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,1
* Animal Agriculture Business Monsanto Co. St. Louis, MO 63198
Department of Animal Sciences University of Illinois Urbana, IL 61801
Centre for Dairy Research The University of Reading Reading, UK RG2 9HX
Department of Animal and Avian Sciences University of Maryland College Park, MD 20742
|| Department of Animal Science Cornell University Ithaca, NY 14853
# Institute of Food Research Food Materials Science Division Norwich Research Park Colney, Norwich, NR4 7UA, UK
Corresponding author:
J. L. Vicini; e-mail:
john.l.vicini{at}monsanto.com.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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Key Words: fibrolytic enzyme • malic acid • feed additive • milk production
Abbreviation key: CM= carboxymethyl, CNY= Cornell, NY, CPM= Cornell-Penn-Miner, DMSO= dimethyl sulphoxide, DNS= dinitrosalicylic acid, IL= Illinois, MD= Maryland, MP= metabolizable protein, SSMA= soluble sugars/malic acid, SNY= Spruce Haven, NY
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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Several approaches have been attempted for incorporating enzymes into diets for dairy cattle. Experiments have been conducted to study effects of adding fibrolytic enzymes to dry forages (Yang et al., 1999) or concentrates (Yang et al., 2000) before feeding and adding amylolytic and proteolytic enzymes targeted towards the starch-protein matrix of sorghum (Chen et al., 1995). In other studies, the effects of direct-fed fibrolytic enzymes applied either to the forage component of the diet (Schingoethe et al., 1999; Kung et al., 2000) or to a TMR (Beachemin et al., 1999) have been examined. Phipps et al. (2002) reviewed recent literature on the effect of direct-fed enzymes on feed intake and milk production. They noted that with few exceptions (Lewis et al., 1999), the direct-fed enzymes produced no significant effect on feed intake. While significant increases in milk production following treatment with direct-fed enzymes have been reported by Lewis et al. (1999), Schingoethe et al. (1999), and Zheng et al. (2000), other workers, including Beauchemin et al. (1999), Rode et al. (1999), and Kung et al. (2000), reported only numerical, nonsignificant increases. Also, Yang et al. (1999, 2000) reported significant changes in milk yield when enzymes were mixed at the time of manufacturing with alfalfa hay cubes or the concentrate portion, respectively. In the studies reviewed, the range in milk production response varied from 0 to 6.3 kg/d. Changes in milk composition due to the application of direct-fed enzymes were variable.
The objective of this study was to examine the effects of feeding one of two enzyme treatments on animal performance. In addition, a treatment group was also included to supply nutrients designed to support microbial growth. The nutrients included nonstructural carbohydrates, which were intended to supply energy to microbes and increase ruminal ammonia utilization and dicarboxylic acids to increase lactate utilization by Selenomonas ruminantium (Martin et al., 2000).
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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A total of 257 primiparous and multiparous Holstein cows were assigned to the study. All cows were between 8 and 166 DIM at the start of the pretreatment period and were in good overall health, including good foot, leg, and udder conformation. The numbers of animals assigned within each site and parity group are presented in Table 1
. Cows were assigned to the study at the beginning of a 14-d pretreatment period within site, parity, and stage of lactation. Stages of lactation were early or mid-lactation. Early-lactation cows were <81 DIM at the initiation of the treatment period at all sites except IL where they were <73 DIM.
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Cows were fed a TMR (Table 2) formulated to meet or exceed NRC (1989) requirements. At each site, the feed for 1 d was mixed once daily, and cows were fed once daily (except Illinois where the daily ration was weighed out every morning but fed in a.m. and p.m. offerings). Also, diets were evaluated using the Cornell-Penn-Miner (CPM) model (Barry et al., 1994) to assure that metabolizable protein (MP) allowable milk was at least 2 kg/d greater than ME allowable milk. Across sites, forages were restricted to no more than 60% of forage DM as corn silage and no less than 40% of forage as alfalfa hay. Cows were fed ad libitum.
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Feed ingredients and control TMR were sampled weekly. An aliquot of each sample was frozen, and forage samples were tested weekly for DM to adjust the as-fed amounts of forage and concentrate in the TMR on a weekly basis. At the completion of the trial, samples were composited for the pretreatment period and for every 4-wk interval during the treatment period. Composite samples were analyzed by wet chemistry methods (Dairy One, Ithaca, NY).
Body weights were measured at the beginning of the treatment period and every 4 wk thereafter. Cows were observed daily and health findings were recorded.
Trial #2
A second trial using the same four treatment groups was conducted at the University of Reading using 122 British Holstein-Friesian cows (Table 1
). The experimental design for trial 2 was similar to trial 1, except there was no blocking by stage of lactation. Instead, all cows began the pretreatment between 11 and 17 DIM, and treatments were initiated 14 d later between 25 and 31 DIM. Also, the diet contained approximately 75% of the forage DM as corn silage and 25% as grass silage (Table 2).
Enzyme Assays
Enzyme preparations A and B were assayed for xylanase, endoglucanase, exoglucanase, ß-glucosidase, ß-xylosidase, 
-L-arabinofuranosidase, -L-arabinopyranosidase, acetyl esterase, and -glucuronidase as described below.
Xylanase activity.
Xylanase was assayed as described by Bailey et al. (1992). A 15-ml test tube containing 1.8 ml of 1.0% (wt/vol) birchwood xylan (Roth 7500) prepared in 50 mM sodium citrate buffer pH 5.3, and 0.196 ml of distilled water were preincubated at 50°C for 10 min. The enzyme reaction was initiated by adding 4 µl of 1:200 diluted enzyme sample, and the reaction continued for 5 min at 50°C. The reaction was terminated by adding 3 ml of dinitrosalicylic acid (DNS) reagent, and the reducing sugars released were determined by the DNS method (Miller, 1959). The unit of enzyme activity was expressed as the amount of enzyme required to release 1 µmol of reducing sugars as xylose equivalent min-1.g-1 of the enzyme sample.
Endoglucanase activity.
Endoglucanase was assayed as described by Wood and Bhat (1988). A 15-ml test tube containing 1 ml of 1.0% (wt/vol) of carboxymethyl cellulose (CM-cellulose, medium viscosity), 0.5 ml of 100 mM sodium acetate buffer pH 5.0, and 0.492 ml of distilled water was preincubated at 50°C for 10 min. At the end of 10 min, 8 µl of 1:200 diluted enzyme sample was added and incubated for a further 15 min at 50°C. The reaction was terminated by adding 3 ml of DNS reagent, and the amount of reducing sugars released was determined as described above. The unit of enzyme activity was expressed as the amount of enzyme required to release 1 µmol of reducing sugars as glucose equivalent min-1.g-1 of the enzyme sample.
Exoglucanase activity.
Exoglucanase was assayed as described by Wood and Bhat (1988). A 15-ml test tube containing 1 ml of 1.0% (wt/vol) of Avicel (Fluka, Sigma-Aldrich, Dorset, UK), 0.5 ml of 100 mM sodium acetate buffer pH 5.0, and 0.492 ml of distilled water was preincubated at 50°C for 10 min. At the end of 10 min, 8 µl of 1:250 diluted enzyme sample was added, and the tubes were incubated for a further 2 h at 50°C. The reaction was terminated by placing the tubes in a boiling water bath for 10 min. The tubes were centrifuged at 1350 x g for 10 min, and the reducing sugars present in the supernatant were determined by the DNS method (Miller, 1959). The unit of enzyme activity was expressed as the amount of enzyme required to release 1 µmol of reducing sugars as glucose equivalent min-1.g-1 of the enzyme sample.
ß-Glucosidase, ß-xylosidase, -L-arabinofuranosidase, -L-arabinopyranosidase, and -D-glucuronidase activities.
ß-Glucosidase, ß-xylosidase, -L-arabinofuranosidase, -L-arabinopyranosidase, and -D-glucuronidase were assayed using microtiter plates in duplicate and using two different enzyme concentrations. A 150-µl reaction volume containing 100 µl of 3 mM substrate, 37.5 µl of 100 mM sodium acetate buffer, pH 5.0, and 12.5 µl of enzyme (1:20 to 1:100 diluted) was incubated at 40°C for 15 min. The substrate for each activity was the respective p-nitrophenyl-glycoside (i.e., p-nitrophenyl-ß-D-glucoside for ß-glucosidase). The reaction was terminated by adding 150 µl of 0.4 M glycine buffer, pH 10.8. The absorbance was read using an Anthos Microplate reader at 405 nm. The unit of enzyme activity was expressed as µmol of p-nitrophenol released min-1.g-1 of enzyme sample.
Acetyl esterase activity.
Acetyl esterase was measured using p-nitrophenyl acetate by microtiter plate method. p-Nitrophenyl acetate (10 mM) was dissolved in 20% (vol/vol) of dimethyl sulphoxide (DMSO), since it was partially soluble in water. However, the above concentration of DMSO had little effect on enzyme activity. Also, the p-nitrophenyl acetate was unstable at temperatures >50°C and pH 7.0. Therefore, the initial rate of the reaction was measured using potassium phosphate buffer at pH 6.8 and at 37°C for 10 min at 405 nm. The reaction was linear up to 10 min and the enzyme activity was determined based on the absorbance at the end of 5 min of incubation. Thus, a typical 200-µl reaction mixture contained 100 µl of 10 mM p-nitrophenyl acetate in 20% DMSO, 50 µl of 50 mM potassium phosphate buffer, pH 6.8 and 50 µl of 1:50 diluted enzyme. The unit of enzyme activity was expressed as µmol of p-nitrophenol released min-1•g-1 of enzyme sample.
Determination of pH optimum of cellulase and hemicellulase activities.
For the determination of pH optimum of xylanase, endoglucanase, and exoglucanase, 1 ml of 1% substrate (birchwood xylan, CM-cellulose, medium viscosity, or Avicel) prepared in distilled water was mixed with 0.9 ml of buffer (50 mM) having varying pH values (citrate-phosphate, pH 2.6 to 7.0; potassium-phosphate, pH 6.6 to 8.0, and boric acid-sodium hydroxide, 8.0 to 9.0) and incubated at 50°C for 10 min. At the end of 10 min, 0.1 ml of suitably diluted enzyme sample was added to the reaction mixture, and the assay was carried out as described above. The reducing sugars released were determined by the DNS method (Miller, 1959), and the pH optimum was determined by plotting the unit of enzyme activity versus pH.
The pH optimum of ß-glucosidase, ß-xylosidase, -L-arabinofuranosidase, and -L-arabinopyranosidase was determined by using microtiter plates as described above and using the above mentioned three buffer systems. In all cases, the pH optimum was determined by plotting the unit of enzyme activity versus pH.
Determination of temperature optimum of xylanase, endoglucanase, and exoglucanase activities.
The optimum temperatures were determined by measuring the xylanase, endoglucanase, and exoglucanase activities present in samples A and B at temperatures between 10 and 70°C at the optimum pH of each enzyme activity. The assay conditions for each activity were as described above. The optimum temperature of each enzyme activity was determined by plotting the activity versus temperature.
Statistical Analyses
Due to differences in the experimental design of the two production trials, each trial was analyzed separately. Data were analyzed by SAS (1997) using the following model:
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where U = overall mean, Ti = treatment effect, Sj = site effect, TSij = treatment by site interaction, Bk = stage of lactation block, TBik = treatment by stage of lactation interaction, Ml = parity group, TMil = treatment by parity interaction, PREijkl = pretreatment covariate, b = regression coefficient for PREijkl, and eijkl = residual error. When P < 0.25, the treatment by site interaction was used as the error term to test significance of the treatment main effect. Treatment effects were deemed significant at P < 0.05. Data for trial 2 were analyzed separately using a similar model excluding the site and stage of lactation terms.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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). The most common reasons for removal were displaced abomasum, mastitis, foot problems, and not adapting to tiestalls. Additionally, data from animals that completed the study but were identified as having health problems that could impact production results were removed. Both the study investigator and the investigator for each site evaluated these cows individually, and if both investigators agreed, data for these animals were excluded from analysis. Data from eight animals in trial 1 were removed due to health problems such as early dry-off and foot health. Data from five additional animals were not included due to not meeting protocol defined DIM at the initiation of treatments. There were no indications that removals were affected by treatments.
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and 5, respectively. For both trials, milk production was not affected (P > 0.10) by treatments. Milk fat and milk protein percentages also were not affected by feeding these additives. Additionally, there were no effects of treatment on SCC, feed intake, or BW change during the 84-d treatment period. In trial 1, there were no treatment x site interactions, indicating that responses were not obtained within any one site. There were no treatment interactions with stage of lactation from trial 1, indicating that responses were not dependent on stage of lactation. Additionally, for both trials 1 and 2, there were no treatment x parity interactions, suggesting that treatments were not effective for either parity group.
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-L-arabinofuranosidase, -L-arabinopyranosidase, acetyl esterase, and -D-glucuronidase) and cellulase (endoglucanase, exoglucanase, and ß-glucosidase) activities (Table 6). In general, the levels of all cellulase and hemicellulase activities in sample A were higher than in sample B, except for -L-arabinofuranosidase (Table 6). Also, both A and B enzyme samples had greater xylanase and endoglucanase activities compared with the other cellulase and hemicellulase activities (Table 6).
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and Table 6. The xylanase from enzyme samples A and B was active between pH 3.0 to 9.0 (Figure 1) and had broad pH optima between 5.0 to 6.0 and 5.5 to 5.8, respectively (Table 6). The highest activity of xylanase from both enzyme samples at optimal pH was similar (Figure 1).
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). The exoglucanase from samples A and B was active between pH 2.6 to 7.0 (Figure 1), but this enzyme had the optimal activity between pH 4.2 to 4.6 and at pH 5.0, respectively. The ß-glucosidase from both enzyme samples was active between pH 2.6 to 8.0, but optimally at pH 4.5 (Table 6). Nevertheless, the ß-glucosidase from sample A had a broader pH optimum and greater activity than that from sample B.
The ß-xylosidase from samples A and B was active between pH 2.6 to 8.0 (Figure 1), but optimally active between pH 2.6 to 4.0. The ß-xylosidase activity in both enzyme samples decreased with the increase in pH from 4.0 to 8.0. Thus, the ß-xylosidase from enzyme samples A and B showed an identical pH versus activity profile except the total activity in sample A was twofold higher than that from sample B.
Both -L-arabinofuranosidase and -L-arabinopyranosidase from enzyme samples A and B were active between pH 2.6 to 7.0 (Figure 1). The -L-arabinofuranosidase from enzyme samples A and B was optimally active between pH 2.6 to 4.0, whereas the -L-arabinopyranosidase from enzyme samples A and B was optimally active at pH 4.3 and at pH between 4.0 to 4.2, respectively. At optimum pH, the activity of -L-arabinofuranosidase from enzyme sample B was five times higher than that from enzyme sample A, whereas the activity of -L-arabinopyranosidase from enzyme sample A was four times higher than that from enzyme sample B.
The xylanase from enzyme sample B was optimally active at 50°C, whereas that from enzyme sample A was optimally active at 60°C (Figure 2). Xylanase from both enzyme samples showed only 10% of the optimal activity up to 20°C (Figure 2). Nevertheless, the activity of xylanase from both samples increased sharply with an increase in temperature up to 50 to 60°C and decreased sharply above 60°C. The endoglucanase from both enzyme samples was optimally active at 60°C (Figure 2). Above 60°C, the endoglucanase activity from sample B decreased rapidly, whereas the endoglucanase activity of sample A decreased more slowly. However, up to 40°C, the endoglucanase from both enzyme samples showed only 20% of the optimal activity. The exoglucanase from enzyme samples A and B was optimally active at 50°C (Figure 2). The pH and temperature activity profiles clearly suggested that the cellulase and hemicellulase activities from these enzyme samples would be less efficient in the rumen because pH and temperature would not be optimum.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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Several mechanisms have been proposed whereby supplemental enzymes could improve animal performance (Beauchemin and Rode, 1996). The most obvious mechanism would be improved digestibility of cellulose or hemicellulose. Improved digestibility has been measured in several studies with dairy cattle (Beauchemin et al., 1999; 2000; Rode et al., 1999; Yang et al., 1999; 2000) and has been associated with increased rate of particle outflow from the rumen, possibly caused by increased rate of digestion and/or lower rumen fluid viscosity, but has not been associated with increased DMI. Under these conditions, if retention time is reduced, the benefits of enhanced fibrolytic activity may be small.
The lack of production responses and the pH optima curves and optimum temperatures suggest that the amount of supplemental enzymatic activity for degradation of cellulose and hemicellulose may be limited in the environment of the rumen of the cows offered the diet used in the current study. Typical pH of the rumen is higher than the pH optima that were obtained, and ruminal temperature is lower than the optimal temperature for enzymatic activity. In general, at pH = 6 most of the enzymatic activities of either enzyme A or B will be reduced by about two-thirds of their optimal activity, and at 38°C, these activities will be further reduced by two-thirds. Therefore, the combined effects of pH and temperature that exist in the rumen would suggest that moderate fibrolytic activity, compared with the amount added, could occur within the rumen with these exogenous enzymes.
Wallace (1997) indicated that two roles supplemental enzymes could fill are either to amplify or complement ruminal activities. If their role was to amplify an existing activity, then the amount of activity added to the rumen would need to significantly add to that already present. Loss of activity at temperatures or pH values that significantly deviate from the optima would make this less likely. However, if complementing an existing activity, the combination of feeding high-energy diets and supplementing an enzyme with a pH optimum in the acidic range might be desirable (Wallace, 1994) and effective. Recent work by Morgavi et al. (2000) has shown the existence of a synergistic effect between ruminal and exogenous enzymes and that this can increase the hydrolytic potential within the rumen environment and could be an important mechanism by which enzyme additives improve fiber digestion. But with any diets and under any conditions, it would be necessary that the enzymes be effective near 39°C in order to have ruminal activity.
Although no effects of the direct-fed enzymes were recorded for milk production in the current study, reports using the same enzymes exist in the literature, which have shown significant milk yield responses (Zheng et al., 2000). Among a range of factors that could affect enzyme efficacy, diet composition may be extremely important as in studies with high concentrate-to-forage ratios rumen pH may be considerably closer to the optima for the enzymes, which could greatly improve fiber digestibility. In addition, a number of other factors such as method of application, specific enzyme complexes, time, and duration of enzyme application before feeding, dryness of the feed, dose rate, and stage of lactation in which the diet is fed can all influence milk production response noted with direct-fed enzyme additives. Wallace (2001) suggested that effects of fungal enzyme preparations are most likely at prefeeding. In the present study, enzymes were added each morning when mixing the TMR. Also, compounds that may or may not have fibrolytic activity in the enzyme mixtures may have acted as adjuvants that stimulated ruminal digestibility. If there were changes in the substrates that were used in producing the enzyme mixtures, these other compounds and/or activities may have changed as well.
Grant and Mertens (1992) have demonstrated that fiber digestion in the rumen is inhibited by low pH, and a more favorable ruminal environment may stimulate growth rate of ruminal fiber-digesting bacteria; however, milk or other production variables were not improved by the addition of SSMA. Few published studies are available where SSMA or malic acid alone were fed to dairy cows (Kung et al., 1982). Most research with malic acid examined effects on ruminal organisms and are reviewed by Martin (1998). Results of previous studies indicate two potential mechanisms of production responses to supplemental SSMA. One potential mechanism is that malic acid may stimulate lactic acid uptake by Selenomonas ruminantium, resulting in a greater ruminal pH. The lack of a production effect may indicate that ruminal acidosis was not a factor that affected milk production for the cows at these sites. Additionally, there was no indication of acidosis based on mean milk fat content (Table 4 and 5). Ruminal pH is affected by the fermentable carbohydrate in the diet. Nocek (1997) recommended that NSC in diets of dairy cows range from 30 to 40% to avoid acidosis. The average NFC at four sites was approximately 33% and at Cornell was 40%. Fiber (ADF and NDF) percentages of the diets were well above the minimum amounts recommended by the current NRC (2001).
The second mechanism would be increased incorporation of ruminal ammonia by the addition of the soluble or nonstructural sugars. The diet may not have been limiting in soluble sugars because NSC was within the recommended range (Nocek, 1997), and diets were formulated at all sites using the CPM model with the restriction that MP allowable milk was 2 kg/d greater than ME allowable milk.
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CONCLUSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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FOOTNOTES |
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2 Current address: Renessen LLC, Bannockburn, IL 60015.
Received for publication March 6, 2002. Accepted for publication May 24, 2002.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONACKNOWLEDGEMENTSREFERENCES |
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