rain”. 4.» 1: I...J.r .32....” .i x -2 :4. . as i. : 3,332...- intizd. u: .143 .. sad ..r..? :r... raga” mum ((95315) llllllllllllllllllll1|Hll||||ll|lllllll 3 1293 01413 2363 This is to certify that the dissertation entitled ISOLATION AND CHARACTERIZATION OF RIMINAL HICROORGANISHS ABLE TO DEGRADE CHITINOUS NASTE presented by MARIO ANTONIO COBOS-PERALTA has been accepted towards fulfillment of the requirements for Ph.D. degree in Animal Science / Majory essor Date 12-7-95 MSU is an Affirmative Action/Equal Opportunity Institution 0-12771 LIBRARY MIchigan State University PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES Mum on or More data duo. DATE DUE DATE DUE DATE DUE T’T‘T’j MSU Is An Nflmottvo Mon/Emu Opportunlty Intuition Wm: —_________————— ISOLATION AND CHARACTERIZATION OF RUMINAL MICROORGANISMS ABLE TO DEGRADE CHITINOUS WASTE By Mario Antonio Cobos-Peralta A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Animal Science 1 994 ABSTRACT ISOLATION AND CHARACTERIZATION OF RUMINAL MICROORGANISMS ABLE TO DEGRADE CHITINOUS WASTE By Mario Antonio Cobos-Peralta Chitin rich waste by-products generated by the seafood industry, such as crab shell meal and shrimp carapaces, contain considerable nutritive value and are low in cost, which have justified exploring their potential use as a feed for ruminants. Literally nothing is known abbut the ruminal microorganisms directly involved in degradation Of chitin. The main objective of this research, was to generate basic information about these ruminal microorganisms. A rapid fluorescent assay for detection of Chitinolytic activity was validated for anaerobic conditions. Using this method, it was demonstrated that certain bacteria from the rumen of dairy cows, do possess chitobiase and chitinase activity, which are necessary for chitin degradation. Using an anaerobic selective prereduced medium with purified chitin as the sole energy source, a Chitinolytic bacteria consortia from ruminal fluid of fistulated dairy cows was maintained. A strictly anaerobic spore-forming bacterium was isolated from this Chitinolytic consortia. This spore-forming bacterium was identified as Clostn'dium paraputn‘flcums var. ruminantium. The bacterium degrades a limited number of substrates including: Chitin, crab Shell meal, chitobiase, N-acetylglucosamine and glucose. The fermentation end products of chitin degradation were acetate, lactate and butyrate. Some optimal conditions for its growth and chitinolytic activity were: pH, between pH 6.5 and 7.0; Ca” concentration, between 2 to 6 mM and presence of yeast extract (0.2 g) in the chitin medium. Between a pH range of 6.5 to 7.0, the bacteria doubled its mass in 13.2 :t 2.4 h, and the percent of pure Chitin dry matter digestion averaged 81.13 1 0.33% after 72 h of incubation at 39°C. Glucose and N-acetylglucosamine at low concentration enhanced bacterial growth, but at high concentration adversely affected growth and bacteria activity. Scanning electron microscopy photomicrographs of shrimp carapaces inoculated with a pure culture of Clostridium paraputrificum var. ruminantium revealed that this bacterium, attachs and colonizes the inner surface of the shrimp carapace. Similar results were Obtained when shrimp carapaces were incubated in the rumen of fistulated dairy cows, but in this case a bacterial community was Observed. Considering the efficiency of chitin degradation exhibited by C. paraputn’ficum. var. ruminantium, it is proposed that this bacterium may be useful as a ruminal inoculum to increase the potential use of shellfish wastes as a feed for ruminants. ACKNOWLEDGEMENTS I wish to express my gratitude to Dr. Melvin T. Yokoyama, my major professor for his guidance, support, and friendship. My appreciation is also expressed to the members of my advisory committee: Dr. Roy S. Emery, Dr. Stanley L. Flegler, Dr. Rawle Hollingsworth and Dr. Steven Rust, for their valuable contributions to this study. My gratitude is extended to Dr. John A. Breznak and Dr. Julius H. Jackson, from the Department of MicrobiOIOQY; Dr. Susan Lowe, from the Department of Biochemistry; Dr. Karen Chou, from the Department of Animal Science; Dr. Karen L. Klomparens and Dr. John W. Heckman from the Electron Optics Center; for their assistance and advice during my stay at Michigan State University. I also want to thank Dr. Fredric M. Husby form the University of Alaska (Fairbanks), for supplying the crab shell meal used in this study. I would like to acknowledge the "Consejo Nacional de Ciencia y Tecnologia" (CONACYT), from México for awarding me a fellowship for the support of my graduate studies; the Office for International Students and Scholars for its support at the end of my program and the Colegio de Postgraduados, especially the Director and friends of the Centro de Ganaderia for their consideration and patience in waiting for my return. I give special thanks to my friends: Dr. Susan Hengemuehle for her helpful advice in different laboratory techniques; Tadd Dawson for his unselfish assistance with the chromatography study, Abner Rodriguez for his important collaboration with the in vitro study, and Dr. Telmo B. Oleas for his very helpful suggestions during my stay at the Department of Animal Science. Finally, I wish to express my deep gratitude to my wife, Araceli Cobos, and my daughters, Catalina and Jessica. Their love and patience made this study possible. TABLE OF CONTENTS LIST OF TABLES .......................................... XI LIST OF FIGURES ........................................ XIII INTRODUCTION ........................................... 1 CHAPTER 1 Literature review ........................................... 3 Potential use of chitinous waste as a food for animals. ........... 8 Pathways of chitin degradation. .......................... 12 Chitinolytic microorganisms. ............................. 15 Bacterial attachment to the substrate. ...................... 17 Assay of chitinase activity. .............................. 18 Literature cited ...................................... 22 CHAPTER 2 Use of 4- methylumbelliferyl chitin-substrates for detection of chitinolytic activity in ruminal microorganisms. .................................. 27 vi Abstract ........................................... 28 Introduction ......................................... 29 Materials and methods ................................. 31 Technique of rumen fluid sampling ................... 31 4-MUF-Chitin stock solutions. ....................... 31 Methodology .................................... 32 Results ............................................ 35 Discussion ......................................... 37 Literature cited ...................................... 40 CHAPTER 3 Isolation and characterization of ruminal bacteria which degrade chitin . . . 42 Abstract ........................................... 43 Introduction ......................................... 44 Material and methods ................................. 45 Enrichment and roll tube media ..................... 45 Animals. ...................................... 45 Sampling and inoculation procedure .................. 47 Isolation and purification ........................... 47 Electron microscopy .............................. 47 Primary identification of rumen bacteria isolated .......... 48 4-Methylumbelliferyl Chitin substrates test .............. 48 vii Characterization of Chitinolytic microorganisms ........... 48 Effect of clarified ruminal liquid on chitinolytic activity ...... 49 Results ............................................ 50 Ruminal bacteria isolated .......................... 50 Results Of the 4-methylumbelliferyl test ................ 53 Chitinolytic strain identification ...................... 53 Discussion ......................................... 57 Literature cited ...................................... 60 CHAPTER 4 Clostn'dium paraputn'flcum var. ruminantium : colonization and degradation of shrimp carapaces in vitro observed by scanning electron microscopy . . . . 62 Abstract ........................................... 63 Introduction ......................................... 64 Materials and methods ................................. 65 Organism and culture conditions ..................... 65 Shrimp carapace preparation for colonization study ....... 65 Scanning electron microscopy ...................... 66 Chitin determination .............................. 66 Results ............................................ 68 Discussion ......................................... 75 Literature cited ...................................... 78 CHAPTER 5 Ruminal bacteria colonization and degradation of shrimp carapaces in situ Observed by scanning electron microscopy ....................... 80 Abstract ........................................... 81 Introduction ......................................... 83 Materials and methods ................................. 84 Shrimp carapace preparation for in situ study ........... 84 In situ technique ................................ 85 Scanning electron microscopy ...................... 85 Results ............................................ 87 SEM of shrimp carapace between 3 to 12 h of ruminal incubafion ..................................... 87 SEM Of shrimp carapace after 1 to 2 d of ruminal incubation 88 SEM of shrimp carapace after 3 to 7 d of ruminal incubation 90 Discussion ......................................... 95 Literature Cited ...................................... 99 CHAPTER 6 Comparison of in vitro degradability of crab shell, pure chitin and alfalfa hay by Clostridium paraputn'ficum var. ruminantium and by mixed bacteria from the rumen Of dairy cattle ....................................... 101 Abstract ........................................... 1 02 Introduction ......................................... 103 Materials and methods ................................. 104 Results ............................................ 106 Discussion ......................................... 112 Literature cited ...................................... 116 CHAPTER 7 Influence of physicochemical factors on the rate of growth and efficiency of chitin hydrolysis by C. paraputrificum var. ruminantium .............. 118 Abstract ........................................... 119 Introduction ......................................... 120 Materials and methods ................................. 121 Results ............................................ 123 Discussion ......................................... 132 Literature cited ...................................... 136 CHAPTER 8 Summary ............................................... 139 BIBUOGRAPHY .......................................... 145 LIST OF TABLES CHAPTER 1 1. Chemical composition of different shellfish wastes (D.M. basis) ....... 9 CHAPTER 2 1. Composition of the CH-medium and GCS-RF roll tube medium . . . . 34 2. Results of MUF-chitin substrates assay with different bacteria inocula 36 CHAPTER 3 1. Composition of the chitin and GCS-RF media used for the isolation and characterization of ruminal chitinolytic bacteria ..... 46 2. Characteristics of the C. paraputrificum isolated from the rumen, as interpreted from the RapID ANA II System results .......... 54 3. Summary of substrates used by Clostn'dium paraputrificum and Peptostreptococcus Sp. isolated from the rumen Of dairy cows . . . . 56 CHAPTER 4 1. Composition of the shrimp carapace anaerobic medium used for observing bacteria attachment and degradation of Chitin ......... 68 xi CHAPTER 6 1. Effect of inocula type on in vitro organic matter degradation of chitin, over time ...................................... 108 2. Effect Of inocula type on in vitro organic matter degradation of crab shell meal, over time .............................. 109 3. Effect of inocula type on in vitro organic matter degradation of alfalfa hay, over time .................................. 110 4. Effect of inocula type on in vitro organic matter degradation of Chitin, crab shell meal and alfalfa hay ..................... 111 CHAPTER 7 1. Composition of the chitin media used to study the effects of pH on growth and chitinolytic activity ......................... 123 xii LIST OF FIGURES CHAPTER 1 1. The Chemical structure of the repeating sugar residues of chitin, chitosan and cellulose .................................. 6 2. Chitin degradation. Chitinolytic and chitosan pathway .......... 14 CHAPTER 3 1. Scanning electron micrographs of the main ruminal bacteria with sustained growth in chitin medium ........................ 52 CHAPTER 4 1. Structural characteristics of the shrimp carapace at its outer surface, inner surface and a lateral view .......................... 72 2. Series of scanning electron micrographs of shrimp carapace incubated with C. paraputn'ficum var. ruminantium ............. 73 3. Scanning electron micrographs of shrimp carapace incubated with C. paraputrificum var. ruminantium representing the major events that takes place between 24 and 72 h of incubation ............... 74 xiii CHAPTER 5 1. Series of scanning electron micrographs of shrimp carapaces after 3 to 48 h of incubation in the rumen ....................... 89 2. Scanning electron micrographs of shrimp carapace incubated in the rumen, representing the major events that takes place between 3 to 7 d of incubation ..................................... 93 3. Series of scanning electron micrographs of the shrimp carapaces showing their degree of colonization and degradation Of the outer surface after 12 to 96 h of incubation ...................... 94 CHAPTER 6 1. Effect of inocula type on in vitro organic matter degradation of chitin ............................................. 108 2. Effect of inocula type on in vitro organic matter degradation of crab shell meal .......................................... 109 3. Effect of inocula type on in vitro organic matter degradation of alfalfa hay ............................................... 110 CHAPTER 7 1a. Effect of pH on growth of C. paraputn'ficum var. ruminantium ..... 125 1b. Effect Of pH on the extent of chitin digestion after 72 h of incubation .......................................... 125 xiv 2a. 2b. 3a. 3b. 4a. 4b. 5a. 5b. Effect Of glucose on growth of C. paraputrr'ficum var. ruminantium . . 127 Effect of glucose on the extent of chitin digestion after 72 h of incubafion .......................................... 127 Effect Of N-acetylglucosamine on growth of C. paraputriflcum var. ruminantium ........................................ 128 Effect of N-acetylglucosamine (NAG) on the extent of chitin digestion after 72 h of incubation ................................ 128 Effect of calcium concentration (Mm) on growth of C. paraputriflcum var. ruminantium ..................................... 130 Effect of calcium on the extent of chitin digestion after 72 h of incubafion .......................................... 130 Effect of deleting yeast extract (YE) or trypticase (Try) from the CH- medium on growth of C. paraputn'flcum var. ruminantium ........ 131 Effect of deleting yeast extract or trypticase (Try) on the extent of chitin digestion after 72 h of incubation ..................... 131 XV INTRODUCTION Shellfish waste, is the generic name used to describe the by-products generated by the fishing industry after meat extraction from marine crustaceans such as crab, shrimp, lobster and prawn, and freshwater crustaceans such as crawfish. Shellfish waste consists of shells, viscera and unextracted meat. The shells of some marine crustaceans are processed (mainly in the USA and Japan), for the extraction of the valuable biopolymer chitin, and the production of its deacetylated derivative, chitosan. However, most of the shellfish waste, which approximates 85% of the fresh catch, is largely disposed Of by its return to the environment either on landfill sites or by direct return to the sea. This presents a waste disposal problem. In 1972, the Environmental Protection Agency (EPA) Classified Shellfish processing waste as a marine environmental pollution problem. A potential use of large quantities of shellfish waste is as a feed for livestock. For example, crab shell meal contains in excess of 40% crude protein and its energy and mineral values are also considerable. The main limitation of shellfish wastes as a feedstuff is its chitin content which can not be efficiently digested by most productive animals. However, ruminant animals, because of the symbiotic microorganisms living in their rumen, are considered better adapted than 2 nonruminants to assimilate the nutrients contained in shellfish waste. Previous research has demonstrated the potential Of chitinous waste as a feed for ruminants which implies that some ruminal microorganisms can degrade Chitin. However, the ruminal microorganisms involved in chitin degradation have never been isolated and identified. The working hypothesis of this research is that the degradation of chitin rich substrates in the rumen is performed by selected number of ruminal bacteria. With the general Objective to test this hypothesis as well as to generate basic information about these ruminal microorganisms, the objectives of the present research thesis were: 1. To validate a fluorescent test for rapid identification Of chitinolytic activity in ruminal microorganisms (Chapter 2). 2. To isolate and characterize ruminal microorganisms which degrade chitin and shellfish waste (Chapter 3). 3. To determine the mode in which ruminal chitinolytic microorganisms colonize and degrade chitin-rich substrates (Chapter 4 and 5). 4. To compare the chitinolytic activity of the ruminal chitinolytic bacteria isolated with mixed bacteria from the rumen of dairy cow (Chapter 6). 5. To determine the effect of some physicochemical factors on the growth and chitinolytic activity of the ruminal bacterium isolated (Chapter 7). CHAPTER 1 LITERATURE REVIEW 4 Chitin is one of the most abundant polysaccharides in the biosphere, probably only second to cellulose (Pei, 1989). A recent estimate for the annual production Of chitin in the world is between 10‘° to 1011 metric tons (Gooday et al. 1991). Chitin is found in very diverse taxonomic groups. With few exceptions, Chitin is ubiquitous in the fungi: in cell walls of hyphochytridiomycetes, chytridiomycetes, zygomycetes, ascomycetes, basidiomycetes, and deuteromycetes (Gay et al., 1993; Bartinicki and Lippman, 1982). Chitin occurs sporadically among the Protista: in cyst walls of some ciliates, flagellates and amoebas (Mulish et al., 1993; Arroyo and Carabez, 1982). It is also very prevalent in the marine environment, where much of its production is in the form of carapaces of zooplankton (e.g. krill, Herwing et al., 1988) and invertebrates, particularly in the exoskeletons and perithropic membranes of arthropods where chitin comprises 58 to 85 % of the dry weight (Pel, 1989). Chemically, chitin is a linear amino-sugar homopolymer composed of 2- acetamidO-2-deoxy-r3-o-glucose (i.e. N-acetyl glucosamine) residues with l3-1-4 linkages (Fig. 1.1). This polysaccharide is insoluble in water because Of the extensive intermolecular hydrogen bonding between adjacent Chains, which excludes all water from the interior of crystalline structures present in the polymer fibers (Hackman, 1987). Chitin occurs in three polymorphic forms which differ in the arrangement of the adjacent chains. a-Chitin has antiparallel chains, B-Chitin has parallel Chains and y-chitin which has two Chains "up” to to every one "down" (Blackwell, 1988). a-Chitin is by far the most common form, which is found in fungi, protistans and invertebrate exoskeletons (Gooday, 1990a). 5 In invertebrates, chitin is always found associated with other structural components such as proteins, specifically arthropodin and sclerotin with both covalent and noncovalent bonding (Gooday, 19903). The green color of many insects and the red color of crustacean shells is due to the linkage of carotenoids with chitin amino groups (Muzzarelli, 1977) and with minerals, mainly with calcium carbonate. Another modification of chitin is its deacetylation to chitosan. The monomer Ofchitosan is 2-amino-2-deoxy-r3-D-glucose (i.e. D-glucosamine) with 8-1— 4 linkages (Fig. 1). Chitosan is the naturally occurring structural component of the cell wall of Mucor rouxii (Davis and Bartnicki-Garcia, 1984). In some yeast, such as Saccharomyces cerevisae, chitosan is the major component of ascospore walls. In arthropods, chitosan has been found in some insects as part of their flexible structures (Gooday, 1990a). It should be mentioned that chitosan rather than chitin is the polymer that has the greatest potential for a wide range of industrial uses due to its polycationic nature (Kienzle et al., 1984). CHWWI CHITOSAN CELLULOSE Fig. 1. The Chemical structures of the repiting sugar residues of chitin, chitosan and cellulose. 7 Chitin is a complex molecule, which is insoluble in water, dilute alkalis, dilute mineral acids and all common organic solvents (Hackman, 1987). Because of its unique Chemical characteristics, plus the availability of large amounts of pure chitin extracted from shellfish waste and estimated as 1.2 x 10'5 tons annually on a worldwide basis (Knorr, 1991), there is considerable interest in research on the possible applications of this material and its derivatives. Actually, novel applications of chitin, chitosan and related enzymes have been reported in the areas of animal feeding, control of pathogenic soil and plant microorganisms, drug delivery, biomedical materials, blood anticoagulants, chelation of metals, pollution control and functional membranes. An extensive review Of these topics can be found in Muzzarelli et al. (1986); Zikakis (1984) and Wright and Retnakaran (1987) and Skjak-Braek, et al. (1989). Presently, chitin is obtained from shrimp carapaces, because of their availability in high amounts from the shrimp fishing activities in Texas and Mexico (Muzzarelli, 1977), but crab shell (Husby et al., 1981) and crawfish (NC, at al., 1989) are also potential sources. These sources are generically referred to as Shellfish wastes or chitinous wastes. However, more easily extracted and purer sources of Chitin have been recently identified. The diatoms Thalassiosira fluviatilis (weissflogii) and Cylotella cryptica produce a unique form of chitin called "chitan" which is highly crystalline, not cross-linked with amino acids or other glycans, and is not complexed with calcium, magnesium, or other cations (Smucker, 1991). In the future, it is possible that the demand for pure chitin and derivatives for medicine, surgery and dentistry, will be supplied by marine 8 plankton and fungi (Shimara et al., 1988). Therefore, the potential use of shellfish wastes will be focused on other practical areas, such as animal feeding. Potential use of chitinous waste as a food for animals. Because of the similarity between the molecular structure of chitin and cellulose, which differ only in the substitution of an acetylamine group for the hydroxyl group on carbon-two of the glucose units; by-products high in chitin such as shellfish waste, are considered a potential source of energy for ruminants. In addition, its nitrogen content (6.89%) which is equivalent to 43.06% of crude protein, together with its high mineral content in the case of crab shell, make shellfish waste not only a potential source of energy, but also of protein and minerals. In Table 1, is shown the main chemical components of different shelfish wastes as reported by different researchers. However, some variations in the concentration of the main chemical components, can be expected, due to the process of meat extraction and species variety. Table 1. Chemical composition Of different shellfish wastes (D.M. basis). — Item Shellfish waste Crab1 Crawfish2 Shrimp‘ peenngs Proximate composition, % Dry matter 96.3 94.3 95.7 Crude protein 32.4 35.5 37.9 Chitin 20.2 15.3 21.7 Ash 44.6 42.1 40.8 Ether extract - - - - - - - - - Mineral composition, % Calcium 22.4 13.1 24.9 Phosphorus 2.0 0.8 2.51 I Husby, et al., 1951. 2 Velez et al., 1991. Species belonging to the order Insectivora and Cheiroptera (bats), are able to digest Chitin or chitinous compounds due to the production of chitin degrading enzymes synthesized and secreted by their gastric mucosa (Jeuniaux, 1993). However, in herbivorous species, chitinases are usually lacking in their digestive system or if the enzymes are present (e.g. abomasum, pancreas and kidney of cow and goat), they are considered unimportant because of their low concentration or non-inducible characteristic (Lu'mbland et al. 1974). In theory, chitinous wastes have potential as a feed for ruminants such as cattle, Sheep and goats, because of the ruminant's symbiotic relationship with the microorganisms which inhabit the rumen. The hypothesis is that some ruminal 1O bacteria species are able to utilize chitin as a substrate. The end products of chitin degradation and also the chitinolytic bacteria themselves, will be subjected to extensive digestion within the alimentary tract Of ruminants. Thus, ruminal microorganisms, enable ruminants to utilize chitinous wastes as a source of energy and nitrogen. Patton and Chandler (1975), evaluated various chitinous sources such as cockroaches, grasshoppers, shrimp meal, crab meal and purified chitin by in vivo ruminal techniques. They found that all the sources showed an increased solubility in the rumen of sufficient magnitude to warrant speculation of true chitin digestion. In a subsequent experiment, Patton et al.(1975) reported a successful metabolic trial in which 10 and 20% crab meal was substituted for soybean meal in a basal ration comprised of hay, corn and soybean meal. The diet was formulated to satisfy the nutritional requirements of rapidly growing calves. Chitin digestibility for the crab meal was measured as 66% but was highly variable from 26 tO 87%. Body weight gain, feed intake, feed efficiency and dry matter digestibility, were not significantly different for the basal and crab meal rations. They concluded that crab meal is a potential energy and protein source. A different result was obtained by Ortega and Church (1979) in a study in which crab meal, pure chitin, chitosan, N-acetyl glucosamine and D-glucosamine were evaluated by in vitro rumen digestion trials. They reported that N-acetyl glucosamine and D-glucosamine are utilized as a nutrients for ruminal bacteria, but crab meal, chitin and chitosan are used to a limited extent. Brundge et al. (1981 and 1984), reported that king crab (Paralithodes 11 cantschatica) meal, and tanner crab (Chionoecetes bairdi) meal, are acceptable protein supplements in concentrates for lactating dairy cattle, when fed at up to 22.5% of the ration, with no negative effects on milk production or flavor. However, crab meal inclusion at levels of 30% of the ration, resulted not only in a decrease in feed intake but also a decrease in body weight. The negative effects on animal performance when crab meal is fed at high levels has been related to low palatability (Brundage et al, 1981 and 1984) and increased mineral levels (Patton et al, 1975); but Laflame (1988), found that the negative effect on feed intake and performance of beef calves fed a ration with 35% crab meal added to the grain ration was eliminated after a 1 to 2 month period Of adaptation. A proper adaptation period may be a key factor related to an animal's response to the intake of chitinous compounds. A good example of adaptation period effect, is given by Velez et al. (1991), who reported negative results when he evaluated crab and crawfish waste meals as protein sources for growing dairy heifers. The animals in their experiment were fed with a reasonable percent of crab shell meal (15%) in the ration, but their respective experimental diets were offered for only a 1 to 2 weeks adjustment period prior to data collection. The key question is, who requires this adaptation period; the ruminant or the ruminal microorganisms? Since the enzyme mainly responsible for chitin degradation (e.g. chitinase) has been characterized as non-inducible and scarce in ruminant tissues (Lumbland et al. 1974), it is reasonable to speculate that it is not the ruminant but the ruminal bacteria which requires the adaptation period to efficiently utilize chitinous compounds. 12 Previous research conducted with ruminants and chitinous compounds, rely on the assumption that certain unidentified ruminal microorganism are able to degrade chitin. NO one has reported which ruminal microorganisms (bacteria, fungi or protozoa) are involved in chitin degradation and their growth Characteristics. The study of these basic areas, will provide a better understanding of the potential and limits of chitinous waste as a feed for ruminants. Pathways of Chitin degradation. An organism must excrete an extracellular enzyme to solubilize and shorten the chitin molecule so that breakdown products can enter the cell to be metabolized. There are two documented pathways of chitin degradation, the chitinolytic and the Chitosan pathways (see, Fig.1.2). The best-studied is the chitinolytic pathway which is also considered the most important. In this system, the complete enzymatic hydrolysis of the glycosidic bonds in chitin to free N-acetyl-D-glucosamine is accomplished by two hydrolases. First, the enzyme chitinase (EC 3.2.1.14, poly-B-1,4--(2-acetamido-2-deoxy)-D- glucoside glycanohydrolase) which may be present as either an exo- or endo- chitinase or both. Exo-chitinase cleaves diacetylchitobiose units from the nonreducing end of the chitin chain. Endo-chitinase cleaves glycosidic linkages randomly along the chitin chain, including tetramers and to a lesser extent trimers (Jeuniaux, 1966), eventually giving diacetylchitobiose as the major product, together with some triacetylchitotriose. The diacetylchitobiose produced, is then hydrolysed by Ill-N-acetylglucosaminidases(EC 3.2.1.30 B-2-acetamido-2-deoxy-D- glucoside acetamidoxyglucohydrolase) which hydrolyses terminal, non-reducing 2- acetamido-2-deoxy-l3-D-glucose residues in chitobiase and chitotriose. Some B-N- 13 acetyl glucosaminidases can also act weakly as exochitinases, cleaving monosaccharide units from the non-reducing ends of chitin chains (Gooday, 1990b). The Chitinolytic system is considered ubiquitous and has been reported in aerobic bacteria (Roberts and Cabib, 1982), anaerobic bacteria (Mountfort and Rhodes, 1991; Pel et al., 1990), fungi (Reyes and Martinez, 1988) and even plant cells (Verburg and Huynh, 1991 ). Although the same enzymatic system has been isolated from different organisms, the chitinolytic enzymes may possess different characteristics. For example, the molecular weight of the enzymes ranges between 30,000 to 50,000 units (Cabib, 1987); the isoelectric point can be either acidic, 3.7(Hara et al., 1989) or basic, 8.7 (Verburg and Huynh, 1990) and optimal activity occurs between pH 5.0 and 7.6 (Hara et al., 1989 and Pedraza and LOpez, 1991 respectively). More recently, an alternative chitinolytic system has been reported; the chitosan pathway. This system consists of three enzymes. Chitin deacetylase, first deacetylates chitin to give chitosan, which in turn can be hydrolyzed to chitobiose by action of the enzyme chitosanase (Gooday et al., 1991; Trudel and Asselin, 1990). The enzyme glucosaminidase then hydrolyses chitobiase to glucosamine. The end products of chitin hydrolysis (e.g. N-acetylglucosamine and glucosamine) can be transported into the cells. Further metabolism of N-acetylglucosamine may be via phosphorylation to N-acetylglucosamine-6-phosphate, or via deacetylation to glucosamine, and then deamination (Gooday, 1990b). The importance of the chitosan pathway under both aerobic and anaerobic conditions is still unknown. 14 CHITIN Chitinase(s) Oligomers V N-acetylchitobiose N-acetylglucosaminidase \f‘ V N-acetylglucosamine Chitin deacetylase r21 Chitosan Chitosanase ,___/ V Chitobiose Glucosaminidase ,2; V Glucosamine Fig. 2. Chitin degradation. Chitinolytic and chitosan pathway. 15 Chitinolytic microorganisms. The microbial degradation of Chitin has been a topic Of intense scientific interest for many years. It is now well documented that chitinases (EC 3.2.1.14), N-acetyl-B-D-glucosaminidases (EC 3.2.1.30) and N- acetyl-B-D-hexosaminidases (EC 3.2.1.52) are widely distributed in bacteria, fungi (Muzarelli, 1993), and even in some ciliate protozoa (VillagOmez et al., 1993). Chitin is the most important structural polysaccharide in the cell wall of fungi and different fungal chitinases are able to breakdown the three main types of cell walls in fungi (chitin-B-glucan of Chytridiomycetes, chitin-cellulose of Hyphochytridiomycetes and B-glucan-cellulose of Oomycetes; Gay, ef al. 1993). Some chitinolytic fungi isolated from forest soils can exceed the chitinolytic activities of bacteria (Gooday, 1990b). However, the importance Of fungal chitinases in the process of chitin mineralization may be considered secondary to the essential role that these enzymes play in the life cycle of these organisms (Pedraza and LOpez, 1991). They are involved in apical growth, cell wall softening during hyphal branching, germination, degradation of septa for the mobilization of nuclei and hyphal fusion (VillagOmez et al. 1993). Chitin also naturally occurs as the major cell wall component Of the ruminal anaerobic fungi Neocallimastix frontalis, Piromonas comunis and Sphaeromonas comunis (Orpin, 1977). However, under pure culture conditions, various strains of Neocallimastix frontalis, Piromonas comunis and Sphaeromonas comunis have not shown the ability to utilize chitin as a substrate for growth (Theodorou et al. 1993). It is important to mention that chitosan, but not chitin, is the major cell wall component of fungi belonging to the family Mucoraceae, such as Absidia, Mucor 16 and Rhizopus (Ohtakara et al., 1990). In these fungal species the enzymes chitin deacetylase and chitosanase are more important in the process of chitin mineralization (Kafetzopoulos et al. 1993), but as mentioned before, these fungal enzymes may be more important in the process of cell growth and division. Chitin degrading bacteria have been isolated from a wide range of habitats representing a variety of Gram positive and Gram negative species, including Vibrio harveyi and V. furnissi from marine environments (Montgomery and Kirchman, 1993); and Myxobacterium, Serratia, Pseudomonas, Achromobacter, Bacillus, Flavobacten'um, Streptomyces from soil and fresh water habitats (Gooday, 1990b). In a survey of nearly 500 environmental samples (soil, fresh water and sea water), the chitinolytic bacteria with the highest chitin liquefying activity were Pseudomonas aeruginosa and Vibn'o angi/larum. These bacteria dissolve decalcified prawn cuticles in 3-4 days (Muzzarelli, 1993). Microbial degradation of chitin under strictly anaerobic conditions has been investigated very little. It is known that anoxic marine sediments have low to very low chitin content (under 100 ug chitin per gram of decalcified sediment), which suggest a very active microbial degradation process of chitinous compounds under anaerobic conditions (Poulicek and Jeuniaux, 1991). Chitinolytic activity by anaerobic bacteria has been detected in estuarine sediments where, Chitin degradation is accomplished by the combined activity of primary Chitin-degrading species (5 x 10° per ml) and secondary sugar-fermenting bacteria (1-3 x 10" per ml) which may enhance the activity of the primary chitinolytic bacteria (Pel and Gottschal, 1986a). These same investigators, using an anaerobic Chitin- 17 enrichment media, isolated eight chitinolytic Clostn'dium strains from an estuarine anoxic-sediment (Pel and Gottschal, 1986b). All strains were strictly anaerobic. Electron scanning microscopy samples from mid-growth phase cultures, showed adherence of bacteria to chitin particles. Another property shared by all of these isolates was the requirement for a very low redox-potential (<-160 mV) for both, optimal growth and chitinolytic activity. A broad pH-optimum for growth on chitin was found from 6.0 to 7.0. It may be noted that these characteristic Closely resemble those found in the rumen environment, and it can be expected that some ruminal bacteria may be able to degrade chitinous compounds under normal environmental conditions. Bacterial attachment to the substrate. Although not well documented, it is believed that the rate of chitin degradation is enhanced by the adherence of chitinolytic microflora (Gooday, 1990b). Ou and Alexander (1974) observed that separation of chitin and chitinolytic bacteria by layers of microbeads greatly reduced the rate of mineralization of chitin particles by pure cultures of a soil Pseudomonas species. Adherence of ruminal microbes to plant digesta is also considered an important factor in the process of plant degradation (Rasmussen et al., 1989). At present, it is known that cellulolytic ruminal species, such as Ruminococcus flavefaciens, R. albus and Fibrobacter succinogenes, adhere to plant cell walls. In the same way, rumen amylolytic bacteria, such as Bacteroides amylophilus, B. ruminicola and Bifidobacten‘um sp. will attach to starch granules (Yokoyama, 1988). The factors involved in the process of bacteria attachment are unknown in most bacteria species, but studies have been done on the cellulase 18 system of C. thermocellum. Lamed et al. (1983) purified a very high molecular weight, multifunctional, multienzyme complex from C. thermocellum which has been coined "cellulosome". The purified "cellulosome" was responsible for the adhesion of C. thermoceI/um to the insoluble cellulosic substrate. Another important role of bacterial attachment to particulate matter is to ensure the bacterial numbers in the rumen. In Open ecosystems such as the rumen, where materials are constantly being introduced and removed, and the bacterial generation time (eg. time for doubling of a population) may be longer that the turnover rate of rumen digesta, they would eventually wash out of the rumen. Since the passage rate Of the particulate phase is much slower than that Of the liquid phase in the rumen, slower growing species are prevented from being washed out by attaching to various surfaces (Yokoyama, 1988). Assay of chitinase activity. The determination of chitinolytic activity presents some serious problems. The first difficulty originates with the substrate itself, because of its insolubility in aqueous solutions. This problem makes kinetic measurements difficult and their interpretation uncertain. In order to solve this problem, water-soluble derivatives of chitin have been used, such as colloidal Chitin which is Obtained from pure chitin or shrimp shells after successively treating with 1 N HCI, 0.5 N NaOH, 0.5% KMnO,, Na28205, hot water, and alcohol, then ground in water and mixed with concentrated HZSO, (Jeuniaux, 1966). Besides the difficulty of preparing colloidal chitin, another drawback is that during the process, chitin may suffer several alterations in its structure that make the end product nonrepresentative of the chitin molecule. A most popular water-soluble 19 derivative of chitin is glycochitin, in which C-6 Of acetylglucosamine residues is joined in an ether linkage with a hydroxyethyl group (Cabib, 1987), but as in the case Of colloidal chitin, glycochitin, present some alterations in its structure with respect to the native chitin (e.g. decrease the molecular weight and degree of polymerization, deacetylation and the introduction of foreign radicals) that may limit its use as a substrate to study chitin hydrolysis. Reacetylation of chitosan with acetic anhydride to regenerate the chitin molecule (Molano, et al, 1977), is considered an excellent substitute for colloidal chitin or glycolchitin as a substrate in chitinase assay. The regenerated chitin offers the advantage that it can be readily prepared in large amounts and in a finely dispersed state, and more importantly, it allows for the use Of [3Hjacetic anhydride for the preparation of tritium-labeled chitin, making this substrate a more sensitive assay for chitinolytic activity. Another difference to other chitin derivatives is the fact that the acetylation of chitosan does not introduce alterations in the chitin molecule other than a decrease in the polymer length. But it must be noted that regenerated chitin is highly hydrated and more susceptible to chitinases, therefore the reaction rate will be faster than with native chitin. Chitin degradation can be measured by the disappearance of substrate or by the appearance of products. In the assay for chitinase activity, the formation of products is frequently measured rather than the disappearance of Chitin, because the second estimation relies on changes in viscosity or turbidity and this measurement is not sensitive enough to establish the reaction rate of the enzyme (Cabib, 1987). In contrast, the formation of products can be detected in several 20 ways. A common assay involves determination of reducing sugars (e.g. N- acetylglucosamine) by the Morgan-Elson test for N-acetyl-2-amino sugars (Robyt and White, 1990). However, this colorimetric test, loses sensitivity if Oligosaccharides or diacetylchitobiose are produced rather than N- acetylglucosamine. The use of thin-layer chromatography for separation of N- acetylglucosamine and chitin oligosaccharides (Powning and Irzykiewicz, 1967), may be the assay of choice when the end products of the chitinolytic enzymes are not really known. Another factor that must be considered, is that chitinases, usually do not produce N-acetylglucosamine and therefore, the reducing sugars estimation is not useful, unless B-N-acetyl-hexosaminidase is added to the reaction mixture (Cabib, 1987). Recently, several new chitin substrates labeled with various dyes have become commercially available and have been used successfully to measure chitinase activity without the expense and hazard of radiolabels. These include, chitin azure (Evrall, et al. 1990), colloidal chitin or carboxymethyl chitin conjugated with remazol brilliant violet 5R (Wirth and Wolf, 1990) and the fluorogenic substrates with 4-methylumbelliferone as aglycone (O'Brien and Colwell, 1987). Although, radiolabeled chitin (3H-chitin) is still the most sensitive assay, with limits of enzyme detection as low as 0.00004 U chitinase per ml or 0.0133 pg per ml, the 4-MUF-chitin assay is also considered very sensitive, being able to detect chitinase concentrations Of 0.00016 U per ml or 0.533 pg per ml. The chitin azure method, is not very sensitive, but can detect chitinase levels of 0.0075 U enzyme per ml or 2.5 pg per ml (Hood, 1991). An important advantage of the 4-MUF assay is the 21 capability to estimate which chitinolytic enzymes are involved. In theory, a positive reaction with the 4-MUF-N-acetyl-B-D-glucosaminide means the presence of the enzyme N-acetylglucosaminidase, a positive reaction with 4-MUF-B-D-N,N'- diacetyl-chitobioside, means the presence of chitinase, as well as a positive reaction with 4-MUF-8-D-N,N',N"-triacetyI-chitotrioside. Although the assays described can be used with either the chitinolytic enzymes or with the chitinolytic microorganisms, there are no reports of their efficacy under anaerobic conditions, which is the characteristic of most of the microorganisms that inhabit the rumen. 1O 22 LITERATURE CITED Arroyo, B. A. and A. Carabez. 1982. Location of chitin in the cyst wall Of Entamoeba invadents with colloidal gold tracers. J. Parasitol. 68:253-258. Bardnlckl, G. S. and E. Uppman. 1982. Fungal cell wall composition. p.229-252. In: A.J. Laskin and HA. Lechevalier (eds). CRC Handbook of Microbiology, 2nd ed., Vol. IV. Microbial composition:carbohydrates, lipids and minerals. CRC Press, Boca Raton. Blackwell, J. 1988. Physical methods for determination of chitin structure and conformation. Meth. Enzymol. 161:435-442. Brundage, A.L., Husby, F.M., Beardsley, G.L and Burton, V.L. 1981. King crab meal in concentrates for lacting cows. J. Dairy Sci. 64: 433-440. Brundage, A.L., Husby, F.M., Henugson, M.L, Simpson,W.L. and Burton, V.L. 1984. Acceptability of Tanner crab meal in concentrates for lactation. J. Dairy Sci. 67: 1965-1970. Cabib, E. 1987. The synthesis and degradation of Chitin. ln: Advances in enzymology. Vol. 59, p. 59-101. Davls, L. L. and BartlnlckI-Garcla. 1984. The co-ordination of Chitosan and Chitin synthesis in Mucor rouxii. J. Gen. Microbiol. 130:2095-2102. Evrall, C. C., R. W. Attwell and C. A. Smith. 1990. A semi-micro quantitative assay for determination of chitinolytic activity in microorganisms. J. Microbiol. Meth. 12:183-187. Gay, L, H. Chanzy, V. Bulone, V. Glrard and M. Fevre. 1993. Biosynthesis of chitin in the fungus Saprolegnia. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 57-66. Gooday, G. W. 1990a. Physiology Of microbial degradation of Chitin and chitosan. Biodegradation, 1:177-190. 11 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 23 Gooday, G. W. 1990b. The ecology of chitin degradation. In: K.C. Marshall, Advances in microbial ecology. N.Y. Plenium Press. Vol. 11, p. 387-429. Gooday, G. W., J. l. Presser, K. Hillman and M. G. Cross. 1991. Mineralization of chitin in an estuarine sediment: The importance of the chitosan pathway. Biochemical Systematics and Ecology. 19:395-400. Heckman, R.H. 1987. Chitin and the fine structure of cuticles. In: Wright and Retnakaran, Chitin and benzoylphenyl ureas. Dr.W Junk Publishers. Dordrecht, The Netherlands. Hara, 5., Y. Yamamura, Y. Fujil, T. Mega and T. lkenaka. 1989. Purification and characterization of chitinase produced by Sterptococcus erythraeus. J. Biochem. 105:484-489. Herwing, R. P., N. B. Pellerln, R. L. lrgens, J. S. Makl and J. T. Staley. 1988. Chitinolytic bacteria and chitin mineralization in the marine waters and sediments along the Antarctic Peninsula. FEMS Microbiol. Ecol. 53:101-112. Hood, M. A. 1991. Comparison of four methods for measuring chitinase activity and the application of 4-MUF assay in aquatic environments. J. Microbiol. Methods. 13:151-160. Husby, F. M., A. L Brundage and R. L White. 1981. Utilization of shellfish meals in domestic feeds. In: W. S. Otwell (Ed). Seefood waste management in the 1980's: Conference proceedings. Report NO. 40 Florida Sea Grant College. Orlando, Fl. Jeuniaux, C. 1966. Chitinases. In: E. F. Neufeld and V. Ginsburg. Methods in enzymology. Vol. 8. p. 644-650. Jeunlaux, C. 1993. Chitinolytic systems in the digestive tract of vertebrates: a review. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 233-244. Kafetzopoulos, D., A. Martinou and V. Bourlotis. 1993. Purification of chitin deacetylase from Mucor roxii. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 147-154. KIenzIe-Sterzer C., D. Rodriguez-Sanchez and C. Rha. 1984. Solution properties of Chitosanzchain conformation. In: J. Zikakis. Chitin, Chitosan and related enzymes. Academic Press, Inc. Orlando Fl. p.383-393. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 24 Knorr, D. 1991. Recovery and utilization of chitin and chitosan in food processing waste management. Food Technol. 45:116-122. Laflame, LF. 1988. Utilization of crab meal fed to young beef cattle. Can. J. Anim. Sci. 68: 1237-1244. Lamed, R., E. Setter and E. A. Bayer. 1983. Characterization of a cellulose-binding, cellulase-containing complex in Clostridium thermoceI/um. J. Bacteriol. 156:828-836. Lumbland, G., B. Hedersted, J. Lind and M. Steby. 1974. Chitinase in goat serum. Preliminary purification and characterization. Eur. J. Bloch. 46:367- 376. Molano, J., A. Duran and E. Cabib. 1977. A rapid and sensitive assay for chitinase using tritiated chitin. Analytical Bichemistry. 83:648-656. Montgomery, M. T. and D. L. Kichman. 1993. Role of chitin-binding proteins in the specific attachment of the marine bacterium Vibn’o harveyi to chitin. Appl. Environ. Microbiol. 59:373-379. Mulisch, M. G. Schermuly and M. Walther. 1993. Chitin synthesis in the ciliated protozoan Eufolliculina uh/igi. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 119-128. Muzzarelli, R. A. 1993. Advances in N-acetyl-B-D-glucosaminidases. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 357-374. Muzzarelli, R. A., Jeuniaux Ch. and Gooday G.W. 1986. Chitin in nature and technology. Plenum Press. N.Y. Muzzarelli, R. A. 1977. Chitin. Pergamon Press, LTD. Oxford. No, H. K., S. P. Meyers and K. S. Lee. 1989. Isolation and characterization of chitin from crawfish shell waste. J. Agric. Food Chem. 37:575-579. 0 'BrIen, M. and R. R. Colwell. 1987. A rapid test for chitinase activity that uses 4- m-ethylumbelliferyl- --N --acetyl B- D-glucosaminide. Appl. Environ. Microbiol. 53: 1718- 1720. Orpln, C. G. 1977. The occurrence of chitin in the cell walls Of the rumen organisms Neocallimastix frontalis, Piromonas communis and Sphaeromonas communis. J. Gen. Microbiol. 99:215-218. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 25 Ortega, E. and Church,.D.C. 1979. In vitro digestion studies with crab meal and related chitinous compounds as feedstuffs for cattle. Proc. West. Sec. Am. Soc. Anim. Sci. 30:302-305. Ohtakara, A., H. Matsunaga and M. Mitsutoml. 1990. Action pattern of Strptomyces gn‘ceus chitinases on partially N-acetylated chitosan. Agric. Biol. Chem. 54:3191-3199. Ou, L and M. Alexander. 1974. Effect of glass microbeads on the microbial degradation of chitin. Soil Sci. 118: 164-167. Patton, RS and Chandler P.T. 1975. In vivo digestibility of chitinous materials. J. Dairy Sci. 58:397-403. Patton, R.S., P.T. Chandler and 0.6. Gonzalez. 1975. Nutritive value of crab meal for young ruminanting calves. J. Dairy Sci. 58:404-409. Pedraza, R. M. and E. Lopez-R. 1991. Chitinase activity in germinating cells of Mucor rouxii. Antonie van Leeuwenhoek. 59:183-189. Pel, R. 1989. Microbial interactions in anaerobic chitin-degrading mixed cultures. Thesis of the Department of Microbiology of the University of Groningen, The Netherlands. Pel, R. and J. C. Gottschal. 1986a. Chitinolytic communities from an anaerobic estuarine environment. In: Chitin in nature and technology. Muzzarelli, R. A. A., C. Jeuniaux and G. W. Gooday (Eds). Plenum Press, N.Y. p. 539-546. Pel, R. and J. C. Gottschal. 1986b. Mesophilic chitin degrading anaerobes isolated from an esturine environment. FEMS Microbiol. Ecol. 38:39-49. Poullcek, M. and C. Jeuniaux. 1991. Chitin biodegradation in marine environments: An experimental approach. Biochemical Systematics and Ecology. 19:385-394. Pownlng, R. F. and H. lrzykiewlcz. 1967. Separation of chitin Oligosaccharides by thin-layer chromatography. J. of Chromatog. 29:115- 119. Rasmussen, M. A., B. A. White and R. B. Hespell. 1989. Improved assay for quantitating adherence of ruminal bacteria to cellulose. Appl. Environ. Microbiol. 55:2089-2091. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 26 Robyt, J. F. and B. J. White. 1990. Biochemical techniques, theory and practice. Waveland Press, Inc. IL. p. 216-217. Skjak-Break, G. T., T. Anthonsen and P. Sandford. 1989. Chitin and Chitosan. Proceedings from the 4th. International Conference on Chitin and Chitosan. Trondeheim, Norway, 1988. Elsevier Applied Science. N.Y. Shlmara, K., Y. Taklguchl and T. Kobayashl. 1988. Screening of Mucoraceae strains for chitosan production. In: G. Skjak, T. Anthonsen and P. Sandforf. Chitin and chitosan. Elsevier Applied Sci. p. 171-178. Smucker, A. R. 1991. Chitin primary production. Biochemical Systematics and Ecology. 5:357-369. Theodorou, M. K., S. E. Lowe and A. P. J. Trinci. 1993. Anaerobic fungi and the rumen ecosystem. Microbiology Group, Department of Cell and Structural Biology (ed). University Of Manchester, UK. p. 45. Velez, S.A., J.C. Allen, C.M.Keery and R.W. Adkinson. 1991. Evaluation Of crab and crawfish waste meals as protein sources for growing dairy heifers. J. Dairy Sci. 74:234-242. Verbug, J.G. and K0. Huynh. 1991. Purification and Characterization of an antifungal chitinase from Arabidopsis thaliana. Plant Physiol. 95:450-455. VIIIagOmez-Castm J. C., M. Pedraza-Reyes, C. Calvo- Mendez and E. LOpez-Romero. 1993. Fungal and amebic chitinases. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 311-323. erth, S. J. and G. A. Wolf. 1990. Dye-labelled substrates for the assay and detection of chitinase and lysozyme activity. J. of Microbial. Methods.12:197-205.. Wright, J.E. and Retnakaran A. 1987. Chitin and benzoylphenyl ureas. Dr.W.Junk Publishers. DordreCh, The Netherlands. Yokoyama, M. T. and K. A. Johnson. 1988. Microbiology of the rumen and Intestine, p.125-144. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Zlkakls, JP 1984. Chitin, chitosan and related enzymes. Academic Press, Inc. Orlando Fl. CHAPTER 2 USE OF 4- METHYLUMBELUFERYL CHITIN-SUBSTRATES FOR DETECTION OF CHITINOLYTIC ACTIVITY IN RUMINAL MICROORGANISMS. 27 28 ABSTRACT A rapid assay for detection of chitinolytic activity based on the hydrolysis of fluorogenic substrates prepared from 4-methylumbelliferone (4-MUF) was tested in ruminal bacteria. The substrates used were; 4-MUF-N-acetyl-[3-D- glucosaminide, 4-MUF-B-D-N-N'-diacetylchitiobioside, and 4-MUF-8—D-N-N‘-N"- triacetylchitotriose. After some modifications for anaerobiosis, the rapid test was simple to perform, quick, reliable and relatively inexpensive. The assay conditions included: substrates concentration, 5 pM per ml; pH, 7.0; incubation temperature, 395°C and incubation time, 20 min. WIth this method, it was demonstrated that certain bacteria from the rumen of dairy cows, do possess chitobiase and chitinase activity. Since the dairy cows used in this experiment, have never been fed with chitinous compounds, it is possible that the growth of Chitinolytic bacteria in the rumen may be stimulated by the presence of yeast and fungi, whose cell walls are rich in Chitin. Considering the accuracy and efficiency of this technique to detect chitinolytic activity even in strict anaerobes, we suggest its use in further studies to elucidate the potential and limits of chitinous waste as a feed source for ruminants. 29 INTRODUCTION The potential use of chitinous wastes (e.g. crab and crawfish waste meals) as a feed for dairy and beef cattle has been recently studied by Brundage et al. (1981, 1984), Laflame (1987) and Velez et al. (1991). Since the production of chitinolytic enzymes by different digestive tissues of ruminants is low (Lumbland et al., 1974), it has been assumed that microorganisms inhabiting the rumen are responsible for chitin digestion rather than the animal. In this way, ruminal microorganisms enable the host animal to convert chitinous wastes into higher quality animal products. However, until now, it has not been determined which microorganisms (e.g. bacteria, fungi or protozoa) are involved in chitin degradation in the rumen and how efficiently they grow and degrade chitin. A better understanding of the potential use and limits of chitinous compounds as feeds for ruminants should begin with the selection of suitable techniques which allow detection and enumeration of ruminal chitinolytic microorganisms. Several screening procedures have been devised for the detection of chitinolytic activity in microorganisms. Some tests depend on the production Of zones of Clearing in solid media previously opaque, due to the presence of soluble forms of chitin such as colloidal chitin (West and Colwell, 1984; WIrth and Wolf, 1990). The main problem with plating methods, is that some anaerobic bacteria do not form colonies, even thought strict anaerobic techniques are followed. Radiolabeling 3O techniques by either reacetylation of chitosan with [3H]acetic anhydride to produce 3H-Chitin (Molano et al., 1977), or using [“Cjacetate to produce 1‘C-chitin (Herwig et al., 1988) are the most rapid and sensitive. However, these methods require special facilities for the preparation of radioactive substrates and special equipment for interpretation of results. Recently, several new chitin substrates labeled with fluorescent or other dyes have become commercially available and have been used successfully to measure chitinolytic activity. For example, chitin azure, a colloidal chitin conjugated with remazol brillant blue, when hydrolyzed releases the blue dye which is then measured spectrophotometrically (Evrall et al., 1990 and Hackman and Golberg, 1964). In a similar way, fluorogenic Chitin derivatives prepared from 4-methylumbelliferone (4-MUF), when hydrolyzed, released 4-MUF which generates fluorescence (Maddocks and Greenan, 1975) that can be measured using a fluorescence spectrophotometer (Hood, 1991) or simple UV light (O'Brien and Colwell, 1987). Although radiolabeled chitin is considered the most sensitive method to use, with limits Of enzyme detection as lowest as 0.00004 U per ml, the 4-MUF-Chitin method is also considered very sensitive being able to detect Chitinase concentrations as low as 0.00016 U per ml (Hood, 1991). The same author, after comparing radiolabeled, colorimetric and fluorescent techniques, concluded that due to their simplicity, sensibility and commercial availability, 4-MUF-Chitin substrates should be considered the first method of choice. This rapid test has been successfully used to measure chitinolytic activity of aerobic microorganisms, but its efficacy with anaerobic microorganisms is not known. 31 The Objective Of this study was to test the 4-MUF-chitin method (O'Brien and Colwell, 1987) for determining chitinase activity in fresh ruminal fluid samples, after modifications of the original methodology to insure bacterial requirements for anaerobiosis. A bacteria culture of a strict anaerobic chitinolytic bacteria previously isolated from the digestive tract of a fish was also used to assess the accuracy of the modified technique. MATERIALS AND METHODS Technique of mmen fluid sampling. Fresh ruminal fluid samples, from two ruminally fistulated dairy cows fed alfalfa hay and never previously fed with shellfish waste, were collected just before the morning feeding. Using sterile gloves, five samples of ruminal fluid contents were aseptically taken from different parts Of the rumen, squeezed by hand and the extracted ruminal fluid was deposited into previously sterilized glass culture tubes (18 x 150 mm). The culture tubes were sealed with sterile rubber stoppers and taken to the laboratory for further sample preparation. 4-MUF-Chltln stock solutions. Three methylumbelliferyl substrates (Sigma, Co.) were examined to determine chitinolytic activity. They include the Chitin monomer, 4-MUF-N-acetyI-B-D-glucosaminide (MUF-NAG); the dimer, 4-MUF-B—D- N-N'-diacetylchitobioside (MUF-ChB) and the trimer 4-MUF-B-D-N-N'-N"- triacetylchitotriose (MUF-ChT). Stock substrate solutions were prepared by the 32 method described by O'Brien and Colwell (1987). Into 2 ml of dimethylformamide (Sigma Co.), 50 pmol (either MUF-NAG, MUF-ChB or MUF-ChT) Of fluorogenic substrate was added. For the test, 0.6 ml of the stock solution was diluted in 9.4 ml of 0.1 M phosphate buffer (test solution). The modification of this technique included: use of 0.1 M phosphate buffer (pH 7.0) instead of pH 7.4, sterilization of the test solutions by membrane filtration using sterile filters (pore size 22 pm), and saturating the test solutions with CO2 gas for 10 min to attain anaerobiosis. Methodology. The ruminal fluid samples were filtered through Whatman paper (No. 4) and recovered in culture tubes under C02. From each sample, 1 ml was transferred to 2 ml centrifuge tubes and centrifuged at 3500 g for 5 min. The supernatant was discarded by decantation and the pellet was resuspended in 1 ml of MUF-NAG, MUF-ChB or MUF-ChT solution by gentle vortexing. To Check the efficiency of the technique for detection of anaerobic chitinolytic bacteria, the MUF- chitin substrates were also incubated with a chitinolytic bacteria culture previously isolated from the intestinal contents of a bluegill (Lepomis macochinus rafinesque), a freshwater fish. This chitinolytic culture was a mixed culture Of strict anaerobic bacteria (nonpublished data). Negative controls consisted of autoclaved samples of rumen fluid and the chitinolytic bacteria cultured from the fish digestive tract incubated with each Of the different MUF-chitin substrates solutions. Also, the distinct MUF-chitin substrates (test solutions) without inoculation were incubated to check for reactive stability. Five replicates from the ruminal fluid, the Chitinolytic culture isolated from the fish and the negative controls were incubated at 39°C and Checked for 33 fluorescence every 10 min. Fluorescence, was checked by exposing the microcentrifuge tubes containing the samples to UV light with a wavelength of 360 nm. Bacteria numbers in filtered ruminal fluid and chitinolytic bacterial culture at the beginning of the test, were estimated by serial dilutions and development of colony forming units (CFU) on glucose, cellobiose, starch-ruminal fluid (GCS-RF) solid anaerobic medium, using the roll tube technique (Hungate, 1969). The bacteria in rumen fluid and the pure culture were also inoculated (1ml) into anaerobic chitin medium (CH-medium), to confirm their ability to degrade Chitin. The composition of GCS-RF medium and CH-medium are given in Table 1. 34 Table 1. Composition of the chitin and GCS-RF media used for the isolation and characterization of ruminal chitinolytic bacteria. Component Medium Chitin GCS-RF Quanty per 100 ml Deionized water 51.6 ml 50.5 ml Clarified ruminal quuid° 30.0 mi 30.0 ml Sodium carbonate 8% solution 5.0 ml 5.0 ml Mineral solution 1° 5.0 ml 5.0 ml Mineral solution 2° 5.0 ml 5.0 ml Cysteine-sulfide solutiond 2.0 ml 2.0 ml Resazurin 0.1% solution 0.1 ml 0.1 ml Trypticase-peptone (BBL®) 0.2 g 0.5 9 Yeast extract (BBL®) 0.1 g 0.2 g Chitin (Sigma) 1.0 g - - - - Agar - - - - 1.5 g Glucose - - - - 60 mg Cellobiose - - - - 60 mg Starch - - - - 60 mg Clarilled ruminal IIQUIU IS prepared as IOIIOWI FIGS“ ruminal ”Ula pI'BVIOUSIy filtered through a cheesecloth is centrifuged at 23,420 g for 15 min at 4°C, autoclaved 20 min at 15 psi-121°C, then recentrifuged as described. ° Mineral solution 1 contains (per 1000 ml): KZHPOM 6.0 9 ° Mineral solution 2 contains (per 1000 ml): KHZPO4, 6 g; (NH4)2SO,, 6 9; NaCl, 12 g; M9804, 2.45 g and CaClo 2 H20, 1.6 9 (Bryant and Robinson, 1962). d Mix L-Cysteine, 2.5 9 (dissolved in 15 ml of 2N NaOH); NaZS.9HZO, 2.5 g and resazurin, 0.1 ml (1% sol. in H20). Then, bring the mix to a final volume of 100 ml with deionized water. Heat or microwave the mix until the resazurin indicator is colorless, seal the container and autoclave (bubbling with CO2 is not required). 35 RESULTS The results of the rapid test using different MUF-chitin substrates are given in Table 1.2. The results obtained confirm the capability of this rapid test to detect chitinolytic activity, even under anaerobic conditions. The rumen fluid and chitinolytic culture from the intestinal contents Of fish, started to give a weak positive reactions after 10 min of incubation, but a strong light-blue fluorescence was observed after 20 min of incubation in all samples and repetitions. The strong fluorescence in positive reactions was maintained 24 to 48 h in samples stored at 4°C. The negative controls (MUF-chitin substrates inoculated with either autoclaved ruminal fluid or chitinolytic culture), as expected, gave a negative reaction, but some of the repetitions gave a weak positive reaction, in particular with the samples tested with MUF-ChT. The false reactions were probably due to the instability of the MUF-substrates or sample contamination. The other MUF- chitin substrates incubated without inoculation were stable and did not give positive reactions at any time. Due to the strong fluorescence Observed in positive reactions, it was not necessary to add sodium bicarbonate as suggested by Slifkin and Gil (1983), to enhance the fluorescence intensity of the released methylumbelliferyl. However, the source of UV light does play an important role, and the use Of UV lamps with a wavelength below 360 nm may not show as much fluorescence. 36 Bacteria concentration in ruminal fluid and pure culture, estimated as colony forming units (CFU) is given in Table 2. The results indicated a good bacteria concentration after sample preparation. After 2 to 3 d of incubation of the anaerobic CH-medium inoculated with filtered rumen fluid or the pure chitinolytic culture from the fish digestive tract, a dense bacterial growth accompanied with high gas production and disappearance of chitin was Observed, confirming the positive results detected by the MUF-tests for chitinolytic activity in ruminal fluid. Table 2. Results of MUF-chitin substrates assay with different bacteria inocula. — lnocula MUFNAG’ MUF-ChBb MUF-ChT° CFU per ml Filtered rumen fluid + + + 1.3 x 10‘° Chitinolytic bacteria + + + 1.3 x 109 culture“ Filtered rumen fluid - - - N/A° autoclaved Chitinolytic bacteria - - - N/A culture autoclaved MUF-substrates - - - N/A without inoculation ' Z-MUF-N-acetyl-B-D-glucosaminide ° 4-MUF-B-D-N-N'-diacetylchitobioside ° 4-MUF-B-D-N-N'-N"-triacetylchitotriose d Isolated from the intestinal contents of a bluegill, Lepomis macochinus rafinesque ° Not applicable 37 DISCUSSION The rapid test method using methylumbelliferyl-chitin substrates described by O'Brien and Colwell (1987) and Hood (1991), was easily adapted for anaerobic bacteria. All three 4-methylumbelliferyl chitin substrates used for detection of Chitinolytic activity were hydrolyzed by the two anaerobic bacteria communities studied. The entire procedure for the positive identification of chitinolytic bacteria requires approximately 20 min. During this short period of time, it is almost impossible that a contaminating bacteria with an ability to degrade Chitin to overcome the bacteria concentration in the sample, resulting in a false-positive reaction. Lack of fluorescence in the negative controls, confirms that the generation of fluorescence was enzymatic in nature. Thus, it was shown that fluorescence was not produced in MUF-substrates noninoculated and the activity in the presence of MUF-substrates was abolished after autoclaving the bacteria from the filtered rumen fluid and the chitinolytic pure culture used as inocula. Another important result was the observation that chitinolytic bacteria are normally present in the rumen microbial population even though the cows were never fed chitinous compounds. The presence of bacteria with chitinolytic activity inside the rumen could be due to the natural availability of the substrate from the ruminal anaerobic fungi whose cell walls are made of chitin (Orpin, 1977). Another 38 possible source of chitin are the chitinous cell wall of yeast and fungi which are usually found in the animal's feed (Middelhoven and van Baalen, 1988). In the present study, no attempt was made to identify which of the ruminal microorganisms degraded chitin, but examination under phase contrast microscope Showed that mainly bacteria were in the sample used as inocula. However, this does not discount the possibility that some ruminal fungi and protozoa may also utilize Chitin, as it has been found in other habitats (Gooday, 1990 and VillagOmez etaL,1993) Although optimal conditions of incubation for the 4-MUF-chitin substrates method are a pH of 7.5 and 35°C (Hood, 1990), there was no apparent negative effect by using a pH of 7.0 and an incubation temperature of 39°C. The later values are similar to those found in the rumen environment and therefore the results obtained may be more reliable. A very useful reason for using different MUF-substrates, is the possibility of detecting the presence Of different chitinolytic enzymes. In theory, 4-MUF-N- acetyl-B-D-glucosaminide (MUF-NAG) represent the dimer, diacetylchitobiose and a positive reaction indicates the presence of the enzyme N-acetylglucosaminidase also referred as diacetylchitobiase (EC 3.2.1.30 B-2-acetamido-2-deoxy-D- glucoside acetamidoxy- glucohydrolase) which hydrolyses terminal, non-reducing 2-acetamido-2-deoxy-I3-D-glucose residues in chitobiose and chitotriose, leaving N-acetylglucosamine units. In the same manner, 4-MUF-B-D-N-N'-N"-triacetyl- chitotriose (MUF-ChT), represent a tetramer of N-acetylglucosamine units and a positive reaction indicates the presence of the enzyme chitinase (EC 3.2.1.14, 39 poly-B-1-4-(2-acetamido-2-deoxy)-D-glucoside glycanohydrolase) which cleaves glycosidic linkages randomly along the chitin chain and diacetylchitobiose units from the nonreducing end of the chitin chain, including tetramers and to a lesser extent trimers (Jeuniaux, 1966), eventually giving diacetylchitobiose as the major product. Since the bacteria in ruminal fluid were positive for both 4-MUF-NAG and 4-MUF-ChT, it can be concluded that ruminal bacteria have the enzymes chitinase and N-acetylglucosaminidase required for complete degradation of chitin to N- acetylglucosamine units. Chitotriose units represented by 4-MUF-B-D-N-N'-diacetylchitobioside (4- MUF-ChB) can be hydrolyzed by either chitobiose or chitinase enzyme. Therefore, a positive reaction with 4-MUF-ChB can not be attributed to any particular enzyme and the use of this fluorescent substrate can be omitted, especially if 4-MUF-NAG and 4-MUF-ChT are included. In summary, the results support the use of the methylumbelliferyl method for rapid detection of chitinolytic ruminal microorganisms. The method presents several advantages such as being rapid, simple to perform, commercially available and reliable. In addition, this rapid method, allows the detection of different enzymes related to the process of chitin hydrolysis. 10. 4O LITERATURE CITED Bryant, M. P. and l. M. Robinson. 1962. Some nutritional characteristics of predominant culturable ruminal bacteria. J. Bacteriol. 64:605-613. Brundage, A.L, Husby, F.M., Beardsley, G.L and Burton, V.L 1981. King crab meal in concentrates for lacting cows. J. Dairy Sci. 64: 433-440. Brundage, A.L, Husby, F.M., Herlugson, M.L, Slmpson,W.L. and Burton, V.L. 1984. Acceptability Of Tanner crab meal in concentrates for lactation. J. Dairy Sci. 67: 1965-1970. Evrall, C. C., R. W. Attwell and C. A. Smith. 1990. A semi-micro quantitative assay for determination of chitinolytic activity in microorganisms. J. Microbiol. Meth. 12:183-187. Gooday, G. W. 1990. The ecology Of chitin degradation. In: K.C. Marshall, Advances in microbial ecology. N.Y. Plenium Press. Vol. 11, p. 387-429. Hackman, R.H. 1987. Chitin and the fine structure of cuticles. In: Wright and Retnakaran, Chitin and benzoylphenyl ureas. Dr.W Junk Publishers. Dordrecht, The Netherlands. Herwing, R. P., N. B. Pellerln, R. L Irgens, J. S. Makl and J. T. Staley. 1988. Chitinolytic bacteria and chitin mineralization in the marine waters and sediments along the Antarctic Peninsula. FEMS Microbiol. Ecol. 53:101-112. Hood, M. A. 1991. Comparison of four methods for measuring chitinase activity and the application of 4-MUF assay in aquatic environments. J. Microbiol. Methods. 13:151-160. Hungate. R. E. 1969. A roll tube method for cultivation Of strict anaerobes. In: J. R. Norris and D. W. Ribbons (Ed). Methods in microbiology. Vol. 3, p. 117-132. Academic Press, Inc. New. York. Jeunlaux, C. 1966. Chitinases. In: E. F. Neufeld and V. Ginsburg. Methods in enzymology. Vol. 8. p. 644-650. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 41 Laflame, LF. 1988. Utilization of crab meal fed to young beef cattle. Can. J. Anim. Sci. 68: 1237-1244. Lumbland, G., B. Hedersted, J. Lind and M. Steby. 1974. Chitinase in goat serum. Preliminary purification and characterization. Eur. J. Bioch. 46:367- 376. Maddocks, J. L and M. J. Greenan. 1975. A rapid method for identifying bacteria enzymes. J. Clin. Phatol. 28:686-687. Middelhoven W. J. and A. H. M. va Baalen. 1988. Development of the yeast flora of whale-crap maize during ensiling and during subsequent aerobiosis. J. Sci. Food. Agric. 42:199-207. Molano, J., A. Duran and E. Cabib. 1977. A rapid and sensitive assay for chitinase using tritiated chitin. Analytical Biochemistry. 83:648-656. O'Brien, M. and R. R. Colwell. 1987. A rapid test for chitinase activity that uses 4-methylumbelliferyl-N-acetyl-B-D-glucasaminide. Appl. Environ. Microbial. 53:1718-1720. OrpIn, C. G. 1977. The occurrence of chitin in the cell walls of the rumen organisms Neocallimastix frantalis, Piromonas communis and Sphaeromonas communis. J. Gen. Microbial. 99:215-218. Slifkin, M. and G. M. Gil. 1983. Rapid biochemical test for identification of group A, B, C, F, and G streptococci from throat cultures. J. Clin. Microbial. 18:29-32. Velez, S.A., J.C. Allen, C.M.Keery and R.W. Adkinson. 1991. Evaluation of crab and crawfish waste meals as protein sources for growing dairy heifers. J. Dairy Sci. 74:234-242. VlllagOmez-Castro J. C., M. Pedraza-Reyes, C. Calvo- Mendez and E. LOpez-Romero. 1993. Fungal and amebic chitinases. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 311-323. West, P. A. and R. R. Colwell. 1984. Identification and classification of Vibrianaceae: an overview. p.285-363. In: Vibrios in the environment. John WIIey & Sons, Inc., New York. erth, S. J. and G. A. Wolf. 1990. Dye-labelled substrates for the assay and detection of chitinase and lysozyme activity. J. of Microbial. Methods.12:197-205. CHAPTER 3 ISOLATION AND CHARACTERIZATION OF RUMINAL BACTERIA WHICH DEGRADE CHITIN 42 43 ABSTRACT Using an anaerobic selective medium with purified chitin as the sole energy source and ruminal fluid from fistulated dairy cows as inocula, four morphologically distinct bacteria were isolated. Scanning electron micrographs of the mixed culture revealed a cocci, a spore—former, a selenamanad and a spirochete. Pure cultures were only obtained for the cocci and spare-former. The spare-former was a Gram positive, motile rod 3 to 5 pm in length with rounded ends. However, in older cultures the bacterium will grow either as a long filament or develop a terminal swollen Spore. This spore-former bacterium is a strict anaerobe, which rapidly degrades chitin, crab shell meal, chitobiase, N-acetylglucosamine, glucose and galactose. However, it is unable to degrade cellulose, a common characteristic of other Clostridial species in the rumen. Using an automated system for identification of anaerobes and due to its indigenous location inside the rumen, the spore-former was identified as Clostridium paraputn'ficums var. ruminantium. This bacterium is stimulated by, but does not require Clarified ruminal fluid for optimum grow and chitinolytic activity. C. paraputn'ficums var. ruminantium is unable to degrade amino acids. The cocci, was Gram positive, growing in pairs or 4-8 units chains. The cocci, a Peptostreptococcus sp. was unable to degrade chitin, but its ability to utilizes lactate which is one of the end-products of chitin degradation, makes this cocci an important secondary bacteria in the process of Chitin degradation. 44 INTRODUCTION Previous studies (eg. Veles, 1991 and Laflame, 1988) show that livestock can derive some nutritional benefit when fed chitinous processing wastes. Crab meal consist of shells, viscera, unextracted meat, and some non-protein nitrogen components. As such, crab meal contains in excess of 40% crude protein. However, a serious limitation in the use of these marine by product as feeds is the recalcitrance of their chitin component to digestion. Ruminants (e.g. cattle, sheep, goats) by virtue of having a large microbial population in their rumen have the capability to partially degrade chitin (Patton, 1975), but the ruminal bacteria involved in chitin degradation have never been isolated and identified. In fact, microbial degradation of chitin and the rates of chitin degradation under strictly anaerobic conditions have been less well investigated not only in the rumen, but also in other anaerobic ecosystems. For example, it is known that anoxic marine sediments have a very active microbial degradation of chitinous compounds (Poulicek and Jeuniaux, 1991), but the anaerobic microorganisms involved have not been isolated. Today, there is only one report about the isolation of strictly anaerobic bacteria from estuarine anoxic sediment, which have been identified as chitinolytic Clostn’dium strains (Pel and Gottschal,1986). In a previous study (chapter 2), it was demonstrated that the rumen of dairy cows contain ruminal bacteria apparently able to degrade chitin. The objective of 45 this research was to isolate and characterize these ruminal bacteria which degrade Chitin. MATERIAL AND METHODS Enrichment and roll tube media. To isolate the ruminal microorganisms involved in chitin degradation, an enrichment medium with purified Chitin as the sole energy source was prepared by the anaerobic techniques of Hungate (1969) as modified by Bryant (1972). The composition of the chitin (CH) medium and of the glucose, cellobiose, starch and ruminal fluid (GCS-RF) agar medium used for anaerobic roll tubes are shown in Table 1. Compared with the GCS-RF medium, the Ch-medium used pure chitin (Sigma), as energy source instead of a combination of carbohydrates (glucose, cellobiose and starch). The trypticase- peptone and yeast extract concentration was also lower in the CH-medium than in GCS-RF medium. Since chitin is insoluble in water, 0.1 g was weighed and individually added to each culture tube and after sterilization, 9 ml of the other medium components were added, under CO2 gas passed though hot copper filings. Animals. Two rumen fistulated Holstein cows from the Dairy Teaching and Research Center (MSU), were used to obtain ruminal fluid. These cows were fed alfalfa hay and had never been previously fed with shellfish waste. 46 Table 1. Composition of the chitin and GCS-RF media used for the isolation and characterization of ruminal chitinolytic bacteria. Component Medium Chitin GCS-RF Quanty per 100 ml Deionized water 51.6 mi 50.5 ml Clarified ruminal liquid“ 30.0 mi 30.0 ml Sodium carbonate 8% solution 5.0 ml 5.0 ml Mineral solution 1° 5.0 ml 5.0 ml Mineral solution 2° 5.0 ml 5.0 ml Cysteine-sulfide solutiond 2.0 ml 2.0 ml Resazurin 0.1% solution 0.1 ml 0.1 ml Trypticase-peptone (BBL®) 0.2 g 0.5 9 Yeast extract (BBL®) 0.1 g 0.2 g Chitin (Sigma) 1.0 g - - - - Agar - - - - 1.5 g Glucose - - - - 60 mg Cellobiose - - - - 60 mg Starch - - - - 60 mg lClarified ruminal “quiz is prepared as lollow: Fresh ruminal Hula previously filtered through a cheesecloth is centrifuged at 23,420 g for 15 min at 4°C, autoclaved 20 min at 15 psi-121°C, then recentrifuged as described. ° Mineral solution 1 contains (per 1000 ml): K,HPO,, 6.0 9 ° Mineral solution 2 contains (per 1000 ml): KH,PO4, 6 g; (NH,),SO4, 6 g; NaCI, 12 g; MgSO4, 2.45 g and CaClo 2 H,O, 1.6 9 (Bryant and Robinson, 1962). d Mix L-Cysteine, 2.5 9 (dissolved in 15 ml of 2N NaOH); Na,S.9H,O, 2.5 g and resazurin, 0.1 ml (1% sol. in H,O). Then, bring the mix to a final volume of 100 ml with deionized water. Heat or microwave the mix until the resazurin indicator is colorless, seal the container and autoclave (bubbling with CO, is not required). 47 Sampling and Inoculation procedure. Ruminal contents were sampled through the ruminal fistula before the morning feeding. Representative samples from different locations of the rumen, were obtained, to extract the ruminal fluid and fill a sterile container (1 liter) to its capacity. At the laboratory, the ruminal fluid was vigorously mixed by hand and 10 subsamples (1ml, each) were inoculated in culture tubes containing 9 ml of anaerobic CH medium. This and all subsequent procedures were performed under anaerobic conditions. Isolation and purification. To stimulate the growth of chitinolytic bacteria over other ruminal bacteria, the ruminal fluid was inoculated (1 ml) into Ch- medium. The cultures were incubated at 39°C and subcultured (1 ml) into fresh Ch-medium at 72 h intervals. After three transfers, the cultures were serially diluted and dilutions inoculated into GCS-RF agar medium, using the anaerobic roll tube technique (Hungate, 1969). Inoculated roll tubes, were incubated at 39°C and after 2 d of incubation, discernible colonies were aseptically picked and reinoculated into CH-medium. After 3 more days of subsequent incubation at 39°C, one ml of each isolate culture, was serially diluted and the dilutions in the 10‘ to 10"° range inoculated in GCS-RF agar medium and rerolled, to check for pure colonies. When more than one colony type was observed, the entire process was repeated. The bacteria types growing in Ch-medium, as well as those recovered from colonies in inoculated roll tubes, were observed and morphologically characterized using a phase contrast microscope. Electron microscopy. The bacteria growing in Ch-medium after three subcultures were observed under scanning electron microscopy. 1 ml of the 48 culture was centrifuged at 35009 for 1 min and the pellet resuspended and fixed in 0.5 ml of 2% glutaraldehyde (in 0.1M phosphate buffer pH 7.0) at room temperature for 1 hr. After fixation, two or three drops of the sample were placed in coverslips previously treated with poly-L-Iysine (1% solution in distilled water) for 5 min and then the coverslips were gently rinsed with distilled water to remove the nonattached sample. Dehydration was achieved using an ethanol series (25, 50, 75, 95 and 100%), 20 min each step, then the samples were critical-point dried and sputter-coated, following the procedures described by Klomparens et al. (1986). The samples were examined in an JEOL JSM-35C scanning electron microscope at the Center for Electron Optics, M.S.U. Primary Identification of rumen bacteria isolated. Pure cultures of the isolated bacteria were characterized as to their Gram stain, shape, motility, catalase reaction, spare formation, as well as, for their ability to ferment pure chitin (Sigma), crab shell meal, N-acetylglucosamine, glucose, fructose and lactate. 4-Methylumbelllferyl Chitin substrates test. The presence of the enzymes Chitobiose and Chitinase, was determined by means of the 4-MUF-N-acetyl-B-D- glucosaminide (4-MUF-NAG) and 4-MUF-B-D-N-N'-N"-triacetylchitiotriose (4-MUF- ChT) substrates respectively, using the method described by O'Brien and Colwell (1987), with modifications as described in Chapter 2. Characterization of chitinolytic microorganisms. The microorganisms which degraded chitin and crab shell were more extensively characterized using the RaplD ANA II System (IDS, Inc.) at the microbiological facilities of the College of Veterinary Medicine. 49 The end products of chitin fermentation produced by the isolated chitinolytic bacteria were determined by high performance liquid chromatography (HPLC). The equipment used was a Waters Chromatography 712 WISP. The column (300 x 7.8) employed was a HPX-87H organic acids analysis column (BIO-RAD). The mobile phase was 0.005 N H,SO, with a flow rate of 0.6 ml per min. The samples injected included: 20 pl of 3 (1 old pure cultures in Ch-medium, sterile Ch-medim, and a standard mixture containing known concentrations of lactate, formate, acetate, propionate, butyrate and N-acetylglucosamine. The ability of the chitinolytic bacterial isolates to degrade other polymers was also tested. Pure cultures were inoculated into media where Chitin was replaced by either chitosan, cellulose or starch and incubated at 39°C for 3 d. The develop of turbidity and gas production in the Inoculated media, was considered as proof of substrate utilization. Effect of clarified ruminal liquid on chitinolytic activity. To determine if clarified ruminal liquid (CRL) had unknown factors that could influence the Chitinolytic activity of the bacterial isolates, Ch-medium was prepared with and without CRL. Culture tubes contained Ch-medium with and without CRL were inoculated with the chitinolytic bacterial isolates, and the percent of Chitin degradation was compared. Ch-medium without CRL was prepared by substituting clarified ruminal liquid with deionized water, while all other Ch-medium components were added in the same quantities (see Table 3.1). The pure Chitin (Sigma) used in this experiment, was previously boiled, placed in a dacron bag with 55 pm pore size (Ankom, Fairport, NY), then vigorously washed with distilled water to remove 50 chitin particles and impurities smaller than 55 pm. Finally the chitin in the dacron bag was dried in an oven for 24 h at 60°C. Culture tubes (7 replicates per treatment), containing exact quantities of pure chitin (between 0.1 to 0.1050 g) were autoclaved, then pipetted with 9 ml of sterile Ch-medium with and without CRL. The different culture tubes, were inoculated with 1 ml of the pure chitinolytic bacteria culture isolated. After 4 d of incubation at 39°C, the remaining chitin, was recovered by passage through Whatman paper filters (No. 541), which was previously weighed. The filters with the undigested chitin were dried 24 h at 60°C, and weighed. Three pieces of the Whatman filter paper were used as a blank. After corrections for paper lost or gain of weight during the process, the weight of the undegraded chitin recovered was statistically analyzed by ANOVA (Steel and Torrie, 1980). RESULTS Ruminal bacteria Isolated. CH-medium inoculated with fresh ruminal fluid from an alfalfa hay fed cow gave rise to a diverse population of anaerobic bacteria. Growth was observed within a day following inoculation. After 48 h of incubation, the high gas production and the turbidity of the cultures suggested strong bacterial growth and activity. Examination by phase contrast microscopy of the diverse population, showed at least four distinctly bacteria types with sustained growth after three subcultures in CH-medium. The different bacteria types observed were: 51 a cocci, a spirochete, a selenamanad and a spore-former (Fig.1a). However, after several attempts to isolate pure colonies, only the cocci and the spore-forming bacterium developed colonies on roll tubes with GCS-RF medium. The cocci was a Gram (+), motile, catalase (-), occurring in pairs or small 4 to 8 unit chains (Fig. 1b). The cocci was classified as a member of the genera Peptostreptococcus, in agreement with the characteristics of the genera as described by Treagan and Pulliam (1982). Pure cultures of the Peptostreptococcus sp. grew well in an anaerobic media with either glucose, fructose, galactose, lactose, starch or N- acetylglucosamine, but it was unable to grow alone in chitin or crab shell. For this reason, the Peptostreptocaccus sp. was considered a secondary bacteria in chitin degradation. The clostridial strain was a Gram (+), straight rod with rounded ends, motile, 3 to 5 pm long. Older cultures showed larger rods (7 to 9 pm long) with terminal oval spore (1pm long, 0.73 pm thick), and long curved filament forms up to 20 to 30 pm long. All different forms were 0.33 pm thick (Fig. 1c). 52 Fig. 1. Scanning electron micrographs of the main ruminal bacteria with sustained growth in Chitin medium. (a) The Chitinolytic community was formed by a cocci, a spirochete, a selenamanad and a spore-forrner. Bar, 1pm. (b) The Gram positive, motile and catalase (-) cocci was tentatively Clasified as a Peptostreptococcus so. and it was unable to degrade chitin. Bar, 1pm. (c) The spore former was identified as Clostridium oaraoutrificum. and it was able to degrade chitin. Bar, 1 pm. $0513.33 53 Chitin degradation was more extensive (P<.05) in the CH-medium with clarified ruminal liquid (CRL) than in CH-medium without CRL (89.58%:334 vs 85.05%1253 respectively). However, the degree of chitin degradation was relatively high with both treatments. Therefore, It can be concluded that the ruminal clostridial isolate did not require ruminal fluid, but chitinolytic activity is stimulated when clarified rumen fluid is present in the CH-medium. Results of the 4-methylumbelliferyl test. Pure cultures of the cocci and the spore-forming isolates were used for these tests. The cocci was positive with 4- MUF-NAG and negative with 4-MUF-ChT, while the spore-forming, was positive with both fluorescent tests. Fluorescence in positive reactions was observed within 15 to 20 min of incubation. In agreement with the discussion made on Chapter 2, the results obtained indicate that the cocci was able to utilize chitobiase but unable to degrade the chitin polymer and the spare former was able to completely degrade the chitin polymer to N-acetylglucosamine units. Chitinolytic strain Identification. The chitinolytic spore former was identified by the RaplD ANA II System, with 99 % of confidence, as Clostridium paraputn'ficum. An interpretation of the results obtained from the RaplD ANA II System is presented in Table 2. The results indicate that the isolated ruminal spore former uses a limited number of substrates, indicating its substrate specificity characteristic. It is also remarkable in its inability to utilize amino acids as a nitrogen source. 54 Table 2. Characteristics of the C. paraputn’ficum isolated from the rumen, as interpreted from the RaplD ANA II System results. Test Reaction Test Reaction Motility + lndole produced - Gram reaction + Arabinose - Urea - Fructose - Lysine - Galactose + Glycine - Glucose + Arginine - Raffinase - Serine - Xylose - Phenylalanine - N-acetylglucosamine + Praline - Mannose - Threonine - Phosphate + Alanine - Leucine - In 10 samples the end products of chitin fermentation were: acetate, butyrate and lactate with an average molar ratio of 64.1:19.5:16.4 respectively. There was no detection of other organic acids or N-acetylglucosamine as end products of chitin fermentation. Pure cultures of C. paraputn'ficum were able to degrade the polymers of chitin, crab shell and starch but do not degrade either cellulose or Chitosan. The growth of C. paraputn'ficum on starch medium as compared to its growth on CH- medium, was much slower with a lag-phase of approximately 2 d before degradation was observed. In addition, it was difficult to maintain the culture pure 55 and usually a secondary bacteria contamination developed or the culture was lost after three transfer to fresh starch medium. These observations suggest that even starch is not very well used as a substrate by this bacterium. After three transfers in starch medium (9 d) the culture (1 ml) was reinoculated into CH-medium, where the bacteria was still able to degrade chitin. A summary of the substrates used by C. paraputn’flcum and Peptostreptococcus sp. isolated in this experiment are shown in Table 3. 56 Table 3. Summary of substrates used by Clostridium paraputrificum and Peptostreptococcus sp. isolated from the rumen of dairy cows. Substrate C. Peptostrepto- paraputriflcum coccus sp. Glucose + + Fructose - + Galactose + + N-acetylglucosamine + + Lactate - + Chitin + - Chitosan - n/ta Starch i + Cellulose - - Crab shell meal + - 4-MUF-N-acetyl-B-D-glucosaminide, + + reaction 4-MUF-B-D-N-N'-N"-triacetylchitiotriose, + - reaction I not tested 57 DISCUSSION The efficiency of utilization of shellfish waste has been recently studied in dairy cattle (Velez, 1991) and beef cattle (Laflame, 1988). However, the results of the present study, are the first report of a specific ruminal bacteria involved in chitin degradation. Since the ruminal fluid used as inocula was from a dairy cow never previously fed chitin-rich feeds it can be assumed that chitinolytic activity is expressed by some of the indigenous microorganisms living in the rumen. The natural source of chitin for these bacteria may be fungi and yeast often found with forages ingested by the animal (Middelhoven and van Baalen, 1988). Another possible source could be the anaerobic ruminal fungi such as Neocallimastix frontalis, whose cell wall is made of chitin (Gay et al. 1989). The metabolic profile of the clostridial isolate, indicate that this bacteria is highly specialized for chitin utilization. It is unable to degrade cellulose, a common characteristic of other clostridial species isolated from the rumen of cattle and sheep such as, C. polysaccharolyticum (Van Gylswyk et al., 1980), C. chartatabidum (Kelly et al., 1987) and C. Iongisporum (Varel, 1989). The fact that the ruminal chitinolytic bacteria are unable to degrade chitosan, indicate that this microorganism uses the chitinolytic pathway (Gooday, 1990) in the process of chitin degradation, and supports the results obtained with the fluorescence technique, which indicated the presence of the enzymes chitinase and N- 58 acetylglucosaminidase in this ruminal bacteria. Using the RaplD ll ANA system, the clostridial isolate was classified as Clostridium paraputrificum, with 99% confidence in agreement with its metabolic profile. However, at least 50 different strains of this Clostridium have been isolated from very diverse environments such as soil, marine sediments, human infant feces, and porcine, avian and bovine feces (Cato, et al. 1986). Therefore, we proposed that the ruminal strain isolated be designated C. paraputrificum var. ruminantium. The coccal isolate, was tentatively classified as a member of the genus Peptostreptococcus. This anaerobic cocci was unable to degrade chitin, but its ability to utilize N-acetylglucosamine and lactate indicate that this bacterium may have an important role in chitin degradation, at least by utilizing the lactate produced by C. paraputn'ficum. The role of the spirochete and the selenamanad in chitin degradation remains uncertain until pure colonies can be obtained, but it may be inferred that the selenamanad may carry out the secondary fermentation of the end-products of C. paraputn'ficum in particular of lactate, due to the characteristic ability of Selenomonas ruminantium species to ferment lactate to either acetate, .propionate or longer chain fatty acids (Yokoyama, 1988). The extent of chitin degradation by C. paraputn’ficum var. ruminantium was very high (e.g. 89.58% 1 3.34) for a 4 d incubation period, in comparison with another study on Chitin fermentation, which by coincidence was also with an anaerobic clostridial species. A pure culture of Clostridium sp. strain 9.1 isolated 59 from estuarine sediments required 12 d to degrade approximately 85% of the chitin in the medium (Pel et al., 1989). The significance of C. paraputriflcum var. ruminantium in the ruminal digestion of chitinous wastes and animal performance, is still difficult to assess. But the relatively fast and efficient degradation of chitin, especially when ruminal fluid is present, suggest that this bacteria may have potential as an inocula when chitinous feeds make a significant part of the ruminant diet. 10. 6O LITERATURE CITED Bryant, M. P. 1972. Commentary on the Hungate technique for culture of anaerobic bacteria. Amer. J. Clin. Nutr. 25:1324-1328. Cato, E. P., W. L. George and S. M. Fine Gold. 1986. Endospore-forming gram-positive rods and cocci. Genus Clostridium. In: P. H. A. Sheath (Ed). Bergey's manual of systematic bacteriology. p1141-1200. Williams & VIfillkins. Baltimore, MD. Gay, L, M. Hebraud, V. Glrard and M. Fevre. 1989. Chitin synthase activity from Neocallimastix frontalis, an anaerobic rumen fungus. J. Gen. Microbiol. 135:279-283. Gooday, G. W. 1990. The ecology of chitin degradation. In: KC. Marshall, Advances in microbial ecology. N.Y. Plenium Press. Vol. 11, p. 387-429. Hungate. R. E. 1969. A roll tube method for cultivation of strict anaerobes. In: J. R. Norris and D. W. Ribbons (Ed). Methods in microbiology. Vol. 3, p. 117-132. Academic Press, Inc. New. York. Kelly, W. J., R. V. Asmundson and D. H. Hoperoft. 1987. Isolation and characterization of a strictly anaerobic, cellulolytic spore former: Clostridium chartatabidum sp. nov. Arch. Microbiol. 147:169-173. Klomparens, K. L, S. L FIegIer and G. R. Hooper. 1986. Procedures for transmission and scanning electron microscopy for biological and medical science (Manual). Center for Electron Optics, Michigan State University. Laflame, LF. 1988. Utilization of crab meal fed to young beef cattle. Can. J. Anim. SCI. 68: 1237-1244. Middelhoven W. J. and A. H. M. va Baalen. 1988. Development of the yeast flora of whole-crop maize during ensiling and during subsequent aerobiosis. J. Sci. Food. Agric. 42:199-207. O'Brien, M. and R. R. Colwell. 1987. A rapid test for chitinase activity that uses 4-methylumbelIiferyl-N-acetyl-B-D-glucosaminide. Appl. Environ. Microbiol. 53:1718-1720. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 61 Patton, R.S. and P. T. Chandler. 1975. In vivo digestibility of chitinous materials. J. Dairy Sci. 58:397-403. Pel, R. and J. C. Gottschal. 1986. Chitinolytic communities from an anaerobic estuarine environment. In: Chitin in nature and technology. Muzzarelli, R. A. A., C. Jeuniaux and G. W. Gooday (Eds). Plenum Press, N.Y. p. 539-546. Pel, R., G. Hessels, H. Aslfs and J. C. Gottschal. 1989. Chitin degradation by Clostridium sp. strain 9.1 in mixed cultures with saccharolytic and sulfate-reducing bacteria. FEMS Microbiol. Ecol. 62:191-200. Poulicek, M. and C. Jeuniaux. 1991. Chitin biodegradation in marine environments: An experimental approach. Biochemical Systematics and Ecology. 19:385-394. Steel, R. G. D. and J. H. Torrie. 1980. Principles and procedures of statistics. McGraw-Hill Book Co., New York. Treagan, L and L Pulliam. 1982. Medical microbiology laboratory procedures. W. B. Saunders Company, Philadelphia, PA. p. 78-89. Varel, V. H. 1989. Reisolation and characterization of Clostridium Iongisporum, a ruminal sporeforming cellulolytic anaerobe. Arch. Microbiol. 152:209-214. Van Gylswyk, N. 0., E. J. Morris and H. J. EIS. 1980. Sporulation and cell wall structure of Clostridium polysaccharo/yticum comb. nov. (Formerly Fusobacten‘um polysaccharo/yticum). J. Gen. Microbiol. 121:491-493. Velez, S.A., J.C. Allen, C.M.Keery and R.W. Adkinson. 1991. Evaluation of crab and crawfish waste meals as protein sources for growing dairy heifers. J. Dairy Sci. 74:234-242. Yokoyama, M. T. and K. A. Johnson. 1988. Microbiology of the rumen and intestine, p.125-144. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. CHAPTER 4 CLOSTRIDIUM PARAPUTRIFICUM VAR. RUMINANTIUM : COLONIZATION AND DEGRADATION OF SHRIMP CARAPACES IN VITRO OBSERVED BY SCANNING ELECTRON MICROSCOPY 62 63 ABSTRACT The degradation of shrimp carapaces by Clostridium paraputrificum var, ruminantium as well as its morphology and growth on this chitinous substrate was observed by means of scanning electron microscopy. Bacteria growing on shrimp carapace showed vegetative cells 3 to 5 pm in length with rounded ends. Cells with terminal spores were larger averaging 7.3 pm in length. The spores were prominent between 1 to 2 pm in length and 0.7 to 1 pm in width. Filamentous forms were observed only when the substrate had been depleted. Incubation of Clostridium paraputn‘ficum var, ruminantium with pieces of shrimp carapace, was characterized by three major events. First, the bacteria attached to the inner surface of the shrimp carapace. Colonization and growth was only observed in the inner surface of the shrimp carapace. This activity was observed within the first 3 h after incubation and extended through 24 h of incubation. Between 24 to 72 h of incubation, bacterial growth was characterized by the degradation of the different multilayers of chitin that comprise the shrimp carapace to finally reach the outer surface. After 96 h of incubation, when the shrimp carapace had been almost totally degraded, the bacteria entered a death phase and latency, marked by the predominance of free spores and lysed bacteria. Since chitin degradation is initiated only after bacterial attachment occurs, a study of factors that influence this adhesion may result in more efficient use of chitinous waste as a feedstuff for ruminants. 64 INTRODUCTION The rumen of cattle has a volume of about 100 to 300 liters (Ogimoto and lmai, 1981) and a dry matter % of ruminal contents ranging between 7% to 14% (Owens and Goetsch, 1988). Therefore, the rumen can be essentially considered a liquid environment. As such, adherence of ruminal microbes to plant digesta is considered an important factor in the process of plant degradation (Rasmussen et al., 1989). At present, it is known that cellulolytic ruminal species, such as Ruminococcus flavefaciens, R. albus and Fibrobacter succinogenes, adhere to plant cell walls. In the same way, rumen amylolytic bacteria, such as Bacteroides amylophilus, B. ruminico/a and Bifldobacten’um sp. will attach to starch granules (Yokoyama, 1988). Presumably, this attachment to the insoluble substrate optimizes contact between enzymes associated with the cell envelope and the substrate (Enart, 1983), allowing for more efficient uptake of the soluble end products of hydrolysis by the bacteria. Previously, a spore-forming bacterium identified as Clostridium paraputn'ficum var ruminantium which has the ability to degrade Chitin was isolated from the rumen of dairy cows. This bacterium has a strong chitinolytic activity and can efficiently degrade almost 90% of the chitin in an anaerobic medium in just 3 d (see chapter 3). Although the intimate association of the bacterium with chitin suggested that it attached to chitin prior to degradation, similar to the attachment 65 of other bacteria to cellulose and starch, not much was known about the actual process. With the objectives to better characterize the morphology of the chitinolytic clostridial isolate and to better understand the process of chitin degradation by this bacterium, the present study was conducted using scanning electron microscopy. MATERIALS AND METHODS Organism and culture conditions. A chitinolytic bacterium previously isolated from the rumen of dairy cows, Clostridium paraputriflcum var. ruminantium, was use in this study. The bacterium was grown at 39°C in chitin (CH) medium (CH-medium components described in table 3.1). A 48 h culture of the bacterium growing in CH-medium was used as inoculum. The bacteria concentration in the inoculum, as estimated by the roll tube technique was about 1010 cells per ml. Shrlrnp carapace preparation forcolonizatlon study. Shrimp carapace (SC) were used as substrate to observe bacterial colonization rather than crab shell meal or pure chitin (Sigma), because in initial attempts, the latter two substrates did not provide enough contrast with the scanning electron microscope. The SC was washed in tap water to separate residues of meat, cut into pieces of about 0.5 cm’, and boiled in tap water for 10 min. Five small pieces of the boiled SC were placed into eight dacron bags (5 x 12.5 cm) with 50 pm pore Size (Ankom, Fairport, NY). Also, in order to observe for possible cellulose colonization and/or 66 degradation, one small piece of Whatman paper No. 4 (0.5 x 2 cm) was added to each bag. The dacron bags were placed into a 1 liter Erlenmeyer flask, containing 500 ml of anaerobic SC-medium. The SC-medium components are given in Table 1, and was prepared anaerobically according to the method of Hungate (1969) as modified by Bryant (1972). The SC-medium with the dacron bags, was inoculated with 5 ml of a 48 h culture of C. paraputrificum var. ruminantium, then incubated at 397°C. The dacron bags were recovered aseptically and consecutively after 0, 3, 6, 12, 24, 48, 72 and 96 h of incubation. The dacron bags were previously found to be very useful for a fast and aseptic recovery method for multiple samplings and that is the main reason for their use. Scanning electron microscopy. At appropriate time intervals, dacron bags were removed aseptically and the residual shrimp carapace were recovered, gently rinsed with distilled water, and immediately fixed for 1 h in 2% glutaraldehyde in 0.1 M phosphate buffer (pH 7.0). After fixation the samples were rinsed with 0.1 M phosphate buffer. Samples were then dehydrated with a graduated ethanol series (25, 50, 75, 90 and 100% ethanol), and critical point dried with C0,. Specimens were mounted on aluminum stubs, sputter-coated and examined with a JEOL JSM-3SC scanning electron microscope at the Center for Electron Optics of the M.S.U., following the procedures described by Klomparens et al. (1986). The same procedures were followed with the pieces of Whatman filter paper added to the same dacron bag for each period. Chitin determination. In order to have an idea of the initial content of chitin 67 in the shrimp carapaces used in this study, the content of chitin in the samples was determined by gas chromatography. The equipment used was a Hewlett Packard 5890A. The column (300 x 7.8) employed was a 100% methyl silicon column (BB1). The carrier was helium with a flow rate of 9.4 ml per min. The samples were prepared by gas chromatography following the procedures established by Dr, Hollingsworth in his laboratory in the Department of Biochemistry. Shrimp carapaces were cut nito small pieces and 3 mg digested with 20 pl of acetic acid and 50 pl of trifluoroacetic acid for 48 h at 70°C under nitrogen gas, to avoid the samples absorption of humidity. Then, N- acetilglucosamine residues and amino acids in the sample were acetylated with trifluoroacetic acid anhydride and methanol. A standard containing pure N-acetyl glucosamine (Sigma), was also prepared for gas chromatography. The chromatography system included an integrator which estimates the different signals from the amino acids and N-acetylglucosamine contained in the sample and translates the results as percent of the total nitrogen compounds. 68 Table 1. Composition of the shrimp carapace anaerobic medium used for observing bacteria attachment and degradation of chitin. Item Quantity per 100 ml Deionized water 52.6 ml Clarified ruminal liquid 30.0 ml Sodium carbonate 8% solution 5.0 ml Mineral solution 1‘I 5.0 ml Mineral solution 2° 5.0 ml Cysteine-sulfide solutionc 2.0 ml Resazurin 0.1% solution 0.1 ml Trypticase-peptone (LBBQ) 0.2 9 Yeast extract (LBBQ) 0.1 g ' Mineral solution 1 contains (per 1000 ml): K,HPO,, 6.5 9 ° Mineral solution 2 contains (per 1000 ml): KH,PO,, 6 g; (NH,),SO,, 6 9: NaCl, 12 g; MgSO,, 2.45 g and CaCI,o 2 H,O, 1.6 9 (Bryant and Robinson, 1962). ° Mix L-cysteine, 2.5 9 (dissolved in 15 ml of 2N NaOH); Na,S.9H,O, 2.5 g and resazurin, 0.1 ml (1% sol. in H,O). Then, bring the mix to a final volume of 100 ml with deionized water. Heat or microwave the mix until the resazurin indicator is colorless, seal the container and autoclave (bubbling with CO, is not required). RESULTS Fig. 1 shows structural characteristics of the shrimp carapace (SC) as to its outer surface, inner surface and lateral view, at time zero of inoculation. The outer surface (Fig. 1a) is an even surface with the eventual appearance of small appendages about 700 pm in length which look like the rhizoid structure of a 69 fungus. In contrast, the inner surface (Fig. 1b) is rough with honeycomb-like compartments present in the curved parts of the exoskeleton. Between the outer and Inner surface, there appears to be several layers of chitin (Fig. 1c), which together with the outer and inner surface layers make up the shrimp carapace. Fig. 2, is a series of scanning electron micrographs representing the major events which take place during the process of colonization and degradation of SC by C. paraputn'ficum var. ruminantium at different incubation periods. The results show that the bacterium attachs on the SC inner surface within the first 3 h of incubation (Fig. 2a), while the outer surface does not Show any evidence of bacteria colonization (Fig. 2b). After 6 h of incubation colonization at the inner surface has spread considerably (Fig. 2c). In contrast, the outer surface still remains uncolonized at the same time, even after the degradation of the external appendages is apparent (Fig. 2d). Strong evidences of the shrimp carapace digestion were observed after 9 h of incubation, as indicated by the presence of typical digestion grooves formed on the inner surface of the shrimp carapace (Fig. 2e). At the same time, colonization and degradation was also present on the different multilayers of chitin at the exposed sides of the shrimp carapace (Fig. 2f) and in addition, the bacteria have formed digestion pits that penetrate into the interior of the chitin layers (Fig. 2f). After 24 h of incubation, the ruminal Clostridium Showed profuse growth and colonization on the inner surface of the shrimp carapace (Fig. 3a). At the outer surface near cracked areas, bacterial growth is also evident just behind the outer layer, while the external surface remains unaltered (Fig. 3b). Between 48 to 72 70 h of incubation the presence of digestion pits that penetrate into the interior of the chitin layers can be observed (Fig. 3c). By 72 to 96 h of incubation, most of the shrimp carapace has been digested with the exception of the thin outer layer, which looks undigested, but very fragile, due to the lack of support (Fig. 3d). By 96 h of incubation, most of the substrate Is depleted, and the bacteria enter a distinct phase marked by the appearance of lysed bacteria and released spores indicating the senescence of the culture (Fig. 3e and f). While most of the shrimp carapace seems to be degraded by 4 d of incubation, the culture was checked to see if the bacteria would switch their enzymatic system to cellulose degradation as an alternative source of energy. However, the Whatman paper added was neither colonized nor degraded even by the end of the incubation period (7d of incubation). Cellular morphology. Vegetative cells were straight or slightly curved rods about 0.3 pm in wide by 3 to 5 pm in length with rounded ends, and occur singly or in pairs. Rods carrying spore were larger than the vegetative cells averaging 7.3 pm in length. Spores are oval, terminal and cause the cells to swell. The spores were about 1 to 2 pm in length and 0.7 to 1 pm in wide. Sporulation seems to be triggered by either the lack of substrate for growth or by the adverse build up of toxic metabolites in the medium. Also filamentous forms were observed mainly when the shrimp carapace had been degraded. The results of the gas chromatography study, indicated that the N-acetyl glucosamine content in the shrimp carapaces represented 37.0% of the organic matter. The ash content of the shrimp carapaces used in this study (on average 71 of 3 samples, determined by muffle furnace ashing) was 38.9%, therefore, the organic matter content of shrimp carapaces represent 61.1 % in dried samples. 72 Fig. 1. Structural characteristics of the shrimp carapace at its outer surface (fig. a; bar, 10 pm), inner surface (fig. b; bar, 10 pm) and a lateral view showing the different chitin layers that comprise its structure (fig. a: bar, 5 pm). 73 Fig. 2. Series of scanning electron micrographs of shrimp carapace incubated with C. paraputrificum var. ruminantium. (a) Morphological details of vegetative and spare former forms attached to the inner side of the shrimp carapace after 3 h of incubation. Bar, 2 pm. (b) View of the outer side at the same incubation period (3 h) with no signs of bacterial growth on its surface. Bar, 5 pm. (C) Degree of colonization after 6 h of incubation. Bar, 10 pm. (d) Outer surface of the shrimp carapace still uncolonized after 6 h of incubation. Bar, 10 pm. (e) Digestive grooves have been formed on the inner surface of the shrimp carapace after 9 h of incubation. Bar, 5 pm. (f) Strong colonization at the different layers that comprise the shrimp carapace was also observed by 9 h of incubation. Degradation of chitin is indicated by the presence of digestion pits that penetrate into the interior of the Chitin layers. Bar, 5 pm. 74 Fig. 3. Scanning electron micrographs of shrimp carapace incubated with ; paracutrificum var. ruminantium representing the major events that takes place between 24 and 72 h of incubation. (a) Profuse bacterial growth and colonization of the inner surface was observed by 24 h of incubation. Bar, 1pm. (b) At the same period of time, in cracked areas, is also evident the prodigal bacterial growth, behind the outer surface which remains undigested. Bar, 10 pm. (C) Between 48 to 72 h of incubation the degree of digestion is evident by the presence of digestion pits in most of the inner surface of the shrimp carapace. Bar, 1 pm. (d) By 72 to 96 h of incubation, most of the chitin layers have been digested with the exception of the cutest layer. Bar, 5 pm. (a and f) represent the senescence of the culture as indicated by the appearance of lysed bacteria and the high proportion of free spores. Bar, 1 pm. \‘W. r’-‘ K." ,I ‘ 1‘; 5‘..- _ I .é ‘\ x \I x ‘h 75 DISCUSSION The results of this study confirms the ability of C. paraputri'flcum var. ruminantium to bind to chitin. This characteristic is not particularly surprising if we consider the strong adhesive characteristic of chitin (Gooday, 1990), and the necessity for this attachment. The adhesion of C. paraputn'ficum var. ruminantium to chitin may be essential in aqueous environments such as the rumen ecosystem, where the substrate must be degraded to dimers or monomers before their efficient absorption can occur. Attachment would optimize contact between the chitinolytic enzymes and the chitin molecules. In addition, the N-acetylglucosamine and/or chitobiase produced from chitin degradation would be more efficiently absorbed. Without attachment, there is also the possibility that the chitinolytic enzymes may be degraded and assimilated by other ruminal bacteria, orthe N-acetylglucosamine released during chitin hydrolysis may be taken up by competing bacteria. The mechanisms of adhesion are not known, but Montgomery and Kirchman (1993) reported that the specific attachment to chitin particles appears to be mediated by chitin-binding proteins associated with the bacterial cell membrane. Another possible mechanism suggested is that lateral and polar flagella are responsible for cell attachment to binding sites on the chitin surface (Belas and Colwell, 1982). The external surface of the shrimp carapace, seems to have a protective 76 film or structural complexity which prevents bacterial attachment and is recalcitrant to degradation. This protective film may play a role in the defense of shrimp against bacterial or fungal infections. It is unknown if this protective film is also present in other shellfish wastes, but it may be wise to grind shellfish wastes when it is used as a feed for ruminants, to increase the attachment surface and the efficiency of degradation. Although not well documented, it is believed that the rate of chitin degradation is enhanced by the adherence of chitinolytic microflora (Gooday, 1990). Ou and Alexander (1974) observed that separation of chitin and chitinolytic bacteria by layers of microbeads greatly reduced the rate of mineralization of chitin particles by pure cultures of a soil Pseudomonas species. This phenomena has also been observed in Clostridium thermoceI/um, an anaerobic bacterium, which efficiently degrades a-crystalline cellulose after the cell adheres very strongly to the insoluble substrate (Bayer and Lamed, 1986). Since bacterial attachment to chitin seems to be essential for chitin degradation to occur, attention has been placed on possible plant components that inhibit or decrease bacterial binding and therefore the degradation of chitinous compounds. For example, wheat germ agglutinin (WGA), a chitin-binding lectin from wheat embryos exhibits binding specificity toward chitin (Bloch and Burger,1974). Wheat germ agglutinin lectin and other chitin-binding Iectins isolated from stinging nettle rhizomes are potent inhibitors of fungal growth. Inhibition is accomplished by crosslinking chitin chains in the fungi cell wall and inhibiting chitinases recognition of performed chitin Chains in the process of apical growth 77 of fungal hyphae (Broekaert, 1989). It is unknown if chitin-binding lectins are regular components of forages used as feedstuffs. Also, it is unknown if lectins are easily degraded by ruminal proteolytic bacteria. However, it is known that the binding of lectins to chitin, blocks proteolytic and B-N-acetyl-glucosaminidase activity on lectins and chitin, respectively (Knorr, 1991). Therefore, if lectins are present in the rumen of cattle fed diets rich in chitin by-products like shellfish waste, it is possible to obtain a negative effect on digestion of chitinous compounds and animal performance. Not much is known about the factors which affect the animal response to the intake of chitinous compounds, especially, those factors which affect the efficiency of the rumen chitinolytic microorganisms. However, the observation that bacterial attachment is essential for chitin hydrolysis, opens up a new research area concerning the possible factors that affect bacterial attachment to chitin and its consequence in the rumen ecosystem. 10. 11. 78 LITERATURE CITED Bayer, E. A. and R. Lamed. 1986. Ultrastructure of the cell surface cellulosome of Clostridium thermocellum and its interaction with cellulose. J. Bacteriol. 167:828-836. Belas, M. R. and R. R. Colwell. 1982. Adsorption kinetics of laterally and polarly flagellated Vibrio. J. Bacteriol. 151:1568-1580. Bloch, R. and M. M. Burger. 1974. Purification of wheat germ agglutinin using affinity chromatography of chitin. Biochem. Biophys. Res. Commun. 58:13-17. Broekaert, W. P., J. V. Parijs, F. Leyns, H. Joos and W. J. Peumans. 1989. A chitin-binding lectin from stinging nettle rhizomes with antifungal properties. Science. 245:1100-1102. Bryant, M. P. 1972. Commentary of the Hungate technique for culture of anaerobic bacteria. Amer. J. Clin. Nutr. 25:1324-1328. Bryant, M. P. and I. M. Robinson. 1962. Some nutritional characteristics of predominant culturable ruminal bacteria. J. Bacteriol. 64: 605-613. Enart, T. M. 1983. Microbial cellulases. In: W. M. Fogarty (Ed.). Microbial enzymes and technology. Applied Science Publishers, London. p. 183-223. Gooday, G. W. 1990. Physiology of microbial degradation of chitin and chitosan. Biodegradation 1: 177-190. Hungate, RE. 1969. A roll tube method for cultivation of strict anaerobes. In: J.R. Norris and D.W. Ribbons (ed). Methods in Microbiology. p.117-132. Academic Press Inc, NY. Klomparens, K. L, S. L Flegler and G. R. Hooper. 1986. Procedures for transmission and scanning electron microscopy for biological and medical science. Center for Electron Optics, Michigan State University. Knorr, D. 1991. Recovery and utilization of chitin and chitosan in food processing waste management. Food Technol. 45:116-122. 12. 13. 14. 15. 16. 17. 79 Montgomery, M. T. and D. L Kirchman. 1993. Role of chitin-binding proteins in the specific attachment of the marine bacterium Vibrio harveyi to chitin. Appl. Environ. Microbiol. 59: 373-379. Oglmoto, K and S. Imai. 1981. Atlas of rumen microbiology. Japan Scientific Societies Press, Tokyo. Owens, F. N. and A. L. Goetsch. 1989. Ruminal fermentation. p.145-171. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Ou, L and M. Alexander. 1974. Effect of glass microbeads on the microbial degradation of chitin. Soil Sci. 118: 164-167. Rasmussen, M. A., B. A. White and R. B. Hespell. 1989. Improved assay for quantitating adherence of ruminal bacteria to cellulose. Appl. Environ. Microbiol. 55:2089-2091. Yokoyama, M. T. and K. A. Johnson. 1988. Microbiology of the rumen and intestine, p.125-144. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. CHAPTER 5 RUMINAL BACTERIA COLONIZATION AND DEGRADATION OF SHRIMP CARAPACES IN SITU OBSERVED BY SCANNING ELECTRON MICROSCOPY. 80 81 ABSTRACT The objective of this study was to determine if ruminal bacteria are able to either attach and/or degrade chitin in the rumen. Dacron bags containing small pieces of shrimp carapace (SC), were placed in the rumen of a fistulated dairy cow and then removed at 0, 3, 6, 12, and 24 h, and then every day for 7 d. After in situ incubation, the retrieved samples were observed by scanning electron microscopy (SEM). Within the first 12 h, a slow succession of bacterial colonization was observed, primarily characterized by a cocci which attached to the SC by means of large appendages, a spirochete and a small rod. The rod appeared to produce digestion grooves in the shrimp carapace. Between 24 to 48 h of incubation, bacterial growth on SC was marked by an aggressive development of coccal colonies and also a rod attached to the inner surface of the SC. The massive development of these colonies limited an assessment of the degree of shrimp carapace degradation during this period, but a rich bacterial community was observed and signs of chitin digestion were evident. The two major bacterial types observed in this community, were found growing inside or in Close proximity to the digestion pits. One was a very thin, but long treponeme with tight coils and the second was a rod with spherical bodies attached polarly. Although, all the bacteria observed had unique morphological characteristics, they do not appear to have been previously described and it is assumed that they are part of a minor bacterial 82 fraction of the ruminal flora. The ruminal Clostridium previously isolated was not observed as part of this community although by the final day of incubation, vegetative cells were observed. The result obtained indicate that several bacterial species are associated with the process of shrimp carapace degradation in the rumen. Ruminal fungi and protozoa were not observed as being part of the chitinolytic community. The in situ technique supports the premise that the rumen of cattle never fed with chitinous feeds, has an indigenous flora able to colonize and degrade chitinous waste. The process seems to involve the concerted activity of a bacterial community rather than a few specific species, but the importance of each bacteria in the process (positive or negative) remains unknown until pure cultures are obtained. 83 INTRODUCTION By-products high in chitin such as shellfish wastes, are considered a potential alternative source of feed for productive animals. In addition, to their energy value for ruminants (2.56 to 2.85 Mcal per Kg of DM), shellfish wastes constitute also an excellent source of protein (30.6 to 43.06% crude protein) and minerals, specially of calcium carbonate (Brundage et al., 1981 and 1985 and Velez, 1991). The ability of ruminants to utilize chitin (Ortega and Church, 1979) or chitinous byproducts such as crab shell meal (Brundage et al., 1984), crawfish meal (Velez et al., 1991) or shrimp shell meal (Patton, 1975), has been studied. However, the ruminal microorganisms directly involved in the process of chitin degradation or digestion have never been studied or isolated. In Chapter 2 the isolation of certain ruminal bacteria which grew in an anaerobic medium with chitin as the only source of energy was reported. One of these bacteria was a spare- former classified as Clostridium paraputn’ficum var. ruminantium, which was a very active and specific Chitinolytic bacteria (see Chapter 3). Another ruminal bacteria isolated was a Peptastreptococcus sp. which was able to hydrolyse diacetylchitobiose and lactate. However, other bacteria in the selective enrichment culture could not be characterized, due to the difficulty of growing them in pure culture. It is conceivable that other ruminal bacteria species may be involved in the degradation of chitinous compounds such as shellfish wastes, as has been 84 reported in other anaerobic environments (Pel and Gotschal, 1986). In Chapter 4 it was demonstrated that bacterial attachment to shrimp carapace is a prerequisite for chitin degradation. The inference being that ruminal bacteria which can bind to chitin should be considered chitinolytic or at least part of a secondary community related to the degradation process. With the objective to determine if other ruminal bacteria unable to be grown in in vitro cultures, have the ability to attach and degrade chitinous compounds, small pieces of shrimp carapaces were introduced in the rumen of a dairy cow, retrieved at different times and observed by scanning electron microscopy. The other objective of this study was to detect if the chitinolytic ruminal Clostridium previously isolated is able to colonize and degrade the shrimp carapace as observed in vitro with pure cultures. MATERIALS AND METHODS Shrimp carapace preparation for in situ study. Shrimp carapaces (SC), were washed in tap water to separate residues of meat, cut into pieces of about 0.5 cm2, and boiled in tap water for 10 min. Five small pieces of the boiled SC were placed into 5 x 12.5 cm dacron bags with 50 pm pore size (Ankom, Fairport, NY). The dacron bags (11) containing the SC samples were sealed with black rubber stoppers and two rubber bands, then placed together into a larger (38 x 46 cm) nylon mesh bag as described by Schlegel (1993). In order to avoid a possible 85 involvement of contaminating bacteria, the mesh bag containing the small dacron bags was enclosed in aluminum foil and autoclaved 20 min at 15 PSI. Also, as was done before in the in vitro experiment (chapter 4), in order to observe for cellulose colonization and/or degradation, 3 small piece of Whatman filter paper No. 4 (0.5 x 2 cm) was added to each dacron bag. In situ technique. A rumen fistulated cow (Holstein breed) housed at the Michigan State University, Dairy Teaching and Research Center, previously used to isolate ruminal chitinolytic bacteria, was used for the present in situ study. The cow had been fed only on alfalfa hay, and had never previously been exposed to shellfish waste feeds. The mesh bag containing the small dacron bags, was placed into the rumen of the cow after the morning feeding. Inside the rumen, to ensure a similar starting time of exposure to ruminal microorganisms, the different dacron bags were carefully soaked with rumen liquid and then situated between solid ruminal contents. The dacron bags were consecutively recovered after 0, 3, 6, 12 h of incubation and then every 24 h for 7 d. To avoid or limit possible changes in the samples due to air exposure, between the period of time required for both sample transportation (eg. from the dairy farm to the laboratory) and sample processing, the removed dacron bag was placed into a sterile glass container (100 ml capacity) filled with rumen fluid just before sample collection. Scanning electron microscopy. In the laboratory, the dacron bags were open and the residual shrimp carapaces were gently rinsed with distilled water and Immediately fixed for1 h in 2% glutaraldehyde in 0.1 M phosphate buffer (pH 7.0). After fixation the samples were rinsed with 0.1 M phosphate buffer. Samples were 86 then dehydrated in graduated ethanol series (25, 50, 75, 90 and 100% ethanol), and critical point dried with CO, Specimens were mounted on aluminum stubs, sputter-coated and examined with a JEOL JSM-35C scanning electron microscope at the Center for Electron Optics of the MSU, following the procedures described by Klomparens et al. (1986). The same procedures were followed with the pieces of Whatman filter paper enclosed in the same dacron bag. 87 RESULTS SEM of shrimp carapace between 3 to 12 h of ruminal incubation. The results obtained show a slow succession of bacterial colonization of samples between 3 to 12 h of incubation. Three main bacteria types were attached to the surface of the shrimp carapace. Bacterial colonization started in the inner surface of the shrimp carapaces, as was observed with the in vitro experiment (Chapter 4), but one pleomorphic coccal bacterium was able to attach to the outer surface of the shrimp carapace by means of a slime-like material that resembles long appendages. The cocci was about 1 to 1.7 pm in diameter. The appendages around its body (Fig.1a) could be an artifact produced during the process of the sample critical pain dried (Dr. Stanley Flegler, 1994, personal comunication, Center for Electron Optics, MSU). Other attached bacterium observed was a small Treponema sp. about 0.15 pm in diameter and 3.5 to 4.5 pm in length. The treponeme had less than four complete turns of its helix (Fig. 1b) and in liquid medium, exhibited a very active serpentine type movement. The third type of bacterium observed during this period was a small rod about 0.7 pm in width and 1.5 to 1.8 pm in length, usually growing in pairs. Scanning electron micrographs of this rod showed the first evidence of chitinolytic activity. It appears that after attachment to the shrimp carapace, these rods secrete Chitinolytic enzymes 5U CE (1'. 88 through their cell walls, given the bacteria an appearance of digging into the inner surface of the shrimp carapace (Fig. 1c). Although other bacteria were observed, the three bacterial species described, were the only ones with a sustained presence in different samples from 3 to 12 hr of incubation. Compared with the in vitro experiment, bacteria colonization in situ was slower and less uniform. SEM of shrimp carapace alter 1 to 2 d of ruminal incubation. The most important feature during this incubation period was the rapid development of two types of branched colonies. One, was the coccal bacteria previously described. Its growth tended to cover most of the inner surface of the shrimp carapace. In very densely colonized regions, the colonies were covered with a fibrillar and thick slime-like substance, which did not allow for an accurate assessment of the degree of chitin degradation behind the attached cells (Fig. 1d). The second colony observed during this incubation period, was a rod shaped bacterium, 0.8 pm in width and 1.5 to 3.0 pm in length, growing single or in pairs. This bacterium showed a very aggressive growth, but only on the inner surface of the shrimp carapace. This rod attached to the shrimp carapace surface by means of thick slime-like substance, that after critical point drying of the samples looks like a thick peritrichous appendages (Fig. 1e). The third bacterium observed was a very coiled, large and thin spirochete about 100 nm in diameter and up to 10 pm in length with very tight spirals (Fig. 1f). In the same photograph a certain degree of shrimp carapace digestion is observed. 89 Fig. 1. Series of scanning electron micrographs of Shrimp carapaces after 3 to 48 h of incubation in the rumen. (a) Morphological details of a coccal bacterium attached to the inner side of the shrimp carapace by means of a thick slime-like substance resembling long appendages. Bar, 1 pm. (b) Morphological details of an small treponeme. Bar, 1 pm. (C) Morphological details of a small rod whit appearance of digging the inner surface of the shrimp carapace. Bar, 10 p. (d) Outer surface of the shrimp carapace with notable degree of bacteria colonization. Bar, 5 pm. (e) Rod shaped bacteria attached to the shrimp carapace by means of peritrichous appendages. Bar, 1 pm. (f) Morphological details of a second treponeme species attached to the inner size of the shrimp carapace. Degradation of Chitin is suggested by the fibrous structure of the shrimp carapace behind the bacteria. Bar, 2 pm. . MW i- I rea‘S 903m 90 SEM of shrimp carapace after 3 to 7 d of ruminal incubation. A regular pattern of shrimp carapace colonization and degradation was not found. As such, samples with only 3 d of incubation showed strong physical evidences of chitin degradation in some areas of the shrimp carapace, while other areas on the same sample, were either never colonized or degraded. In addition, it was impossible to determine which ruminal bacteria was predominant at a particular time. The same irregular pattern was observed throughout the entire study. Therefore, the results are focused on the presence and characteristics of the different bacteria observed during the process of chitin degradation, without implying that a particular species was predominant or determining the time at which the shrimp carapace colonization and degradation reached a peak. When discernible, a rich bacterial community was observed behind the slime material produced by the cocci. Forming this community were primarily two spirochetes. One, was a very thin and large with very tight spirals described previously. The second spirochete was smaller about 0.15 pm in diameter and 3.5 to 4.5 pm in length (Fig. 2a). This spirochete resembled the morphological characteristics of the genus Treponema and was found in much lower concentration. This spirochete was previously observed among the bacteria seen between 3 to 12 h of incubation (Fig. 1b), and also among the ruminal bacteria isolated in chitin medium (Chapter 3). The larger spirochete was observed in the digestive pits that penetrate directly into the interior of the different layers of chitin that comprise the shrimp carapace (Fig. 2b). 91 Another bacterium associated with the digestive pits was a rod about 2 pm in length and 0.6 pm in width, lightly curved with one end rounded and the other tapered. This bacterium exhibited a unique characteristic, the presence of two or three spherical bodies at the tapered end (Fig. 2c and 2d), but the genus to which it belongs is unknown. The spirochetes and the rod with spherical bodies in Close association with the digestion pits as well as the coccal and the rod with peritrichous appendages which usually cover the shrimp carapace, seems to coexist as a mixed colony on the inner surface of the shrimp carapace. The small rod detected at the beginning of the incubation period (Fig. 10) was again observed, in this occasion, their rich growth on the inner surface of the shrimp carapace give the appearance to start degrading the slime material produced by the cocci type bacterium, in which was enclosed (fig 3e). Although by the end of the experiment were observed rods exhibiting morphological characteristics similar to those of the vegetative cells of C. paraputri'fi'cum var. ruminantium (Fig. 3f), the ruminal clostridium was not observed as part of the diverse community. In summary, a series of scanning electron micrographs (Fig. 3a, b, C and d) show the degradation of the outer surface of shrimp carapaces during their residence in the rumen. In contrast with the inner surface, the outer surface was not heavily colonized, even with the coccal type, but signs of degradation were observed by the first 12 h of incubation (Fig. 3a, b). After 24 to 48 h of Incubation, several large holes which were colonized by bacteria, could be observed (Fig. 3c), but there was no colonization of the outer surface. Finally, after 3 to 5 d of 92 incubation the outer surface showed evidence of both colonization and degradation (Fig. 4d). The small pieces of Whatman filter paper placed together with the shrimp carapaces, were totally degraded or shredded in such way that after 24 h of incubation, they were impossible to find, indicating the normal capability of some ruminal bacteria to degrade cellulose. 93 Fig. 2. Scanning electron micrographs of shrimp carapace incubated in the rumen, representing the major events that takes place between 3 to 7 d of incubation. (a) A rich bacterial community grew and colonized the inner surface. Bar, 1pm. (b, C and d) Usually, covered by slime-like material two treponemes species and a rod with spherical bodies were observed inside digestive pits. Bar, 1 pm. (e) The small rod observed since the beginning of the in situ study, seems to growth apart from the bacterial community. Bar, 2 pm. (i) After 7 d of incubation, a bacteria with morphological Characteristics similar to those of the vegetative cells of C. paraoutrificum var. rumina_ntium were detected. Bar, 2 pm. ' A” -__.q-—-— qr- .- "‘3’? \«W 2‘ 2.”: A}? il‘ ‘ Pa : 3V ' 9. 3“"? ‘ . .. . If}? - ’ - . ‘ o r . 94 Fig. 3. Series of scanning electron micrographs of the shrimp carapaces showing their degree of colonization and degradation of the outer surface after: (a) 12 h of incubation. Bar, 2 pm; (b and c) 24 to 48 h of incubation. Bar, 5 and 10 pm. and (d) 72 to 96 h of incubation. Bar, 5 pm. In general, irregular and relatively low bacterial colonization was observed through out the entire study. 95 DISCUSSION It is clear from this investigation that ruminal bacteria are able to attach to chitin particles introduced into the rumen of cattle, and furthermore that these bacteria are engaged in the shrimp carapaces degradation. However, considering the nature of this experiment, which was based solely on SEM observations, the ability to degrade chitin can only be attributed to the small rod growing in pairs, which gave strong physical evidences of their capability (Fig. 1c). With the other observed bacteria it will be necessary to obtain them in pure cultures to determine their chitinolytic capability. Anaerobic bacteria from marine environments, have the ability to adhere to chitin particles and utilize N-acetylglucosamine, but do not degrade chitin (Kaneko and Colwell, 1978). This characteristic may apply to the coccal and the rod shaped bacteria which were attached to the shrimp carapace by means of a slime- like material. Also, it is possible that this material was an artifact produced by the scanning electron microscopy. With respect to the occurrence of spirochetes, this is the second report in which spirochetes have been observed in anoxic Chitinolytic communities. In anaerobic sediments of Lake Erie, spirochetes have been observed as part of the succession microorganisms involved in the process of chitin degradation (Warnes, et al., 1977). According to current taxonomic criteria (Canale-Parola, 1986), the 96 two spirochetes observed in this experiment, are appropriately assigned to the genus Treponema inasmuch as they are strict anaerobes and host-associated. Treponemes, are common inhabitants of the bovine rumen and their occurrence in large numbers (108 cells per ml of ruminal fluid) is considered an indication that the contributions of these bacteria to the metabolic processes in the rumen are quantitatively significant (Stanton and Canale, 1979). With exception of one large ruminal treponeme which is able to degrade pectin (Ziolecki, 1979), ruminal treponemes are unable to degrade polymers, and will only ferment a limited number of soluble carbohydrates. As such, treponemes depend on the metabolic activities of the animal host and of other microorganisms present in the rumen to obtain carbon dioxide, B-vitamins and short chain fatty acids which are essential for their growth (Canale-Parola, 1986). Although there is no supporting evidences for the ability of ruminal treponemes to degrade polymers such as cellulose, starch or chitin, it is known that Treponema bryantii (a ruminal species) enhances cellulose breakdown by Fibrobacter succinogenes (Stanton and Canale, 1980). The mechanism responsible for the enhancement of cellulose breakdown is not understood, but it is known that cellulose fibers are more accessible to F. succinogenes cells in the presence of spirochetes than in their absence. Therefore, it is plausible to expect a similar role for the treponemes observed on the shrimp carapaces during the process of chitin degradation. It is possible that the end products of chitin fermentation produced by chitinolytic species, serve as Chemoattractants for the treponemes. The constant movement of treponemes species searching for these and products, may Increase the width and depth of the 97 digestive pits produced by the chitinolytic bacteria as observed in the chitin layers of the shrimp carapace (Fig. 2b). The treponemes may also randomly push chitinolytic bacteria away from the remains of partially hydrolyzed chitin chains and move them toward uncolonized layers of chitin, resulting in an enhancement of chitinolytic activity. The rod with spherical bodies was the last bacteria observed in association with the digestion pits, but as with the spirochetes, it is unknown if this rod plays a primary or secondary role in the process of chitin degradation. Despite their peculiar morphological characteristics, and with the exception of the small spirochetes (Fig. 1b) which resemble the genus Treponema, all the other bacteria observed do not appear to have been previously reported or described. Bacterial diversity within the rumen ecosystem is very great with more than 22 genera and 63 species (Yokoyama and Johnson, 1988). However, only about 20 species with numbers greater than 107 per g of ruminal contents are considered the major players in the population (Russell, 1986). It is, very possible that the microorganisms observed colonizing the shrimp carapaces, are part of the minor fraction of the rumen bacterial population. Assuming that the Chitinolytic bacterial community is part of the minor ruminal bacteria population, the time required for this community to develop as the major bacterial species in the rumen, would represent the long adaptation period (up to 2 months) required by ruminants fed with chitinous waste, to start showing a positive response (Laflame, 1988; Brundage et al., 1984 and White, 1981). In contrast with the earlier scanning electron microscopy studies with pure cultures, presented in Chapter 4, in this experiment: (1) it was impossible to 98 establish a sequence of events due to the inherent limitations of the experiment, (2) C. paraputri'fi'cum var. ruminantium was not among the dominant bacterial species detected in the community, (3) bacterial colonization and degradation of the different layers of chitin that comprise the shrimp carapace proceeded much slower and irregularly than in pure cultures. The identification of the ruminal bacteria forming the chitinolytic community is still pending, but their unusual morphological details like large spirochetes with very tight coils and rods with spherical bodies, leads us to believe that they are part of a minor ruminal bacteria fraction. 10. 99 LITERATURE CITED Brundage, A.L, Husby, F.M., Herlugson, M.L, Simpson,W.L and Burton, V.L. 1984. Acceptability of Tanner crab meal in concentrates for lactation. J. Dairy Sci. 67: 1965-1970. Brundage, A. L, F. M. Husby, G. L Beardsley and V. L Burlin. 1981. King crab meal in concentrates for lacting cows. J. Dairy Science. 64:433- 440. CanaIe-Parole, E. 1986. The Spirochetes. p. 38-70. In: N. R. Krieg and J. G. Holt. Bergey's manuaal of systematic bacteriology. Williams & Vtfilkins. Baltimore. MD. Kaneko, T. and R. R. Colwell. 1978. The annual cycle of Vibrio parahaemo/yti'cus in Chesapeake Bay. Microb. Ecol. 4:135-155. Laflame, LF. 1988. Utilization of crab meal fed to young beef cattle. Can. J. Anim. Sci. 68: 1237-1244. Ortega, E. and Church,.D.C. 1979. In vitro digestion studies with crab meal and related chitinous compounds as feedstuffs for cattle. Proc. West. Sec. Am. Soc. Anim. Sci. 30:302-305. Patton, R.S. and Chandler P.T. 1975. In vivo digestibility of chitinous materials. J. Dairy Sci. 58:397-403. Pel, R. and J. C. Gomschal. 1986. Mesophilic chitin degrading anaerobes isolated from an esturine environment. FEMS Microbiol. Ecol. 38:39-49. Stanton, T. B. and E. CanaIe-Parola. 1980. Treponema bryantii sp. nov., a rumen spirochete that interacts with cellulolytic bacteria. Arch. Microbiol. 127:145-156. Russell, J. B. 1986. Ecology of rumen microorganisms: energy use. p.74- 98. In: A. Dobson and M. J. Dobson (ed). Aspects of digestive physiology in ruminants. Comstock Publishing Associates. Cornell University Press, Ithaca. 11. 12. 13. 14. 15. 16. 17. 100 Stanton, T. B. and E. CanaIe-Parola. 1979. Enumeration and selective isolation of rumen spirochetes. Appl. Environ. Microbiol. 38:965-973. Schlegel, M. L 1993. Grazing systems for direct-seeded alfalfa pastures. p. 57. MS Thesis, Animal Science Department, Michigan State University. Velez, S.A., J.C. Allen, C.M.Keery and R.W. Adkinson. 1991. Evaluation of crab and crawfish waste meals as protein sources for growing dairy heifers. J. Dairy Sci. 74:234-242. Wames, C. A. and C. l. Randles. 1977. Prelyminary studies on chitin decomposition in Lake Erie sediments. Ohio J. Sci. 77:224-230. White, R. L. 1981. Ruminant utilization of chitin and other components of tanner crab meal. M.S. thesis, Univ. of Alaska, Fairbanks, Alaska. Yokoyama, M. T and K. A. Johnson. 1988. Microbiology of the rumen and intestine, p.125-144. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Ziolecki, A. 1979. Isolation and characterization of large treponemes from the bovine rumen. Appl. Environ. Microbiol. 37:131-135. CHAPTER 6 COMPARISON OF IN VITRO DEGRADABILITY OF CRAB SHELL, PURE CHITIN AND ALFALFA HAY BY CLOSTRIDIUM PARAPUTRIFICUM VAR. RUMINANTIUM AND BY MIXED BACTERIA FROM THE RUMEN OF DAIRY CATTLE. 101 102 ABSTRACT The chitinolytic activity of bacteria from the ruminal fluid of a dairy cow never previously exposed to any shellfish waste feed, was compared to that of C. paraputn'ficum. var. ruminantium, a chitinolytic bacterium isolated from the ruminal fluid of the same cow. Pure chitin and crab shell meal were used as substrates. Also, alfalfa hay was used to test the bacteria capability to degrade a common substrate of ruminant's ration. All comparison were based on the in vitro organic matter degradability of the substrates after 2, 4, 6, 12, 24, 36 and 48 h of incubation. Differences (P<.05) in the organic matter degraded were observed after 4, 12 and 24 h of incubation in alfalfa hay, crab shell and chitin respectively, but they were more evident (p<.05) by the end of the incubation period. Pure chitin was more extensively degraded by the bacteria culture than by the mixed bacteria in ruminal fluid (63.25 vs 30.55 %). Similar differences were observed with the crab shell meal ( 75.35 vs 44.71%). In contrast, alfalfa hay was better degraded (P<.05) by the bacteria in ruminal fluid than by C. paraputn'ficum. var. ruminantium (62.04 vs 44.86%). The pure culture resulted in very efficient degradation of chitin and crab shell with a mean degradation rate after 48h of incubation of 316 and 377 mg per gram of substrate per day (respectively). Considering its efficiency, C. paraputrificum. var. ruminantium may have potential as a ruminal inoculum to increase the efficiency of shellfish waste degradation in the rumen. 103 INTRODUCTION Shellfish processing wastes (including crab, shrimp, crawfish, lobster, etc.) are a rich source of chitin which is currently underutilized. Shellfish wastes are the main source of pure chitin, chitosan and other chitin-derivatives, which have found many industrial, medical and environmental applications (Muzzarelli, 1993; Skjak- Braek et al., 1989; and Wright and Retnakaran, 1987). Despite the numerous uses of chitin, the amount of shellfish wastes generated by the marine industry far exceeds demand, and proper disposal is considered a marine environmental pollution problem. Considering their nutritive value (Brundage et al., 1981 and 1984; and Velez, 1991), the use of these shellfish wastes as a feedstuff for animal production represents a potential alternative for the surplus supply. In Chapter 3 the isolation of an anaerobic chitin-degrading ruminal bacteria, belonging to the genus Clostridium was reported. This bacterium was isolated from a chitinolytic community, which had been enriched from the ruminal fluid of dairy cows using Chitin as the sole energy substrate. Based on its characteristics, this bacterium was tentatively identified as C. paraputn‘ficum var. ruminantium. Scanning electron microscopy studies indicated that a pure culture of this bacterium would attach and degrade the native Chitin in shrimp carapace, more efficiently than a mixed bacterial community from the rumen (Chapter 4 and 5). However, there was no quantitative support for this conclusion. The purpose of 104 the present study was to quantitatively compare the differences in chitin degradation between mixed ruminal bacteria and a pure culture of C. paraputrifi'cum var. ruminantium. In addition, their capability to degrade alfalfa hay was also compared. The comparison was based on the extent of the substrates organic matter (OM) degraded, using the in vitro procedure described by Tilley and Terry (1963). This procedure is commonly used in forage evaluation systems to appraise ruminal digestibility. In vitro organic matter degraded (IVOMD) measured by this procedure is highly correlated with in vivo digestibility. The results of this study will provide a better understanding about the potential use of the ruminal clostridial isolate to increase the utilization of chitinous waste by the animal. MATERIALS AND METHODS The substrates used in this study were: pure chitin powder (Sigma Chemical Co.), crab shell meal from Alaskan Tanner crab (Chionocetes bairi'di) kindly supplied by Dr. Fred Husby (University of Alaska, Agricultural Experiment Station, Fairbanks), and alfalfa hay. Either, ruminal fluid or a pure culture of C, paraputrificum var. ruminantium were used as inocula. The ruminal fluid was collected from a rumen fistulated dairy cow just prior to the morning feeding. The cow was being fed alfalfa hay and had never been exposed to any chitin-rich feeds. Ruminal fluid was strained through 4 layers of cheesecloth, sparged with 105 carbon dioxide. A pure culture of C. paraputrificum var. ruminantium in exponential phase (10° cells per ml) was used as inoculum In a second series. In vitro incubation of substrates followed the first stage of digestion technique as described by Tilley and Terry (1963) which corresponds to the ruminal degradation phase. Prior to inoculation, 0.5 9 samples of pure chitin, crab shell meal, and alfalfa hay substrates were weighed in triplicate and placed in 100 ml round bottom glass tubes. One set of samples was weighed for each incubation time period (2, 4, 6, 12, 24, 36 and 48 h). McDougall's saliva (McDougalI, 1948), containing urea (0.5 g per liter) and held at 39°C in a water bath, was mixed with either ruminal fluid or the pure culture of C. paraputriflcum var. ruminantium (1:1 v/v) and saturated with carbon dioxide. The mixes were added to each tube in 40 ml aliquots (eg. 20 ml McDougall saliva and 20 ml of ruminal fluid or pure culture of the clostridial species). The samples were stoppered with rubber plugs fashioned with gas escape valves to allow for gas production. At completion of the incubation periods, the residual organic matter was calculated for each sample as described by Tilley and Terry (1963). The inocula tested were compared using the in vitro organic matter degraded rather than the in vitro dry matter degraded, due to the high differences on ash content among chitin, alfalfa hay and crab shell meal (1.78, 9.23 and 36.6%, respectively). Statistical analysis were performed by least-squares analysis of variances in a completely randomized design (Steel and Torrie, 1980). 106 RESULTS In vitro organic matter degradation (IVOMD) of pure chitin was not significantly different (P>0.5) between the ruminal fluid inocula and the clostridial inocula for the first 12 h of incubation. However, after this period the percent of organic matter disappearance was significantly (P<.05) increased with the clostridial inocula in comparison to the ruminal fluid inocula (Table 1 and Fig. 1). On the average, after 48 h of incubation, the degradation of chitin was twice as high with samples inoculated with C. paraputrificum var. ruminantium than with ruminal fluid as inocula (63.25 vs 30.55%, respectively). With the crab shell meal, there were no differences (P>0.5) due to inocula during the first 4 h of incubation, but starting with the 6 h through the end of the incubation period (48 h), samples inoculated with C. paraputrificum var. ruminantium were more effectively degraded (P<.05) than samples inoculated with ruminal fluid. On average, after 48 h of incubation, the IVOMD from crab shell meal inoculated with C. paraputrificum var. ruminantium was about 1.7 times higher than the IVOMD reached with the samples inoculated with ruminal fluid (Table 2 and Fig. 2). In contrast to pure chitin, crab shell meal inoculated with either ruminal fluid or the clostridial culture, required only 6 h of incubation to reach an IVOMD similar to that reached on pure chitin after 36 h of incubation, and the extent of organic matter disappearance was higher after 48 h of incubation (see Table 1 and 2). The IVOMD results obtained 107 with alfalfa hay as substrate are presented in Table 3 and Fig. 3. In contrast to pure chitin, the IVOMD of alfalfa hay inoculated with ruminal fluid was higher (P.05) were observed between alfalfa hay and crab shell during the first 12 h of incubation. Chitin was the less efficiently utilized substrate (P<.05). Both inocula were more efficient in degrading crab shell than pure chitin (P<.05). 108 Table 1. Effect of inocula type on in vitro organic matter degradation of chitin, over time. — l n o c u l a lncubafion _ . __ time, h Ruminal flU|d Clostridial SEM Isolate 2 1.87 2.14 0.21 4 2.02 3.38 0.73 6 2.65 4.11 0.70 12 6.58 7.49 0.67 24 8.24‘I 14.63b 1.20 36 18.653 26.88b 2.17 48 30.55“ 63.25” 0.44 ll Means with unlike superscripts differ (P<.53) 70 n - Ruminalfluid °\° 60 F + c“ Clostridium 53? —e— E 50 c O) 8 L 40 ’— G) 3:: E .2 30 F / C / // 15 20 _ /, ,-/ // 2 / //1 g ///*“’ / E 10 F 1/1/::£;’/’_///'/ 0 Kip:l '// l l 1 l 2 4 6 12 24 36 48 Incubation time, h Fig. 1. Organic matter degradation of chitin. In vitro organic matter degradation, % 109 Effect of inocula type on in vitro organic matter degradation of crab shell meal, over time. — Table 2. I n o c u I a lncubafion , _ ,. time, h Ruminal flund Clostridial SEM Isolate 2 2.32 4.06 0.56 4 7.85 10.04 1.80 6 15.513 19.27” 0.96 12 17.66“ 34.06” 2.54 24 20.918 43.83” 0.94 36 41 .42a 55.40b 2.43 48 44.71“‘ 75.35b 0.77 ans with unlike superscripts differ ( <.0 ) 80 v Q Ruminal fluid .// + // Clostridium 60 — +3“ [,0 ///l/ a l \ \\ \‘h 20 — //..////‘ %/ 0 l l l l l l 2 4 6 12 24 36 48 Incubation time, h Fig. 2. Organic matter degradation of crab shell meal In vitro organic matter degradation, °/o 110 Table 3. Effect of inocula type on in vitro organic matter degradation of alfalfa hay, over time. _ l n o c u I a lncubafion time, h Ruminal fluid Clostridial SEM Isolated 2 3.52 1.87 1.10 4 10.05“ 3.80” 0.95 06 15.12 9.93 3.08 12 18.86 13.67 3.22 24 41.62“ 29.50b 1.51 36 53.84“ 26.67b 0.72 48 62.04“ 44.86b 2.48 Means with unlike superscripts differ (P<.0 ) 70 F Ruminal fluid >- /I + 60 / Clostridium /r —e— 50 ~ / t // /Q) , . 40 +— / / 30 7 // 7 // // / / 20 ~ /./ / _ /I’// / 10 r //./ fl/L/ L 177/ O l l l l l l 2 4 6 12 24 36 48 Incubation time, h Fig. 3. Organic matter degradation of alfalfa hay. 111 Table 4. Effect of inocula type on in vitro organic matter degradation of chitin, crab shell meal and alfalfa hay. Incubation C. paraputrl'ficum var. ruminantium time h Chitin Crab shell Alfalfa hay SEM 2.14“ 4.06b 1.87“ 0.43 3.38“ 10.04b 3.80“ 0.76 4.11“ 19.27“ 9.93 b 1.29 12 7.49“I 34.06“ 13.67 b 1.47 24 14.63“ 43.83“ 29.50b 1.31 36 26.88“ 55.40b 26.67“ 1.87 48 63.25b 75.35“ 44.86“ 4.15 Bacteria from ruminal fluid 1.87“ 2.32“ 3.52b 0.72 2.02“I 7.85“ 10.05“ 1.25 2.65“ 15.51“ 15.12b 1.90 12 6.58“ 17.66b 18.86b 2.40 24 8.24“ 20.91“ 41.62“ 1.24 36 18.65“ 41 .22b 53.84“ 1.93 48 30.55“ 44.71b 62.04“ 1.52 “Means Wlthln the same inocula type with diaerent supescn'pts, difier ”<55; 112 DISCUSSION The type of inocula used (Fig. 6.1, 6.2 and 63) gave the expected result and confirmed the higher efficiency of C. paraputn'ficum var. ruminantium for chitin or shellfish waste degradation as compared to ruminal fluid (P<.05). In contrast, alfalfa hay was better degraded (P<.05) by the bacteria in ruminal fluid than by the pure culture of C. paraputn'ficum var. ruminantium (62.04 vs 44.86%, respectively) suggesting that alfalfa hay is more susceptible to microbial degradation than chitin or crab shell in the rumen. The more extensive degradation of crab shell meal over chitin has been reported for in vitro (Ortega and Church, 1979) and in vivo (Patton and Chandler, 1975) experiments with dairy cattle. The reason seems to be related to their chitin content. Crab shell meal consists not only of chitin, but also viscera, unextracted meat, and minerals (White, 1981). As such, chitin content in crab shell meal is only between 13.5% to 20.3% of the dry matter (Velez et al., 1991 and Laflame, 1988, respectively), while with pure chitin (Sigma) about 98 to 99% of the dry matter is chitin. The major component of the mineral fraction in crab shell is calcium carbonate and, to a lesser extent, calcium phosphate (Muzzarelli, 1977). These salts are considered to be negatively correlated with the extent of crab shell degradation (Patton et al., 1975). However, the results of this study, suggest that 113 the degradation rate of crab shell meal was not adversely affected by its high mineral concentration (37% of DM). The poor degradation of alfalfa hay in relation to chitin or crab shell meal, supports the previous conclusion made in Chapter 3 with respect to the high specificity of C. paraputriflcum var. ruminantium to degrade chitin-rich substrates. This ruminal chitinolytic bacteria efficiently degraded crab shell meal with a mean degradation rate (considering the IVOMD after 48 h of incubation) of 377 mg per gram of substrate per day. This value seems to be much higher in comparison to other chitin degrading bacterial species. For example, Boyer and Kator (1985) estimated the in vitro rate of degradation of crab shell by marine bacteria, as being a maximum rate of 207 mg per gram of substrate per day. In the preceding chapter it was shown that in the rumen of dairy cattle never having been fed any chitinous feeds, there exist a ruminal bacteria consortia which is able to attach and degrade shrimp carapaces which major component is chitin. In this chapter, the chitinolytic ability of these bacteria are quantitatively demonstrated by the results obtained from the IVOM D of pure chitin. However, these indigenous chitin degrading ruminal microbes, are not as efficient as the chitin degrading strain of C. paraputrificum var. ruminantium previously isolated from the rumen. Apparently the poor competitiveness of these ruminal bacteria for chitin degradation, may reflect a better adaptation to the strongly fluctuating food conditions experienced in the rumen. This hypothesis is supported by the fact that the previously isolated ruminal clostridium was not one of the dominant species associated with the shrimp carapaces incubated in the rumen 114 (Chapter 5). The clostridium, being highly selective for chitin, may be unable to compete in the rumen with other bacteria which can utilize other alternative substrates besides chitin. It should be noted that ruminal clostridial species represent a minor fraction of the ruminal microflora (Kelly, et al., 1987). It has been postulated (Gottschalk et al., 1981) that clostridia may be at a disadvantage in environments with a constant nutrient supply, such as the rumen, because of the energy required to maintain the genetic information for spore production. At this moment it is not known if C. paraputn'ficum var. ruminantium is one of the predominant chitinolytic bacteria in the rumen of animals adapted to the intake of chitinous feeds. Based on in vivo and in vitro experiments with dairy cattle, White (1981) concluded that the chitinous feed must be present in the animal's diet for at least 6 weeks before the full potential of the chitinolytic ruminal microflora can be demonstrated. Considering the efficiency to degrade pure chitin and crab waste meal, C. paraputn'ficum var. ruminantium may have potentials as a ruminal inoculum to increase the efficiency of shellfish waste degradation in the rumen. However, more research about this potential is required and future research needs to focus on, (1) the determination and isolation of microflora which produce chitinase in animals adapted to the intake of shellfish waste, (2) the ability of C. paraputrificum var rumiantium to become a dominant chitinolytic bacteria in animals adapted to chitinous feeds in their diet, (3) the potential use of this clostridia as an inoculum to enhance ruminal chitin-rich substrates degradation, and (4) the effect of C. 115 paraputrificum var. ruminantium inoculum on animal performance during and after adaptation to the feeding of shellfish waste. 10. 116 LITERATURE CITED Boyer, J. N. and H. I. Kator. 1985. Method for measuring microbial degradation and mineralization of 1‘C—labelled chitin obtained from the blue crab, Cal/inectes sapidus, Microb. Ecol. 11:185-192. Brundage, A.L, Husby, F.M., Henugson, M.L, Simpson,W.L. and Burton, V.L. 1984. Acceptability of Tanner crab meal in concentrates for lactation. J. Dairy Sci. 67: 1965-1970. Brundage, A. L, F. M. Husby, G. L. Beardsley and V. L Burfln. 1981. King crab meal in concentrates for lacting cows. J. Dairy Science. 64:433- 440. Godsehalk, G., J. R. Anderson and H. HIpe. 1981. The genus Clostridium (nonmedical aspects), p. 1767-1803. In: Starr M. P., H. Stolp, H. G. Truper, A. Balows and H. G. Schlegel (eds). The prokaryotes, vol II. Springer, BerlinHeilderberg, New York. Husby, F. M. 1993. Professor, University of Alaska, Agricultural Experiment Station, Fairbanks 99701. Kelly, W. J., R. V. Asmundson and D. H. Hoperoft. 1987. Isolation and characterization of a strictly anaerobic, cellulolytic spore former: Clostridium chartatabidum sp. nov. Arch. Microbiol. 147:169-173. Laflame, L F. 1988. Utilization of crab meal fed to young beef cattle, Can. J. Anim. Sci. 68:1237-1244. McDougaII, E. J. 1948. Studies of ruminal saliva. 1. The composition and output of sheep's saliva. Biochem. J. 43:49. Muzzarelli, R. A. 1993. Advances in N-acetyl-B-D-glucosaminidases. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 357-374. Muzzarelli, R. A. 1977. Chitin. Pergamon Press, LTD. Oxford. 11. 12. 13. 14. 15. 16. 17. 18. 117 Ortega, E. and D. C. Church. 1975. In vitro digestion studies with crab meal and related chitinous compounds as feedstuffs for cattle. Proc. West. Sec. Am. Soc. Anim. Sci. 30:302-305. Patton, R. S. and P. T. Chandler. 1975. In vivo digestivility evaluation of chitinous materials. J. Dairy Sci. 58:397-403. Skjak-Break, G. T., T. Anthonsen and P. Sandford. 1989. Chitin and Chitosan. Proceedings from the 4th. International Conference on Chitin and Chitosan. Trondeheim, Norway, 1988. Elsevier Applied Science. N.Y. Steel, R. G. D. and J. H. Torrie. 1980. Principles and procedures of statistics (2nd Ed). McGraw-Hill Book Co., New York. Tilley, J. M. A. and R. A. Ten. 1963. A two-stage technique for the In vitro digestion of forage crops. J. Brit. Grass. Soc. 18:104-111. Velez, S. A., J. C. Allen, C. M. Keery and R. W. Adkinson. 1991. Evaluation of crab and crawfish waste meals as protein source for growing dairy heifers. J. Dairy Sci. 74: 234-242. White, R. L. 1981. Ruminant utilization of chitin and other components of tanner crab meal. MS. thesis, Univ. of Alaska, Fairbanks, Alaska. erght, J.E. and Retnakaran A. 1987. Chitin and benzoylphenyl ureas. Dr.W.Junk Publishers. Dordrech, The Netherlands. CHAPTER 7 INFLUENCE OF PHYSICOCHEMICAL FACTORS ON THE RATE OF GROWTH AND EFFICIENCY OF CHITIN HYDROLYSIS BY C. PARAPUTRIFICUM VAR. RUMINANTIUM. 118 119 ABSTRACT The chitinolytic ruminal bacteria C. paraputn'ficum var. ruminantium was examined for its ability to grow and degrade chitin on chitin (CH) medium under different physicochemical factors including: different pH (5.7, 6.1, 6.5, 7.0 and 7.3); presence of either glucose, N-acetylglucosamine or starch (5.5 and 55 mM); presence of CaCl2.2H20 (2, 4, 6, and 8 mM), and deletions of yeast extract or trypticase in the CH-medium. The optimal pH for bacterial growth and chitinolytic activity was between pH 6.5 and 7.0. Between this pH range, the bacteria doubled its mass in 13.2 h t 2.4, and the percent of chitin dry matter digestion averaged 81.13% :I: 0.33 after 72 h of incubation at 39°C. Glucose and N—acetylglucosamine at low concentration (5.5 mM) enhanced bacterial growth but chitin digestion was unaffected. In contrast, high concentrations (55 mM) of these substrates, affected growth and bacteria activity negatively. Additions of starch, did not affect bacterial growth and activity. C. paraputn’ficum var. ruminantium required Ca2+ for growth and chitinolytic activity. There were no noticeable differences, when the concentration of CaCI2-2HZO was between 2 to 6 mM, but a decrease in growth and activity was observed with 8 mM. Omission of trypticase from the CH-medium did not affect bacteria performance, but the deletion of yeast extract drastically affected the growth and chitinolytic activity. 120 INTRODUCTION The ruminal clostridium C. paraputn'ficum var. ruminantium was further examined as to the effects of various physicochemical factors on its growth rate and chitinolytic activity. Factors that were examined in a series of studies included: variations in the chitin (CH) medium pH; deletion of yeast extract and trypticase in the CH-medium; additions of glucose, N-acetylglucosamine or starch and the effect of different calcium concentration. The different factors studied were chosen because of their potential role under actual ruminal feeding conditions using shellfish wastes. Inhibition of ruminal bacteria growth and the consequential reduction of structural carbohydrate degradation has been partially attributed to declines in pH (Hiltner and Dehority, 1983 and Russell and Dombroski, 1980). Also, cellulose degradation and metabolism in the rumen can be adversely affected by the presence of soluble sugars (Thurston et al., 1993). N-acetylglucosamine and glucose were used in this study, because they are among the few substrates that C. paraputn‘ficum var. ruminantium can utilize for growth (Chapter 3) and secondly, because these substrates are repressors of chitinase synthesis in fungi (Reyes et al, 1989 and Raymond et al., 1986) and yeast (Cruz, et al., 1993). However, in some chitinolytic bacteria, N-acetylglucosamine does not seems to inhibit chitinolytic activity (Pel, et al., 1986). 121 Shellfish wastes, in particular crab shell meal, contains a considerable proportion of calcium carbonate (White, 1981). Calcium carbonate may not affects ruminal bacteria growth or activity, but if the Ca” ion is released from the salt, it can have an influence on bacteria performance. MATERIALS AND METHODS The pH optima was determined by measuring the growth and chitinolytic activity of C. paraputrificum var. ruminantium in chitin medium, adjusted in pH to 5.7, 6.1, 6.5, 7.0 and 7.3 using a potassium phosphate buffer system (KH2P04/K2HPO,). Since the cultures were incubated in a media enriched with 002, the final pH of the media was corrected using the carbon dioxide-bicarbonate buffer system (Breznak and Costillow, 1994). The composition of the different CH- media is shown in Table 1. The media were prepared by the anaerobic techniques of Hungate (1969) as modified by Bryant (1972). Quantities of pure chitin, close to 0.19, were added to culture tubes and after sterilization, 9 ml of the media components were added under CO2 gas previously passed through hot copper column. Sets of triplicate culture tubes containing the different CH-media (9 ml), were aseptically inoculated with 100 pl of a 48 h culture of C. paraputn'ficum var. ruminantium grown in CH-rumen fluid medium. The pure culture had a calculated cell concentration of 1010 per ml. After inoculation, the culture tubes were incubated at 39°C. 122 The effect of pH on bacterial growth was monitored by absorbance (600 nm) readings taken every 12 h after the bacterial cultures reached an absorbance value of 0.2. The readings were taken for 72 h of incubation. Sterile uninoculated CH- media with the different pH were used as controls. Chitinolytic activity was measured, after 72 h of culture incubation. Wlth the completion of the incubation, the undigested chitin was recovered in previously weighed filter paper (Whatman No. 4), dried 24 h in a 60°C oven, then weighed to calculate the amount of digested chitin in each sample. CH-medium with the optimal pH determined for bacterial growth and activity was used in the subsequent studies. The effect on bacteria growth and chitinolytic activity due to deletions of yeast extract, or trypticase from the CH-medium was studied as described for the pH assay. In a similar manner, additions of N-acetylglucosamine, glucose or starch to the CH-medium (5.5 or 55 mM) and their effect on bacteria growth and activity, were determined. Finally, the effect of calcium concentration on bacterial growth and chitinolytic activity was determined by additions of 0, 2, 4, 6, and 8 mM of CaCI-ZHZO to the CH-medium. Statistical analysis for each factor were performed by least-squares analysis of variances in a completely randomized design (Steel and Torrie, 1980). 123 Table 1. Composition of the chitin media used to study the effects of pH on growth and chitinolytic activity. Component pH of chitin media 5.7 6.1 6.5 7.0 7.3 per 100 ml KHZPO4 (0.1 M sol.), ml 83.6 73.0 52.9 31.4 17.2 K2HPO4 (0.1 M sol), ml 7.6 17.1 35.2 49.2 57.4 Sodium carbonate (8% sol), ml 0.4 1.5 3.5 11.0 17.0 Mineral solution “, ml 5.0 5.0 5.0 5.0 5.0 Cysteine-sulfide sol“, ml 2.0 2.0 2.0 2.0 2.0 Resazurin 0.1% sol. 0.1 0.1 0.1 0.1 0.1 Trypticase-peptone (BBL®), g 0.2 0.2 0.2 0.2 0.2 Yeast extract (BBL®), g 0.1 0.1 0.1 0.1 0.1 Chitin (Sigma), 9 1.0 1.0 1.0 1.0 1.0 “ Mineral solution contains (per 1000 ml): (NH,)2SO,, 5 g: NaCI, 12 g; “95:34, 2.45 g and CaCIo 2 H,O, 1.6 9 (Bryant and Robinson, 1962). “ Mix L-Cysteine, 2.5 9 (dissolved in 15 ml of 2N NaOH); Na28.9H20, 2.5 g and resazurin, 0.1 ml (1% sol. in H20). Then, bring the mix to a final volume of 100 ml with deionized water. Heat or microwave the mix until the resazurin indicator is colorless, seal the container and autoclave (bubbling with CO2 is not required). RESULTS When C. paraputn'ficum var.ruminantium was grown in CH-media with differing pH, it attained its highest optical density between pH 6.5 and 7.0 (Fig. 1a). The extent of pure chitin digestion was incomplete after 72 h of incubation for all five pH values examined, but the cultures with a pH value between 6.5 and 7.0 124 had the highest (P<.05) percent of chitin digestion (Fig. 1b). The percent of chitin degraded ranged from a high of 81.20% for CH-medium (pH 7.0), to just 27.64% for CH-medium (pH 5.7). Using the optical density (OD6OOnm) values between 0.2 and 0.4, the doubling times (T2) of C. paraputn'ficum var. ruminantium at pH of 5.7, 6.1, 6.5, 7.0 and 7.3 were 48, 23.4, 10.8, 15.6 and 36 h, respectively. The shortest T2 at pH 6.5 and 7.0, confirmed that the optimal pH for growth and chitinolytic activity of the ruminal clostridia was between 6.5 and 7.0. Somewhat surprising was the gradual increase in bacterial density of the culture growing in CH-medium at pH 7.3. It is possible that at this pH C. paraputn'ficum var. ruminantium will still utilize and ferment chitin at a slow rate, resulting in a gradual decrease of the medium pH, resulting in better condition for growth and chitinolytic activity. In contrast, the negative effect of pH 6.1 and 5.7 on bacterial growth and activity was not offset by the time and the lower chitin digestion obtained at these pH values indicate a strong adverse influence on the chitinolytic activity and growth of C. paraputn'ficum var. ruminantium. 125 1.2 5.7 «fr-MM + 1 FM 6.1 / /l\_\ _e_ E \‘x . 6.5 g 0.8 . \ _._ V :57 PM - 7,0 g 0.6 \' 7.3 '8 l- //w‘:‘ * 7~—‘f*—t:;~;\\\ .4 . g 0 4 , l “I \\‘\.\ .5 / / / / //." 7H;¢‘\\ ’\ 8 ., / TN 7 \63 0.2 O L i I I I j 1 0 12 24 36 48 60 72 Medium, pH Fig. 1a. Effect of pH on growth of C. paraputnficum var. ruminantium. \l 00 (.O O O O O) 0 Dry matter digestion (DMD, %) A O 00 O 20 “o” MediumiH 5.7 6.1 6.5 7.0 7.3 DMD, % "I- 27.64 40.71 78.65 81.20 61.62 Fig. 1b. Effect of pH on the extent of chitin digestion after 72 h of incubation. 126 When C. paraputn'ficum var. ruminantium was incubated with supplemental glucose (55 mM) added to the CH-medium (pH 6.9 to 7.0), the growth rate was enhanced (Fig. 2a) with a doubling time of 8 h, 24 min, while the doubling time was 11 h, 24 min for the control. Also it seemed that the clostridia preferentially utilized glucose over chitin as indicated by the substantially lower chitin degradation (Fig. 2b). The same pattern was observed when 55 mM of N- acetylglucosamine (NAG) was present in the CH-medium (Fig. 3a), with T2= 8 h, 12 min, but the adverse effect on chitin digestion was less marked (P<.05) than with glucose (Fig. 30). Additions of 5.5 mM of either glucose or NAG, also stimulated the rate of growth (T2: 10 h, 36 min for glucose and 10 h, 48 min for NAG versus T2=11 h, 24 min for the control), but there were no differences (P>.05) on the percent of chitin digested after 72 h of incubation with respect to the control (Fig. 2b and 3b). Inclusion of starch (5.5 or 55 mM) to the CH-medium (pH 6.9 to 7.0), did not affect bacterial growth and chitinolytic activity with respect to the control (P>.05), indicating that starch is poorly utilized by the ruminal chitinolytic clostridium. 127 1.6 . Control + 1.4 1- . ‘7’“.“2 5.5 E 12 ‘ ’ \\ 55 C r \ \\\ C \._ \z: + a 1 7 I" A.\\\\;J>(.\. g 0.8 *- Q.’ I\ l a: . , \\ , U ,/ “x g 0.6 ~ ./ J \i 0 04 _ . ‘ I 172” 0 2 a” 0 I i I I I I I 0 12 24 36 48 60 72 Medium, pH Fig. 2a. Effect of glucose on growth of C. paraputn'ficum var. ruminantium. (I) (D O O r I \l O I Dry matter digestion (DMD, %) m o: O O I l .5 O l 30 Glu mM concentration 0.0 5.5 55.0 DMD, % “I 80.23 80.80 37.34 Fig. 2b. Effect of glucose (Glu) on the extent of chitin digestion after 72 h of incubation. Optical density, (600nm) 128 1.2 Control /-\\ _.— 1 z /./" 5.5 ,I" ~9— .7 55 0.8 P / (gt/"J + 0.6 ”I... I / O/ 0.4 r f). 0.2 a’ 0 I I I I I I I 0 12 24 36 48 60 72 Medium, pH Fig. 3a. Effect of N-acetylglucosamine on growth of C. paraputn'ficum var. ruminantium. 85 ... A 80 4 '- °“ i g 75 —— e, - .§ 70 ~— Cl) 0, . O) ‘6 65 6 a3 . 8 E 60 c a . O “I 55 ‘- 50 NAG mM concentration 0.0 5.5 55.0 DMD, % ~-~ 80.23 81.46 55.97 Fig. 3b. Effect of N-acetylglucosamine (NAG) on the extent of chitin digestion after 72 h of incubation. 129 When CaCl2-2HZO was deleted from the CH-medium (pH 6.9 to 7.0), bacterial growth and chitinolytic activity were negatively affected (Fig. 4a and b). Under these conditions, the time required for doubling its mass (T2) was increased to 31 h, 12 min, and the culture never reached the densities observed when CaCl2-2HZO was present in the CH-medium. Bacterial growth and extent of chitin degradation was similar when the concentration of CaCI2 2HZO was between 2 and 6 mM. However, bacterial growth and chitin degradation tended to decrease (P>.05) when the molar concentration of CaCI2-2HZO in CH-medium reached 8 mM. In average, T2 and percent of chitin digestion were: 10 h, 44 min and 80.40% versus 17 h, 48 min and 76.10% in the cultures growing in CH-medium with 2 to 6 mM or 8 mM of CaCl2 2HZO, respectively. When yeast extract was deleted from the CH-medium (pH 6.9 to 7.0), bacterial growth was drastically lower (Fig. 58), and the percent of chitin which was degraded (Fig. 5b) decreased more than 7 fold compared with the control (80.23% versus 11.37% respectively). In contrast, when trypticase was deleted from the CH-medium, bacterial growth was slightly affected and the doubling time was slightly lower when trypticase was present than when it was deleted from the CH- medium (14 h, 24 min versus 15 h, 24 min). However, the percent of chitin digestion was similar between the clostridia cultures growing in CH-medium with and without trypticase. 130 0.9 I 0 08~ [/84 f . 1r . / / .2.- / 2" \ 2 '\\.\.-._. _ I" ,5 0.7 2 1.12,-.- / 222662;? a} 8 . I ..... .... .. a 0.6 ~- / ‘ 5 3: f ,2.“ M — @062 /' , 8 504~ x“ ” 003— , , //o 0.2 {72/ 0 1 I I I I I I I ' 0 12 24 36 48 60 72 Medium, pH Fig. 4a. Effect of calcium concentration (mM) on growth of C. paraputn‘ficum var. ruminantium. 90 fieoi ' ,l- ”‘ s . ~. §70~ g . §60~ (n m I- 0) "§ 50 F 8 E 40 ~ 3* 0 .2 30~g _ I 20 Calcium, mM 0 2 4 6 8 DMD, % 2.. 26.18 81.09 78.31 81.81 76.10 Fig. 4b. Effect of calcium on the extent of chitin digestion after 72 h of incubation. 131 1.2 Control + 1 — /.\ w'o YE E Ix/ . . ., § 0.8 2 ' 9 l £5 3 06 ~ 1 d) U [,- _ ’//l. g 0.4 — . 2" o . /7¥ ' / 0 2 c; 22 2 a. H EEK 4”er HE. H. - .2222» O I . I I I I I 12 24 36 48 60 72 Medium, pH Fig. 58. Effect of deleting yeast extract (YE) or trypticase (Try) from the CH-mediu on growth of C. paraputn'ficum var. ruminantium. \I 00 co 0 O O ' l ‘ l a I O) O T A 0 Dry matter digestion (DMD, %) 00 01 O O N O l 10 CH-medium Control w/o Try w/o YE DMD, % .... 80.23 80.01 16.37 Fig. 5b. Efl‘ect of deleting yeast extract (YE) or trypticase (le) on the extent of chitin digestion after 72 h of incubation. 132 DISCUSSION The optimal pH for growth and chitinolytic activity (6.5 to 7.0), is within the normal pH range usually found in the rumen environment, and C. paraputn'ficum var. ruminantium may not be limited in its chitinolytic activity by the ruminal pH. However, the slow doubling times (T2) exhibited by this bacteria, even at the best rate of 10.8 h (CH-medium, pH 6.5) may limit its capability to compete with other ruminal bacteria with faster growth rates. The rate of growth in clostridial species, seems to be a function of the degree of bacterial specialization to degrade polymers. Most saccharolytic species may double their mass within 30 to 40 min (Andreesen et al., 1989). However, chitinolytic clostridia from aquatic environments double their mass in about 72 h (Pel, et al., 1986), a value much higher than that determined for the ruminal clostridium studied. The inhibition of growth below pH 6.1 (Fig. 7.28) is consistent with the known sensitivity of other ruminal bacteria to lower pH (Russell and Dombroski, 1980). The ruminal pH of dairy cattle fed with shellfish waste has not been reported and it would be interesting to determine if the negative effect on animal performance observed when cows are fed with high levels of shellfish waste (Brundage et al. 1984 and Laflame, 1987), is due or related to the develop of an acidic pH in the rumen. It should be noted that a major end product of chitin fermentation is lactate (Chapter 133 3), whose production may decrease the ruminal pH and thereby the extent of chitin digestion. When glucose or N-acetylglucosamine were added to the CH-medium at high concentration (55 mM), the ruminal clostridia grew faster, but the percent of chitin degraded was negatively affected (Fig. 73b and 7.4b). Soluble carbohydrates like NAG and glucose are utilized and fermented faster than structural carbohydrates such as chitin. The significant (P<.05) decrease in chitin degradation could be a consequence of a faster fermentation rate of the soluble sugars over chitin, resulting in a decrease in the medium pH to levels which adversely affect bacteria growth and activity before any appreciable degradation of chitin occurs. Small additions of glucose or NAG to the CH-medium (5 mM) also stimulated growth rate, but the extent of chitin degradation was not affected in comparison to the control. These results, suggest that the rate of chitinolysis is unaffected by small additions of NAG or glucose into the CH-medium. Similar results were reported by Pel et al. (1986) who found that N-acetylglucosamine (0.5 mM) had a noninhibitory effect on chitin degradation by chitinolytic clostridia species isolated from aquatic sediments. Exoenzymes such as amylases and proteases are calcium proteins, and calcium dipicolinate is an important component of endospores (Gottschalk, 1979). Calcium ions, are also required to maintain the stability of the Iipopolysaccharide layer of the outer membrane of Gram negative bacteria (Schlegel, 1988). In addition, CaCl2 has been shown to stimulate the secretion of SP32, a chitinase from pumpkin cells (Esaka et al., 1990). Therefore, it is not too surprising that this 134 cation is essential for the growth and chitinolytic activity of C. paraputn'ficum var. ruminantium. Based on the results obtained, Caz“ is clearly required for the optimal growth of C. paraputn'ficum var. ruminantium and that concentrations of the cation up to 6 mM, in CH-medium does not effect bacterial growth or chitinolytic activity. Another possible beneficial role of Ca” might be to increase the uptake of N-acetylglucosamine by the ruminal clostridia, as demonstrated with glucose, galactose and mannose in other Gram negative bacteria (Wong, 1993). At high concentrations, Caz‘forms complexes with carboxyl groups in peptidoglycan chains (Lewis, 1975). At high concentrations, Caz“ also interacts with the acid proteins of the 8 layer of Azotobacter vine/andl'i and other uncharged sugar moieties. As result, the cell reduces its permeability to sugars (Wong, 1993). This premise could explain the decrease in bacterial growth and percent of chitin degradation observed when the concentration of CaCl2 2HZO in CH-medium was 8 mM. The main reason for studying the effects of calcium ion on growth and chitinolytic activity of C. paraputn'ficum var. ruminantium, was concerned with the possible adverse effects of the high concentration of calcium carbonate in shellfish wastes. However, White (1981), demonstrated that independent of the extent of crab shell meal degradation in the rumen, the degradation of calcium carbonate is minimal, being digested in the abomasum, where the acidic gastricjuice reacts with the calcium carbonate matrix of the crab shell, releasing carbon dioxide. Therefore, the main source of Ca2+ in the rumen will be the animal's saliva, which has a Ca2+ ion concentration of 0.4 mEq per liter (Church, 1988). It seems that 135 calcium carbonate may not interfere with the chitinolytic activity of C. paraputn'ficum var. ruminantium in cows fed with shellfish wastes. The low growth of C. paraputn‘ficum var. ruminantium in CH-medium without yeast extract, indicates the essential requirement for B-complex vitamins by this bacterium. However, a study replacing yeast extract by a solution of B- complex vitamins in the CH-medium, and the effect of deleting each vitamin one at time on the bacterial growth and activity is required to confirm this presumption. At this moment, it is unknown if one or more B-complex vitamins are required, but it is known that these compounds are biosynthesized de novo by many other rumen bacteria (Yokoyama, 1988). This means that under ruminal conditions, C. paraputrl'ficum var. ruminantium may have to interact with other ruminal bacteria in order to exist and express its chitinolytic activity. 10. 136 LITERATURE CITED Andreesen J. R., H. Bahl and G. Gottschalk. 1989. Introduction to the physiology and biochemistry of the genus Clostridium. p.27-62. In: N. P. Minton and D. J. Clarke (ed). Clostridia. Biotechnology handbooks No. 3. New York. Breznak, J. A. and R. N. Costillow. 1994. Physicochemical factors in growth. p. 137-154. In: P. Gerhardt, R.G.E. Murray, W. A. Wood and N. R. Krieg (ed). American Society of Microbiology, Washington, D. C. Brundage, A.L, Husby, F.M., Herlugson, M.L, SImpson,W.L. and Burton, V.L. 1984. Acceptability of Tanner crab meal in concentrates for lactation. J. Dairy Sci. 67: 1965-1970. Bryant, M. P. 1972. Commentary on the Hungate technique for culture of anaerobic bacteria. Amer. J. Clin. Nutr. 25:1324-1328. Bryant, M. P. and I. M. Robinson. 1962. Some nutritional characteristics of predominant culturable ruminal bacteria. J. Bacteriol. 64: 605-613. Church, D. C. 1988. Salivary function and production, p.117-124, In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Cruz, J., M. Rey, J. M. Lora, A. H. Gallego, F. Dominguez, J. A. P. Tom, A. Llobell and T. Benites. 1993. Carbon source control on B-glucanases, chitobiose and chitinase from Trichoderma harzianum. Arch. Microbiol. 159:316-322. Esaka, M, K. Enokl, B. Kouchl and T. Sasaki. 1990. Purification and characterization of abundant secreted protein in suspension-cultured pumpkin cell. Plant Physiol. 93:1037-1041. Gottschalk, G. 1979. Bacterial metabolism, p.1-11. Springer-Verlag, New York. HIItner, P., and B. A. Dehority. 1983. Effect of soluble carbohydrates on digestion of cellulose by pure cultures of rumen bacteria. Appl. Environ. Microbiol. 46:642-648. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 137 Hungate. R. E. 1969. A roll tube method for cultivation of strict anaerobes. In: J. R. Norris and D. W. Ribbons (Ed). Methods in microbiology. Vol. 3, p. 117-132. Academic Press, Inc. New. York. Laflame, LF. 1988. Utilization of crab meal fed to young beef cattle. Can. J. Anim. Sci. 68: 1237-1244. Lewis, J. C. 1975. Calcium-peptidoglycan interactions, p.547-549. In: P. Gerhardt, R. N. Costillo and H. L. Sadoff (eds). Spores VI. American Society for Microbiology, Washington, DC. Pel, R.,G. Hessels,H.AaIfs and J. C. Gottschal. 1986. Chitin degradation by Clostridium sp. strain 9.1 in mixed cultures with saccharolytic and sulfate-reducing bacteria. FEMS Microbiol. Ecol. 62:191-200. Raymond, J., S. T. Leger, R. M. Cooper and A. K. Chamley. 1986. Cuticle-degrading enzymes of entomophatogenic fungi: Regulation of production of chitinolytic enzymes. J. Gen. Microbiol. 132:1509-1517. Reyes, F., J. Calatayud and M. J. Martinez. 1989. Endochitinase from Aspergl'I/us nidulans implicated in autolysis of its cell wall. FEMS Microbiol Lett. 60:119-124. Russell, J. B. and D. B. Dombroski. 1980. Effect of pH on the efficiency of growth by pure cultures of rumen bacteria in continuous culture. Appl. Environ. Microbiol. 39:604-610. Schlegel, H. G. 1988. General microbiology. 014-76, 6th ed. Cambridge University Press. Cambridge. Steel, R. G. D. and J. H. Torrie. 1980. Principles and procedures of statistics (2nd Ed.). McGraw-Hill Book Co., New York. Thurton, 8., K. A. Dawson and H. J. Strobel. 1993. Cellobiose versus glucose utilization by the ruminal bacterium Ruminococcus albus. Appl. Environ. Microbiol. 59:2631-2637. White, R. L. 1981. Ruminant utilization of chitin and other components of tanner crab meal. M.S. thesis, Univ. of Alaska, Fairbanks, Alaska. Wong, T. Y. 1993. Effects of calcium on sugar transport in Azotobacter vine/andii. Appl. Environ. Microbiol. 59:89-92. 138 23. Yokoyama, M. T. and K. A. Johnson. 1988. Microbiology of the rumen and intestine, p.125-144. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. CHAPTER 8 SUMMARY 139 140 Shellfish wastes are rich in chitin, an insoluble polymer of the amino-sugar N-acetylglucosamine, and large amounts are generated by the fishing industry after meat extraction from marine crustaceans. Present knowledge of the potential use of shellfish wastes as a feed for ruminants have primarily been focused on the animal responses to the ingestion of these by-products. However, the ruminal microbes directly involved in the process of chitin degradation have never been studied. This study has focused on the isolation and characterization of chitinolytic ruminal bacteria and the implication of these bacteria in the assimilation of the nutrients present in shellfish waste by the animal. The 4-methylumbelliferyl (MUF)-chitin substrates technique used for identification of chitinolytic activity, was successfully validated under anaerobic conditions. In future studies, this technique will be very useful as a rapid, reliable and simple assay for identification of chitinolytic activity in anaerobic bacteria. It should be noted that the fluorescent test will be useful, not only for studies with ruminal bacteria, but also for identification of chitinolytic activity of anaerobic bacteria from other sections of the digestive tract of either ruminant or nonruminant animals. Using the fluorescent test, it was demonstrated that chitinolytic bacteria are naturally occurring in the rumen of dairy cows never exposed to chitinous feeds. Also, it was determined that the chitinolytic pathway (involving the enzymes chitinase and N-acetylglucosaminidase), is prevalent in the ruminal bacteria community. 141 The MUF-chitin substrates studies confirmed the previous observations of several researchers about the capability of certain ruminal bacteria to degrade chitinous by-products. However, it was shown that chitin degradation is not related or similar to the degradation of cellulose. Different bacteria with no cellulolytic activity are responsible for the ruminal degradation of chitin. Scanning electron microscopy photomicrographs of shrimp carapaces incubated in the rumen of fistulated cows showed, that several bacteria are part of a chitinolytic consortia and that attachment to the substrate is very important in the process of chitin-rich substrates degradation. Considering the unique morphological characteristics of the bacteria observed, and the absence of reports describing them, it is believed that these chitinolytic bacteria are part of a minor fraction of the ruminal microflora. The increase in numbers of these bacteria in the rumen is important before substantial benefits of feeding ruminants with chitin-rich feeds will be observed. Perhaps the most significant achievement of the present research, was the isolation and characterization of a ruminal chitinolytic bacteria. The bacterium isolated was a spore-former, identified as Clostridium parapufrificum var. ruminantium, a non-toxigenic, non-hemolytic clostridial species. The metabolic profile of this clostridial isolate, indicate that this bacterium is highly specialized for chitin utilization. It is unable to degrade cellulose and utilizes starch poorly. C. paraputn'ficum var. ruminantium, does not degrade chitosan, the deacetylated form of chitin. The former characteristics, confirm the results obtained with the fluorescent technique, which indicated that this bacterium uses the chitinolytic 142 pathway. Scanning electron microscopy photomicrographs of shrimp carapaces incubated with a pure culture of C. paraputrificum var. ruminantium, indicated that this bacterium attaches to the inner surface of the shrimp carapaces but does not attach to the outer surface, which seems to be protected with a biofilm of unknown composition. Since bacterial attachment and colonization of the shrimp carapace seems to be essential for chitin degradation, it is suggested that the chemical composition of this biofilm and its relation to the process of bacterial attachment and chitin degradation be further studied. The aggressive colonization and degradation of the shrimp carapaces suggested by the scanning electron microscopy study, was quantitatively supported by the high percent of chitin digested (69. 89.58% 1; 3.34) when C. paraputrificum var. ruminantium, was incubated (4 d at 39°C) with pure chitin. The fermentation end products of chitin fermentation were acetate, lactate and butyrate. In vitro organic matter degradation (IVOMD) studies indicated crab shell meal waste is efficiently degraded by Clostridium paraputn'ficum var. ruminantium. However, degradation of chitin-free substrates (e.g. alfalfa hay) is poor compared with mixed bacteria in ruminal fluid. In general, the results of varying the pH, and the composition of the chitin medium, indicate that C. paraputrificum var. ruminantium, will express its maximum growth and chitinolytic activity within the pH range (6.5 to 7.0) usually found in the rumen environment, but it may be enhanced by its requirement of B-complex vitamins (as suggested by it requirement of yeast extract) which can be supplied 143 by other ruminal bacteria. Glucose and N-acetylglucosamine at low concentration (5.5mM) seem to increase its growth, while at high concentration (55mM), seem to adversely affect its growth and chitinolytic activity. However, considering that inside the rumen, there are other bacteria which ferment soluble sugars more efficiently, it might be argued that the influence of sugars on chitin degradation under ruminal conditions would be inconsequential. Strong enhancement of chitin degradation were observed in pure cultures of C. paraputn'ficum var. ruminantium supplemented with CaCI2 indicating the essential requirement of calcium ion in the growth and chitinolytic activity of this ruminal bacterium. It is assumed that the high concentration of calcium carbonate in crab shell meal, will not represent a limit for the expression of the bacterium chitinolytic activity. Although, C. paraputriflcum var. ruminantium, seems to be very effective in degrading chitinous compounds under ruminal conditions, the slow doubling times exhibited by this bacterium, even at the best rate, may limit its capability to compete with other ruminal bacteria with faster growth rates. Although it is proposed that C. paraputrificum var. ruminantium may be useful as an ruminal inocula to increase the efficiency of digestion of chitin-rich compound, it is unknown if this bacterium will become one of the predominant chitinolytic bacteria in the rumen of animals adapted to the intake of chitinous feeds. It has been postulated that clostridia may have a disadvantage in environments with a constant nutrient supply, such as the rumen, because of the energy required to maintain the genetic information for spore production. Future research needs to include, ( 1) the determination and isolation of microflora which 144 produce chitinase in animals adapted to the intake of shellfish waste, (2) the ability of C. paraputriflcum var ruminantium to become a dominant chitinolytic bacteria in animals adapted to chitinous feeds in their diet, (3) the potential of this clostridia as an bioinoculant to enhance ruminal chitin degradation, and (4) the effect of C. paraputrificum var. ruminantium inoculum on animal performance during and after adaptation to the feeding of shellfish waste. BIBLIOGRAPHY 145 BIBLIOGRAPHY Andreesen J. R., H. Bahl and G. Gottschalk. 1989. Introduction to the physiology and biochemistry of the genus Clostridium. p.27-62. In: N. P. Minton and D. J. Clarke (ed). Clostridia. Biotechnology handbooks No. 3. New York. Arroyo, B. A. and A. Carabez. 1982. Location of chitin in the cyst wall of Entamoeba invadents with colloidal gold tracers. J. Parasitol. 68:253-258. Bartinicki, G. S. and E. Uppman. 1982. Fungal cell wall composition. p.229-252. In: A.J. Laskin and HA. Lechevalier (eds). CRC Handbook of Microbiology, 2nd ed., Vol. IV. Microbial cornpositionzcarbohydrates, lipids and minerals. CRC Press, Boca Raton. Bayer, E. A. and R. Lamed. 1986. Ultrastructure of the cell surface cellulosome of Clostridium thermocellum and its interaction with cellulose. J. Bacteriol. 167:828-836. Belas, M. R. and R. R. Colwell. 1982. Adsorption kinetics of laterally and polarly flagellated Vibrio. J. Bacteriol. 151:1568-1580. Blackwell, J. 1988. Physical methods for determination of chitin structure and conformation. Meth. Enzymol. 161:435-442. Bloch, R. and M. M. Burger. 1974. Purification of wheat germ agglutinin using affinity chromatography of chitin. Biochem. Biophys. Res. Commun. 58:13-17. Boyer, J. N. and H. I. Kator. 1985. Method for measuring microbial degradation and mineralization of 1‘C-labelled chitin obtained from the blue crab, Callinectes sapidus, Microb. Ecol. 11:185-192. Breznak, J. A. and R. N. Costillow. 1994. Physicochemical factors in growth. p. 137-154. In: P. Gerhardt, R.G.E. Murray, W. A. Wood and N. R. Krieg (ed). American Society of Microbiology, Washington, D. C. 10. 11. 12 13 14. 15. 16. 17. 18. 19. 20. 21. 146 Broekaert, W. F., J. V. Parijs, F. Leyns, H. Joos and W. J. Peumans. 1989. A chitin-binding lectin from stinging nettle rhizomes with antifungal properties. Science. 245:1100-1102. Brundage, A.L, Husby, F.M., Beardsley, G.L and Burton, V.L 1981. King crab meal in concentrates for lacting cows. J. Dairy Sci. 64: 433-440. Brundage, A.L, Husby, F.M., Herlugson, M.L, Simpson,W.L and Burton, V.L. 1984. Acceptability of Tanner crab meal in concentrates for lactation. J. Dairy Sci. 67: 1965-1970. Bryant, M. P. 1972. Commentary on the Hungate technique for culture of anaerobic bacteria. Amer. J. Clin. Nutr. 25:1324-1328. Bryant, M. P. and I. M. Robinson. 1962. Some nutritional characteristics of predominant culturable ruminal bacteria. J. Bacteriol. 64:605-613. Cabib, E. 1987. The synthesis and degradation of chitin. In: Advances in enzymology. Vol. 59, p. 59-101. CanaIe-Parole, E. 1986. The Spirochetes. p. 38-70. In: N. R. Krieg and J. G. Holt. Bergey's manual of systematic bacteriology. Williams & Wilkins. Baltimore. MD. Cato, E. P., W. L. George and S. M. Fine Gold. 1986. Endospore-forming gram-positive rods and cocci. Genus Clostn'dium. In: P. H. A. Sneath (Ed). Bergey's manual of systematic bacteriology. p.1141-1200. Williams & WllIkins. Baltimore, MD. Church, D. c.1988. Salivary function and production, p.117-124. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Cruz, J., M. Rey, J. M. Lora, A. H. Gallego, F. Dominguez, J. A. P. Toro, A. Llobell and T. Benites. 1993. Carbon source control on B-glucanases, chitobiose and chitinase from Trichoderma harzianum. Arch. Microbiol. 159:316-322. Davis, L L and Bartinicki-Garcia. 1984. The co-ordination of chitosan and chitin synthesis in Mucor rouxii. J. Gen. Microbiol. 130:2095-2102. Enart, T. M. 1983. Microbial cellulases. In: W. M. Fogarty (Ed). Microbial enzymes and technology. Applied Science Publishers, London. p. 183-223. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 147 Esaka, M, K. Enold, B. Kouchl and T. Sasaki. 1990. Purification and characterization of abundant secreted protein In suspension-cultured pumpkin cell. Plant Physiol. 93:1037-1041. Evrall, C. C., R. W. Attwell and C. A. Smith. 1990. A semi-micro quantitative assay for determination of chitinolytic activity in microorganisms. J. Microbiol. Meth. 12:183-187. Gay, L, H. Chanzy, V. Bulone, V. Girard and M. Fevre. 1993. Biosynthesis of chitin in the fungus Saprolegnia. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 57-66. Gay, L., M. Hebraud, V. Girard and M. Fevre. 1989. Chitin synthase activity from Neocallimastix frontalis, an anaerobic rumen fungus. J. Gen. Microbiol. 135:279-283. Gooday, G. W. 1990a. Physiology of microbial degradation of chitin and chitosan. Biodegradation,1:177-190. Gooday, G. W. 19900. The ecology of chitin degradation. In: K.C. Marshall, Advances in microbial ecology. N.Y. Plenium Press. Vol. 11, p. 387-429. Gooday, G. W., J. I. Presser, K. Hillman and M. G. Cross. 1991. Mineralization of chitin in an estuarine sediment: The importance of the chitosan pathway. Biochemical Systematics and Ecology. 19:395-400. Gottschalk. G. 1979. Bacterial metabolism, p.1-11. Springer-Verlag, New York. Gottschalk. G., J. R. Anderson and H. Hipe. 1981. The genus Clostridium (nonmedical aspects), p. 1767-1803. In: Starr M. P., H. Stolp, H. G. Truper, A. Balows and H. G. Schlegel (eds). The prokaryotes, vol II. Springer, Berlin, Heilderberg, New York. Hackman, R.H. 1987. Chitin and the fine structure of cuticles. In: Wright and Retnakaran, Chitin and benzoylphenyl ureas. Dr.W Junk Publishers. Dordrecht, The Netherlands. Hara, 8., Y. Yamamura, Y. Fujii, T. Mega and T. Ikenaka. 1989. Purification and characterization of chitinase produced by Streptococcus erythraeus. J. Biochem. 105:484-489. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 148 Herwing, R. P., N. B. Pellerin, R. L lrgens, J. S. Maid and J. T. Staley. 1988. Chitinolytic bacteria and chitin mineralization in the marine waters and sediments along the Antarctic Peninsula. FEMS Microbiol. Ecol. 53:101-112. Hiltner, P., and B. A. Dehority. 1983. Effect of soluble carbohydrates on digestion of cellulose by pure cultures of rumen bacteria. Appl. Environ. Microbiol. 46:642-648. Hood, M. A. 1991. Comparison of four methods for measuring chitinase activity and the application of 4-MUF assay in aquatic environments. J. Microbiol. Methods. 13:151-160. Hungate. R. E. 1969. A roll tube method for cultivation of strict anaerobes. In: J. R. Norris and D. W. Ribbons (Ed). Methods in microbiology. Vol. 3, p. 117-132. Academic Press, Inc. New. York. Husby, F. M., A. L. Brundage and R. L White. 1981. Utilization of shellfish meals in domestic feeds. In: W. S. Otwell (Ed). Seefood waste management in the 1980's: Conference proceedings. Report No. 40 Florida Sea Grant College. Orlando, Fl. Husby, F. M. 1993. Professor, University of Alaska, Agricultural Experiment Station, Fairbanks 99701. Jeuniaux, C. 1966. Chitinases. In: E. F. Neufeld and V. Ginsburg. Methods in enzymology. Vol. 8. p. 644-650. Jeuniaux, C. 1993. Chitinolytic systems in the digestive tract of vertebrates: a review. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 233-244. Kafetzopoulos, D., A. Martinou and V. Bourlotis. 1993. Purification of chitin deacetylase from Mucor roxil'. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 147-154. Kaneko, T. and R. R. Colwell. 1978. The annual cycle of Vibrio parahaemolyticus in Chesapeake Bay. Microb. Ecol. 4:135-155. Kelly, W. J., R. V. Asmundson and D. H. Hoperoft. 1987. Isolation and characterization of a strictly anaerobic, cellulolytic spore former: Clostridium chartatabl'dum sp. nov. Arch. Microbiol. 147:169-173. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 149 KIenzIe-Sterzer C., D. Rodriguez-Sanchez and C. Rha. 1984. Solution properties of chitosan2chain conformation. In: Zikakis, J.P. Chitin, chitosan and related enzymes. Academic Press, Inc. Orlando Fl. p.383-393. Klomparens, K. L, S. L Flegler and G. R. Hooper. 1986. Procedures for transmission and scanning electron microscopy for biological and medical science (Manual). Center for Electron Optics, Michigan State University. Knorr, D. 1991. Recovery and utilization of chitin and chitosan in food processing waste management. Food Technol. 45:116-122. Laflame, LF. 1988. Utilization of crab meal fed to young beef cattle. Can. J. Anim. Sci. 68: 1237-1244. Lamed, R., E. Setter and E. A. Bayer. 1983. Characterization of a cellulose-binding, cellulase-containing complex in Clostridium thermocellum. J. Bacteriol. 156:828-836. Lewis, J. C. 1975. Calcium-peptidoglycan interactions, p.547-549. In: P. Gerhardt, R. N. Costillo and H. L. Sadoff (eds). Spores VI. American Society for Microbiology, Washington, DC. Lumbland, G., B. Hedersted, J. Lind and M. Steby. 1974. Chitinase in goat serum. Preliminary purification and characterization. Eur. J. Bioch. 46:367- 376. Maddocks, J. L and M. J. Greenan. 1975. A rapid method for identifying bacteria enzymes. J. Clin. Phatol. 28:686-687. McDougaIl, E. J. 1948. Studies of ruminal saliva. 1. The composition and output of sheep's saliva. Biochem. J. 43:49. Middelhoven W. J. and A. H. M. va Baalen. 1988. Development of the yeast flora of whole-crop maize during ensiling and during subsequent aerobiosis. J. Sci. Food. Agric. 42:199-207. Molano, J., A. Duran and E. Cabib. 1977. A rapid and sensitive assay for chitinase using tritiated chitin. Analytical Biochemistry. 83:648-656. Montgomery, M. T. and D. L chhman. 1993. Role of chitin-binding proteins in the specific attachment of the marine bacterium Vibrio harveyi to chitin. Appl. Environ. Microbiol. 59:373-379. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 150 Mulisch, M. G. Schenhuly and M. Walther. 1993. Chitin synthesis in the ciliated protozoan Eufo/Iiculina uh/igi. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 119-128. Muzzarelli, R. A. 1993. Advances in N-acetyI-B-D-glucosaminidases. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 357-374. Muzzarelli, R. A., Jeuniaux Ch. and Gooday G.W. 1986. Chitin in nature and technology. Plenum Press. N.Y. Muzzarelli, R. A. 1977. Chitin. Pergamon Press, LTD. Oxford. No, H. K., S. P. Meyers and K. S. Lee. 1989. Isolation and characterization of chitin from crawfish shell waste. J. Agric. Food Chem. 37:575-579. O'Brien, M. and R. R. Colwell. 1987. A rapid test for chitinase activity that uses 4-methylumbelliferyI-N-acetyI-B-D-gIucosaminide. Appl. Environ. Microbiol. 53:1718-1720. Oglmoto, K and S. Imal. 1981. Atlas of rumen microbiology. Japan Scientific Societies Press, Tokyo. Orpln, C. G. 1977. The occurrence of chitin in the cell walls of the rumen organisms Neocallimastix frontalis, Piromonas communis and Sphaeromonas communis. J. Gen. Microbiol. 99:215—218. Ortega, E. and Church,.D.C. 1979. In vitro digestion studies with crab meal and related chitinous compounds as feedstuffs for cattle. Proc. West. Sec. Am. Soc. Anim. Sci. 30:302-305. Othakara, A., H. Matsunaga and M. Mitsutomi. 1990. Action pattern of Streptomyces gn‘ceus chitinases on partially N-acetylated chitosan. Agric. Biol. Chem. 54:3191-3199. Ou, L and M. Alexander. 1974. Effect of glass microbeads on the microbial degradation of chitin. Soil Sci. 118: 164-167. Owens, F. N. and A. L Goetsch. 1989. Ruminal fermentation. p.145-171. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Patton, R.S. and Chandler P.T. 1975. In vivo digestibility of chitinous materials. J. Dairy Sci. 58:397-403. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 151 Patton, R.S., P.T. Chandler and 0.6. Gonzalez. 1975. Nutritive value of crab meal for young ruminanting calves. J. Dairy Sci. 58:404-409. Pedraza, R. M. and E. L6pez-R. 1991. Chitinase activity in germinating cells of Mucor rouxii. Antonie van Leeuwenhoek. 59:183-189. Pel, R. 1989. Microbial interactions in anaerobic chitin-degrading mixed cultures. Thesis of the Department of Microbiology of the University of Groningen, The Netherlands. Pel, R., G. Hessels, H. Aslfs and J. C. Gottschal. 1989. Chitin degradation by Clostridium sp. strain 9.1 in mixed cultures with saccharolytic and sulfate-reducing bacteria. FEMS Microbiol. Ecol. 62:191-200. Pel, R. and J. C. Gomchal. 1986a. Chitinolytic communities from an anaerobic estuarine environment. In: Chitin in nature and technology. Muzzarelli, R. A. A., C. Jeuniaux and G. W. Gooday (Eds). Plenum Press, N.Y. p. 539-546. Pel, R. and J. C. Gottschal. 1986b. Mesophilic chitin degrading anaerobes isolated from an estuarine environment. FEMS Microbiol. Ecol. 38:39-49. Poulicek, M. and C. Jeuniaux. 1991. Chitin biodegradation in marine environments: An experimental approach. Biochemical Systematics and Ecology. 19:385-394. Powning, R. F. and H. Irzykiewicz. 1967. Separation of chitin oligosaccharides by thin-layer chromatography. J. of Chromatog. 29:115- 119. Rasmussen, M. A., B. A. White and R. B. Hespell. 1989. Improved assay for quantitating adherence of ruminal bacteria to cellulose. Appl. Environ. Microbiol. 55:2089-2091. Raymond, J., S. T. Leger, R. M. Cooper and A. K. Chamley. 1986. Cuticle-degrading enzymes of entomophatogenic fungi: Regulation of production of chitinolytic enzymes. J. Gen. Microbiol. 132:1509-1517. Reyes, F., J. Calatayud and M. J. Martinez. 1989. Endochitinase from AspergiI/us nidulans implicated in autolysis of its cell wall. FEMS Microbiol Lett. 60:119-124. Robyt, J. F. and B. J. White. 1990. Biochemical techniques, theory and practice. Waveland Press, Inc. IL. p. 216-217. 81. 82. 83. 84. 85. 86. 87. 88. 89. 90. 91. 92. 152 Russell, J. B. 1986. Ecology of rumen microorganisms: energy use. p.74- 98. In: A. Dobson and M. J. Dobson (ed). Aspects of digestive physiology in ruminants. Comstock Publishing Associates. Cornell University Press, Ithaca. Russell, J. B. and D. B. Dombroski. 1980. Effect of pH on the efficiency of growth by pure cultures of rumen bacteria in continuous culture. Appl. Environ. Microbiol. 39:604-610. Schlegel, H. G. 1988. General microbiology. p.14-76, 6th ed. Cambridge University Press. Cambridge. Schlegel, M. L 1993. Grazing systems for direct-seeded alfalfa pastures. p. 57. MS Thesis, Animal Science Department, Michigan State University. Skjak-Break, G. T., T. Anthonsen and P. Sandford. 1989. Chitin and Chitosan. Proceedingsfrom the 4th. International Conference on Chitin and Chitosan. Trondeheim, Norway, 1988. Elsevier Applied Science. N.Y. Shlmara, K., Y. Taklguchl and T. Kobayashi. 1988. Screening of Mucoraceae strains for chitosan production. In: G. Skjak, T. Anthonsen and P. Sandforf. Chitin and chitosan. Elsevier Applied Sci. p. 171-178. Slifkin, M. and G. M. Gil. 1983. Rapid biochemical test for identification of group A, B, C, F, and G Streptococci from throat cultures. J. Clin. Microbiol. 18:29-32. Smucker, A. R. 1991. Chitin primary production. Biochemical Systematics and Ecology. 52357-369. Stanton, T. B. and E. CanaIe-Parola. 1980. Treponema bryantii sp. nov., a rumen spirochete that interacts with cellulolytic bacteria. Arch. Microbiol. 127:145-156. Stanton, T. B. and E. CanaIe-Parola. 1979. Enumeration and selective isolation of rumen spirochetes. Appl. Environ. Microbiol. 38:965-973. Steel, R. G. D. and J. H. Torrie. 1980. Principles and procedures of statistics. McGraw-Hill Book Co., New York. Theodorou, M. K., S. E. Lowe and A. P. J. Trinci. 1993. Anaerobic fungi and the rumen ecosystem. Microbiology Group, Department of Cell and Structural Biology (ed). University of Manchester, UK. p. 45. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 103. 104. 105. 153 Thurton, B., K. A. Dawson and H. J. Strobel. 1993. Cellobiose versus glucose utilization by the ruminal bacterium Ruminococcus albus. Appl. Environ. Microbiol. 59:2631-2637. Treagan, L and L Pulliam. 1982. Medical microbiology laboratory procedures. W. B. Saunders Company, Philadelphia, PA. p. 78-89. Tilley, J. M. A. and R. A. Terry. 1963. A two-stage technique for the In vitro digestion of forage crops J. Brit. Grass. Soc. 18:104-111. Van Gylswyk, N. 0., E. J. Morris and H. J. Els. 1980. Sporulation and cell wall structure of Clostridium polysaccharolyticum comb. nov. (Formerly Fusobacten‘um polysaccharolyticum). J. Gen. Microbiol. 121:491-493. Varel, V. H. 1989. Reisolation and characterization of Clostridium Iongisporum, a ruminal sporeforming cellulolytic anaerobe. Arch. Microbiol. 152:209-214. Velez, S.A., J.C. Allen, C.M. Keery and R.W. Adkinson. 1991. Evaluation of crab and crawfish waste meals as protein sources for growing dairy heifers. J. Dairy Sci. 74:234-242. Verbug, J.G. and KO. Huynh. 1991. Purification and characterization of an antifungal chitinase from Arabidopsis thaliana. Plant Physiol. 95:450-455. Villagémez-Castro J. C., M. Pedraza-Reyes, C. CaIvo- Mendez and E. LOpez-Romero. 1993. Fungal and amebic chitinases. In: Muzzarelli (ed). Chitin enzymology. European Chitin Society. Ancona, Italy, p. 311-323. Wames, C. A. and C. I. Randles. 1977. Prelyminary studies on chitin decomposition In Lake Erie sediments. Ohio J. Sci. 77:224-230. West, P. A. and R. R. Colwell. 1984. Identification and classification of Vibn'onaceae: an overview. p.285-363. In: Vibrios in the environment. John Wiley & Sons, Inc., New York. White, R. L 1981. Ruminant utilization of chitin and other components of tanner crab meal. MS. thesis, Univ. of Alaska, Fairbanks, Alaska. erth, 5. J. and G. A. Wolf. 1990. Dye-labelled substrates for the assay and detection of chitinase and lysozyme activity. J. of Microbiol. Methods.12:197-205.. Wong, T. Y. 1993. Effects of calcium on sugar transport in Azotobacter vine/andii. Appl. Environ. Microbiol. 59:89-92. 106. 107. 108. 109. 154 Wright, J.E. and Retnakaran A. 1987. Chitin and benzoylphenyl ureas. Dr.W.Junk Publishers. Dordrech, The Netherlands. Yokoyama, M. T. and K. A. Johnson. 1988. Microbiology of the rumen and intestine, p.125-144. In: D. C. Church (Ed). The ruminant animal, digestive physiology and nutrition. Prentice Hall. New Jersey. Zikakis, J.P. 1984. Chitin, chitosan and related enzymes. Academic Press, Inc. Orlando Fl. Ziolecki, A. 1979. Isolation and characterization of large treponemes from the bovine rumen. Appl. Environ. Microbiol. 37:131-135. MICHIGAN STATE UNIV. LIBRARIES llIIIIIIIIIIIHIIIIIIIIIIIIIIIIIIIIIIIIIIIIIHIIIIIIHIIIUIIII 31293014132363