Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Determination of tannin levels in multi-purpose Kenyan trees and fodder crops, their variation and effect… Kangara, John N. N. 1993

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Notice for Google Chrome users:
If you are having trouble viewing or searching the PDF with Google Chrome, please download it here instead.

Item Metadata


831-ubc_1993_fall_kangara_john.pdf [ 3.57MB ]
JSON: 831-1.0098826.json
JSON-LD: 831-1.0098826-ld.json
RDF/XML (Pretty): 831-1.0098826-rdf.xml
RDF/JSON: 831-1.0098826-rdf.json
Turtle: 831-1.0098826-turtle.txt
N-Triples: 831-1.0098826-rdf-ntriples.txt
Original Record: 831-1.0098826-source.json
Full Text

Full Text

DETERMINATION OF TANNIN LEVELS IN MULTI-PURPOSEKENYAN TREES AND FODDER CROPS, THEIR VARIATIONAND EFFECT ON PROTEIN DIGESTIBILITY IN RUMINANTS.John Nduati Ngugi Kanga'raB. Sci. (Agr.), University of Nairobi, 1985A THESIS SUBMITTED IN PARTIAL FULFILMENT OFTHE REQUIREMENTS FOR THE DEGREE OFMASTER OF SCIENCEinTHE FACULTY OF GRADUATE STUDIESDEPARTMENT OF ANIMAL SCIENCEWe accept this thesis as conformingto the requir•standardTHE UNIVERSITY OF BRITISH COLUMBIAJuly 1993© John Nduati Ngugi Kang'araIn presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.^(SignatureDepartment of ^6-(1/11141—^gC/EAICThe University of British ColumbiaVancouver, CanadaDate ^1/471-114)^3 DE-6 (2/88)ABSTRACTEnergy and protein are the major limiting nutrients in dairy production on thesmall scale mixed farms in Kenya. Commercial feed supplements are expensive andtherefore multipurpose fodder trees (MPT) and forage crops are advocated as thealternative supplements, because they are inexpensive, able to provide green forageeven in dry season and have high protein content. These trees have tannins whoselevels, seasonal and altitude distribution have not been established. Previous studieshave indicated that tannins may have either beneficial effects like bloat control andincreased protein bypass, or deleterious effects like the reduction feed intake anddigestibility of protein in animals fed on tanniferous feed. The objectives of this studywere to determine the tannin levels in the MPT, as influenced by altitude and season,and the effect of these tannins on ruminal degradation and intestinal digestion of thediet.Samples of four multipurpose fodder trees and four forage crops viz. leucaena,sesbania, gliricidia, calliandra, velvet bean, green leaf and silver leaf desmodium andcassava, were collected at Mombasa (low altitude, below 300 m ASL.) in the wetseason and at Embu (high altitude, 1500 m ASL.) in both wet and dry season. Tanninand protein content were determined using gravimetric and wet oxidation nitrogendetermination methods respectively. The effect of tannin on protein degradability wasdetermined by comparing the polyethylene glycol (PEG) treated with untreated foragesamples using the mobile nylon bag technique. The treated and untreated samples wereincubated separately in the rumen of four Holstein cows with both rumen andduodenal cannula for 0, 6, 12, 24, 48 and 96 hours. Afterwards two sample of eachspecies from time 12 and 24 hours were inserted into the intestine through duodenalcannula, and recovered from the feces. The dry matter (DM) and protein of samplesrecovered from the rumen and feces were determined and these values were fitted in anon linear regression equation P = a + b(1— ")The results indicated that the MPT had ytterbium-precipitatable tanninsranging from 16.08 (±2.39)% of the DM in Gliricidia sepium to 30.31 (±2.42)% inDesmodium. intortum. The tannin content varied significantly (P<0.05) with species.The altitude did not have significant effect on tannin content, but tannins withinspecies behaved differently with season. The protein content differed significantly(P<0.05) with species. Proteins were significantly (P<0.05) higher in wet than in thedry season. The altitude had no effect on protein content. The tannin : protein ratioalso varied significantly (P<0.05) with species. Species also had a significantinteraction with season.Tannins significantly (P<0.05) reduced the rumen effective degradability ofboth DM and crude protein in all species, resulting in large quantities of undegradeddietary nitrogen(N). Tannin also significantly (P<0.05) decreased the total tractdigestion of the DM (DMD) and depressed the digestible crude protein (DCP) ofleucaena, calliandra, cassava and aintortum, but had no effect on gliricidia, sesbaniaand velvet bean DCP. A large proportion of the rumen undegraded dietary protein thatreached the intestines was degraded in most species except calliandra. Tanninsignificantly (P<0.05) altered the degradability constants a, b, and c for both DM andCP by reducing fraction a and the rate of degradation c and increasing the b fraction.From the study it was concluded that the MPT and forage crop species, despitetheir high protein content, are not good protein supplements. Their tannins reduceboth the CP digestibility and the DM degradability in the rumen. Lastly, potentiallyviable treatments that reduce the effect of tannin on digestibility are suggested.Table of contentsABSTRACT^LIST OF TABLES viiLIST OF APPENDIX TABLES^ viiiLIST OF FIGURES^ ixLIST OF APPENDIX FIGURES^ACKNOWLEDGEMENT xiGENERAL INTRODUCTION^ 1CHAPTER 1. DETERMINATION OF TANNIN LEVELS AND THEIRVARIATION^ 21.1 INTRODUCTION ^ 21.2. LITERATURE REVIEW 21.2.1 DEFINITION AND SIGNIFICANCE OF TANNIN^41.2.2 CHEMICAL CLASSIFICATION^ Hydrolysable tannins Condensed tannins^ 61.2.3 OCCURRENCE AND ECOLOGICAL SIGNIFICANCE OFTANNINS ^ 61.3 MATERIALS AND METHODS^ 81.3.1 SAMPLE COLLECTION 81.3.2 CHEMICAL ANALYSIS^ 91.3.3 STATISTICAL ANALYSIS 111.4 RESULTS^ 121.4.1 TANNIN CONTENT^ 121.4.2 PROTEIN CONTENT 121.4.3 TANNIN: PROTEIN RATIO^ 161.4.4 DISCUSSION^ 18iv1.6 CONCLUSION^ 20CHAPTER 2. DETERMINATION OF TANNIN EFFECT ON PROTEINDEGRADATION IN THE RUMEN AND INTESTINAL DIGESTION212.1 INTRODUCTION^ 212.2 LITERATURE REVIEW 222.2.1 FORMATION OF TANNIN-PROTEIN COMPLEX^ Mechanisms^ Specificity of tannins^ 232 .2.1.3 Factors influencing tannin-protein interaction^242.2.2 EFFECT OF TANNIN ON ANIMALS^ Intake^ Digestibility and animal performance^ In the monogastric^ Effect of tannins in ruminants^282.3. MATERIALS AND METHODS^ 312.3.1 SAMPLE PREPARTION 312.3.2 IN VIVO INCUBATION^ 312.3.3 ANALYSIS AND CALCULATION^ 332.4 RESULTS AND DISCUSSION 352.4.1 DM AND CP DISAPPEARANCE IN THE RUMEN^ 352.4.2 INTESTINAL DM AND CP DIGESTIBILITY^ 422.5 SUMMARY AND CONCLUSION^ 52GENERAL CONCLUSION^ 54BIBLIOGRAPHY^ 55VAPPENDICES^ 68A. The neutral detergent fiber and acid detergent fiber of the MPT foragespecies^ 68B. The effect of tannins on rumen degradation and intestinal digestion ofDM and CP in period one (40g.PEG per gram tannin) and period two PEG(100g PEG/ g tannin)^ 70C. A bio-assay for the optimum level of polyethylene glycol (PEG) ,that wouldinhibit the effect of MPT and forage tannins on the DM and CP^79viList of TablesTable 1 The sample collection and analysis summyary^ 10Table 2 Mean variation of tannin and protein with species 13Table 3 Variation of tannin:protein with species^ 16Table 4 Effect of tannin on DM effective degradability in the rumen at assumed outflowrate k=0.04 and 0.06^ 36Table 5 DM and CP effective degradability in the rumen values of different speciesat k=0.04 and 0.06 37Table 6 Effect of tannin on degradability constants of DM and CP^38Table 7 Animal variation in the effective degradability of the DM at two rumen outflowrates.^ 39Table 8 Effect of period on % DM and CP effective degradability at k =0.06^39Table 9 The effect of tannin on effective degradability of CP at assumed outflow ratek=0.04 and 0.06^ 40Table 10 The mean total tract DMD and DCP of different species^43Table 11 Mean proportion of the total tract digestion of the DM contributed by intestinaldigestibility (IDMD) of different species ^ 44Table 12 Mean intestinal DM digestibility (ID MD) of different species^ 44Table 13 Effect of period on mean total tract % digestibilty of untreated forage^45Table 14 The effect of tannin on IDMD of different species^ 46viiList of appendix tablesTables^ PageI. NDF % in MPT and forages based on DM^ 68II. ADF % in the MPT and forages based on the DM 69III. Effect of tannins on ruminal degradation and intestinal digestibility of proteinin different preiod^ 70IV. Effect of tannin on effective degradability of the CP^ 71V. Effect of tannin on degradation constants of the DM 72VI. Effect of tannin on degradation constants of protein^ 73VII. Effect of tannin on total tract digetibility of the DM (DMD)^74VIII. The effect of tannin on the total tract digestibility of CP (DCP)^75IX. The effect of tannin on intestinal DM digetibility(IDMD)^76X. the effect of tannin on intestinal digestibility of CP (EDCP) 77viiiList of figures1. Fodder species and season interaction^ 142. The interaction effects of species and seasons on protein levels ^ 153 The interaction effects of fodder species and seasons on tannin levels^ 174 Proportional contribution of intestine to total tract DM digestibility^475. The interaction effects of species and treatment on total tract digestion^486. The interaction effects of fodder species and treatments on intestinal proteindigestibility^ 507. Proportional contribution of intestine to total tract CP digestibility^ 51ixList of appendix FiguresFigure.^ PageI-IV Percentage microbial activity change with increase in PEG concentrationin the MPT species ( leucaena, sesbania, Calliandra and gliricidia)^ 82V-WI Percentage microbial activity change with increase in PEG concentrationin forage crops (velvet bean, D. intortum and cassava)^83xACKNOWLEDGEMENTI would like to express my sincere gratitude to my research supervisor Dr. J.A. Shelford for his capable guidance and invaluable advice throughout the progress ofmy research and thesis preparation.I am most grateful to members of my graduate committee to DR. L. J. Fisherof Agriculture Canada Agassiz, Dr. R. M. Tait of Animal Science Dept. for readingand commenting on this thesis, and Dr. M. Pitt Associate Dean Faculty of Agriculturefor invaluable advice during the preparation of this thesis.I am also grateful to my fellow students, the technical staff (both laboratoryand farm), and members of staff Kenya Agricultural Research Institute (K.A.R.I)Embu, Muguga and Mtwapa for their support and assistant at different stages of myresearch.My gratitude also goes to K.A.R.I and Canadian International DevelopmentAgency (CIDA) for awarding the scholarship that made this study possible.Finally my special thanks goes to my dear wife Mary and children; Wanjiru,Kang'ara, Wanyoike and Mucina for their inspiration, patient and understandingduring that long period of my absence. You are indeed a special family.xiGENERAL INTRODUCTIONThe soil and water conservation technology involving the incorporation of MPT trees intoarable lands is gaining acceptance in the farming community in Kenya (Scherr, 1992). This isbecause these trees not only provided fuel and conserve soil and water, but also are a source ofhigh protein forage for livestock feeding. Using the current knowledge, the dairy cattle fed onthese forages have not been able to exploit fully the potential of these crops. The milk yieldsof cows fed on these fodders have been lower than expected.Investigation on the level of feeding of these fodders to dairy cows have been going onalthough most of them are based on the protein content of the feed. Some concluded studies oncattle supplemented on tanniferous feed indicate a marginal increase in milk yield (Mwinga etal., 1992). Tannins, common to most MPT and some forages have been implicated as the causefor the low animal performance (Reed, 1986; Lefroy, 1992). There is therefore, a need toprovide more information on the levels and distribution of these tannins in different altitudes andseasons. Farmers should be able to identify the species that are suitable to their area consideringthe elevation of their land, and when to harvest the MPTs considering that tannin levels arelikely to change with the seasons. There is also a need to determine the effect of tannins inthese MPT species and the conventional forages on digestion and utilization of nutrients by dairycows so that the dairy farmers can be advised accordingly.This study therefore identified two areas requiring immediate attention. Mainlydetermination of levels of tannin and their distribution and the nutrient digestibility. The twowere investigated in two phases and reported separately in two chapters. Phase one was mainlya survey into existing forages, which was partly based in Kenya for field work, and partly atUBC for laboratory work. Phase two was mainly digestibility trials, all conducted at UBC.1CHAPTER 1DETERMINATION OF TANNIN LEVELS AND THEIR VARIATION1.1 INTRODUCTIONEnergy and protein are the two major nutrients limiting dairy cattle production onsmall scale mixed farms in Kenya. A large proportion of these nutrients are providedby fodder grass, pastures or their combination. The level of supplementation withcommercial concentrates currently being practiced is inadequate and provides only asmall portion of nutrients required. Commercial dairy feed supplements areexpensive. Consequently, farmers prefer feeding relatively cheap alternative sourcesto supply these nutrients.During the dry periods, growth of fodder and pastures after cutting is usuallynegligible. To counter these dry period shortages, cattle are fed with farm cropresidues, namely: maize stovers, sorghum and other cereal straws. These residueshave a low protein content ranging from 3-7% of the dry matter (DM), but have highof structural carbohydrates content whose digestibility partially depends on the levelof crude protein (CP) intake (Kossila, 1984).Inexpensive sources of protein have been leguminous pasture species namely:-Kenya white clover (Trifolium semipilosum.)  silver leaf desmodium (Desmodiumuncinatum), green leaf desmodium (Desmodium intortum) and alfalfa (Medicagosativa). These legumes were conventionally sown with grass as a mixed sward in agrazing system of management up till the early 1970's. Since then dairy cattlemanagement has been changing gradually to semi-zero and zero-grazing systems ofmanagement. In these systems, forage is cut and fed to the animals in stalls. Thechange from grazing was prompted by the gradual decrease in land available forpastures; a product of land subdivision due to high population pressure.2Under the zero-grazing system, Napier grass has been widely adopted due to itshigh dry matter yield per unit area. Alfalfa is sown in some areas where the soils arenot very acidic. Dual purpose sweet potato (Ipomea batatas) varieties are grown inmost areas to provide tubers for the family and vines for animal consumption. In thedry period both alfalfa and sweet potato vine yields are adversely affected by moisturestress.Since the majority of the farmers depend on rain water, and the land size limitsavailability of forage for conservation, protein deficiency in zero-grazed animals isrampant. There is a need for the appropriate protein sources compatible with zero-grazing type of management, especially in areas where alfalfa can not thrive.The increase in population density has also resulted in increased tree felling forfuel, construction and cropping. As early as 1970 environmental degradation wasevident in the deforested parts (Huggan and Westly, 1989). In 1978, somegovernment ministries, local and international institutions, started working togetherin rural reafforestation and research in agroforestry in order to conserve theenvironment (Huggan and Westly, 1989). Since 1980, agroforestry practices weredisseminated by both government and non-governmental agents, and are gaining wideacceptance (Scherr, 1992). These technologies involved integration of multipurposetrees with the conventional food and cash crops.The purpose of agroforestry is to conserve soil and water, provide wood forconstruction and fuel, and forage for livestock feeding. The introduction ofmultipurpose trees (MPT) in the dairy producing areas therefore provided anacceptable alternative source of protein. Most MPT are rich in protein (Jones, 1979),and their deep rooting system enables them to remain green even in dry period. Thischaracteristic makes them the only source of green foliage when grasses and3herbaceous plants are dry, and probably the only inexpensive source of protein duringthis period (Lefroy et al., 1992).Feeding regimes which incorporate MPT and herbaceous forages likeDesmodium sp. and cassava (Manihot esculenta) foliage are based mainly on theirprotein levels. (Leucaena leucocephala) is an exception to this, because in addition toprotein content, mimosine levels limit its inclusion in dairy cattle rations to 25% ofthe DM (Jones, 1979). It has been established that the majority of these MPT havehigh levels of tannins (Lefroy et al., 1992).In East Africa, browse species were found to contain phenolics which includetannins to levels of up to 50% of organic matter (OM) (Reed, 1986). Tannins fromdifferent plant species have either adverse or beneficial effects on animal performancedepending on their chemical structure (Waghorn et al., 1986; Kumar andVaithiyanathan, 1990). Quantities of tannin in these MPT should also be taken intoconsideration when formulating cattle rations. Therefore, there is need for moreinformation on the type and quantities of tannin in MPT and their role in the digestionof major nutrients.The objective of phase one in this study was:-To determine the levels of tannin and their variation in multipurpose foddertrees and herbacious crops of Kenya.1.2 LITERATURE REVIEW1.2.1 Definition and significance of tanninTannins are secondary plant metabolites whose functions in plants are not wellknown (Waterman, 1992). Plants respond to attack by herbivores, fungi, insect andbacteria by increasing tannin levels (Bate-Smith 1973; Feeny 1976). Barry and4Duncan (1984) reported a decline in feed intake for sheep fed on high-tannin forage.Van Hoven (1984) also reported that greater kudus (browsers) died of starvationdespite plenty of tanniferous browse. In birds, high selectivity for low-tannin sorghumwas evident in a mixed field of low- and high-tannin cultivars (Butler, 1989). Suchfindings have led some workers to conclude that tannins serve as a chemical defense.These views however, are being challenged because some animals show somepreference for tannic foliage (Robbins et al., 1987a). It has now been established thatanimals avoid excessive intake of high levels of tannins, but the tolerance of tanninsvaries from species to species (Mole and Waterman, 1987a). The significance oftannin in animal nutrition arises from the fact that it binds proteins to form a tannin-protein complex (Swain, 1979; Butler, 1989). The complex so formed is resistant orslows degradation by animal or microbial digestive enzymes (Swain, 1979; McLeod,1974; Kumar and Singh, 1984)The ability of tannin to bind with protein has been exploited for centuries toconvert hides and skin into leather. Tannin cross links with collagen chains of the hideto make durable, bacteria resistant leather (Mangan, 1988; Haslam, 1989). Thesetannins were extracted with water from macerated plant parts (Haslam, 1989).Tannins were then defined as water-soluble plant extracts with the ability to tan hidesinto leather. Since then it has been found that these extracts were composed of largemolecules of phenolics. Tannin is therefore defined as water soluble plant phenolicmetabolites with molecular weight ranging between 300 and 3000 Daltons, andability to bind and/or precipitate proteins.51.2.2 Chemical classification of tannins.Tannins are classified into two categories (Haslam, 1989) namely: hydrolysableand condensed tannins. Hydrolysable tannins.Hydrolysable tannins consist of a carbohydrate moiety in which the hydroxylgroups are esterifial to gallic, digallic and hexahydroxydiphenic acids. Thehydrolysable tannins are easily hydrolyzed by heating with weak acid or enzymes suchas penicillin tannase (McLeod, 1974). The ability of hydrolysable tannin to bind andprecipitate protein is controversial. Some workers state that hydrolysable tannins forma stronger precipitate with protein than condensed tannin (Cooper and Owen-Smith,1985), while others maintain that hydrolysable tannin bonds are weak and have noeffect on digestibility of protein in ruminants (Hagerman et al., 1992). Thehydrolysable tannin protein complexes are hydrolyzed in the rumen to gallic acidwhich is absorbed and excreted in the urine. Condensed tannins.These are polymeric compounds of flavan-3-ols or flavan-3-4 diol and relatedderivatives. About 50% of condensed tannin, when heated with mineral acid,produces a red color typical of anthocyanidins such as cyanidins and pelargonidin(Mangan, 1988). They are therefore generally referred to as proanthocyanidins.Unlike hydrolysable tannins, condensed tannins do not have a carbohydrate core. Twomajor examples of condensed tannins are procyanidins and leucoanthocyanins.61.2.3 Occurrence and ecological distribution of tanninsTannins occur in almost all plant genera, but mainly in dicotyledons andparticularly woody or tree legumes (McLeod 1974). Mangan (1988) reported that by1954 over 500 species had been shown to have varying levels of condensed tannin.The levels of tannin vary with plant species (Ford, 1978). Although tannin can befound in all parts of the plant, some parts have higher tannin content than others. InGramineae, such as sorghum and finger millet, tannins are found in grains only andnone in the vegetation (Barney et al., 1989). Swain (1965) found that tanner's sumactree (Rhus coraria) leaves, bark and wood had 27, 6, and 4% tannin in the DMrespectively while quebracho tree (Schinopsis quebracho) had 7% in the bark 2.5% inthe soft wood and 20% in the hard wood (Leinmuller et al., 1991). Within the samespecies tannin may vary with varieties (Akbar and Gupta, 1985). Usually the dark orbrown coated seed of sorghum and finger millet have higher tannin content than thelight coated (Bullard et al., 1981).The concentration of tannins or polyphenols changes in the same plant partwith maturity. In sorghums, tannins increases sharply after pollination up till doughstage and decline as the grain dries (Bullard et al., 1981). In plum fruit, the tannincontent is higher when green and declines as it ripens (Goldstein and Swain, 1963).Changes in tannin content with age or maturity is attributed to polymerizationof phenolic monomers which are predominant in the immature stage. Under in vitroconditions, bovine serum albumin (BSA) precipitation increased with degree ofpolymerization (Horigome et al., 1988). This is probably because monomers are toosmall to cross link effectively with protein in young plants (Hagerman et al. 1992).After maturity and at ripening, tannins decline due to polymerization, becoming too7large to complex protein effectively (Goldstein and Swain, 1963; Bullard et al.,1981).Deficiency in major soil nutrients such as nitrogen (N), phosphorus (P) andpotassium (K) results in higher levels of tannins in growing plants (Barry and Forss,1983; Bryant et al., 1987). Soil pH affects the availability of some major nutrientssuch as P through fixation in acidic soil, thus affecting tannin distribution (Gartlan etal., 1980).Tannin levels in plants also vary with season due to changes in soil nutrientavailability as a result of precipitation. This moisture change affects growth and altersthe DM:tannin ratio (Coley, 1988). Increase in daylight, moisture, and temperatureenhance plant growth in summer or the rainy season in tropical climates. Warmth andmoisture also favor insect and microbial multiplication resulting in increasedinfestation and herbivory on plants (Coley, 1988). The plant responds to these attacksby further production of tannin, forming galls and callus in the affected parts (Feenyand Bostock, 1968; Haslam, 1989).1.3 MATERIALS AND METHODS1.3.1 Sample collectionFoliage samples of eight Kenyan multi-purpose fodder trees and crops, werecollected at Mombasa (low 300 m above sea level (ASL)) and Embu (high 1500mASL) as summerised in Tablel. In Embu, samples were collected in both dry and wetseason of February and June 1991 respectively, while in Mombasa only wet seasonsamples were collected in early July 1991. The species collected were: Leucaenaleucocephala, Sesbania sesban. velvet bean Mucuna pururiens, Calliandra calothyrsus,8Gliricidia sepium, cassava foliage Manihot esculenta, green leaf desimodiumDesmodium intortum  and silver leaf desimodium Desmodium uncinatum. Freshsamples were oven dried at 60° C for 3 days weighed, ground and shipped foranalysis at the University of British Columbia.1.3.2 Chemical analysisThe dry matter was determined by placing 1.2 to 2.0 g of the sample in a pre-heatedand weighed aluminum dish, and both were oven dried at 105° C for 24 hours andweighed. Two samples per species were used to calculate the percentage DM, usingthe following formula (W3-W1/ W2-W1) x 100where W1 = dish weightW2 = original sample and dish weightW3 = cooled dry sample and dishThe CP was determined through N determination using the wet oxidation procedureof Parkinson and Allen (1975).Tannin levels were determined using a gravimetric method described by Reed at al.(1985). This method exploits rare earth elements, affinity for phenolics at neutral pH.These elements are used to precipitate total soluble phenolics extracted from plantparts. The method uses ytterbium acetate to precipitate phenolics extracted fromforage samples(100mg) using 70% acetone. The precipitates are refrigerated for 24hours. After refrigeration, the ytterbium-tannin precipitates are then filtered,subjected to chlorophyll removal by washing with acetone, oven dried, weighed,ashed and reweighecl to determine the ytterbium-precipitable tannin. This method isinexpensive and avoids the problems associated with colorimetric methods of tannindetermination.samples were analysed in triplicate and the mean of the two samples used for analysis.9Tablel .The sample collection and analysis summaryAltitude Dry season Wet seasonHigh collected analyd/sam collected analyd/samLeucaena 2^3 2 3Sesbania 2 3 2 3Gliricidia nil^- 2 3calliandra 2^3 2 3Velvet bean nil 2 3D. intortum 2^3 2 3D. uncinatum 2 3 2 3Cassava 2^3 2 3LowLeucaena nil^- 2 3Sesbania nil 2 3.Gliricidia nil 2 3Calliandra nil^- 2 3Velvet bean nil - 2 31.3.4 Statistical analysisThe analysis of variance and means comparisons (least significant difference) werecarried out using the general linear model procedure from the SAS package version6.04(SAS/STAT software, 1985), as an 8x2x2 factorial experiment.The model used was;Yijk =1-t ±0i+Ij+70-07ix+ Otij +Eiji(where YijK = measured variable= mean13i = species effectii = altitude effectYK = seasonal effectf3yhc= species x seasonal effect13tii = species x altitude effectcjj = error effect111.4 RESULTS1.4.1 Tannin contentThe tannin content is presented as ytterbium precipitatable (Y-ppt) phenolics.The Y-ppt phenolics estimate the percentage total phenolics in the DM, which includehydrolysable tannins, condensed tannins and their monomers.The percentage Y-ppt phenolics varied significantly with species (P <0.05).Their least square means (LSM) ranged from 16.08 (± 2.39) % of the DM inGliricidia sepium to 30.31 (±2.42) in Desmodium intortum  (Table 2). The altitudedid not have significant effect on Y-ppt phenolics contents,nor did altitude interactionwith species. Although seasons do not indicate significant differences on tannin levels,their interaction with species were significant (P< 0.05). Species behaved differentlyfrom each other in different seasons resulting in varying changes of tannin content(Figure 1). Calliandra tannin content showed a sharp increase (from 17% in dryseason to 30% in wet season) in the wet season and decline in the dry season. Cassavatannins on the other hand were higher in dry season compared to the wet seasondecreased in wet season and increased in dry season. In D. uncinatum, changes intannin were gradual but increased in wet season and declined in dry season. Leucaena,sesbania and D. intortum  tannin remained almost unchanged regardless of season.1.4.2 Protein contentThe protein content presented in Table 2 showed significant differences(P <0.05) between species and seasons. The altitude had no significant effect onprotein content. There was a significant interaction between species and season (P <0.05)(Figure 2). All the species showed a positive protein increase in the wet seasonwith leucaena having the highest protein increase.12Table 2 Mean variation of tannin and protein with speciesTannin(s.e. m)% of the DMProtein (s. e. m)% of DMCalliandra 25.96cd ±1.88 23.42cde +1.11Cassava 26.24cd ±2.42 25.77de ±1.38D. intortum 30•31d ±2.41 18.20a^±1.38D.uncinatum 23.53bc ±2.41 19.06ab ±1.38Gliricidia 16.08a ±1.39 20.69bc ±1.36Leucaena 24.16c ±1.88 25.86e^±0.99Sesbania 27.97cd ±1.88 23.49cd ±1.06Velvet Bean 17.33ab ±2.39 24.90cde ±1.36Values followed by similar superscript in a column are not significantlydifferent(P <0.05).131 1--_-35.---,.-o 30C/1Z>CD^25ff.-4r.ZSE- 2015Dry^ WetRainfall SeasonFigure 1. Fodder species and season interaction.• Cassava, 7 Desimodium, intortum • Sesba,nia,O Leuca,ena„ V Desmodiwm, uncinatu-m„ 0 Callia,ndra14 32302826242?20181611215DRY^WETRainfall SeasonFigure 2. The interaction effects of species andseasons on protein levels.• Cassava, 7 Dessmodium tin,tortum. • Sesbania,0 Leucaena, V Desmodium uncinatum. 0 Calliandra151.4.4 Tannin : protein ratio.The tannin:protein ratio Table 3. was obtained by dividing the percentage Y-pptphenolics of a species by the percentage protein content of the same species. Theseratios indicated the amount of tannin per unit of protein, and varied significantly(P <0.05) with species. Tannin:protein ratio also varied significantly with seasons.The interaction between species and season ffigure 3) was also significant (P <0.05).Table 3. Variation of tannin : protein ratio with speciesSpecies Tannin:protein (s.e.m)Calliandra 1.081) ±0.09Cassava 1.15b ±0.11D. intortum 1.70d ±0.11D. uncinatum 1.21c ±0.11Gliricidia 0 . 88ab ±0 . 11Leucaena 1.02ab ±0.08Sesbania 1.28 ±0.08Velvet bean 0.78a ±0.11Values with similar superscripts in the column are not significantly different(P<0.05)16Figure 3. The interaction effects of fodder speciesand seasons on tannin levels.• Cassava, 7 Desmoclium tintortum, • Sesbania 0 Leucaena,V Desmodium uncinatum, 0 Calltiandra175 DISCUSSIONIn this study, tannins in the multi-purpose fodder trees and crops indicatedsignificant (P < 0.05) differences between species which corroborates the observationsmade by Martin and Martin (1982), Reed (1986), and Burritt et al. (1987). However,even where similar species were studied, the methods of analyses were different fromthe one used in this study and therefore the values may not be comparable. In thisstudy, tannin levels for gliricidia were lower than those reported by Vadiveloo andFade! (1992), but those for leucaena were similar to theirs.The tannin levels did not vary with altitude. This contrasted with the findings ofGartlan et al. (1980) who reported significantly higher tannin content foliage numberof species at lower altitude (<650 m above sea level (ASL)) compared to similarspecies at high altitudes (>1300 m ASL) in the tropics. The results from the currentstudy agree with those obtained by Baldwin et al. (1987), who found no significanttannin variation with geographical location.Contrary to the findings of Feeny and Bostock (1968) in oaks, Shultz et al.(1982) and Baldwin et al. (1987) in sugar maple and birch, seasons had no significanteffect on tannin content, which corroborated the findings of Cooper et al. (1988) inseveral South African savanna browse species. However, the significant interactionobserved between species and season in this study, is an indication that MPT and cropspecies respond differently to moisture and temperature changes in tannin production.Cassava tannins were higher in the dry season and lower in the wet season, which wasopposite to the other species. With the other species investigated, tannin levels werefound to be higher in the immature stage of growth, which is often in the wet season,and then declined with maturity in the late wet and dry period (Gartlan et al., 1980;Schultz et al., 1982; Provenza and Malechek, 1984). The magnitude of decline in18tannin content varied in each species. Calliandra and D. uncinatum showed aremarkably sharp decline from wet to dry season, while Sesbania and D intortum hadshowed little change. Different species will therefore have altered levels of tannindepending on the season even in the same environment.If leaf life span expectation and light intensity result in increased production oftannin (Coley, 1988), then these tropical tree species would be expected to have hightannin content since they are evergreen, and light is not limiting in the tropics. Inmost studies involving tannin distribution (Feeny and Bostock, 1968; Gutlan et al.,1980; Baldwin et al., 1987; Cooper et al., 1988), trends of changes in tannin contentfrom bud to mature leaf were monitored in all the stages of development. In this studythe samples were taken once in each season, and probably that is why seasonalchanges in tannin content could not be detected, hence the need for more samplingtimes.Protein content differed significantly (P <0.05) with species and season. This istypical of most tropical forages, whose crude protein content is higher during thegrowing period, and declines with maturity. The CP levels are at the lowest in the dryperiod (Minson, 1988). The CP contents obtained in this study were similar to thosereported in other studies (Jones, 1979; Reed et al., 1982; Akbar and Gupta, 1985;Van Eyes et al., 1986).The observed differences (P <0.05) in tannin: protein ratio between species andseason in this study were similar to those reported by Tangendjaja et al. (1986).Elevating the protein content in tanniferous forages reduces the tannin : protein ratio,and results in increased acceptability of forages and performance of the animal(Provenza and Malechek, 1984; Cooper et al., 1988). Therefore the observed changesin tannin: protein ratios resulted from the significant seasonal changes in protein19content, while seasonal tannin changes were negligible. The animal performanceattributable to tannin content would vary with the changing ratios. This may be truewhether the CP changes are seasonal or there is a deliberate addition of protein intothe animal ration. This is because tannins are competitive inhibitors of proteases(Mole and Waterman, 1987b). Therefore increasing the protein content results inmore unoccupied binding sites on the protein for enzymes to attach. Proteolysis willthen proceed though at a reduced rate relative to low tannin forages.1.6 CONCLUSIONIn this study all the MPT and crops investigated had tannins characteristic ofthe specie. The average tannin levels did not vary much with the season, but theprotein content varied with the season. This affected the tannin per unit protein(tannin : protein ratio), which may lead to seasonal change in animal performance.Frequent cutting of these fodders under zero-grazing type of dairy cattlemanagement results in forage composed of young leaves and shoots which are rich inprotein . This is likely to reduce the tannin: protein ratio. On the other hand, stress dueto continuous disturbance through frequent defoliation, may lead to increased tannincontent and the result would be an increase in tannin: protein ratio.There is need for further investigation to elucidate: (1) the effects of frequentcutting of MPT and crops on the tannin: protein ratio, (2) the effect of changingthese ratios on animal performance (3) and the seasonal tannin distribution pattern.20CHAPTER 2DETERMINATION OF TANNIN EFFECT ON PROTEIN DEGRADATION INTHE RUMEN AND INTESTINAL DIGESTION2.1 INTRODUCTIONThe survival of ruminant animals on fibrous feeds is due to the symbioticrelationship with rumen micro-organisms. Of the rumen microbes, bacteria are themost abundant. The other micro-organisms include protozoa and fungi (Orskov,1982).Once the animal is fed, a large portion of the dietary protein is hydrolyzed intoshort chain peptides or amino acids. These products are either incorporated directlyby the microbes or deaminated to ammonia (NH3) by the bacterial deaminases. Theresulting NH3 becomes the primary nitrogen (N) source for microbial proteinsynthesis (Armstrong and Weekes, 1983). About 75% of rumen microbial N isderived from NH3 (Oldham and Parker, 1981).Excess ammonia diffuses through the rumen wall into the blood stream, fromwhere it finds its way to the liver, is converted into urea and excreted in urine, orrecycled back into the rumen through saliva. The dietary protein and the recycledNH3 are not the only sources of N for microbial protein synthesis. The non-proteinnitrogen (NPN) inherent in some feeds such as silage or urea are important sourcesof N. The NPN can provide sufficient N needs for maintenance and substantial milkproduction (NRC 1988). However, for high levels of production, some additional trueprotein would result in even better performance, because it not only maximizesmicrobial protein production but also increases the undegraded protein available forthe host enzymatic digestion.21Some dietary protein escapes degradation in the rumen and together withmicrobes, are flushed into the lower gut (abomasum and intestines) where they aredigested to provide the host animal with amino acids. The extent of proteindegradation ranges from 35-80 % (Oldham, 1977), and depends on the chemicalnature of the feed, mode of processing, and rumen outflow rate. The outflow rate isin turn affected by the frequency of feeding. The higher the frequency, the higher theoutflow rate and the lower the extent of degradation, hence more protein bypass.The presence of some chemicals such as tannins and formaldehyde has beenshown to reduce the extent of protein degradation and increase the dietary proteinbypass. Digestion of the bypass proteins in the intestines is not 100% efficient assome, depending on their chemical composition, are voided with feces undigested.Overprotection of protein can lead to higher fecal nitrogen loss.In the previous chapter the levels of tannins and their variation with species,altitudes and seasons were reported. The objective of this study was;To determine the effect of these tannins on ruminal degradation and intestinaldigestibility of protein.2.2 LITERATURE REVIEW2.2.1 Formation of tannin-protein complexes2.2.1.1 MechanismThe ability of tannin to bind and precipitate protein is inherent in its chemicalstructure. Hydrolysable tannins are polymers of seven to nine carbon ring (C7-C9)phenolic acids such as gallic and coumaric acids, while condensed tannins arepolymers of flavanoid which is a C15 ring structure. Both hydrolysable and22condensed tannins have numerous phenolic hydroxyl groups exposed and unreacted ontheir surfaces. It is believed that the phenolic hydroxyl groups forms a hydrgen bondwith the carboxyl group of the peptide to form a complex (Leinmuller et al. 1991). Atone time this hydrogen bonding was claimed to be the main binding mechanismbetween tannins and proteins (Gustayson, 1954). Since then hydrophobic, electrostaticand covalent interactions have been suggested (Butler et al., 1984). Oh et al. (1980)following an intensive study involving cytochrome C tannins immobilized in asepharose column and several eluents, provided strong evidence in favor ofhydrophobic interaction over hydrogen bonding and the other interactions.Hydrophobic interaction is possible because both tannin and protein havehydrophobic regions, mainly the aromatic nuclei of tannin and the aliphatic andaromatic side chain of amino acids (Oh et al. 1980). Oh et al. (1980) also ruled outthe possibility of electrostatic interaction for condensed tannin in acidic and neutralmedia. Butler et al. (1984) working with sorghum tannins found no evidence insupport of covalent bonds. As for electrostatic interaction, Butler et al. (1984) foundthat binding was possible in a very high pH medium, and even then most proteinswould be negatively charged, hence repulsion forces rather than attraction would bein effect. They however came out strongly in favor of both hydrogen andhydrophobic bonding. In this case then small phenolic molecules (monomers, dimers,trimers etc.) would be less effective in protein binding because of their smallhydrophobic region. If present in large quantities, though not polymerized, smallphenolic molecules can bind protein just like phenolic polymers such as tannins(McManus et al., 1981). Specificity of Tannins23Tannins were assumed to be indiscriminate binders of all proteins (Goldstein andSwain, 1963). However studies by Butler et al. (1984) using affinity chromatographyand a competitive binding assay, found that the proline rich proteins had higheraffinity for tannins than the low proline proteins, large proteins with loose openstructure tended to bind strongly with tannins, and presence of proline in the mediatended to open the protein structures. An earlier report (Hagerman and Butler, 1981)showed that proanthocyanidins selectively precipitated out one protein in the presenceof a large excess of another protein. Glycosylated proteins were reported to have lowaffinity for tannins (Butler et al., 1984; Strumeyer and Malin, 1970) although thesefindings are contrary to those of Asquith et al. (1987). It can then be concluded thattannins do not bind all protein equally. Some proteins are bound more strongly andfaster than others. Gelatin for example, binds with tannin faster and more stronglythan some proteins (Calderon et al., 1968).Condensed tannins from various plant species show structural differences. Someminor differences such as linkage isomerism at C3 and C4 may result in differentaffinity for proteins by relatively similar tannins (Clausen et al., 1990). This and thedegree of polymerization could probably account for the differences in ability to bindprotein, observed between condensed tannin from different plant species (Asquith andButler, 1985). Factors influencing tannin-protein interactionThe interaction of protein and tannin is influenced by metallic ions in the media.Divalent cations such as Ca2 + and Mg2 + enhances the precipitation of protein bytannin more than monovalent cations such as Na+ and K+ (Martin et al., 1985).The pH of the media also influences the ability of tannin to bind with protein. Jonesand Mangan (1977) found that leaf protein fraction 1 formed stable complexes with24tannins between pH 3.5 and 7.0. Below pH 3.0 and above pH 8.0 the complex wasunstable and easily dissociated.The interaction between pH and metallic ions has a complementary effect onprecipitation of protein by tannin. Martin et al. (1985) found that at pH 6.15 and6.90, the presence of sodium and potassium ions favored the precipitation of ribulose-1,5 bisphosphate carboxylase (RUBPC) by tannic acid. At pH 6.15 in the presence ofsodium and potassium salts, 49% of RUBPC in the media was precipitated and only16% in the absence of salts. At pH 6.90 in the presence of salts, 84% of RUBPC wasprecipitated compared to 10% without salts. Higher pH above 7.55 resulted in littleor no precipitate even in the presence of salt (Martin et al., 1985).Presence of detergents in the media result in the reduction of tannin-proteinprecipitation. Lysolecithin, a surfactant found in gut fluid of some insects, completelyprevented precipitation of RUBPC by tannins at pH 6.15, 6.90, and 7.55, even in thepresence of salts (Martin et al., 1985). Sodium dodecyl sulfate (SDS) is an effectivesolubilizer of the tannin-protein complex (Oh et al., 1980). Mammalian bile cholicacid dissolves precipitates of bovine serum albumin (BSA) and tannin when added atconcentration similar to that found in the small intestine of humans (8mM) (Mole andWaterman, 1985).Polymers such as polyvinylpyrrolidone (PVP) and polyethylene glycol (PEG)have been known to have very high affinity for tannin. They have therefore been usedin purification of plant enzymes to prevent tannins from interfering with enzymes(Glen et al., 1972; Rayudu et al., 1970). PEG (mol. wt. 4000) effectively solubilizeda complex of tannin and leaf protein fraction 1 under in vitro conditions (Jones andMangan, 1977).Under in vivo conditions, spraying high tannin Lotus pedunculatuswith PEG reduced the reactive tannin from 63g to 7g per kg DM and this resulted inenhanced intake and digestibility of DM, and total nitrogen digestibility (Mangan,1988). Studies involving animals indicated that PEG applied through spraying or25drenching, reduced the effect of tannin on protein utilization (Barry and Duncan,1984; Barry and Manley, 1986; Nunez-Hernandez et al., 1991).2.2.2 Effect of tannin on animals2.2.2.1 IntakeThe ability of tannins to bind and precipitate protein is believed to be the cause ofthe decreased voluntary feed intake observed in animals fed on some tannincontaining feeds (Barry and Duncan 1984, Van Hoven 1984, Cooper and Owen-Smith1985, Kumar and Vaithiyanathan 1990). The intake reduction is attributed to bindingof dietary protein, salivary mucoprotein and mucosal epithelial cells (Provenza andMalechek 1984). This causes a diffused feeling of extreme dryness and bitter taste inthe mouth and throat of the animal commonly referred to as astringent (Mole andWaterman 1987a), prompting the animal to avoid tanniferous feeds (Haslam 1989).However, studies by Clausen et al.(1990) indicated that the snowshoe harepreferred bitterbrush foliage over blackbrush despite the fact that bitterbrush tanninshave higher protein precipitating capacity than blackbrush tannin. Cooper et al.(1988) found that, despite the fact that Acacia nilotica has very high tannin content(about 300 mg/g) which taste highly astringent, it was highly favored by impalas overother less astringent species. Robbins et al. (1987a) could not find conclusiveevidence that protein precipitating ability of tannin is an effective defense againstherbivory. The three studies therefore question the validity of tannin as an inhibitor ofvoluntary intake.Jones and Mangan (1977) found that bovine sub-maxillary mucoprotein couldbind with tannins only at temperatures below 250 C, and dissociated at temperaturesabove 250 C. This ruled out binding of salivary mucoprotein at normal bodytemperatures. Robbins et al. (1987a) suggested that some depolymerisal tannins areabsorbed into the blood stream and most likely inhibit intake through toxicity in the26liver. A combination of factors appears to be involved in inhibition of voluntary feedintake and tannin is one of them (Cooper et al., 1988; Clausen et al., 1990). Digestibility and animal performance2. In the monogastricTannins ingested with food have been found to inhibit digestion of food bothin monogastric and ruminant animals. In pigs, feed intake and digestibility values arelower in pigs fed on high than low tannin sorghums diets (Cousins et al., 1981,Mitaru et al., 1984). Digestibility and growth reduction have also been reported forrats fed on tannin containing feeds (Joslyn and Glick, 1969; Jambunathan and Mertz,1973; Horigome et al., 1988) and in chickens fed on sorghum based diets (Armstronget al., 1973). The reduction in digestibility was attributed to irreversible binding ofdietary protein by tannins, forming a tannin-protein complex that resists the effect ofdigestive enzymes (Mole and Waterman, 1987a; Horigome et al., 1988) or to bindingdigestive enzymes and inactivating them (Goldstein and Swain, 1965) or to bindingboth enzyme and substrate (Hagerman and Butler, 1978; Horigome et al., 1988). Thetannin-protein complex is then passed out in feces undigested, hence the increasedfecal nitrogen observed in both humans and animals (Kumar and Singh, 1984).Prolonged feeding of animals with tanniferous feed leads to a decline in growth,weight and egg production in the case of chickens (Armstrong et al., 1973; Schaffertet al., 1974; Sell and Rogler, 1983; Mitaru et al., 1984; Myer et al., 1986).The binding of enzymes by tannin as an inhibitor of protein digestion hasbeen disputed following the finding that most enzymes isolated from rats feeding ontanniniferous diet are fully active (Mitjavila et al., 1977). Mole and Waterman(1987b) also found that tannic acid does not bind with tryptic enzyme but rathercompetes with it for protein substrate. In some cases tryptic hydrolysis of BSA wasincreased in the presence of condensed tannins (Mole and Waterman, 1985; Mole and27Waterman, 1987b; Oh and Hoff, 1986). Therefore tannin inhibits protein digestibilityby depriving enzymes of the substrate in a tannin-protein complex.The tannin-protein complex can form only under suitable pH and charge, in adetergent-free environment, thus showing some environmental selectivity andspecificity to some proteins. The selectivity of tannin through a high affinity forproline rich proteins, the pH, and the surfactants have been exploited by animals tocounter the effects of dietary tannin on the dietary protein. The raised midgut pH insome insect herbivores dissociates the tannin-protein complex, freeing protein fordigestion in the gut (Berenbaum, 1980). Humans, some rodents, domestic goats andsome wild browsers after a period of exposure to tannic diets show some degree oftolerance to tannin due to the production of salivary proline-rich protein (PRPs)(Mehansho et al., 1983; Robbins et al., 1987b). These PRPs bind with tannin leavingthe dietary protein to digestive enzymes (Mehansho et al., 1983). The elevated fecalnitrogen observed in some animals feeding on tanniferous feed could have originatedfrom the PRPs, and do not reflect the apparent digestibility of the dietary protein.The PRP production and other adaptative mechanisms enables animals to thrive ontanniferous feed.The effect of tannin therefore will vary with species, age and experience ofthe animal, the nutrient composition and other secondary metabolites in the feed.Tannin therefore should not be generally regarded as detrimental to animal nutrition. Effect of tannins in the ruminantIn ruminants, tannins have been shown to have both positive and negative influenceon protein digestibility and animal performance. The negative attributes are similar tothose of monogastrics such as reduced apparent digestibility with tanniferous feeds.Robbins et al. (1987a) reported lower digestibility for the high-tannin containingfoliage fed to mule deer compared to low tannin foliage. Increased fecal nitrogen and28reduced DM digestibility were reported in cattle fed on high tannin Sericea lespecleza(Donelly and Antony, 1969). In goats (Nastis and Malechek, 1981) and elk (Robbins,1983; Provenza and Malechek, 1984), the growth rate was not affected, althoughfecal nitrogen was increased by feeding on tanniferous feed. This suggests that theincreased N loss in the feces may be non-dietary in origin since the animalperformance was not affected, but result from increased consumption of tannins. Itprobably originates from endogenous PRPs.There are several positive effects of tannin on protein digestion and animalperformance that have been reported. These include 1) bloat control, 2) production ofgrowth hormone (GH), and 3) protein protection.McArthur et al. (1964) implicated the fraction one (F1) leaf protein found infresh alfalfa and clover as the foam causing agent in the bloat animals. Tanninscontrol bloat through binding this highly soluble protein to form a stable complex inthe rumen (Jones and Mangan, 1977).Plasma GH in sheep was found to increase linearly with varying levels ofcondensed tannin of Lotus pedunculatus  (Barry, 1984). Growth hormone stimulates Nretention and lipolysis, and results in a leaner meat carcass (Barry et al., 1986b).Essential amino acids are needed by dairy cattle to meet the dailymaintenance, growth and lactation requirements. These essential amino acids areabsorbed from the small intestine of the animal. Their sources are dietary protein thatescapes rumen degradation, and the microbial protein. Usually microbial protein willmeet maintenance and a substantial part of production needs. For a high milkproducer and a fast growing heifer, the microbial source alone may not be adequate,and bypass protein is advocated (NRC 1989).Bypass protein is provided by supplementing the animal with low rumendegradable protein such as meat and bone meal or by protecting high quality highlydegradable protein from microbial degradation in the rumen (Orskov, 1982). Heat,29formaldehyde and tannin treatments have been found to have a protective effect onprotein (Nishimuta et al., 1974, Ferguson 1975; Chalupa, 1975). Ferguson (1975)reported increased wool production when sheep were fed on formaldehyde-protectedprotein. The problem with formaldehyde is that it affects the availability of threeessential amino acids, namely, lysine, tyrosine and cystine (Ashes et al., 1984). Heattreatment was found to increase the quantity of amino acids reaching the abomasum(Nishimuta et al., 1984). The problem with heat treatment is timing and balancing ofthe heat to ensure protection of protein without affecting intestinal digestion throughformation of maillard products (Knipfel, 1981).Dietary tannin binds with Fl leaf protein to form a stable tannin-proteincomplex at the rumen pH 5.5-6.8 (Jones and Mangan 1977). The complex so formedis not degradable by the rumen microorganism. Therefore the bound protein bypassesthe rumen to the abomasum and anterior duodenum where the pH is about 2.5. In thisvery acidic medium, the complex dissociates and protein is released for digestion bythe intestinal enzymes (Jones and Mangan, 1977). Subsequent studies, using Lotusspecies and PEG to reduce the effect of tannin, reported reduced ammoniaconcentration in the rumen, increased duodenal N flow and intestinal essential aminoacid (EAA) flux (Egan and Ulyatt, 1980; Barry and Manley, 1984; Barry et al.,1986a; Waghorn et al., 1987; McNabb et al., 1993). Unlike formaldehyde, tanninsdo not affect the availability of EAAs tyrosine and lysine (Waghorn et al. 1987). Thisis one the advantages of using tannin over formaldehyde.In this study it was assumed that the multi-purpose fodder trees and foragecrops have different types and quantities of tannins.It was hypothesized that :a) when these tannins are consumed by ruminant animals, they will bind to form acomplex with dietary protein and protect it from microbial degradation in the rumen.30b) On reaching the duodenum the complex breaks down and proteins are digested inthe small intestine.The effect of tannin on protein degradation in the rumen could therefore bedetermined through use of the mobile nylon bag technique (Arieli et al., 1988; Faldetet al., 1991). In this technique, feed materials are incubated in the rumen for a pre-determined period in small bags, and the same bags are removed, washed andinserted into the intestines through the duodenal cannula to estimates the rumendegradation and intestinal digestibility and the total tract digestibilty.2.3 MATERIALS AND METHODS2.3.1 Sample preparationSeven of the species samples obtained from the high altitude during the wet seasonwere ground enough to pass through a 5 mm sieve in a Wiley mill, and each dividedinto two portions, to be used in periods one and two. The portions were furthersubdivided into two halves. To one of the halves PEG was added at the rate of 40 mgper g of tannin as used by Jones and Mangan (1977). Tannin content was determinedin part one of this study. PEG was added to suppress the effect of tannin.Approximately 60 g of feed and PEG mixture were placed in 200 ml of water in abeaker and mixed thoroughly with a magnetic stirrer. The other half also was placedin water and thoroughly mixed. All the samples were individually freeze-dried.Approximately 1 g of the dried feed sample was weighed into a pre-weighed, labelednylon bag and heat sealed. The bag dimensions were 3.5 x 5.5 cm. Pore size was 40lam, averaged from six samples observed under Jenameci 2 microscope 400Xmagnification.2.3.2 In vivo incubation31The mobile nylon bag technique as described by Arieli et al. (1988) was used in thisexperiment. Four Holstein cows fitted with both ruminal and duodenal cannulas, werefed in individual stalls on a mixture of orchard grass and alfalfa hay for 14 days priorto experimentation in June 1992.The PEG treated samples were placed in different animals from the untreatedsamples. The bags were introduced into large net bags suspended in the rumen, andconnected with a nylon cord to the cap of the rumen cannula. Duplicate samples ofeach forage species were introduced at a time and remained in the rumen for 96, 48,24, 12, 6, 0 hours, except in 24 and 12 hours where four bags at a time were placed.The placements into different cows were delayed for 20 minutes to give allowance forother activities to be done in between placements. The time 0 samples were incubatedat 37° C in a borate-phosphate buffer (pH 6.8) with a gentle shake (in ashaker/incubator) for 30 minutes. All the samples were removed at once and washedin cold tap water for 15 minutes. Except for four samples of each species, from 12and 24 hours (2 each), all the samples were oven-dried at 60° C for three days andweighed to determine the DM of the remains.The two wet bags per species from 12 and 24 hours were further incubated at 37°C for 2 hours in a pepsin solution composed of 2 g pepsin per liter in 0.1 N HC1. Thesamples bags were rinsed again and inserted into the intestine through duodenalcannula of the same cow they had been incubated in. The bags were recovered fromthe feces, washed, oven dried at 60° C for 3 days and weighed for DM determination.The experiment was repeated again in the second period with the same animals,but the treatments were switched so that those animals that received PEG addedforages received forages without PEG and vice-versa. The level of PEG was alsoincreased from 40 mg to 100 mg per g tannin (i.e. from 0.04% to 0.10%) . Thedecision to increase the PEG was due to little response observed in period one, whenPEG was applied at the Jones and Mangan (1977) rate of 40 mg per g tannin, which32had proven effective with sainfoin (Onobrychis viciifolia Scop) tannins. An in vitrobioassay was conducted separately to determine the optimum PEG levels for eachMPT species giving the highest rumen microbial degradation activity over theuntreated samples. 100g PEG per g tannin was found to be effective and thusadopted for period two (Appendix C).2.3.3 Analysis and calculationThe DM of the residue was calculated by subtracting the bag weight from the weightof the dried residues and bag. Residue N was determined using wet oxidation (Microkjeldahl method) described by Parkinson and Allen (1975). The rumen DMdisappearance was calculated by subtracting the residue weight from the initial DMof the sample (determined in part 1). The results of these calculations were fitted tothe non linear regression equation P = a +b(1—e-') (Orskov and McDonald 1979).where p is the nutrient disappearance from the rumen over time t, a is the fraction ofnutrient that rapidly disappears from the rumen, b is the potentially degradableamount of nutrients with time, and c is the fractional rate of degradation of the bfraction. The parameters a, b, and c, were estimated using a SAS program proceduredeveloped by Agriculture Canada, Research Center, Lethbridge. The effectivedegradability was also estimated with the same Lethbridge SAS procedure using thebc^e (-(c+kr)equation p= a +^c+kwhere k is the assumed rumen flow rates (0.04 and 0.06) and to is the lag. Thisequation takes into consideration the lag phase.The intestinal DM and CP disappearance was taken as the difference between thewhole tract digestion and the rumen disappearance.All the statistical comparisons for effective degradation of DM and CP in therumen, and the digestible DM and CP reaching the intestines, were done using thegeneral linear model (least square means) of SAS package release 6.04 (SAS/STAT33software, 1985). These were analyzed as 7x4x2x2 factorial experiments, with thefollowing model= ,u +1-, + + B k + Cm ± 131q +^+ TBik TCnn etikingwhere Y= Effective degradability or digestionA= Mean= Treatment effectA= Animal effectB= Species effectC= Time effectD= period effectTA= Animal x treatment interactionTB= Treatment x species interactions = random errorThe period treatment effect was tested by comparing the controls in both periodsseparately from the treated because of confounding with treatment effect.2.4 RESULTS AND DISCUSSION2.4.1 Dry matter and Crude protein disappearance in the rumenThe effects of tannins on effective degradability of DM at two rumen outflow rates (k) arepresented in Table 4. Tannin significantly (P <0.05) reduced the effective degradability ofDM of all forages in the rumen. Differences in response to PEG treatment ranging from 41%to 200% in DM effective degradability, for gliricidia and calliandra, respectively wereobserved between PEG treated and untreated forages. The differences in effectivedegradability of the DM between species were also significant (P <0.05) (Table 5). CalliandraDM, with effective degradability of 13.8% , was found to be lowly degradable in the rumenfollowed by D. intortum (21.76%) at k= 0.04, while Gliricidia had the highest DMdegradability (46.49%).These DM disappearance results for untreated samples were similar to thosereported by Jones et al. (1992) for Leucaena and Calliandra. The DM disappearancevalues for Leucaena and Gliricidia (untreated) were higher than those obtained byVadiveloo and Fade! (1992), which could probably be attributed to lower phenoliccontent in this study compared to theirs. Sesbania DM disappearance values weresimilar to those reported by Khalili and Varvikko (1992). Except for velvet bean andcassava whose in situ values were not available, the DM disappearance valuescompared well with the values obtained by Van Eyes et al.(1986). These resultscorroborate the findings of Barry and Manley (1986) and Waghorn et al. (1987), thatPEG reduces the effect of tannin on the DM and enhances the degradation of DM inthe rumen.35Table 4. Effect of tannin on mean DM effective degradability in the rumen at assumedrumen outflow rate k=0.04 and 0.06k=0.04 Without PEG (s.e.m)% of initial DMwith PEG (s.e.m)% of initial DM%increase afterPEG additionLeucaena 38.51 ±1.53 63.71 ±1.53 65.54Sesbania 44.79 ±1.53 71.58 ±1.53 59.81Gliricidia 46.49 ±1.54 65.44 ±1.53 40.76Calliandra 13.80 ±1.53 41.51 ±1.53 200.80Velvet bean 38.62 ±1.53 57.92 ±1.53 49.97D. intortum 21.76 ±2.26 55.55 ±1.53 155.28Cassava 44.82 ±1.54 65.07 ±1.53 45.18Meank=0.0635.54a ±0.64 60.11b ±0.57 69.13Leucaena 30.70^±1.53 58.97^±1.54 92.08Sesbania 35.83^±1.53 66.50^±1.54 85.75Gliricidia 39.76^±1.56 60.45^±1.54 52.01Calliandra 9.93^±1.54 37.80^±1.54 281.05velvet bean 32.59^±1.54 53.20^±1.54 63.24D. intortum 15.65^±2.29 51.23^±1.54 227.56Cassava 37.73^±1.56 59.61^±1.54 58.03MEAN 28.88a ±0.64 55.40b ±0.58 91.82Values with different supersripts on a row are significantly different (P <0.05)36Table 5. DM and CP effective degradability in the rumen mean values of differentspecies at k=0.04 and 0.06Speciesk=0.04% Effdgrd DM (s.e.m)k=0.06% Effdgrd DM (s.e.m)Leucaena 51.11c ±1.08 44.83c ±1.09Sesbania 58.18f ±1.08 51.16f ±1.09Gliricidia 55.97e ±1.09 50.10e ±1.10Calliandra 27.66a ±1.08 23.86a ±1.09Velvet bean 48.27c ±1.08 42.89c ±1.09D. intortum 38.65b ±1.37 33.43b ±1.38Cassava 54.94d ±1.09 48•67d ±1.10Effdgrd CP Effdgrd CPLeucaena 72.12c ±1.49 77.18c ±1.42Sesbania 76.37c ±1.49 80.97c ±1.42Gliricidia 76.30c ±1.62 80.76c ±1.55Calliandra 53.05a ±1.62 57.18a ±1.54Velvet bean 72.58c ±1.62 77.21c ±1.55D.intortum 59.2213 ±1.98 64•34b ± 1.89Cassava 72.67c ±1.62 78.23c ±1.55Values with similar superscripts in a column are not significantly different (P <0.05)Effdgrd= effective degradability37Tannin affected the rumen DM degradability constants a, b and c differently asshown in Table 6. Tannin significantly (P <0.05) lowered the rapidly degradablefraction a and increased the time dependent degradable fraction b, and at the same timedecreased the rate c at which b was degraded. These alteration resulted in the low DMeffective degradability shown in (Table 4).Table 6. Effect of tannins on the degradation constants of DM and CPFraction^Without PEG (s.e.m)^With PEG (s.e.m)DMLag^4.6a^±0.50^3.20a ±0.47a 8.51a ±1.41 37.65b ±1.2863•59b ±2.44^42.37a ±2.224.66a ±0.53 6.41b ±0.48CPLag^3.07a ±0.75^4.46a ±0.69a 39.99 ±1.19 60.29b ±1.10^53.37b ±1.57^34.77a ±1.455•90a ±0.63 8.71b ±0.58Values with similar superscripts in a row are not significantly different(P<0.05)Similar low rumen DM disappearances has been reported by other workers ondifferent tanniferous feeds (Chiquette et al., 1988; Mangan, 1988; Kumar andVaithiyanathan, 1990). Tannin inhibits the carbohydrate degradation and activity ofcellulolytic bacteria by binding with structural carbohydrates (Jung, 1985; Barry etal., 1986a). Hence the lower DM degradation values observed this study. In his38studies, Jung (1985) found that the cellulose digestibility potential was not affected bycinammic acid (phenolic), but the rate of degradation was significantly affected.Table 7. Animal variation in effective degradability of the DM at two rumen outflow ratesCOWNo.% of initial DMk=0.04 (s.e.m)% of initial DMk=0.06 (s.e.m)1 58.22c ±0.82 53.47c ±0.832 36.44a ±0.88 29.59a ±0.883 39•65b ±0.88 33.10b ±0.884 56.99c ±0.82 52.38c ±0.83Values with similar superscripts in a column are not significantly different(P<0.05).Table 8. Effect of period on % DM and CP effective degradability at k=0.06period 1 (s.e.m.)^Period 2 (s.e.m.)DM^36.81a ± 0.57^47.46b ± 0.67CP^66•97a ± 0.877083b ± 0.90Values in a row bearing similar superscripts are not significantly different(P <0.05).The effective DM degradabilities were significantly different (P <0.05)between animals (Table 7). Significant animal variation in degradability has beendocumented (Orskov et al., 1980; Lindberg, 1985; Nocek, 1988). Effective DMdegradability differed significantly (P< 0.05) with the period (Table 8). Althoughvariation with period of incubation is known to occur (Orskov et al., 1980), the39variation in this study was attributed to the increase in PEG from 0.04 to 0.1 . Thiswas likely the case because there was no significant difference between controls inperiod one and two (P <0.05) on both DM and CP degradability in the rumen and thetotal tract digestibility.Table 9. The effect of tannin on effective degradability of the CP at assumed rumenoutflow rate k=0.04 and 0.06k=0.04SpeciesWithout PEG (s.e.m) with PEG (s.e.m) %increase afteraddition of PEGLeucaena 71.75 ±2.00 82.63 ±2.00 15.16Sesbania 74.91 ±2.00 87.04 ±2.00 16.19Gliricidia 75.12 ±2.36 86.40 ±2.35 15.00Calliandra 50.01 ±2.00 64.34 ±2.35 28.63Velvet bean 72.65 ±2.36 81.77 ±2.00 12.54D. intortum 53.12 ±2.98 75.57 ±2.35 22.44Cassava 72.41 ±2.36 84.06 ±2.00 16.09MEAN 67.35a ±0.88 80.04b ±0.82 19.34k=0.06Leucaena 65.75 ±2.10 78.50 ±2.10 12.75Sesbania 69.18 ±2.10 83.57 ±2.10 20.80Gliricidia 69.76 ±2.47 82.83 ±2.10 18.75Calliandra 45.54 ±2.10 60.56 ±2.47 32.98Velvet bean 67.62 ±2.47 77.53 ±2.10 14.66D.intortum 46.49 ±3.12 71.96 ±2.47 54.79Cassava 65.67 ±2.476 79.65 ±2.10 21.23MEAN 61.43a ±0.91 76.37b ±0.84 24.32Values with similar superscripts in a row are not significantly different (P < 0.05)40Tannin significantly (P <0.05) reduced the effective degradability of the CP(table 9). The forages treated with PEG had 19.3% and 24.3% higher effectivedegradability than those without PEG, at k=0.04 and 0.06 respectively. Thedifferences between treated and untreated values were likely to be the contribution oftannin. Desmodium intortum had the largest mean effective degradability reduction of54.8 %, followed by calliandra with 33.0% reduction.The effective degradability of CP in different MPT species at different rumenoutflow rates are presented in Table 5 Only calliandra and D. intortum showedsignificant differences (P <0.005) from each other and from the other five species,which had no significant differences among them. Even with the slower outflowrates, calliandra and D. intortum effective degradabilities were less than 65% of theCP. The CP degradability values obtained in this study for Calliandra (45.5% and50.0%) were higher than those of Jones et al.(1992), who reported only 9% Ndisappearance after 48 hours of incubation. However, Leucaena degradability valuescompared well with their findings.The values of Khalili and Varvikko (1992) forSesbania were close but lower than the ones obtained in this experiment which couldbe attributed to the similar phenolic levels in both studies.Tannin reduced the effective degradability of CP in the rumen by binding withprotein, thus significantly (P <0.05) reducing the rapidly degradable soluble fraction a(Table 6), and turns it into the slowly degradable time dependent fraction b. Tanninalso significantly reduced the rate at which fraction b was degraded i.e. the degradationrate constant c. This means that, if the ruminant animal maintains or increases thefeeding frequency and quantities, then higher quantities of protein would be expectedto leave the rumen undegraded. This therefore, results in the low ruminal degradationreported by other workers on other tanniferous forages (Barry and Manley, 1984;Barry et al., 1986a; Robbins et al., 1987b).4 1Effective degradability of CP varied (p <0.05) significantly with animal andperiod. Such variations have been documented in different feeds (Orskov et al., 1980;Nocek, 1988). However, the periods (Table 8) variation in this experiment was to agreat extent due to increase in PEG content in period two.The decrease in effective degradability of the DM observed in this experimentand others (Chiquette et al., 1988; Muellar- Harvey et al., 1988) could also beattributed to a reduction in rumen degradable nitrogen (RDN) for optimal microbialactivity in tannin containing feeds. Adding urea to tanniferous feeds has been found tocater to microbes and improve in vitro DM digestibility (Schaffert et al., 1974). Thissupports the RDN deficiency hypothesis.2.4.2 Intestinal DM and CP digestibilityThe whole gastro-intestinal tract digestibilities of DM and CP are presented inTable 10. Both digestible dry matter (DMD) and digestible crude protein (DCP)differed significantly (P <0.05) with species. Table 11 presents the intestinal drymatter digestibility (IDMD) of different MPT species and forage crops. The IDMDwas obtained by subtracting the percentage rumen dry matter degradation values fromwhole-tract percentage DM digestion and reflects the intestinal flux and digestibility ofthe DM. It also represents the portion of the total tract percentage digestibilitycontributed by intestinal digestion. Three out of seven species namely; calliandra,cassava foliage and velvet bean had significantly lower IDMD than the rest of thespecies.The effect of tannins on the whole tract DMD, simply referred to as DMD arepresented in Table 12. Tannin significantly (P <0.05) reduced the DMD in all thespecies as evidenced by addition of PEG. The highest differences were in calliandrafollowed by D. intortum . These results are consistent to those reported by Nastis and42Malechek (1981), who found decreased DM, cell wall and protein digestibility whentanniferous feeds were added to goat feeds. Period of incubation had a significant(P <0.05) effect on DMD, which was attributed to the increase of PEG in period 2,because the controls in both periods were not significantly different from each other.The effect of tannins on the IDMD are presented in Table 14. Tanninssignificantly (P <0.05) increased the post rumen dry matter digestibility. Cassavafoliage, velvet bean and Sesbania had over 100% increase in IDMD.Table 10. The mean total tract DMD and DCP of different speciesSpecies DMD^(s.e.m)% of the initial DMDCP^(s.e.m)% of initial CPLeucaena 64.65cd ±1.25 92.48cd ±0.85Sesbania 77.23e ±1.23 97.83d ±0.83Gliricidia 73.88e ±1.23 97.70d ±0.83Calliandra 38.43a ±1.25 73.67a ±0.85Velvet bean 61.33c ±1.23 93.31d ±0.83D.intortum 52.14b ±1.25 87.11b ±0.85Cassava 67•38d ±1.23 92.22c ±0.83Values with similar superscripts in a column are not significantly different (P <0.05).43Table 11. Mean propotion of the total tract digestion of the DM contributed by theintestinal DM digestibility(IDMD) of different speciesSpecies IDMD s.e.mLeucaena 14.71b ± 1.41Sesbania 19•86b ± 1.38Gliricidia 17.55b ± 1.38Calliandra 12.17a ± 1.41Velvet bean 13.48a ± 1.38D. intortum 16•82b ± 1.41Cassava 11.81a ± 1.38Values with similar superscripts in a column are not significantly different (P <0.05).Table 12. The effect of tannin on total tract DMD in different speciesSpecies Without PEG(s.e.m)With PEG(s.e.m) %changeLeucaena 54.30a ±1.70 75 . 00b ±1.82 38.12Sesbania 68.57a ±1.76 85•67b ±1.70 24.93Gliricidia 67.04a ±1.70 80•71b ±1.76 20.39Calliandra 25•40a ±1.70 51•83b ±1.82 104.05Velvet bean 55•17a ±1.70 67•49b ±1.76 22.33D. intortum 38.69a ±1.70 65•60b ±1.83 69.55Cassava 59•53a ±1.70 75•23b ±1.76 26.37MEAN 52.62a ±0.65 71•68b ±0.67 36.22Values with similar superscripts in a row are not significantly different (P <0.05).44Table 13 . Effect of period on mean total tract % digestibility of untreated foragePeriod 1^Period 2DMDDCP56•25a^±2.02^60.55 a ±2.0988.84a^±2.09^g5 77a ±2.07Values with similar superscripts on the same row are not significantly different(P <0.05)Calliandra IDMD was not significantly different between PEG treated(control) and untreated, implying either, PEG was not sufficient or could not reversethe effect of tannin on dry matter that has escaped rumen degradation. It could as wellbe that another unknown factor may be contributing to the low IDMD. Jones et al.(1992) found DMD of Calliandra to be about 23% and suspected the low DMD to havebeen caused by the large alummina and silica content in calliandra. Similar DMDvalues for calliandra were reported by Bamualin et al. (1980).Figure 4 presents the proportional contribution of IDMD to the DMD. TheIDMD increased significantly (P <0.05) as tannin level increased except in calliandrawhile the DMD in all the species decreased.The whole digestion tract digestibility values for CP (DCP) are presented inTable 9. There were significant differences between species in DCP. Except forcalliandra and D. intortum which had 73.7% and 87.1%, most species had over 90%DCP. There was a significant interaction between species and treatment, suggestingthat each species behaved differently to different levels of tannin. Alternatively,tannins from different plant species respond differently to PEG. Figure 5 depicts thisinteraction. A look at the figure indicates that calliandra DCP declines sharply as45leucaena, D. intortum, and cassava without PEG had significantly (P <0.05) lowerDCP than with PEG, while in velvet bean, sesbania and gliricidia did not respond toPEG treatment. Period did not have any effect on DCP probably due to compensationby the intestinal digestion of bypass protein.These results are consistent to those reported by other workers (Jones andMangan, 1977; Barry and Manley, 1984; Barry et al., 1986a; Waghorn et al., 1987),who found high tannin to result in a decline of total tract apparent digestibility, butincreased intestinal N.Table 14. The effect of tannins on EDMD of different speciesSpecies without PEG(s.e.m)with PEG (s.e.m) % increase onPEG additionLeucaena 18•90b^±1.92 10.52a^±2.00 79.65Sesbania 27.20b^±1.99 12.53a ±1.92 117.15Gliricidia 22•59b^±1.92 12.51a ±1.99 80.58Calliandra 13.57a^±1.92 10.77a ±2.05 26.00Velvet bean 18•81b^±1.92 8.14a^±1.99 131.08D.intortum 19•31b^±1.92 14.32a ±2.07 34.84Cassava 16•38b^±1.92 7.23a^±1.99 126.56MEAN 19.5410^±1.92 10.86a ±0.76 79.92Values with similar superscripts in a row are not significantly different (P <0.05).46A^BLeu Ses Gil Cal Vel Desi CasWithout PEGMN DMD M IDMI)Figure 4 Proportional contribution of intestine to totaltract DM digestibility.Leu Ses Gli Cal Vel Desi CasWith PEGMI DMD M IDMD1009590858075706560---_----I 1Without PEG^With PEGFigure 5. The interaction effects of species andtreatment on total tract digestion.0 Leucaena, • Sesbania, 7 Gliricidia, V Calliandra,0 Velvet bean, • Desmodium intortum, LI, CassavaThe intestinal digestible protein (IDP) values, which were obtained fromsubtracting the rumen-degradable CP, indicate the portion of the total tract digestibilitycontributed by the intestinal digestion. The IDP reflects the intestinal dietary N fluxand its availability. IDP varied significantly (P <0.05) with species, treatment andtheir interactions (Figure 6). All species behaved differently with increasing tanninlevels. Calliandra IDP decreased at higher tannin levels, while in all other species,higher levels of tannin resulted in varying increases in IDP. This means that Calliandraproteins are bound irreversibly by tannins and are resistant to rumen microbialdegradation as well as intestinal enzymes. In such a case very high fecal nitrogen anddeclining animal performance would be expected (Lohan et al., 1983). This means thatcalliandra protein must be treated in order to enhance its digestibility.In the other species, the results indicated that undegraded dietary proteins thatentered the intestine were digested. Therefore the results of this study agree withJones and Mangan (1977) and Barry and Manley (1984) that presence of tanninincreased the post ruminal protein digestion compared to low or no tannin.Figure 7 indicates the proportional contribution of IDCP to the total tractdigestibility of CP. The IDCP contribution significantly (P <0.05) increases withincrease in tannin content but lowers The DCP although not to the same extent as inthe dry matter.492 '72001 8biJ16C-1a)^1 401 2oJ1 08 6Without PEG^With PEGFigure 6. The interaction effects of fodder speciesand treatments on intestinal protein digestibility.0 Leucaena, • Sesbania, 7 Gliricidtia, V Calliandra,0 Velvet bean, • Desmodium intortum,^Calliandra50P 00 -erc 70 -eHt 60 -agCAVel^Desi^CasB100 --/90 -p 80 -eIc 70 -eHt 60 -agCUfVel^Desi^CasWithout PEG With PEGDCP M IDCP^11111 DCP M 1DCPFigure 7. Proportion of contribution of intestine to totaltract CP digestibility.2.5 SUMMARY AND CONCLUSIONThis study investigated the effect of tannins in multipurpose fodder trees andforage crops of Kenya, on the rumen degradation and the whole GI tract digestibilityof DM and CP.The results indicated varying but significant reduction in DM and CPdegradation in the rumen, with large and varying quantities of the dietary DM andprotein escaping the rumen undegraded depending on the plant species. These thereforeincreased the dietary DM and N flow into the intestine. Except in the case ofcalliandra, a significantly larger proportion of the escaped DM and CP was digested inthe intestines. Generally both DM and CP digestibilities in the total tract were lower inforages without PEG than with PEG treatment. This implied that tannins reduced theapparent digestibility of both DM and CP. These results support Barry et al. 1986afindings that increasing the dietary tannin concentration linearly increases the duodenalN. flow, but linearly decreases the apparent digestibilty of energy and organic matter,and rumen digestion of hemicelluose,but not of cellulose. At the same time this alsoconfirms Jones and Mangan (1977), finding that PEG is an effective suppresser oftannin effects, and is useful for future animal feeding experiments involving tannins.However, there is need to assay the optimum levels of PEG to be applied to eachfodder species.The results also identified two categories of forages in the multipurpose foddertrees and crops. Category one composed of Calliandra calothyrsus  and Desmodium intortum  with less than 55% DMD and less than 90% DCP, which must be treatedbefore feeding, otherwise it may not be beneficial, or even may be deleterious to theanimal. Category two: composed of forages with more than 55% DMD and 90% DCPwhich may be fed untreated or treated. These include leucaena, gliricidia, sesbania,52velvet bean and cassava, whose treatment may be beneficial to animals, especially inthe periods when the tannin:protein ratios are likely to have an adverse effect.Most of these MPT and forage crops are fed as protein supplements to dairycattle especially in periods of feeds shortage, when the bulk of the ration is mainlycomposed of fibrous farm crop residues and poor quality grasses. In most cases thesemay be deleterious to the cattle in that tannin decreases the DM degradation and theenergy available in addition to a protein reduction in digestibility. There is therefore aneed to treat some of these MPT species in order to achieve better animalperformances. In cases where bypass protein is advocated, tannin may increase thebypass protein.The polyphagous tanniferous feeders have developed mechanisms to counterthe effects of tannins in order to thrive. The mechanism, such as use of surfactantswhich dissolves the protein-tannin complex as used by herbivorous insects (Martin etal., 1985; Mole and Waterman, 1985), may be used as a pre-feeding treatment. Thisarea needs further investigation.Tannins have high affinity for proline-rich proteins and PEG, because of theirstrong hydrophobic and hydrogen bonding characters. These characteristicss have beenexploited by tanniferous browsers to suppress the effect of tannins. The PRPs (withlow protein value) are produced in salivary glands of animals adapted to high tanninfeeds. The PRPs strongly and preferentially bind with tannin, allowing the high qualityprotein in the diet to be digested (Hagerman and Butler, 1981; Mehansho et al., 1983).The PRPs and other cheap protein of low nutritive value and with similar charactersneeds further investigation as treatment agents.There is need to study the cultural practices relating to the fodder trees andcrops that may modify the tannin:protein ratio. These may involve the fertilizing andharvesting regimes.53GENERALCONCLUS ON54BIBLIOGRAPHYAkbar, M.A. and P.C. Gupta, 1985. Proximate composition, and tannin mineralcontents of different cultivars and of various plant parts of subabul (Leucaena leucocephala).Indian J. Anim. Sci. 55:808-812.Arlen, A., A. Ben-Moshe, S. Zamwel, and H. Tagari, 1988. In situ evaluation of theruminal digestibility of heat-treated whole cottonseeds. J Dairy Sci. 72:1228-1233.Armstrong, W.D, W.R. Featherston, and Rogler, 1973. Influence of methionine andother dietary additions on the performance of chicks fed bird resistant sorghum grain diets.Poultry Sci. 52:1592-1599.Armstrong, D. G. and T.E.C. Weekes, 1983. Mini-Review. Recent advances inruminant biochemistry: Nitrogen digestion and metabolism. J. Biochem 15:261-266Ashes, J.R., J. Mangan, and G. S. Sidhu, 1984. Nutritional availability of aminoacids from protein cross-linked to protect against degradation in the rumen. Br. J. Nutr.52:239-247.Asquith, T.A, and L. G Butler. 1985. Use of dye-binding protein asspectrophotometric assay for protein precipitants such as tannin. J. Chem. Ecol. 11:1535-1544.Asquith, T.N. J. Uhlig, H. Mehansho, L. Putman, D.M. Carlson, and L. Butler,1987. Binding of condensed tannins to salivary proline-rich glycoproteins: The role ofcarbohydrate. J. Agric. Food Chem. 35:331-334.Baldwin, LT., J.C. Shultz, and D. Ward, 1987. Patterns and sources of leaf tanninvariation in yellow birch (Betula alleghensis) and sugar maple (Acer saccharum). J. Chem.Ecol. 13:1069-1078.55Bamualim, A., R.J. Jones, and R.M. Murray, 1980. Nutritive value of someintroduced tropical browse legumes in the dry season. Proc. Aust. Anim. Prod., 13:229 -232.Barneys, E.A., G.C. Driver and M. Bilgener. 1989. Herbivores and plant tannins.Advances in Ecol. Research 19: 263-302.Barry, T.N. 1984. The role of condensed tannins in the digestion of fresh Lotuspedunculatus by sheep. Can. J. Anim. Sci. 64 (suppl.): 181-182.Barry, T.N., and D.A. Forss, 1983. The condensed tannin content of vegetable Lotuspendunculatus, its regulation by fertilizer application, and effect upon protein solubility. J.Sci. Food Agric. 34:1047-1056.Barry, T.N.and S.J. Duncan, 1984. The role of condensed tannins in the nutritionalvalue of Lotus pedunculatus for sheep. 1. Voluntary intake. Br. J. Nutr. 51:485-491.Barry, T.N., and T.R. Manley, 1984. The role of condensed tannins in the nutritionalvalue of Lotus pedunculatus  for sheep 2. Quantitative digestion of carbohydrates and proteins.Br. J. Nutr. 51: 493-503.Barry T.N and T.R. Manley, 1986. Interrelationships between the concentrations oftotal condensed tannin, free condensed tannin and lignin in Lotus sp. and their possibleconsequences in ruminant nutrition. J. Sci. Food Agric. 37:248-254Barry, T.N., T.R Manley, and S.J. Duncan, 1986a. The role of tannins in thenutritional value of Lotus pedunculatus  for sheep 4. Sites of carbohydrates and proteindigestion as influenced by dietary reactive tannin concentration. Br. J. Nutr. 55:123-137.Barry, T.N., T.F. Allsop, and C. Redekopp, 1986b. The role of condensed tannins inthe nutritional value of Lotus pedunculatus for sheep. 5. Effect on the endocrine system andon adipose tissue metabolism. Br. J. Nutr. 56:607-614.56Bate-Smith, E.C. 1973. Haemoanalysis of tannins: The concept of relative astringency.Phytochemistry 12: 907-912Berenbaum, M. 1980. Adaptive significance of midgut pH in larval lepidoptera. Am.Nat. 115:138-146.Bryant, J.P., T.P. Clausen, P.B. Reicharbt, M.C. McCarthy, and R.A. Werner,1987. Effect of nitrogen fertilization upon the secondary chemistry and nutritional value ofquaking aspen (Populus tremuloides Michx.)leaves for the large aspen tortrix (Choristoneuraconflictana (Walker)). Oecologia 73:513-517.Bullard, R.W., J.O. York, and S.R. Kilburn, 1981. Polyphenolic changes inripening bird-resistant sorghums. J. Agric. Food Chem. 29:973 -981.Burnt, E.A., J.C. Malechek, and F.D. Provenza, 1987. Changes in concentrationsof tannins, total phenolics, crude protein, and in vitro digestibility of browse due tomastication and insalivation by cattle. J. of Range Management 40(5):409-411.Butler, L.G., 1989. Sorghum Polyphenols. Toxicants of plant origin Vol. 4. Ed P.E.Cheeke. pub. CRC. Press Inc. Boca Raton, Florida.Butler, L.G., D.J. Riedl, D.G. Lebryk, and H.J.Blytt. 1984. Interaction of proteinswith sorghum tannin: Mechanism, specificity and significance. J. American Oil chem. Soc.61:916-920.Calderon, P., J. Van Buren, and W. B. Robinson, 1968. Factors influencing theformation of precipitates and hazes by gelatin and condensed and hydrolyzable tannins. J.Agric. Food Chem. 16:479-482.Chalupa, W. 1975. Rumen bypass and protection of proteins and amino acids. J.Dairy. Sci. 58:1198-1219.57Chiquette, J., K.-J. Cheng, J.W. Costerton, and L.P. Milligan, 1988. Effect oftannins on the digestibility of two isosynthetic strains of birdsfoot trefoil (Lotus corniculatusL.) using in vitro and in sacco techniques. Can. J. Anim. Sci. 68:751-760.Clausen, T.P., F.D. Provenza, E.A. Burnt, P.B. Reichardt, and J.P. Bryant,1990. Ecological implication of condensed tannin structure: A case study. J. Chem. Ecol. 16(8):2381-2391.Coley, P.D. 1988. Effects of plants growth rate and leaf lifetime on the amount andtype of anti-herbivore defense. Oecologia 74: 531-536.Cooper, S.M. and N. Owen-Smith, 1985. Condensed tannins deter feeding bybrowsing ruminants in a South African savanna. Oecologia 67:142 - 146.Cooper, S.M., N. Owen-smith, and J.P. Bryant, 1988. Foliage acceptability tobrowsing ruminants in relation to seasonal changes in the leaf chemistry of woody plants in aSouth African savanna. Oecologia 75:336-342.Cousins, B.W., T.D. Tannicley, Jr., D.A. Knabe, and T. Zebrowska, 1981.Nutrient digestibility and performance of pigs fed sorghums varying in tannin concentration.J. Anim. Sci. 53:1524-1536.Donnelly, E.D. and W.B. Anthony, 1969. Relationship of tannin, dry matterdigestibility and crude protein in Sericea lespedeza . Crop Sci. 9:361-362.Egan, A.R., and M.J. Ulyatt, 1980. Quantitative digestion of fresh herbage by sheep6. Utilization of nitrogen in five herbages. J. Agric. Sci., Camb. 94:47-56.Faldet, MA., V.L. Voss, G.A. Broderick, and L.D. Satter, 1991. Chemical, invitro, and in situ evaluation of heat-treated soybean proteins. J. Dairy Sci. 74:2548-2554.Feeny, P.P., and H. Bostock, 1968. Seasonal changes in the tannin content of oakleaves. Phytochem. 7:871-880.58Feeny, P.P., 1976. Plant apparancy and chemical defence. In: Biochemical interactionsbetween plants and insects. (Eds. J.W. Wallace and R.L. Mansell), Plenum Press New York,pp. 1-40.Ferguson, K.A. 1975. The protection of dietary proteins and amino acids againstmicrobial fermentation in the rumen. In: digestion and metabolism in the ruminant ( ed. I.McDonald and A.C.I. Warner) pp448-464. The Univ.of New England, Publ.Unit.Ford, C.W. 1978. In vitro digestibility and chemical composition of three tropicalpasture legumes, Desmodium intortum cv. Greenleaf, D.tortuosum and Macroptiliumatropurpureum cv. siratro. Aust.J. Agric.Res.,29:963-74Forsberg ,C.W. 1978 Some effects of arsenic on the rumen microflora; an in vitrostudy. Can. J. Microbiol. 24: 36-44.Gartlan, J.S., D.B. Mckey, P.G. Waterman, C.N. Mbi, and T.T. Struhsakers,1980. A comparative study of the phytochemistry of two African rain forests. Biochem.Systematics and Ecol. 8:401-422.Glen, J.L., C. C. Kuo, R.C.Durley, and R.P. Pharis, 1972. Use of insolublepolyvinylpyrrolidone for purification plant extracts and chromatography of plant hormones.Phytochem. 11:345-351.Goldstein, J.L., and T. Swain, 1963. Changes in tannins in ripening fruits.Phytochem. 2:371-383.Goldstein, J.L., and T. Swain, 1965. The inhibition of enzymes by tannins.Phytochem. 4:185-192Goering, H.K. and Van Soest, P.J., 1970. Foragge fiber analysis. Agriculture Handbook No379, USDA, Washington, DC, pp. 1-20Gustayson, K.H. 1954. Journal of Polymer science 12:317. Cited by M.N.Mcleod1974, Nutritional Abstracts and reviews 44:803-815.59Hagerman, A.E. and L.G. Butler, 1978. Protein precipitation method for thequantitative determination of tannins. J. Agric. Food Chem. 26:809-812.Hagerman, A.E., and L.G. Butler, 1981. Specificity of proathocyanidin-proteininteraction. J. Biol. Chem. 256:4494-4497Hagerman, A.E., C.T. Robbins, Y. Weerasuriya, T. C. Wilson, and C. McArthur,1992. Tannin chemistry in relation to digestion. J. Range Manage 45:57-62.Haslam, E. 1989. Plant polyphenols:vegetable tannins revisited. Publ. CambridgeUniv. Press Cambridge. pp. 1-187.Horigome, T., R. Kumar, and K. Okomoto„ 1988. Effect of condensed tanninsprepared from leaves of plants on digestive enzymes in in vitro and and intestines of rats. Br.J. Nutr. 60:275-285.Huggan, R.D. and S.B. Westly, 1992. Agroforestry in Kenya. Fitting the piecestogether. Agroforestry Today 1:1-4Jambunathan, R., and E.T. Mertz, 1973. Relationship between tannin levels, ratgrowth, and distribution of proteins in sorghum. J. Agric. Food Chem. 21:692-697.Jones, R.J.,1979. The value of Leucaena leucocephala as a feed for ruminants in the inthe tropics. World Anim. Rev. 31:13-23.Jones, R.J., R.P. LeFeuvre, and M.J. Playne, 1992. Loss of dry matter, nitrogen,minerals and fiber fractions from nylon bags containing Leucaena leucocephala and twocalliandra species in the rumen. Anim. Feed Sci. Tech. 37: 297-307.Jones, W.T., and J.L.Mangan, 1977. Complexes of condensed tannins of sainfoin(Onobrychis viciifolia) with fraction 1 leaf protein and with submaxillary mucoprotein andtheir reversal by polyethylene glycol and pH. J. Sci. Food Agric. 28:126-136.Josylin, M. A. and Z. Glick, 1969. Comparative effects of gallotannic acid and relatedphenolics on the growth of rats. J. Nutr.98:119-126.60Jung, H.G. 1985. Inhibition of structural carbohydrate fermentation by foragephenolics. J. Sci. Food Agric. 36:74-80.Khalili, H., and T. Varvikko, 1992. Effect of replacement of concentrate mix bywilted sesbania (Sesbania sesban) forage on diet digestibility, rumen fermentation and milkproduction in Friesian X Zebu (Boran) crossed cows fed low quality native hay. Anim. FeedSci. Tech. 36:275-286.ICnipfel, J.E., 1981. Nitrogen and energy availabilities in foods and feeds subjected toheating. Prog. Food Sci. 5:177-192.Kossila, V.L.,1984. The location and potential feed use. In straw and other fibrous by-products as feed. Eds. F.Sundstol and E. Owen. Elsevier, Amsterdam. pp 4-22.Kumar, R. and M. Singh, 1984. Tannins: Their adverse role in ruminant Nutrition. J.Agric. Food Chem. 32:447-453.Kumar, R. and S. Vaithiyanathan. 1990. Occurrence, nutritional significance and theeffect on animal productivity of tannin in trees leaves. Anim. Food Sci. Fech. 30:21-38.Lefroy, E.C., P.R. Dann, J.H. Wildin, R.N. Wesley-smith, and A.A. McGowan,1992. Trees and shrubs as sources of fodder in Australia. Agroforestry ystems 20:117-139.Leinmuller, E., H. Steingass, and K. Menke, 1991. Tannins in ruminant Feedstuffs.Anim. Res. Dev. 33:9-62.Lindberg, J.E. 1985. Estimation of rumen degradability of feed proteins with in saccotechnique and various in vitro methods. A Review. Acta. Agric. Scand. Suppl. 25:64-97.Lohan, 0.P., D. Lail, J. Vaid, and S.S. Negi, 1983. Utilization of oak tree (Quercusincana) fodder in cattle ration and fate of leaf tannins in the ruminant system. Indian J. Anim.Sci. 53:1057-1063.Mangan, J.L. 1988. Nutritional effects of tannins in animal feeds. Nutr. Res.Reviews. 1:209-231.61Martin, J.S. and M.M. Martin, 1982. Tannin assays in ecological studies : Lack ofcorrelation between phenolics, proanthocyanidins and protein-precipitating constituents inmature foliage of six oak species. Oecologia (Berl.) 54:205-211.Martin, J.S., and M.M. Martin, 1983. Tannin assay in ecological studies.Precipitation of ribulose-1,5-bisphosphate carboxylase/oxygenase by tannic acid,quebracho,and oak foliage extracts. J. Chem. Eco1.9:285-294.Martin, M.M., D.C.Rockholm, and J.S.Martin, 1985. Effect of surfactants, pH, andcertain cations on precipitation of proteins by tannins. J. Chem. Ecol. 11:485-493.McArthur, J.M., J.E. Miltimore, and M.J. Pratt, 1964. Bloat investigations. Thefoam stabilizing protein of alfalfa. Can. J. Anim. Sci. 44:200-206.McLeod, M.N. 1974. Plants tannins -Their role in forage quality. CommonwealthBureau of Nutrition . Nutr. Abstr. and Revs. 44 (11):803-815.McManus, J.P., K.G. Davis, T.H. Lilley, and E. Haslam, 1981 The association ofproteins with polyphenols. J. Chem. Soc. Commun. 7:309 -311.McNabb, W., G. C. Waghorn, T. Barry, and I. Shelton, 1993. Effects ofcondensed tannins in Lotus pedunculatus on the digestion and plasma metabolism ofmethionine and cystine in sheep. Abstr. 17th International grassland congress. PalmestonNorth, New Zealand 8-12th Feb 1993.Mehansho, H., A. Hagerman, S. Clements, L. Butler, J. Rogler, and D.M.Carlson, 1983. Modulation of proline-rich protein biosynthesis in rat parotid glands bysorghums with high tannin levels. Proc. Natl. Acad. Sci. USA. 80:3948-3952.Mitaru, B.N., R.D. Reichert, and R. Blair, 1984. The binding of dietary protein bysorghum tannins in the digestive tract of pigs. J.Nutr. 114:1787-1796.62Mitjavila, S., C. Lacombe, G. Carrera, and R. Derache, 1977. Tannic acid andoxidized tannic acid on the functional state of rat intestinal epithelium. J. Nutr. 197:2113-2118.Mole, S. and P.G. Waterman, 1985. Stimulatory effects of tannins and cholic acid ontryptic hydrolysis of Proteins:Ecological implication. J. Chem. Ecol. 11:1323-1332.Mole. S, and P.G. Waterman, 1987a. Tannins as antifeedants to mamalianherbivores-still an open quetion?. In: Allelochemicals role in agriculture and forestry. Ed.G.R. Wailer. American Chemical Society, Washington DC. pp572-587.Mole, S. and P.G. Waterman, 198M. Tannic acid and proteolytic enzymes: Enzymeinhibition or substrate deprivation? Phytochem. 26:99-102.Mueller-Harvey, A.B. McAllan, M.K. Theodorou and D.E. Beever, 1988.Phenolics in fibrous crop residues and plants and their effects on the digestion and utilizationof carbohydrates and proteins in ruminants. Plant breeding and the nutritive value of cropresidues conf. Proc. ILCA, Addis Ababa Dec. 1987.Muinga, R. W. , W. Thorpe and J.H. Topps 1992. Voluntary food intake, Live-weight change and lactation performance of crossbred dair cows given ad libitum Pennisetumpurpereum (napier grass var. Bana) supplemented with leucaena forage in the low land semi-humid tropics. Anim. Prod. 55:331-337Myer, R.O., D.W. Gorbet, and G.E. Combs, 1986. Nutritive value of high- andlow-tannin grain sorghums harvested and stored in the high-moisture state for growingfinishing swine. J. Anim. Sci. 62:1290-1297.Nastis, A.S., and J.C. Malechek, 1981. Digestion and utilization of nutrients in oakbrowse by goats. J. Anim. Sci. 53:283-290Nishimuta, J.F., D.G. Ely, and J.A. Boling, 1974. Ruminal bypass of dietarysoybean protein treated with heat, formalin and tannic acid. J. Anim. Sci. 39:952-957.63Nocek, J.E., 1988. In situ and other methods to estimate ruminal protein and energydigestibility. A review. J. dairy Sci. 71:2051-2069.NRC (National Research Council), 1989. Nutrient requirements of dairy cattle.National academy Press Washington, D.C. 1989.Nunez-Hernandez, G., J.D. Wallace, J.L. Holechek, M.L. Galyean, and M.Cadenas, 1991. Condensed tannins and nutrient utilization by lambs and goats fed low qualitydiets. J. Anim. Sci. 69:1167-1177.Oh, ILI., J.E. Hoff, G.S. Armstrong, and L. A. Haff, 1980. Hydrophobicinteraction in tannin-protein complexes. J. Agric. Food Chem. 28:394-398.Oh, H.I., and J.E. Hoff, 1986. Effect of condensed grape tannins on the in vitroactivity of digestive proteases and activation of their zymogens. J. Food Sci. 51:577-580.Oldham, J.D. 1977. Protein degradation in ruminants in relation to protein need.Process Biochem. 12:9-13.Oldham, J. D.,and D.S.Parker, 1981. Metabolism in the high-yielding dairy cow.Process Biochem Dec./Jan. 1980/81 pp. 30-46.Orskov, E.R., 1982. In: protein nutrition in ruminant. pp.19-134. Publ. Academic.Press (London) Ltd.Orskov, E.R. and I. McDonald, 1979. The estimation of protein degradability in therumen from incubation measurements weighted according to rate of passage. J. Agric. Sci.Camb. 92:499-503.Orskov, E.R., F.D Deb Hovell, and F. Mould, 1980. The use of the nylon bagtechnique for the evaluation of feedstuffs. Tropical Animal Production 5:195-213.Parkinson, J.A and S.E. Allen 1975. A wet oxidation procedure suitable for thedetermination of N and mineral nutrients in biological materials. Comm. Soil Sci. Plant Anal.6:1-11.64Provenza, F.D., and J.C. Malechek, 1984. Diet selection by domestic goats inrelation to blackbrush twig chemistry. J. Appl. Ecol. 21:831 -841.Rayudu, G.V.N., R. Kadirvel, P. Vohra, and F.H. Kratzer, 1970. Effect of variousagents in alleviating the toxicity of tannic acid for chickens. Poultry Sci. 49:1323-1326.Reed, J.D. 1986. Relationship among soluble phenolics, insoluble proathocyanidins andfiber in East African browse species. J. Range Management 39:5-9.Reed, J.D., R.E. McDowell, P.J. Van Soest, and P.J. Horvath, 1982. Condensedtannins: A factor limiting the use of cassava forage. J. Sci. Food Agric. 33:213 -220.Reed, J.D., P.J. Harvath, M.S. Allen and P.J. Van Soest, 1985. Gravimetricdetermination of soluble phenolics including tannins from leaves by precipitation withtrivalent Ytterbium. J. Sci. Food Agric. 36:255-261.Reed, J.D., H. Soller, and A. Woodward, 1990. Fodder tree and straw diets forsheep: intake, growth, digestibility and the effects of phenolics on nitrogen utilization. Anim.Feed Sci. Tech. 30:39-50.Robbins, C. T., 1983. Wildlife feeding and nutrition. Academic Press, New York,USA.Robbins, C.T., T.A. Hanley, A.E. Hagerman, 0. Hjeljord, D.L. Baker, C.C.Schwartz, and W.W.Mautz, 1987a. Role of tannins in defending plants against ruminants:reduction in protein availability. Ecology, 68(1):98-107.Robbins, C.T., S. Mole, A.E. Hagerman, and T.A. Hanley, 1987b. Role oftannins in defending plants against ruminants: reduction in dry matter digestion?. Ecology,68(6):1606-1615.Rosen , H. 1957 A modified ninhydrin colorimetric analysis for amino acids. Arc.Biochem. and Biophysics 67: 10-15.65SAS/STAT Software. Version 6.04. General linear model procedure. SAS instituteInc. N. Carolina.Schaffert, R.E., D.L. Oswalt, and J.D. Axtell, 1974. Effect of supplemental proteinon the nutritive value of high and low tannin (Sorghum bicolor L.) moench grain for thegrowing rat. J. Anim. Sci. 39:500-505.Scherr, S. J. 1992. The role of extension in agroforestry development: evidence fromWestern Kenya. Agroforestry systems 18:47-68.Schmid, P.P.S., and W. Feucht, 1986. Angew Bot. 60:365-375. Cited by Leinmulleret al., 1991 . Tannins in ruminant feedstuffs. In: Animal Research and development 33:9-62.Schultz, J.C., P.J. Nothnagle and I.T. Baldwin, 1982. Seasonal and individualvariation in leaf quality of two northern hardwoods tree species. American J. of Bot. 69:753-759.Sell, D.R and J.C. Rogler, 1983. Effect of sorghum grain tannins and dietary proteinon the activity of liver UDP-Glucuronyltransferase. Proc. Soc. Exp. Biol. and Med., 174:93-101Strtmieyer, D.H., and M.J. MalM, 1970. Resistance of extracellular yeast invertaseand other glycoproteins to denaturation by tannins. Biochem. J. 909-914.Swain, T. 1965. The tannins. In: Plant biochemistry (J. Bonner and Varner,(eds)).Academic Press., New York pp.552-580Swain, T. 1979. Tannins and lignins. In A. Rosenthal and D.H. Janzen (eds).Herbivore: their interaction with secondary metabolites. Academic Press New, York.Tangendjaja, B., J.B. Lowry, and R.B.H. Wills, 1986. Changes in mimosine,phenol, protein and fiber content of Leucaena leucocephala leaf during growth anddevelopment. Aust. J. Exp. Agric. 26:315-31766Vadiveloo, J., and J.G. Fadel, 1992. Compositional analyses and rumen digestibilityof selected tropical feeds. Anim. Feed Sci. Tech. 37:265-279.Van Eys, J.E., I.W. Mathius, P. Pongsapan, and W.L.Johnson, 1986. Foliage ofthe tree legumes gliricidia, leucaena and sesbania as supplements to napier grass diets forgrowing goats. J. Agric. Sci. Camb. 107:227-233.Van Hoven, W., 1984. Tannins and digestibility in greater kudu. Can. J. Anim. Sci64:177-178.Waghorn, G.C., M. Ulyatt, A. John, and M.T.Fisher, 1987. The effect ofcondensed tannins on the digestion of amino acids and other nutrients in sheep fed on Lotuscorniculatus. Br. J. Nutr. 57:115-126.Waldern, D.E. 1971. A rapid micro-digestion procedure for neutral and detergent fiberCan. J. Anim. Sci. 51: 67Waterman, P. G. 1992. Roles for secondary metabolites in plants. In, Secondarymetabolites: Their function and evolution pp.255-269. D.J. Chadwick and J. Whelan.(Eds).Publ. Wiley Interscience. Chichester New York.67APPENDIX AThe neutral detergent fiber (NDF) and the acid detergent fiber (ADF) for the MPT andforage species were determined using the Goering and Van Soest method (1970) with amodification by waldern (1971).Table I NDF % in MPT and forages based on DM High altitudeSpecies^Dry season^Wet seasonLeucaena 34.68^39.86Sesbania^41.68 35.48Gliricidia 35.78Calliandra^47.19^40.87Velvet bean 49.05Desmodium intortum^42.46^40.62D. uncinatum^51.88 47.34cassava foliage 36.62^44.34Low altitudeLeucaena^ 28.38S esbania -^37.61Gliricidia^ 38.62Calliandra 27.93Velvet bean^ 42.63Table II. ADF % in the MPT and forages based on the DM68High altitudeSpecies^Dry season^Wet seasonLeucaena 20.26^20.1Sesbania^26.62 18.96Gliricidia 21.09Calliandra^29.79^23.88Velvet bean 33.85D. intortum^30.55^32.82D. uncinatum 43.01 24.99Cassava foliage^22.81^27.13Low altitudeLeucaena^ 25.85Sesbania 26.13Gliricidia^ 24.83Calliandra 21.91Velvet bean^ 29.3469APPENDIX BEffect of tannins on ruminal degradation and intestinal digestibility of protein in differentperiodsTable III. Effect of tannin on effective digestibility degradability of DM in the rumenSpecies^ % of DM with PEG % increase with% of DM withouttanninPEGsuppressionPeriod 1 (40 mgPEG/ lg tannin)Leucaena 43.78a. ±3.64 46.45a ±3.64 6.00Sesbania 51.26a ±3.64 58.08a ±3.64 13.30Gliricidia 45.37a ±2.97 52.08a +3.64 15.30Calliandra 16.86a ±3.64 25•67b +3.64 52.20Velvet bean 45.05a +3.64 46.80a ±3.64 3.90D. intortum 27.10a ±3.64 38.08b ±3.64 40.0Cassava 44.93 a ±2.97 51.98a ±3.64 15.70Period 2 100gPEG/ g tanninLeucaena 33.24a +1.52 80.96b +1.52 143.60Sesbania 38.30a +1.52 85.08b +1.52 122.10Gliricidia 40.95a ±2.16 78.56b +1.52 91.80Calliandra 10.72a ±1.52 57.35b +1.52 435.00Velvet bean 32.18a +1.52 69.05b +1.52 114.60Cassava 35.58a ±2.15 78.161 +1.52 119.70Values with similar superscripts in a row do not differ significantly (P<0.05)70Table IV Effect of tannin on effective degradability of the CPPeriod 1 Without PEG With PEG % ChangeLeucaena 73.65a ±1.96 78.27a ±1.96 6.30Sesbania 81.28a ±1.96 84.11a +1.96 3.50Gliricidia 80.02a ±2.77 82.21a ±1.96 2.70Calliandra 46.59a ±1.96 59.5013 +1.96 27.70Velvet bean 80.21a ±2.77 78.74a ±1.96 -1.80D. intortum 55.40a ±1.96 67.30b ±1.96 21.50Cassava 74.60a ±2.77 78.67a ±1.96 5.50Period 2Leucaena 69.83a ±2.61 86.98b ±2.61 24.60Sesbania 68.54a ±2.61 89.96b ±2.61 31.30Gliricidia 72.18a ±2.61 90.58b ±2.61 25.50Calliandra 53.42a ±2.61 70.57b ±2.61 32.10Velvet bean 68.37a ±2.61 84.79b ±2.61 24.00Cassava 70.80a ±2.61 89.45b ±2.61 26.30Values with similar superscripts in a row do not differ significantly (P<0.05)Table V. Effect of Tannin on degradation constants of DMFraction Without PEG With PEGPeriod 1Lag 3.12 a^±0.80 3•35a ±0.84a 7.45 a^±0.79 11.62b ±0.8360.57a^±1.69 59.60a ±1.775.78a^±0.90 8.63b +0.95Period 2Lag 5•67b^±0.53 3.21a +0.45a 6.38a^±0.28 64.18a ±0.2470.00b^+4.95 24.32a +4.294.17a^±0.51 4.30a +0.44Values with similar superscripts in a row do not differ significantly (P<0.05)Table VI. Effect of tannin on degradation constants of  proteinFraction Without PEG With PEGPeriod 1Lag 1.33a ±0.72 4.04b ±0.60a 38.30a +1.78 45.05b ±1.49b 53.30a ±2.64 49.02a ±2.21c 7.05a ±0.72 11.20b ±0.60Period 2Lag 4.67a +1.14 5.43a ±1.23a 44.89a ±0.52 74.6913 ±0.56b 52•49b +1.42 21.66a +1.58c 5.14a +0.98 7.36a ±1.06Values with similar superscripts in a row do not differ significantly (P<0.05)TableVII . Effect of tannin on total tract digestibility of the DM (DMD)73SpeciesWithout PEG With PEGPeriod 1Leucaena 60.15a ±2.59 63.74a ±2.78Sesbania 71.60a ±2.59 79.26b ±2.78Gliricidia 71.68a ±2.59 73.22a ±2.78Calliandra 27.63a ±2.59 36.10b ±2.78Velvet bean 59.16a ±2.59 57.73a 4_2.78D. intortum 40.16a ±2.59 47.78b ±2.78Cassava 65.57a ±2.59 63.07a ±2.78Period 2Leucaena 48.45a ±1.92 87.12b ±2.07Sesbania 65.37a ±1.92 92.50b ±1.92Gliricidia 62.40a ±1.92 88.93b ±1.92Calliandra 22.44a ±1.92 67.60b -±2.07Velvet bean 51.18a ±1.92 76•81b ±1.92D. intortum 37.21a ±1.92 83•87b ±2.07Cassava 53.48a ±1.92 86.72b ±1.92Values with similar superscripts in a row do not differ significantly (P <0.05)Table VIII. The effect of tannin on the total tract digestibility of CP (DCP).74Species Without PEG With PEGPeriod 1Leucaena 92.8a ±1.53 94•70a ±1.64Sesbania 97.56a ±1.64 98.80a ±1.53Gliricidia 97.91a ±1.53 98.20a ±1.64Calliandra 67.501 ±1.53 79. 89b ±164Velvet bean 94.28a ±1.53 94.70a ±1.53D. intortum 78.33a ±1.53 86.39b ±1.64Cassava 94.15a ±1.53 92.62a ±1.64Period 2Leucaena 86.65a ±1.44 95. 81b ±1.54Sesbania 95.74a ±1.44 99.12a ±1.54Gliricidia 95.86a ±1.44 98.84a ±1.54Calliandra 60.37a ±1.44 86.82b ±1.54Velvet bean 89.48a ±1.44 94.52b ±1.54D. intortum 85.86a ±1.44 97.94b ±1.54Cassava 86.90a ±1.44 95.20b ±1.54Values with similar superscripts in a row do not differ significantly (P <0.05)Table IX. The effect of tannin on intestinal DM digestibility (IDMD)75Species Without PEG With PEGPeriod 1Leucaena 17.88 a ±2.09 12.33a^±2.25Sesbania 17.96a ±2.25 16.34a^±2.09Gliricidia 18.98a ±2.09 14.94a^±2.25Calliandra 11.76a ±2.09 9.93a^±2.25Velvet bean 14.71b ±2.09 7.55a^±2.09D. intortum 14.46a ±2.09 8.86a^±2.25Cassava 13.13b ±2.09 6.19a^±2.25Period 2Leucaena 19.92b ±3.08 8.30a ±3.31Sesbania 35.40b ±3.08 8.72a^±3.08Gliricidia 26.20b ±3.08 10.20a^±3.08Calliandra 15.39a ±3.08 11.46a^±3.31Velvet bean 22.91b ±3.08 8.9a^±3.31D. intortum 24.1V ±3.08 19.63a^±3.31Cassava 19.6313 ±3.08 8.77a^±3.08Values with similar superscripts in a row do not differ significantly (P <0.05)76Table X. The effect of tannin on the intestinal digestibility of crude protein (IDCP)SpeciesWithout PEG With PEGPeriod 1Leucaena 17.49b ±1.47 11.58a^±1.58Sesbania 10.73a ±1.58 9•59a^±1.47Gliricidia 15.92b ±1.47 9.90a^±1.58Calliandra 20.59a ±1.47 21.86a^±1.58Velvet bean 13.43a ±1.47 10.68a^±1.47D. intortum 22.54b ±1.47 15.17a^±1.58Cassava 13.46b ±1.47 8.87a^±1.58Period 2Leucaena 18.20b ±1.98 10.90a^±2.12Sesbania 30.45b ±1.98 9.58a^±1.98Gliricidia 23.53b ±1.98 7.59a^±1.98Calliandra 7.64a ±1.98 19.54b^±2.12Velvet bean 23.90b ±1.98 8.75a^±2.12D. intortum 16.53a ±1.98 12.48a^±2.12Cassava 16.77b ±1.98 4.89a^±1.98Values with similar superscripts in a row do not differ significantly (P <0.05)77The results of individual periods, indicate that; at 40 mg PEG per 1 g tannin (lowPEG) used in period 1 the tannin effect on the DM disappearance in the rumen was notclearly manifested in most MPT and forage species. Calliandra calothyrsus and Desmodiumintortum are the only species that responded to the addition of 40 mg PEG per 1 g tanninThe degradation constants in low PEG behaved differently with those of high PEG(100 mg PEG/1 g tannin). Generally the high PEG enhanced the DM and the, CPdisappearance in the rumen and the total tract digestibility , but reduced the intestinalcontribution of the total DMD and DCP.78APPENDIX CA bio-assay for the optimum level of polyethylene glycol (PEG): that would inhibitthe effect of MPT tannins on proteinWhen protein feeds enter into the rumen or are incubated insacco or in vitro, therumen microbes actively hydrolyse the protein into short chain peptides and amino acidswhich they utilize directly for microbial protein synthesis or deaminate them intoammonia which is then used as nitrogen (N) source for microbial protein synthesis(Armstrong and Weekes 1983).Tannin in feed binds with the dietary protein and forms a tannin protein complexwhich is resistant to microbial enzymes hydrolysis. PEG has been found to suppress theeffect of tannin and in some cases dissociating the formed complex (reversing the effectof tannin) (Jones and Mangan, 1977).The rate and mode of PEG application has been variable (Jones & Mangan, 1977;Barry and Duncan, 1984; Waghorn et al., 1987). In period one of this study, the rateof 40 mg/g tannin as used by Jones and Mangan (1977) on Lotus pedunculatus wasadopted. This rate was found to be less effective on MPT species. Therefore, there wasa need to change this level to most appropriate one for these tropical forages. WhenPEG is added to tanninferous feeds, and incubated, the microbial activity would behigher and more amino acids released than untreated feed indicating a low microbialactivity. The released amino acids can be quantified through ninhydrin method describedby Rosen (1957).Objective of this assay was to determine the PEG levels that would result in highmicrobial activity.79MATERIALS AND METHODSSample Preparation:A 0.1 g samples from each species were drawn and diluted with phosphatesulphur (pH 6.8) to make up a suspension of 100 g/l.Preparation of rumen fluid:Rumen fluid obtained from two rumen cannulated cows maintained on orchardgrass and alfalfa hay mixture was strained on a 4 layer cheese cloth to fill a 2 1 thermos.The strained fluid was then prepared as described by Forsberg (1978), which involvedcentrifuging the rumen fluid at 600 xg for 5 minutes at 30°C to remove the debris andprotozoa.The resulting supernatant was further centrifuged for another 15 minutes at 1600xg at 30°C under CO2 atmosphere. The pellet was reconstituted with phosphate bufferto 600 nil. An aliquot of lml feed suspension was drawn and placed in 10 test tubes.The test tubes were added 1 ml PEG solution with different concentration of PEG viz.0, 50, 100, 150, 200, 250, 300, 400, 500 mg/g tannin.One ml of rumen fluid was added to all test tubes. Two more test tubes wereadded to this set. One of the two had fluid only and the other feed suspension only. Alltwelve were incubated in anaerobic environment at 39° C for 4 hours. these werereplicated 3 times.Immediately after incubation, fermentation was stopped using 1 ml icy cold 10%trichloroacetic acid (TCA) and let to stand on ice for 30 minutes. Tubes were thencentrifuged at maximum (27000 xg) and 1 ml of supernatant drawn and treated asdescribed by Rosen (1957) for absorbance reading.80The absorbance was read using the Shimantzu uv-spectrophotometer at 570 muand at 440 mu for proline and hydroxyproline.Calculations:True amino acids absorbance was obtained by subtracting from individual feedhydrolysate absorbance, the absorbance value for rumen fluid alone and substratesuspension alone (i.e without rumen fluid).ResultsThe microbial activity versus PEG level graphs were plotted (figure I -VII).The optimum PEG level was taken as that activity where additional PEG resultedin little or no increase the microbial activity (or was at asymptote). However, a lowervalue was considered where PEG levels were exceptionally high (or may not bereasonable). The 100 mg PEG was selected because in most species the microbial activitywas high.200 300 400 5000300 400 500Figures I — VII. Percentage microbial activitychange with increase of PEG concentrationin the MPT species. 0 Run 1, • Run2T 100 71 80 L Loucaena60 -4.0 •E-7- -20 - •C....) -40<100100^200^300^400^500II '70 - Sesbania^10 -^•/C^-10-20 -C.7E-z^-40 ^40E^30 [10 r;^0*^-10 -Callia,ndra.4.-y0-.30 --40• &,„100^200Concentration of PEG (mg,/g tannin)32^•20CVII D e s-rno &um, int ortu-m,5k: -10c.-5C r-Civ -ON20 -- 70>-E-> V120 -Velvet BeanE--^10080 760 -o<^:to 720- 1 0 ^100^200^300^400^5000 •T 600 -C.,^CassavaI 500400 7-Cz; 300 -200100-100^100^200^300^100^500100^200^300^400^500Concentration of PEG(mg/g tannin)S3100^200^300^400^500


Citation Scheme:


Citations by CSL (citeproc-js)

Usage Statistics



Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            async >
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:


Related Items