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ANIMAL NUTRITION |
Department of Animal Sciences, University of Illinois, Urbana 61801
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Abstract |
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 Top Abstract INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Key Words: dog • immunity • microbiota • nutrient digestibility • yeast cell wall
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INTRODUCTION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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The mannan moiety in YCW (as much as 31%, DM basis, Northcote and Horne, 1952
) has been associated with increased (~57%) ileal IgA secretion in rats (Buts et al., 1990). Ileal immunoglobulin A (IgA) increased by 44% in dogs supplemented with a combination of YCW and fructooligosaccharides (FOS; Swanson et al., 2002). Secretory IgA in the intestine limits the absorption of protein antigens and may neutralize viruses (Elson and Mestecky, 1995), and increased secretory IgA may, therefore, be beneficial to gut health.
Mannanoligosaccharides (from YCW) are fermented by dogs (Vickers et al., 2001) and may increase fecal lactobacilli (Swanson et al., 2002) and bifidobacteria (Grieshop et al., 2004) and are, therefore, suggested to have prebiotic effects. Presently, however, too few data are available on characteristics of mannanoligosaccharides for them to be classified as prebiotics (Gibson et al., 2004).
The potential effect of mannanoligosaccharides on the intestinal immune system, combined with the high concentration of mannan components in dry YCW, may make YCW preparations useful functional dietary ingredients in pet foods by improving intestinal health and resistance against intestinal upset. The ideal dose of YCW in pet foods is currently unknown, but high supplementation levels (5% of diet) decrease nutrient digestibility (Zentek et al., 2002). The aim of this study was to evaluate possible dose-response effects of the YCW preparation, Safmannan (Lesaffre Yeast Corporation, Milwaukee, WI), on nutrient digestibility, immunological indices, and fecal microbiota in adult dogs.
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MATERIALS AND METHODS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Five purpose-bred, adult female dogs (Marshall Bioresources, North Rose, NY) with hound bloodlines, an average initial BW of approximately 23 kg, and an average age of 4 yr were surgically prepared with an ileal T-shaped cannula according to the methods described by Walker et al. (1994). After surgery, the dogs were closely monitored daily for clinical abnormalities and given a 2-wk recovery period prior to the experiment. Dogs were housed individually in kennels (2.4 x 1.2 m) in a temperature-controlled room with a 16 h of light:8 h of dark cycle at the animal care facility of the Edward R. Madigan Laboratory on the University of Illinois campus.
Oligosaccharide-free ingredients were used in diet formulation, with brewer’s rice, poultry by-product meal, and poultry fat constituting the main ingredients of the dry, extruded kibble diet (Table 1). The diet formulation was milled at Lortscher Agri Service Inc. (Bern, KS) and extruded at Kansas State University’s BIVAP facility (Manhattan, KS) under the direction of Pet Food and Ingredient Technology Inc. (Topeka, KS). Dogs were offered 140 g of the diet twice daily (0800 and 2000). Chromic oxide was used as a digestion marker. On d 6 through 14 of each experimental period, dogs were dosed with 0.5 g of Cr2O3 at each feeding via a gelatin capsule, for a total of 1.0 g of the marker/d. Fresh water was available at all times.
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Sample Collection
A 5 x 5 Latin square design with 14-d periods was used. A 10-d adaptation phase preceded a 4-d collection of feces and ileal effluent. Ileal effluent was collected 3 times/d, with an interval of 4 h between collections. Individual collections were 1 h in length. Sampling times were rotated 1 h from the previous day’s collection time. For example, sampling times on the first collection day were 0800, 1200, and 1600; on the second day, samples were collected at 0900, 1300, and 1700, etc. Ileal samples were collected by attaching a sterile sampling bag (Whirlpack, Fisher Scientific, Pittsburgh, PA) to the cannula barrel with a rubber band. Prior to attachment of the bag, the interior of the cannula was scraped clean with a spatula and dried digesta was discarded. During collection of ileal effluent, dogs were encouraged to move freely. To prevent the dogs from pulling the collection bag from the cannula, Bite-Not collars (Bite-Not Products, San Francisco, CA) were used during collection times. After ileal effluent collection, the cannula plug was inserted and the cannula site was cleaned with a dilute betadine solution.
Although nutrient digestibility was calculated based on digestion marker recovery, total feces excreted during the collection phase of each period were removed from the floor of the pen, weighed, composited, and frozen at –20°C. All fecal samples during the 4-d collection phase were scored for consistency according to the following system: 1 = hard, dry pellets, small hard mass; 2 = hard, formed, dry stool, remains firm and soft; 3 = soft, formed and moist stool, retains shape; 4 = soft, unformed stool, assumes shape of container; 5 = watery, liquid that can be poured.
On d 14 of each period, a blood sample (5 mL) was collected via jugular puncture into nonheparinized, evacuated tubes to obtain serum Ig (IgA, IgG, and IgM) concentrations. Another 5 mL blood sample was collected in evacuated tubes with EDTA for a complete blood count (total white blood cells, neutrophils, eosinophils, lymphocytes, and monocytes).
Sample Handling
Ileal samples were frozen at –20°C in their individual bags. At the end of the experiment, all ileal effluent samples were composited for each dog for each period and refrozen at –20°C. Prior to analysis, ileal effluent was lyophilized in a Dura-Dry MP, microprocessor-controlled freeze-drier (FTS Systems, Stone Ridge, NY). Feces and diets were dried at 55°C in a forced-air oven. After drying, diets, fecal samples, and ileal samples were ground through a 2-mm screen in a Wiley mill (model 4, Thomas Scientific, Swedesboro, NJ).
On d 14 of each period, fresh fecal samples were collected within 15 min of defecation and an aliquot was immediately transferred to a preweighed Cary-Blair transport media container (Para Pak C&S, Meridian Bioscience Inc., Cincinnati, OH) for subsequent bacterial enumeration (total anaerobes, total aerobes, bifidobacteria, lactobacilli, C. perfringens, and E. coli). Aliquots of fresh feces were transferred to sterile cryogenic vials (Nalgene, Rochester, NY) and frozen at –80°C until DNA extraction for microbial analysis. Additional aliquots were used for pH measurement and fresh fecal DM determination.
Chemical Analyses
Diet, YCW, feces, and ileal samples were analyzed for DM, OM, and ash using AOAC (2000) methods. Crude protein was calculated from Leco total N values (AOAC, 2000). Total lipid content (acid hydrolyzed fat, AHF) was determined by acid hydrolysis followed by ether extraction according to the AACC (1983) and Budde (1952). Gross energy was measured using an oxygen bomb calorimeter (Model 1261, Parr Instruments, Moline, IL). Dietary fiber concentrations [total (TDF), soluble (SDF), and insoluble (IDF) fractions] were determined according to Prosky et al. (1984, 1992). Chromium concentrations in digesta and fecal samples were analyzed according to Williams et al. (1962) using atomic absorption spectrophotometry (Model 2380, Perkin-Elmer, Norwalk, CT).
Monosaccharide composition of YCW was analyzed according to Bourquin et al. (1990). Briefly, an internal standard of 1 mg of inositol/mL in 72% (wt/wt) sulfuric acid was prepared. One milliliter of the internal standard was added to a screw-cap tube, containing 50 mg of finely ground sample, and vortexed gently. After 30 min, the samples were diluted to 2 N sulfuric acid by adding 11 mL of distilled, deionized water. Samples were hydrolyzed for 2 h in a boiling water bath. The hydrolyzed samples were filtered through Whatman GF/D glass fiber filters (Whatman Inc., Florham Park, NJ) and then neutralized by passing through a preparation column containing 15 g of AG 4-X4 anion exchange resin (BioRad, Hercules, CA). Effluents were collected in 200-mL volumetric flasks and brought up to volume with distilled, deionized water. Monosaccharide concentrations in the effluent were determined using HPLC (Bourquin et al., 1990).
Immunological Analyses
Ileal IgA concentrations were determined according to Nara et al. (1983). Briefly, 2 g of lyophilized sample were crushed using a mortar and pestle and suspended in 20 mL of PBS (pH 7.2). Samples were mixed at room temperature for 30 min and then centrifuged at 20,000 x g for 30 min at 4°C. Concentrations of IgA in the supernatant were measured using a radial immunodiffusion kit (MP Biomedicals, Aurora, OH).
After blood was collected in nonheparinized evacuated tubes, samples were centrifuged at 2,000 x g for 20 min at 4°C and the serum was collected. Serum IgA, IgG, and IgM concentrations were measured using selective radial immunodiffusion kits (MP Biomedicals, Aurora, OH). For each kit, the accuracy was ± 10% within the limits of the standard curve, and for each immunodiffusion plate an individual standard curve is created to account for interplate variability. The methodology has been validated by Fahey and McKelvey (1965), and specifically for dog immunoglobulins by Reynolds and Johnson (1970). This assay is for dog-specific Ig with the following detection limits:
IgA 15 to 125 mg/dL,
IgG 250 to 2,400 mg/dL, and
IgM 23 to 230 mg/dL.
The blood collected in the evacuated tubes containing EDTA was used for CBC determination, which was performed on a Cell-Dyn 3500 hematology analyzer (Abbott Laboratories, Abbott Park, IL).
Microbial Analyses
Microbial populations were determined by serial dilution (10–1 to 10–7) of fecal samples in anaerobic diluents before inoculation onto petri dishes of sterile agar, as described by Bryant and Burkey (1953). Total anaerobe and total aerobe agars were prepared according to Bryant and Robinson (1961) and Mackie et al. (1978). The selective medium for bifidobacteria (BIM-25) was prepared using reinforced clostridial agar (BBL Microbiology systems, Cockeyville, MD) according to Munoa and Pares (1988). Lactobacilli were grown on Rogosa SL agar (Difco Laboratories, Detroit, MI). Escherichia coli were grown on EMB agar (Difco Laboratories). Agar used to grow Clostridium perfringens was prepared according to the FDA Bacteriological Analytical Manual (1992). Samples for total aerobes, bifidobacteria, lactobacilli, and C. perfringens were diluted, inoculated, and incubated anaerobically (73% N2, 20% CO2, and 7% H2) at 37°C. Total aerobes and E. coli were incubated aerobically at 37°C. Plates were counted after 12 to 48 h of incubation, depending on the bacterial species. Colony forming units were defined as being distinct colonies measuring at least 1 mm in diameter.
In addition, microbial populations were measured by DNA extraction from fecal samples, followed by quantitative PCR (qPCR) and denaturing gradient gel electrophoresis (DGGE) techniques. Briefly, fecal DNA was extracted from freshly collected samples that had been stored at –80°C until analysis, using a QIAamp DNA stool mini kit (Qiagen, Valencia, CA) according to manufacturer’s instructions. Extracted DNA was quantified using a NanoDrop ND-1000 spectrophotometer (Nano-Drop Technologies, Wilmington, DE). Isolated DNA was amplified by PCR, using primers targeting variable region 3 (V3) of the 16S rDNA, according to Muyzer et al. (1996) and Simpson et al. (1999). Primers used to amplify DNA for DGGE were "eubacterial primer" 341F and 534R (E. coli 16S rDNA; Muyzer et al., 1996). Primer 341F prevents complete dissociation of double stranded DNA during PCR because of its 40-nucleotide GC clamp at the 5'-end. Deoxyribonucleic acid was amplified in a 100 µL volume containing 10 ng of extracted fecal DNA, 25 pmol of each primer, and 50 µL of Taq PCR master mix (Taq PCR Master Mix Kit, Qiagen), and sterile deionized water up to 100 µL. Touchdown cycling was used in the PCR amplification, which lowered the annealing temperature from 65°C by 1°C every second cycle until it reached 55°C; 10 more cycles then were completed with a 55°C annealing temperature. The resulting PCR product was approximately 200 bp in length, and the remaining single-stranded DNA was removed using mung bean nuclease (Stratagene, La Jolla, CA), as described by Simpson et al. (1999).
Separation of PCR products was achieved by parallel DGGE (Muyzer et al., 1993) on a BioRad D-Code System (BioRad). Briefly, DGGE was performed for V3-16S rDNA products (~200 bp) using gels containing 29 to 48% denaturant [100% denaturant:40% (vol/vol) formamide and 7 M urea]. Gels were cast on support film (FMC, Rockland, ME), and gradients were formed using a BioRad Gradient Former Model 475 (BioRad). A 0% denaturant, stacking gel was cast on top of the gradient gel to allow for loading of samples. Electrophoresis was performed at 50 V for 10 min followed by 3 h at 150 V. Bacterial DNA reference ladders were loaded next to each fifth sample lane to allow for standardization of the bands and gel curvature (Simpson et al., 1999). After electrophoresis, DNA was fixed on the gel by soaking in a 10% ethanol/0.5% acetic acid (vol/vol) solution for 2 h. Gels were silver-stained with 0.1% AgNO3 solution (wt/vol) and scanned using a BioRad Densitometer (BioRad). Within each animal, dice’s similarity coefficients between the banding patterns (Van der Gucht et al., 2001) were calculated on the treatment lanes compared with the lanes corresponding to the control lane to indicate how each treatment changed the microbial populations (Dsc = [2j/(a + b)] x 100, where a = the number of DGGE bands in lane 1, b = the number of DGGE bands in lane 2, j = the number of common DGGE bands, and Dsc = 100% if complete similarity is demonstrated).
Quantitative PCR was performed for the bifidobacteria, lactobacilli, and E. coli genera, as well as C. perfringens. Specific primers were used for bifidobacteria (Matsuki et al., 2002), lactobacilli (Collier et al., 2003), E. coli (Malinen et al., 2003), and C. perfringens (Wang et al., 1994). Amplification was performed according to DePlancke et al. (2002). Briefly, a 10-µL final volume contained 5 µL of 2x SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA), 15 pmol of the forward and reverse primers for the bacterium of interest, and 10 ng of extracted fecal DNA. Standard curves were obtained by harvesting pure cultures of the bacterium of interest in the log growth phase in triplicate, followed by serial dilution. Bacterial DNA was extracted from each dilution using a QIAamp DNA stool mini-kit and amplified with the fecal DNA to create triplicate standard curves using an ABI PRISM 7900HT Sequence Detection System (Applied Biosystems). Colony forming units in each dilution were determined by plating on specific agars; lactobacilli MRS (Difco) for lactobacilli, reinforced clostridial medium (bifidobacteria, C. perfringens), and Luria Bertani medium (E. coli). The calculated log cfu/mL of each serial dilution was plotted against the cycle threshold to create a linear equation to calculate cfu/g of dry feces.
Calculations
Dry matter (g/d), recovered as ileal effluent, was calculated by dividing the Cr intake (mg/d) by ileal Cr concentrations (mg of Cr/g of ileal effluent). Ileal nutrient flows were calculated by multiplying DM flow by the concentration of the nutrient in the ileal DM. Ileal nutrient digestibilities were calculated as nutrient intake (g/d) minus ileal nutrient flow (output, g/d), this value was then divided by nutrient intake (g/d). Similar calculations were performed on fecal samples to determine total tract nutrient digestibilities.
Statistical Analysis
Data for continuous variables were analyzed by the MIXED procedure (SAS Inst. Inc., Cary, NC) and data for discontinuous variables by the GLIMMIX procedure of SAS. The experimental design was a 5 x 5 Latin square. The statistical model included the random effects of animal and period and the fixed effect of treatment. All treatment least squares means were compared using preplanned contrasts that tested for linear, quadratic, and cubic effects of YCW supplementation, and comparing no YCW supplementation (control) to all levels of supplementation. A probability of P < 0.05 was accepted as being statistically significant, although effects with P-values between 0.06 and 0.10 were accepted as trends.
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RESULTS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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. The diet contained 29.6% CP and 21.4% AHF on a DM basis. Total dietary fiber concentration was 4.16%, lower than usual so that the full effect of the YCW supplement might be expressed.
The complete chemical composition of the YCW preparation (Safmannan) is presented in Table 2. The YCW contained 48.2% TDF that was mostly insoluble (91% of TDF was IDF), 17.9% CP, and 19.8% acid hydrolyzed fat. Analysis of the sugar composition yielded undetectable concentrations of fucose, arabinose, xylose, sucrose, and fructose as free sugars, whereas mannose made up approximately 89% of total free sugars. After hydrolysis of sugars to monosaccharides, only glucose and mannose were detected and the ratio of glucose to mannose was about 1.3:1. Ninety-one percent of the 94.5% of OM contained in Safmannan was accounted for by protein, fat, and carbohydrate components.
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. The animals were limit-fed to prevent undesirable BW gain. The amount of diet offered daily (280 g/d) was consumed virtually completely by all dogs and met the energy needs of the largest dog in this study.
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. Dry matter intake was not affected by YCW supplementation. Similarly, individual nutrient intakes (OM, CP, AHF, TDF, IDF, and SDF) were not affected by supplementation of YCW.
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Total tract DM digestibility responded cubically (P < 0.05) to YCW supplementation, as did total tract OM, CP, AHF, GE, and IDF digestibility. Compared with the control diet, supplementation of YCW tended (P = 0.09) to result in lower digestibility of IDF. Digestibility of the SDF fraction was not affected by YCW supplementation and was high (83.9 to 101.1%) across treatments. In contrast to ileal digestibility values, total tract digestibility values were lowest for the 0.25% YWC supplementation level.
Fecal scores ranged from 1.8 to 2.1 and were not affected by YCW supplementation. Fecal pH ranged from 6.0 to 6.3 and responded cubically (P < 0.05) to YCW supplementation. Fecal output per gram of feed intake responded quadratically (P < 0.05) to YCW supplementation, with the lowest fecal production at the 0.45% supplementation level.
Blood cell count, cell differential, and serum immunoglobulin concentrations are presented in Table 5. Total white blood cell count tended (P < 0.09) to decrease quadratically with YCW supplementation. Neutrophil and lymphocyte counts were not affected. Monocyte concentrations, however, decreased linearly (P < 0.02), whereas eosinophil concentrations tended (P < 0.07) to decrease quadratically with increasing YCW supplementation. Of the serum immunoglobulins, IgA responded cubically (P < 0.09) to YCW supplementation, but IgG and IgM concentrations were not affected. Ileal IgA concentrations tended (P = 0.09) to increase quadratically with YCW supplementation, with the greatest value (4.6 mg/g of DM) at the 0.25% YCW supplementation level.
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. According to the serial dilution and plating method, bifidobacteria, lactobacilli, total anaerobe, and total aerobe populations were not affected by YCW supplementation. Clostridium perfringens responded cubically (P < 0.09) to YCW supplementation, whereas E. coli populations decreased (P = 0.01) linearly with YCW supplementation.
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Analysis of total fecal microbial DNA using Dice’s similarity index based on DGGE results indicated an increasing quadratic (P < 0.05) effect of YCW supplementation. Compared with the control treatment, the 0.25% supplementation level had the greatest coefficient (85.5), indicating that total fecal microbial DNA present was more similar to the control treatment than was the case for the other supplementation levels (75.7 to 82.1).
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DISCUSSION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Ileal nutrient digestibility tended to be greater with YCW supplementation than without. This tendency is of biological significance as ileal digestibility was up to 11.2, 8.6, 11.6, and 7.4 percentage points different for DM, OM, CP, and GE, respectively, between the highest and the lowest treatment means. Increased ileal nutrient digestibility will likely limit bacterial fermentation in the large intestine due to decreased available fermentable substrate arriving at that site.
Yeast cell wall supplementation perhaps affects viscosity of digesta and, subsequently, ileal rate of passage. Ileal flow was as 30 g/d lower at the 0.25% level compared with the other treatments (data not shown). Nguyen et al. (1998) reported that the alkali-soluble glucans in a Saccharomyces cerevisiae preparation form a semisolid gel. Gel formation could lead to a slower rate of passage in the small intestine and a longer exposure time of the chyme to the enzyme-rich environment of the small intestine. However, it would be expected that greater YCW supplementation levels result in grater viscosities (Dikeman et al., 2006), thus decreasing rate of passage, which was not noted in this study. Nevertheless, supplemental YCW appears to affect digesta flow through the intestinal tract. Previous research (Strickling et al., 2000; Swanson et al., 2002) with YCW preparations in dogs reported no significant effects on ileal nutrient digestibility. Generally, digestibility coefficients obtained here are comparable to those reported by Strickling et al. (2000) and Swanson et al. (2002) when they supplemented ileal cannulated dogs with YCW products (DP607 and BioMos; Alltech Inc., Nicholasville, KY) at approximately 0.5% of the diet offered.
Total tract nutrient digestibilities were lowest at the 0.25% supplementation level. Across treatments, there was a cubic effect on total tract digestibility with increasing YCW supplementation for all nutrients analyzed, except for SDF. Mannanoligosaccharides from YCW may alter lactic acid-producing bacterial populations in the canine gut (Swanson et al., 2002; Grieshop et al., 2004). If bacterial populations are increased, apparent total tract CP digestibility may be decreased. Increased bacterial growth may lower apparent CP digestibility and, in turn, lower OM and DM digestibility, and possibly digestible energy. This latter effect is possible because the energy used for the synthesis of bacterial cells is lost to the animal but is recovered in feces.
The IDF and SDF fractions varied widely in digestibility as compared with other nutrients. The diet used in this study was low (4.16%) in TDF, and small amounts of endogenous compounds that may analyze as TDF impact the amount of TDF recovered in feces. It has been suggested that fermentable carbohydrates such as short-chain FOS increase intestinal mucin secretion as a response to short-chain fatty acid production by lactic acid-producing bacteria. A portion of mucin will analyze as TDF. The effect of FOS supplementation on mucin secretion has been demonstrated in rats (Bovee-Oudenhoven et al., 2003; Ten Bruggencate et al., 2003, 2004, 2005) at 3 to 6% dietary FOS, and in humans at 20 g/d of FOS supplementation (Ten Bruggencate et al., 2006). The mannan moiety in YCW is thought to increase lactic and short-chain fatty acid-producing bacteria similar to FOS, which could affect mucin secretion. In the current study, however, supplements were not purified mannanoligosaccharides, and the supplementation levels were below 1%. Yeast cell wall supplementation at less than 1% of the diet is not typically associated with effects on total tract digestibility in dogs (Strickling et al., 2000; Swanson et al., 2002), but Zentek et al. (2002) reported lowered total tract digestibilities at approximately 5% (of diet offered) YCW supplementation.
Fecal pH responded cubically to supplementation, with the lowest pH noted at the 0.45% supplementation level. This supplementation level, according to qPCR bacterial enumeration (discussed later), had the greatest concentrations of Lactobacillus spp. and Bifidobacterium spp., which could explain the lower pH because these bacteria produce lactic acid and short-chain fatty acids. Fecal output per gram of food intake (as-is basis) was lowest at the 0.45% supplementation level with 0.30 g of feces produced per gram of food intake. Because fecal DM concentrations were not different among treatments (data not shown), these results suggest that a quadratic relationship exists between YCW supplementation and stool bulk. If this observation can be confirmed in further research, it could be useful in the production of pet foods formulated to produce less fecal matter (e.g., for animals that are mainly housed indoors).
Total white blood cell counts tended to decrease quadratically with YCW supplementation, with the lowest numbers (10.6 x 103 cells/µL) at the 0.25% level. The white blood cell counts noted here are similar to those reported by Swanson et al. (2002), who used similar dogs, but supplemented them with a different YCW supplement (BioMos, Alltech Inc., Nicholasville, KY). White blood cell counts reported here are slightly greater than those reported in Beagles and Pointers fed a YCW (BioMos)-supplemented diet (Grieshop et al., 2004). Nevertheless, white blood cell counts reported herein are within physiological range for adult dogs.
Of the specific white blood cell populations, the monocyte count decreased linearly with increasing YCW supplementation, although all values are within normal ranges for dogs. Monocytes are responsible for the phagocytosis of pathogens at a site of infection (as macrophages) but also are involved with the antibody-mediated cellular cytotoxicity that kills host cells infected with pathogens. The linear decrease in monocyte counts noted in this study could suggest a decreased infectious load in the intestine. The mannan moiety found in YCW is able to prevent adherence to the intestinal wall of bacteria expressing type-1 fimbrae (Ofek et al., 1977; Neeser et al., 1986). Mannose is able to occupy man-nose-binding sites on the peptidoglycan layer protruding from the cell wall of probacteria and thus prevents those binding sites from latching on to mannose-containing epitopes of host mucosal cell ligands (Mann and Petri, 1995). As a result, fewer bacterial adhesions would lead to fewer translocated bacterial cells in the intestinal mucosal layer, thereby lowering the need for macrophages in this tissue. This decreased need for macrophages could be reflected in lower peripheral monocyte counts. Despite their importance in immune function, monocyte counts in excess of established physiological values (monocytosis) are generally regarded as a sign that the animal is in a pathogenic state. Lymphocyte counts were not altered in this study, but previous research with dogs reported both increased (Swanson et al., 2002) and decreased (Grieshop et al., 2004) lymphocyte counts.
Eosinophil counts tended to respond quadratically to YCW supplementation. Eosinophils play a role in allergic responses including food allergies (e.g., eosinophilic gastroenteritis; Kelly, 2000). Although patients with clinical eosinophilic gastroenteritis may have normal peripheral eosinophil counts (Talley et al., 1990), the trend in eosinophil counts with YWC supplementation deserves further attention because food hypersensitivity is an increasing problem in dogs as well as in humans. Increases in peripheral white blood cell count above normal levels would not be expected in immunologically unchallenged animals as used in this study. It has been suggested, however, that YCW does not necessarily alter cell counts, but may improve proliferative response of lymphocytes (Darroch et al., 1994) and increase cytokine production by macrophages (Adachi et al., 1994). This means that although immune status may not appear to be improved judging by cell counts, an actual immune response could be enhanced in a YCW-supplemented animal compared with a nonsupplemented animal.
Serum immunoglobulin concentrations were not altered by YCW supplementation with the exception of IgA, which tended to respond cubically with the lowest concentration (16.0 mg/dL) at the 0.25% supplementation. It is not clear why the 0.25% level resulted in 22 to 35% lower IgA concentrations compared with the other treatments and the control. Swanson et al. (2002) reported a 20% increase (P = 0.14) in serum IgA when dogs were supplemented with ~0.5% YCW, which is comparable to the 21% increase (compared with control) noted in this study with the 0.45% YCW supplementation. The 0.25% treatment had the greatest ileal IgA concentration, and was the peak in a quadratic trend (P < 0.09). Increased ileal IgA concentrations in dogs have been reported by Swanson et al. (2002), and also were noted in the cecum of rats (Kudoh et al., 1999). Increased mucosal IgA indicates a greater local resistance to antigen invasion because secretory IgA binds antigens, preventing them from passing mucosal membranes.
The full potential of YCW to affect the immune system may not become clear unless an immune challenge is presented. Therefore, an optimal administration level must be established because studies in rat and humans using FOS supplements suggest that the intestinal tract loses integrity and allows for increased translocation of pathogens (Salmonella typhimurium) in the mucosa (Bovee-Oudenhoven et al., 2003; Ten Bruggencate et al., 2003, 2004, 2005), possibly due to the increased production of short-chain fatty acids. If mannanoligosaccharides from YCW elicit a similar effect as has been noted with FOS, a balance between increased mucosal immune function and intestinal permeability must be sought.
Of the bacterial species quantified using serial dilution and plating, C. perfringens responded cubically to YCW supplementation, with the greatest counts at the 0.25% supplementation level. The high protein content of the diet may increase C. perfringens proliferation as this is a preferentially proteolytic species. Numerically, the enumerated C. perfringens follow the differences in ileal CP digestibility. Perhaps the digesta passing into the large intestine contained more accessible protein for bacteria as the hydrolytic degradation had progressed further than for the treatments with lower ileal CP digestibility. The linear decrease noted in E. coli with increased YCW supplementation is probably due to the ability of the mannose moiety to bind to type-1 fimbrae as they are expressed by E. coli and Salmonella spp. This binding prevents these bacteria from adhering to the intestinal wall and colonizing the intestine. These mechanisms are perhaps the cause of the trend indicating that YCW supplementation appears effective in lowering E. coli compared with the control treatment.
Bacterial counts as determined using qPCR showed a cubic trend for Lactobacillus spp. that was not detected by the plating method. The 0.25% treatment had the lowest lactobacilli counts, whereas plating detected that treatment as being second highest. The 2 treatments with the greatest lactobacilli counts according to qPCR were the lowest according to the plating method. Bifidobacterium spp. and C. perfringens counts according to qPCR did not agree completely with those obtained using plating, but in general, the overall results were similar, with no clear effect of YCW supplementation on these bacterial concentrations. The linear effect on E. coli observed with the plating technique was not detected using qPCR.
The somewhat different results in bacterial concentrations obtained with the 2 enumeration techniques used here could be due to the fundamental differences between the techniques. The serial dilution and plating technique measures live bacteria present in the fecal sample and if bacteria are affected in any way (e.g., strict anaerobes exposed to only a few ppm of oxygen), this could lead to a decrease in viable cells. Although every effort is taken to make agars specific to a certain bacterium or bacterial species, no guarantee exists that other bacteria (or yeasts or fungi) present in the intestine will not be able to grow on a certain specific agar. Moreover, even with swift and accurate sample handling, not all live bacteria present at time of defecation are still alive at the time of plating, thereby affecting the number of cfu detected in a sample. In contrast, qPCR detects the presence of DNA sequences (based on the specific primer) of specific bacteria or a genus of bacteria. Quantitative PCR will quantify intact DNA from dead cells and, subsequently, overestimate the number of live cells in a fecal sample. Nevertheless, qPCR may more accurately estimate the total number of bacterial cells in the intestinal tract. Substrates that are rapidly fermented once they enter the large bowel (e.g., FOS) may be completely used up in the proximal section of the large bowel. Bacteria that can use these types of substrates proliferate rapidly in the proximal large intestine, but may be starved in the distal large intestine, resulting in lower fecal live cell concentrations or less viable cells. Therefore, plating techniques may result in lower bacterial concentrations based on fecal analysis, whereas qPCR is able to account for, at least in part, the bacterial proliferation in the proximal large intestine, and find greater cfu counts.
The greater bifidobacteria concentrations measured here using the plating methodology as compared with qPCR could be due to the complexity of the agar used for this species in the plating technique. Specificity for bifidobacteria is accomplished by adding several antibiotics that may not completely prevent growth of other bacteria given the diversity of the intestinal microbial ecology. Serial dilution and plating methods are useful to demonstrate changes in microbial populations on a genus level in studies with prebiotic compounds; however, they are not well suited to identify changes in individual species (laborious and time-consuming). Molecular techniques such as qPCR are able to effectively detect changes in individual species, as long as the proper (validated) primer is used. With technology for molecular study of (intestinal) microbiota becoming more readily available at lower cost, limitations for (animal) nutrition research to utilize these techniques to their full extent are dissipating.
Total bacterial DNA as compared using Dice’s similarity coefficients on DGGE responded quadratically to YCW supplementation. All treatment coefficients are relative to the control treatment and the 0.25% supplementation level was most similar to the control. This result is interesting because according to bacterial enumeration using serial dilution and plating techniques, the 0.25% supplementation level appears to result in bacterial counts that are least similar to the control treatment. Bacterial enumeration using qPCR agrees better with DGGE results in terms of the counts of the bacterial species quantified in this study. It should be noted, however, that the enumerated bacterial species only represent a small fraction of the total microbiota in the intestinal tract. Changes noted with DGGE analysis could, therefore, be due to alterations in bacterial species other than the ones quantified here. In conclusion, supplementation with YCW appears to affect nutrient digestibility in dogs at the terminal ileum and in the large bowel. The noted differences in digestibility, however, are unlikely to impact the overall health status of the animal. Immunological indices were minimally affected by YCW supplementation in healthy dogs, but the full benefit of YCW supplementation may not be detectable unless the immune system is challenged during supplementation. Supplementation with YCW was able to slightly modulate bacterial concentrations, with a linear decrease in E. coli as the most striking effect. For most outcome variables measure here, the 0.25% supplementation concentration numerically had the highest or lowest mean. Therefore, YCW supplementation at approximately this level appears useful in possible further research with YCW supplementation in dogs. The different microbial enumeration methods did not completely agree in terms of number of cfu measured and trends detected, but this could be due to the fundamental differences between the 2 methods.
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2 Corresponding author: gcfahey{at}uiuc.edu
Received for publication February 5, 2007. Accepted for publication July 16, 2007.
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