Antibiotics As Growth Promotants: Mode Of Action

Recent concerns about the use of growth-promoting antibiotics in pig diets have renewed interest in the immunologic and growth-regulating functions of the gastrointestinal (GI) tract. The numerically dense and metabolically active microbiota of the pig GI tract represents a key focal point for such questions. The intestinal microbiota is viewed typically as a beneficial entity for the host. Intestinal bacteria provide both nutritional and defensive functions for their host. However, the host animal invests substantially in defensive efforts to first sequester gut microbes away from the epithelial surface, and second to quickly mount immune responses against those organisms that breach epithelial defenses. The impact of host responses to gut bacteria and their metabolic activities require special consideration when viewed in the context of pig production in which efficiency of animal growth is a primary objective. Here, we summarize the working hypothesis that antibiotics improve the efficiency of animal growth via their inhibition of the normal microbiota, leading to increased nutrient utilization and a reduction in the maintenance costs of the GI system. In addition, novel molecular ecology techniques are described that can serve as tools to uncover the relationship between intestinal microbiology and growth efficiency.

 

Numerous physiological, nutritional, and metabolic responses to feed-grade antibiotics have been reported, as summarized in Table 1 (Commission on Antimicrobial Feed Additives, 1997). From a production standpoint, feed antibiotics have been consistently shown to improve pig weight gain and feed efficiency as summarized in Figure 1. The data demonstrate that growth and feed efficiency are improved to a greater extent in young pigs than in older animals. Also, the enhancement of body weight gain is greater than the improvement in feed efficiency, indicating that feed intake is increased in animals receiving antibiotic-supplemented feed. However, antibiotics increase gain and feed efficiency even at constant feed intake (Jones and Tarrant, 1982), consistent with a direct effect on growth which is independent of feed intake. These growth responses are associated with improved nitrogen metabolism, including an increase in apparent nitrogen digestibility (3.0%), increased nitrogen retention (5.8%), and reduced nitrogen excretion (10%) in pigs fed tylosin (10 ppm to 50 ppm; Table 2). Growth-promoting antibiotics also benefit protein metabolism irrespective of protein concentration in the diet (Roth and Kirchgessner, 1993). Mechanisms by which antibiotics enhance growth should thus be consistent with their demonstrated effects on growth, feed efficiency, nitrogen metabolism, and with the relatively greater responses observed in younger animals.

 

Table 1. Summary of reported physiological, nutritional and metabolic effects of growth promoting antibiotics (Modified from Commission on Antimicrobial Feed Additive, 1997).

 

Proposed Mechanisms for Antibiotic-Mediated Growth Enhancement
At least four mechanisms have been proposed as explanations of antibiotic mediated growth enhancement: 1) inhibition of sub-clinical infections, 2) reduction of growth-depressing microbial metabolites, 3) reduction of microbial use of nutrients, and 4) enhanced uptake and use of nutrients through the thinner intestinal wall associated with antibiotic-fed animals (Francois, 1962; Visek, 1978; Anderson et al., 2000). These share the common postulate that intestinal bacteria, whether commensal or pathogenic, depress animal growth, either directly or indirectly, through their metabolic activities. Perhaps the best support of this hypothesis is the observation that oral antibiotics do not enhance the growth of germ-free animals (Coates et al., 1963), while inoculating germ-free animals with GI bacteria depresses growth (Coates, 1980). A clear difference between germ-free and conventional animals is a thinner wall of the small intestine, with a reduction in connective tissue and lymphoid elements (Coates, 1980). Microscopic evaluation of germ-free intestine reveals a more regular and slender villus structure, with a thinner lamina propria. Further, the rate of renewal of epithelial cells is slower in germ-free animals (Coates, 1980), which may have a beneficial effect on basal energy expenditure and the efficiency of nutrient utilization. These observations are consistent with the view of Reeds and coworkers (1993) that in rapidly growing young animals, the GI tract and skeletal muscle draw from the same limited supply of nutrients and, in effect, compete for nutrients.

 

Table 2. Effect of antibiotics on nitrogen metabolism in swine.¹

¹Weldon (1997)

 

The Intestinal Microbiota is Competitive with the Host in the Small Intestine but Cooperative in the Large Intestine
Most of the attention given the pig intestinal microbiota has focused on the large intestine. Indeed, the large intestine (cecum and colon) is a major site of microbial colonization because of slow digesta turnover, and it is characterized by large numbers of bacteria (1010 – 1011 per g or ml of content), low redox potential, and relatively high short chain fatty acid (SCFA) concentrations. The composition of the hindgut microbiota is both diverse and stable. Several hundred anaerobic bacterial species and strains appear to coexist without one or a few becoming dominant (Fewins et al., 1957; Smith and Jones, 1963; Allison et al., 1979; Robinson et al., 1981; Robinson et al., 1984; Varel et al., 1987; Anugwa et al., 1989; Butine and Leedle, 1989; Pryde et al., 1999). Microbial activity in the cecum and colon appears to benefit the host (Hedde and Lindsay, 1986), with estimates up to 5-20% of the pig’s total energy being provided by fermentation end-products (Friend et al., 1963). However, the small intestine is the principal site of nutrient and energy absorption, and thus is the region in which bacterial activity is likely to have the greatest influence on the efficiency of growth (Figure 2; Anderson et al., 2000; Gaskins, 2001). In the proximal small intestine, the rate of digesta flow and thus the rate of bacterial washout, exceeds the maximal growth rates of most bacterial species. Accordingly, this intestinal region is colonized typically by bacteria that adhere to the mucus layer or epithelial cell surface. Acid-tolerant lactobacilli and streptococci are thought to predominate in the upper small intestine (Fewins et al., 1957). The distal small intestine (ileum) maintains a more diverse microbiota and higher bacterial numbers (108 per g or ml of contents) than the upper intestine, and is considered a transition zone preceding the large intestine. In a study of microbial gas production in various gastrointestinal regions, Jensen and Jørgensen (1994) reported that the highest H2 concentrations and production rates were found in the distal small intestine in pigs. This indication for significant microbial activity in the pig ileum was substantiated in their study by dense populations of culturable anaerobic bacteria and the detection of high ATP concentrations.

 

Culture-based studies have shown that small intestinal bacteria tend to compete with the host for energy and amino acids (Hedde and Lindsey, 1986). For example, as much as 6% of the net energy in pig diets can be lost due to bacterial utilization of glucose in the small intestine (Vervaeke et al., 1979). Bacterial use of glucose to produce lactic acid reduces the energy available to the host epithelium (Saunders and Sillery, 1982). Lactic acid also enhances peristalsis, thus increasing the rate of nutrient transit through the intestine (Saunders and Sillery, 1982). Amino acids, which are also degraded by small intestinal bacteria, are made unavailable to the pig and produce toxic metabolites such as amines, ammonia, phenols and indoles (Macfarlane and Macfarlane, 1995). Microbial deconjugation and dehydroxylation of bile acids impair lipid absorption and produces toxic degradation products (Hylemon, 1985). Mucolytic activities of intestinal bacteria are likely key for intestinal colonization but compromise the mucosal barrier and indirectly affect the efficiency of growth via stimulation of additional mucus production (Deplancke and Gaskins, 2001).

 

It is interesting to note that although different types of bacteria may generate one or more of the metabolites mentioned, the Gram-positive facultative anaerobes, which are oxygen-tolerant and predominant in the small intestine, often contribute more than one toxic catabolite (Gaskins, 2001). Further, the small intestinal microbiota consists predominantly of Gram-positive bacteria (Stewart, 1997), and most growth-promoting antibiotics target Gram-positive organisms (Table 3). These observations are consistent with involvement of small intestinal Gram-positive bacteria in growth depression.

 

It is curious that a class of organisms that appear to depress growth, namely Gram-positive facultative anaerobes including strains of Lactobacillus and Enterococcus, are also often used as probiotic organisms for enhancing health and promoting growth in livestock (reviewed in Jonsson and Conway, 1992). The growth-promoting effect of probiotics in livestock is less consistent than that observed with antibiotic supplementation (Jonsson and Conway, 1992). Supplementation of animals and humans with certain probiotic bacteria has been shown to provide protection against intestinal, diarrhea-producing pathogens (reviewed in McCracken and Gaskins, 2000). Therefore, probiotics may promote growth under situations in which certain pathogens are present; however, these same organisms in a cleaner facility may suppress growth via the mechanisms discussed above.

 

Table 3. Growth-promoting antibiotics and their antibacterial mode of action.¹

¹Modified from O’Connor (1980), Commission on Antimicrobial Feed Additives (1997), and Lawrence (1998)
²Commonly accepted property. “No” does not necessarily mean 0 percent absorption nor is “Yes” equal to 100 percent absorption.

 

The combined potential of microbial activities to negatively impact intestinal functions clearly supports the hypothesis that certain bacterial populations commonly inhabiting the pig small intestine, though not necessarily pathogenic, cause a depression in growth which is reversed when the responsible organism(s) are metabolically inhibited or eliminated by feed-grade antibiotics (Figure 3). However, the basic questions relating to the effects of microbial production of toxic catabolites in the pig intestine remain mostly unanswered, including the taxonomy, ecology, and metabolic properties of target bacteria. Because of the profound bias introduced by cultivation-based techniques, these questions will be defined most efficiently through the use of novel molecular ecology techniques.

 

Novel Methods for Intestinal Microbial Ecology Studies
Several limitations are associated with cultivation-based microbiological techniques, particularly for surveying the intestinal ecosystem (Vaughan et al., 2000). In addition to being time- and labor-intensive, the use of selective media for different types of bacteria imposes an a priori bias on the types of bacteria that can be enumerated. Further, only 20-40% of bacterial species from the mammalian GI tract can be cultured and identified using current cultivation techniques. In other words, 60-80% of intestinal bacterial species may be overlooked (Vaughan et al., 2000; Langendijk et al., 1995; Suau et al., 1999). In view of this, we have defined the utility and limitations of a cultivationindependent technique that uses the phylogenetic information contained in 16S rDNA to objectively study the intestinal microbiota (Simpson et al., 1999; Simpson et al., 2000; McCracken et al., 2001).

 

Denaturing gradient gel electrophoresis (DGGE) is a polymerase chain reaction (PCR)-based technique in which DNA from a mixed sample is amplified using conserved 16S rDNA bacteriadomain primers (Muyzer et al., 1998). Although all PCR products are of approximately equal size, when electrophoresed on a polyacrylamide gel containing an increasing gradient of DNA denaturants, individual amplicons cease to migrate as the double-stranded products denature according to their G+C content (Freter, 1983). Thus, this approach allows separation of individual sequences based on 16S rDNA G+C content, corresponding to the different microbial species within the sample. Banding patterns from mixed samples are compared to evaluate the relative similarity of microbial communities from different habitats or treatments. Further, following electrophoresis, individual bands can be excised from the gel for sequencing and phylogenetic identification. This allows the characterization of complex microbial populations independent of bacterial cultivation.

 

Effects of Antibiotics on the Normal Microbiota
We have recently used 16S rDNA PCRDGGE to examine the effects of feed grade antibiotics on the intestinal microbiota in ilealcannulated pigs. Two antibiotic regimes were compared with a standard nonmedicated diet: 1) continuous administration of a single antibiotic and 2) an antibiotic rotation sequence in which a different antibiotic was used each week. Newly weaned pigs were fed control (no antibiotic) or medicated diets for five weeks. Ileal microbiota profiles were compared by DGGE-PCR analysis as demonstrated in the preliminary data in Figure 4 (Collier et al., unpublished observations). Similar gels were generated from all pigs. Through the use of statistical measures of community diversity, we observed that the ileal microbiota is more homogenous in anti-biotic-treated pigs. This finding reflects the fact that bacterial community profiles generally varied among individual animals and did so to a greater extent in the control pigs that were weaned to a non-medicated diet. Other molecular-based ecology studies, by us and others, in mice, pigs, and humans also demonstrate that, although the intestinal bacterial community within a single individual is relatively stable over time, bacterial populations from different individuals vary significantly (Simpson et al., 1999; McCartney et al., 1996; Kimura et al., 1997; Zoetendal et al., 1998). Individual variation in bacterial community profiles is consistent with the possibility that genetically encoded chemical epitopes expressed in mucus may dictate which bacteria colonize specific intestinal segments. This has significant implications for intestinal health issues and may also partly explain individual differences in the realization of growth potential in livestock species.

 

The inclusion of antibiotics also eliminated particular bacterial groups as evidenced by the disappearance of certain 16S rDNA PCR bands on the DGGE gels (Fig. 5). Individual bands that were eliminated by antibiotics were excised from DGGE gels and cloned and sequenced to identify the bacteria affected. Thus far, six Lactobacillus and two Streptococcus species have thus far been identified through this process as being selectively depleted in antibiotic-treated pigs. Interestingly, lactobacilli are thought to be one of the main contributors to microbial bile acid biotransformation in the small intestine (Tannock et al., 1989). Indeed, Feighner and Dashkevicz (1987, 1988) proposed that an important mechanism of growth-promoting antibiotics is the inhibition of microbial bile acid biotransformation in the intestine. Microbial deconjugation and dehydroxylation of bile impairs lipid absorption by the host animal (DeSomer et al., 1963; Eyssen, 1973) and produce toxic degradation products that can impair growth (Eyssen and DeSomer, 1963a). Bile acids are not deconjugated in the germfree intestine, demonstrating the important role of commensal bacteria in this process (Madsen et al., 1976). The particular contributions of the lactobacilli to bile acid biotransformation in the intestine is demonstrated by evidence that ileal bile salt hydrolase activity in conventional mice is reduced 86% by the elimination of lactobacilli from the microbiota, and by greater than 98% when both lactobacilli and enterococci are eliminated (Tannock et al., 1989). Using chicks, Eyssen and DeSomer (1963b) first suggested that bile acid transformation products might be responsible for the growth depression caused by intestinal bacteria. Additional evidence from chick studies showed that bile acid deconjugation by intestinal bacteria causes growth depression that is reversible by antibiotic supplementation (Fuller et al., 1984). Further, Feighner and Dashkevicz (1987, 1988) have shown an inverse relationship between the level of cholyltaurine hydrolase activity in the small intestine and growth rate in broiler chickens fed antibiotics. Antibiotictreated pigs also demonstrated decreased concentrations of lithocholic acid and corresponding increases in average daily gain and feed efficiency, consistent with work reported in poultry (Tracy and Jensen, 1987). While additional studies are required to verify the contribution of bile biotransformation to growth depression, these data clearly demonstrate the utility of molecular ecology to study the effects of antibiotics on the intestinal microbiota.

 

CONCLUSIONS

 

It is proposed that antibiotic modification of the small intestinal microbiota of swine permits more efficient intestinal and therefore whole animal growth. Recent advances in swine growth research have been directed primarily at affecting growth, feed efficiency, and body composition by altering the metabolism of muscle and fat (Moody et al., 2000; Beermann and DeVol, 1991). An understanding of the inter-relationship of intestinal physiology, microbiology, and immunology to swine growth will become increasingly important to critically evaluate the impact of commensal bacteria on animal growth. In that regard, it is particularly exciting that molecular techniques are now available that allow a better understanding of how the intestinal microbial profile is changed with the use of antibiotics. These advances will allow the development of novel technologies, management systems, and modified nutrition to optimize intestinal health and animal growth.

 

REFERENCES

Allison, M. J., Robinson, I. M., Bucklin, J. A. and Booth, G. D. 1979. Comparison of bacterial populations of the pig cecum and colon based upon enumeration with specific energy sources. Appl. Environ. Microbiol. 37: 1142-1151.
 
Anderson, D.B., McCracken, V.J., Aminov, R.I., Simpson, J.M., Mackie, R.I., Verstegen, M.W.A. and Gaskins, H.R. 1999. Gut microbiology and growth-promoting antibiotics in swine. Nutrition Abstracts and Reviews. Series B: Livestock Feeds and Feeding. 70: 101-188.
 
Anugwa, F.O.I., Varel, V.H., Dickson, J. S., Pond, W.G. and Krook, L. P. 1989. Effects of dietary fiber and protein concentration on growth, feed efficiency, visceral organ weights and large intestine microbial populations of swine. J. Nutr. 119: 879-886.
 
Beermann, D.H. and Devol, D.L. 1991. Effects of somatotropin, somatotropin releasing factor and somatostatin on growth. In: A.M. Pearson and T.R. Dutson (Eds.). Growth Regulation in Farm Animals. pp. 373-426. Elsevier Applied Science, London, Butine, T.J., and Leedle, J.A.Z. 1989. Enumeration of selected anaerobic bacterial groups in cecal and colonic contents of growing-finishing pigs. Appl. Environ. Microbiol. 55: 1112-1116.
 
Coates, M.E., Fuller, R., Harrison, G.F., Lev, M., and Suffolk, S.F. 1963. A comparison of the growth of chicks in the Gustafsson germ-free apparatus and in a conventional environment, with and without dietary supplements of penicillin. Brit. J. Nutr. 17: 141-151.
 
Commission on Antimicrobial Feed Additives. 1997. Antimicrobial Feed Additives. Government Official Reports, SOU, Ministry of Agriculture, Stockholm, p. 132.
 
Coates, M.E. 1980. The gut microflora and growth. In: T.L.J. Lawrence (Ed.). Growth in Animals. pp. 175-188. Butterworths, Boston, pp.
 
Deplancke, B. and Gaskins, H.R. 2001. Microbial modulation of innate defense: Goblet cells and the intestinal mucus layer. Am. J. Clin. Nutr. 73 (suppl): 1131S-1141S.
 
De Somer, P., Eyssen, H. and Evard, E. 1963. The influence of antibiotics on fecal fats in chickens. In: A.C. Frazer (Ed.). Biochemical Problems of Lipids. pp. 84-90. Elsevier/North Holland Publishing Co., Amsterdam,
 
Eyssen, H. and DeSomer, P. 1963a. Toxicity of lithocholic acid for the chick. Poultry Sci. 42: 1020-1022.
 
Eyssen, H. and DeSomer, P. 1963b. The mode of action of antibiotics in stimulating growth in chicks. J. Exp. Med. 117: 127-138.
 
Eyssen, H. 1973. Role of gut microflora in metabolism of lipids and sterols. Proc. Nutr. Soc. 32: 59-63.
 
Feighner, S.D. and Dashkevicz, M.P. 1987. Subtherapeutic levels of antibiotics in poultry feeds
and their effects on weight gain, feed efficiency, and bacterial cholyltaurine hydrolase activity.
Appl. Environ. Microbiol. 53: 331-336.
Feighner, S.D. and Dashkevicz, M.P. 1988. Effect of dietary carbohydrates on bacterial
cholyltaurine hydrolase in poultry intestinal homogenates. Appl. Environ. Microbiol. 54: 337-
342.
Fewins, B.G., Newland, L.G.M. and Briggs, C.A.E. 1957. The normal intestinal flora of the pig. III. Qualitative studies of lactobacilli and streptococci. J. Appl. Bact. 20: 234-242.
 
Francois, A.C. 1962. Mode of action of antibiotics on growth. World Review of Nutrition and Dietetics. 3: 21.
 
Friend, D.W., Cunningham, H.M. and Nicholson, J.W.G. 1963. The production of organic acids in the pig. Can. J. Anim. Sci. 43: 156-168.
 
Fuller, R., Cole, C. B. and Coates, M. E. 1984. The role of Streptococcus faecium in antibioticrelieved growth depression in chickens. In: M. Woodbine (Ed.). Antimicrobials and Agriculture. pp. 395-403. Butterworths, London, Gaskins, H.R. 2001. Intestinal bacteria and their influence on swine growth. In: A.J. Lewis, LL Southern (Ed.). Swine Nutrition. pp. 585-608. CRC Press, Boca Raton, FL.
 
Hays, V.W. 1969. Biological basis for the use of antibiotics in livestock production. The Use of Drugs in Animal Feeds, National Academy of Sciences, National Research Council Publication 1679, pp. 11-30.
 
Hedde, R.D. and Lindsey, T.O. 1986. Virginiamycin: A nutritional tool for swine production. Agri-Practice. 7, Numbers 3 and 4.
 
Hylemon, P.B. 1985. Metabolism of bile acids in intestinel microflora. In: H. Danielson and J. Svovall (Eds.). Steroids and bile acieds: new comprehensive biochemistry, vol. 12. pp. 331-334. Elsevier Publishing Inc. Amsterdam, The Netherlands.
 
Jensen, B.B., and Jørgensen, H. 1994. Effect of dietary fiber on microbial activity and microbial gas production in various regions of the gastrointestinal tract of pigs. Appl. Environ. Microbiol. 60: 1897-1904.
 
Jones, P.W. and Tarrant, M.E. 1982. The effect of various factors on the efficacy of tylosin as a growth promoter in clinically healthy pigs. Anim. Prod. 34: 115-121.
 
Jonsson, E. and Conway, P. 1992. Probiotics for pigs. In: R. Fuller (Ed.) Probiotics: The Scientific Basis. pp. 260-316. Chapman and Hall, London.
 
Kimura, K., McCartney, A.L., McConnell, M.A. and Tannock, G.W. 1997. Analysis of fecal populations of bifidobacteria and lactobacilli and investigation of the immunological responses of their human hosts to predominant strains. Appl. Environ. Microbiol. 63: 3394-3398.
 
Langendijk P.S., Schut, F., Janse, G.J., Raangs, G.C., Kamphuis, G.R., Wilkenson, M.H. and Welling, G.W. 1995. Quantitative fluorescence in situ hybridization of Bifidobacterium spp. with genus-specific 16S rRNA-targeted probes and its application in fecal samples. Appl. Environ. Microbiol. 61: 3069-3075.
 
Lawrence, K. 1998. Growth promoters in swine. Proc 15th International Pig Veterinary Society, Birmingham, England, July, 1998. pp. 337-343.
 
Madsen, D., Beaver, M., Chang, L., Bruckner-Kardoss, E. and Wostmann, B. 1976. Analysis of bile acids in conventional and germfree rats. J. Lipid Res. 17: 107-111.
 
Macfarlane, S., and G. T. Macfarlane. 1995. Proteolysis and amino acid fermentation. In: Gibson, G. R., and G. T. Macfarlane (Eds.). Human Colonic Bacteria: Role in Nutrition, Physiology, and Pathology. pp. 75-100. CRC Press, Boca Raton.
 
McCartney, A.L., Wenzhi, W. and Tannock, G.W. 1996. Molecular analysis of the composition of the bifidobacterial and Lactobacillus microflora of humans. Appl Environ Microbiol. 62: 4608-4613.
 
McCracken, V.J. and Gaskins, H.R. 1999. Probiotics and the immune system. In: Tannock, G.W. (Ed.). Probiotics: A Critical Review. pp. 85-112. Horizon Scientific Press, Norfolk, England.
 
McCracken, V.J., Simpson, J.M., Mackie, R.I. and Gaskins H.R. 2001. Molecular ecological anlaysis of dietary and antibiotic-induced alterations of the mouse intestinal microbiota. J. Nutr. 131: 1862-1870.
 
Moody, D.E., Hancock, D.L. and Anderson, D.B. 2000. Phenethanolamine repartitioning agents. In: J.P.F. D’Mello (Ed.). Farm Animal Metabolism and Nutrition. pp. 65-96. CAB International, Wallingford, Oxon, UK.
 
Muyzer, G., Brinkhoff, T., Nübel, U., Santegoeds, C., Schäfer, H. and Wawer, C. 1998. Denaturant gradient gel electrophoresis in microbial ecology. In: Akkermans A., van Elsas JD, de Bruijn F. (eds.). Molecular Microbial Ecology Manual. Vol. 3.4.4. p. 1-27. Kluwer Academic Publishers,
 
O’Connor, J.J. 1980. Mechanisms of growth promoters in single-stomached animals. In: T.L.J. Lawrence (Ed.). pp. 207-227. Growth in Animals. Butterworths, London.
 
Pryde, S.E., Richardson, A. J., Stewart, C.S. and Flint, H. J. 1999. Molecular analysis of the microbial diversity present in the colonic wall, colonic lumen, and cecal lumen of a pig. Appl. Environ. Microbiol. 65: 5372-5377.
 
Reeds, P.J., Burrin, D.G., Davis, T.A. and Fiorotto, M.L. 1993. Postnatal growth of gut and muscle: competitors or collaborators. Proc. Nutr. Soc. 52: 57-67.
 
Robinson, I.M., Allison, M.J. and Bucklin, J.A. 1981. Characterization of the cecal bacteria of normal pigs. Appl. Environ. Microbiol. 41: 950-955.
 
Robinson, I.M., Whipp, S.C. Bucklin, J.A. and Allison, M.J. 1984. Characterization of predominant bacteria from the colons of normal and dysenteric pigs. Appl. Environ. Microbiol. 48: 964-969.
 
Roth, F.X. and Kirchgessner, M. 1993. Influence of avilamycin and tylosin on retention and excretion of nitrogen in finishing pigs. J. Anim. Physiol. Anim. Nutr. 69: 245-250.
 
Saunders, D.R. and Sillery, J. 1982. Effect of lactate on structure and function of the rat intestine. Dig. Dis. 27: 33-41.
 
Simpson, J.M., McCracken, V.J., Gaskins, H.R. and Mackie, R.I. 2000. Denaturing gradient gel electrophoresis analysis of 16S rDNA amplicons to monitor changes in fecal bacterial populations of weaning pigs following introduction of Lactobacillus reuteri strain MM53. Appl. Environ. Microbiol. 66: 4705-4714.
 
Simpson, J.M., McCracken, V.J., White, B.A., Gaskins, H.R. and Mackie, R.I. 1999. Application of denaturant gradient gel electrophoresis for the analysis of the porcine gastrointestinal microbiota. J Microbiol Methods 36: 167-179.
 
Smith, H.W. and Jones, J.E.T. 1963. Observation on the alimentary tract and its bacterial flora in healthy and diseased pigs. J. Path. Bact. 86: 387-395.
 
Stewart, C. S. 1997. Microorganisms in hingut fermentors. In: (Mackie R. I., B. A. White, and R. E. Isaacson (Eds.). Gastrointestinal Microbiology. Vol. 2. pp. 142-175. Chapman and Hall, New York.
 
Suau, A., Bonnet, R., Sutren, M., Godon, J-J, Gibson, G.R., Collins, M.D. and Doré, J. 1999. Direct analysis of genes encoding 16S rRNA from complex communities reveals many novel molecular species within the human gut. Appl. Environ. Microbiol. 65: 4799-4807.
 
Tannock, G. W., Dashkevicz, M. P. and Feighner, S. D. 1989. Lactobacillie and bile salt hydrolase in the murine intestinal tract. Appl. Environ. Microbiol. 55: 1848-1851.
 
Tracy, J.D. and Jensen, A.H. 1987. Effects of a dietary antimicrobial (carbadox) on liver cholesterol 7 alpha-hydroxylase activity and bile acid patterns in the young pig. J. Anim. Sci. 65: 1013-1018.
 
Varel, V.H., Robinson, I.M. and Pond, W.G. 1987. Effect of dietary copper sulfate, Aureo SP250, or clinoptilolite on ureolytic bacteria found in the pig large intestine. Appl. Environ. Microbiol. 53: 2009-2012.
 
Vaughan E.E., Schut, F., Heilig, H.G.H.J., Zoetendal, E.G., de Vos W.M. and Akkermans A.D.L. 2000. A molecular view of the intestinal ecosystem. Curr. Issues Intest. Microbiol. 1: 1- 12.
 
Vervaeke, I.J., Decuypere, J.A., Dierick, N.A., and Henderickx, H.K. 1979. Quantitative in vitro evaluation of the energy metabolism influenced by virginiamycin and spiramycin used as growth promoters in pig nutrition. J. Anim. Sci. 49: 846-856.
 
Visek, W.J. 1978. The mode of growth promotion by antibiotics. J. Anim. Sci. 46: 1447-1469.
 
Weldon, W.C. 1997. Tylosin: Effects on nutrient metabolism. In: Proceedings of World Pork Exposition Swine Research Review Elanco Animal Health, Greenfield, IN.
 
Zoetendal, E.G., Akkermans, A.D.L. and de Vos, W.M. 1998. Temperature gradient gel electrophoresis analysis of 16S rRNA from human fecal samples reveals stable and host-specific communities of active bacteria. Appl Environ Microbiol. 64: 3854-3859.
 
 

H. R. Gaskins, Ph.D., University of Illinois
Rex Gaskins, a native of coastal North Carolina, obtained B.S. and M.S. degrees in animal science from North Carolina State University and a Ph.D. in nutritional sciences from The University of Georgia. From 1989-92, he completed postdoctoral studies in immunology at The Jackson Laboratory in Bar Harbor, Maine. Dr. Gaskins has been on the faculty of the University of Illinois since 1992 where he is now Professor of Immunobiology in the Departments of Animal Sciences and Veterinary Pathobiology. Research in his laboratory focuses on host-intestinal microbe interactions and is supported by the National Institutes of Health, the USDA-NRI Competitive Grants Program, and private industry.