of starter-fermented usi at 72 hours 198 HPLC chromatogram of spontaneously-fermented usi at 24 hours 199 HPLC chromatogram of spontaneously-fermented usi at 48 hours 200 HPLC chromatogram of spontaneously-fermented usi at 72 hours 201 202