Do Salivary Proline-Rich Proteins Counteract Dietary Hydrolyzable Tannin in Laboratory Rats?

Journal of Chemical Ecology, Vol. 30, No. 9, September 2004 (©C 2004)

DO SALIVARY PROLINE-RICH PROTEINS COUNTERACTDIETARY HYDROLYZABLE TANNIN IN

LABORATORY RATS?

MICHELE M. SKOPEC,1,∗ ANN E. HAGERMAN,2

and WILLIAM H. KARASOV1

1Department of Wildlife Ecology, University of WisconsinMadison, Wisconsin 53706, USA

2Department of Chemistry & Biochemistry, Miami UniversityOxford, Ohio 45056, USA

(Received December 11, 2003; accepted May 26, 2004)

Abstract—We hypothesized that dietary hydrolyzable tannins would not act asdigestibility reducing substances but would be excreted in the feces if the tanninwere ingested by rats producing salivary proline-rich proteins (PRPs). To testthat hypothesis we used two groups of Sprague–Dawley rats: tannin-naı̈ve ratsthat were secreting basal levels of salivary PRPs and tannin-habituated rats thatwere secreting elevated levels of PRPs. The animals were fed for 10–18 d ondiets containing 3% (w/w) purified hydrolyzable tannin [pentagalloyl glucose(PGG)] that was periodically spiked with chemically synthesized, radiolabeled1,2,3,4,6-penta-O-galloyl-[U-14C]-D-glucopyranose (1 µCi per gram diet). ThePGG-habituated rats excreted three times more of the consumed 14C in theirfeces than did the PGG-naı̈ve rats (11.4% for PGG-habituated rats vs. 3.5% forPGG-naı̈ve rats, P < 0.05). The addition of 3% PGG to the diet of the PGG-naı̈verats had no significant effect on apparent dry matter or nitrogen digestibility(P > 0.05). However, dry matter digestibility and nitrogen digestibility weresignificantly decreased by PGG in the diets of the PGG-habituated rats (7 and25%, P < 0.001, respectively). Production of PRPs increased the amount ofPGG excreted intact in the feces but at the cost of diminishing apparent drymatter and nitrogen digestibility.

Key Words—Tannin, pentagalloyl glucose, proline-rich protein, apparentdigestibility, laboratory rat.

∗ To whom correspondence should be addressed. E-mail: [email protected]

1679

0098-0331/04/0900-1679/0 C© 2004 Springer Science+Business Media, Inc.

1680 SKOPEC, HAGERMAN, AND KARASOV

INTRODUCTION

Tannins are polyphenolic natural products that readily bind protein under con-ditions that are present in the gastrointestinal tract of many species (Hagermanet al., 1998). The presence of tannins in a foodstuff may reduce palatability, limitdigestibility, and increase toxic load (Mole et al., 1990). There are two mainsubgroups of tannins, condensed and hydrolyzable tannins (Haslam, 1989). Con-densed tannins are polymers of flavan-3-ols that are linked via interflavan bondsand are not readily susceptible to degradation. Hydrolyzable tannins have a polyolcore (oftentimes glucose) esterifed to simple phenolic acids such as gallic acid.Hydrolyzable tannins are susceptible to acid, base, or enzyme-catalyzed hydroly-sis, yielding small phenolic acids, such as gallic acid and glucose. Hydrolysis inthe mammalian digestive tract (Murdiati et al., 1991; Bravo et al., 1994) releasesphenolic acids that may be locally toxic to gut microflora (Lowry et al., 1996) ormay be absorbed from the intestine and cause systemic toxicity within the animal(Niho et al., 2001). Even if the amount of phenolic acid absorbed from the intestineis below the toxic dose, a metabolic cost may be paid by the animal to detoxifyand excrete the absorbed phenolics (Thomas et al., 1988; Illius and Jessop, 1995).

The fundamental characteristic of tannins is their propensity to precipitateproteins, and it has long been hypothesized that tannins reduce nitrogen digestibil-ity by binding to dietary protein and digestive enzymes in the gut (Austin et al.,1989; Robbins et al., 1991; McArthur et al., 1995). This idea was challenged afterit was found that tannic acid—a mixture of hydrolyzable tannins and low molecu-lar weight phenolics (Hagerman et al., 1992)—reduces nutrient assimilation by adifferent mechanism than reducing digestibility (Bernays et al., 1989; Mole et al.,1990, 1993). However, because tannic acid is a mixture containing both tanninsand low molecular weight phenolics (Hagerman et al., 1992), it is unclear whetherthe reductions in nutrient assimilation are due to tannins in the tannic acid or dueto the toxicity of the low molecular weight phenolics.

When consuming tannins, some mammals secrete salivary proline-rich pro-teins (PRPs) that bind to the tannins with high affinity (Hagerman and Butler,1981; Mehansho et al., 1983, 1992). It has been proposed that the salivary PRPsbind to the tannins in the gastrointestinal tract, forming indigestible complexes.The complexed tannins may escape both enzymatic and microbial degradation andbe excreted in the feces, thus protecting the animal from both digestibility reduc-tion and toxicity (Austin et al., 1989). However, the lack of a suitable analyticalmethod to quantitate the amount of tannin in fecal samples has hindered tests ofthis hypothesis (Hagerman and Butler, 1989; Mole et al., 1993; Mueller-Harvey,2001).

Laboratory rats normally secrete only low levels of salivary PRPs but tannin-containing diets induce production and secretion of salivary PRPs (Mehanshoet al., 1983, 1992). Because PRP secretion is an inducible response in laboratory

PROLINE-RICH PROTEINS AND HYDROLYZABLE TANNIN CONSUMPTION 1681

rats, they make an ideal test species to determine the role of PRP secretion incounteracting dietary tannin because it is possible to evaluate the physiologicaleffects of low vs. high PRP production in not only the same species, but in thesame individuals. Also laboratory rats are reasonable models for the many speciesthat share their digestive strategy of little pregastric fermentation, digestion ofcell solubles primarily in the small intestine, followed by hindgut fermentationof some undigested residues. To take advantage of the inducibility of PRP secre-tion, we used two groups of rats: PGG-habituated rats, which ingested PGG for8 d pretrial in order to induce PRP production, and PGG-naı̈ve rats, which hadnever consumed PGG and were not secreting PRPs when the experiment started.We used a pure, homogeneous hydrolyzable tannin 1,2,3,4,6-penta-O-galloyl-D-glucopyranose [pentagalloyl glucose (PGG)], spiked with chemically synthesized,radiolabeled pentagalloyl glucose (14C-PGG) to directly investigate the fate andconsequences of ingested hydrolyzable tannins in laboratory rats.

METHODS AND MATERIALS

Animals and Housing. Ten 6-wk-old, male Sprague–Dawley rats(Harlan Teklad, Madison, WI) weighing 145–160 g were housed individually inplastic metabolic cages (Nalge Nunc International, Rochester, NY) that allowedcomplete separation of urine and feces. The rats were placed on a reverse lightcycle of 12D:12L with an ambient temperature of 25◦C and a relative humidityof 40%. All procedures described followed established guidelines for the care andhandling of laboratory animals and were approved by the Research Animal Re-source Center Animal Care Committee at the University of Wisconsin, Madison.

Diet. AIN-76A powdered diet was used as the basal diet (see Table 1 forcomposition; American Institute of Nitrition, 1977). PGG, the test tannin, was

TABLE 1. COMPOSITION OF DIETS

Ingredientsa Basal diet (g/kg) 3% PGG diet (g/kg)

Casein 200 200DL-Methionine 3 3Sucrose 500 500Corn starch 150 150Corn Oil 50 50Cellulose 50 20Pentagalloyl glucose 0 30Mineral Mix, AIN-76 35 35Vitamin Mix, AIN-76A 10 10Choline bitartrate 2 2

a All ingredients except pentagalloyl glucose were from TekladInc., Madison, WI.


added to the diet on a 3% per dry weight basis in place of cellulose (hereafterreferred to as the 3% PGG diet). The PGG was added daily to the diet beforefeeding to minimize oxidation.

Pentagalloyl Glucose. PGG was purified from commercially available tannicacid (Sigma-Aldrich, St. Louis, MO) as described by Hagerman et al. (1997). Theradiolabeled 1,2,3,4,6-penta-O-galloyl-[U-14C]-D-glucopyranose was chemicallysynthesized from [U-14C]-D-glucose (Perkin–Elmer Life Science, Boston, MA)and tri-O-benzylgallic acid (Sigma-Aldrich, St. Louis, MO) and had a radiopurityof at least 95% (Chen et al., 2003). The structure and purity of the natural andsynthetic PGG were established by HPLC, MS, and NMR (Chen et al., 2003)before the feeding trials started and after the trials were completed to confirmcompound integrity throughout the experiment. Both the unlabeled as well asradiolabeled PGG were stored at –80◦C when not in use.

Experimental Procedure. Rats that had been consuming the basal diet for10 d were randomly assigned to two groups of five rats each, the PGG-habituatedor PGG-naı̈ve group. For the first 8 d of the experiment, the five rats assigned to thePGG-habituated group consumed the 3% PGG diet, while the five rats assigned tothe PGG-naı̈ve group were maintained on the basal diet (Figure 1). Food intake andbody weight were measured daily. Feces uncontaminated by urine were collectedevery 24 hr and frozen.

After 8 d of habituation, the experimental period was initiated. During theexperimental period, all rats received the 3% PGG diet. Rats reduce their feedintake when they are initially introduced to a PGG-containing diet (Table 2), so inorder to ensure an equivalent PGG dose between the two groups we utilized a pair-feeding design during the experimental period. Each rat in the PGG-habituatedgroup was paired with a rat in the PGG-naı̈ve group and was fed only the amountof 3% PGG diet consumed by the PGG-naı̈ve rat with which it was paired. Ond 1, 3, 5, and 7 of the experimental period, the rats were fed a 5-g meal of the3% PGG diet with 1 µCi 14C-PGG per gram of food. In a preliminary study, themaximum residence time of 14C-PGG fed to either PGG-naı̈ve or PGG-habituatedrats was 48 hr, so the rats were dosed with the radiolabeled PGG every otherday. Rats were fed the 14C-PGG diet in shoebox cages instead of the metabolic

FIG. 1. Experimental timeline. Days highlighted in gray were used to determine the 3-daverages of food intake, weight gain, proline excretion, and apparent dry matter and nitrogendigestibility.


TABLE 2. FOOD INTAKE AND WEIGHT GAIN

Days Group Diet Food intakea (g/d) Weight gaina (g/d)

1a–3a PGG-habituated 3% PGG 7.6 ± 0.6d −2.1 ± 0.6c

1a–3a PGG-naı̈ve Basal 14.4 ± 0.7a 5.9 ± 0.1a

1–3 PGG-habituated 3% PGG 9.3 ± 0.4c 1.2 ± 0.7b

1–3 PGG-naı̈ve 3% PGG 10.2 ± 0.3c −0.1 ± 0.2c

8–10 PGG-habituated 3% PGG 12.9 ± 0.2b 2.8 ± 1.6b

8–10 PGG-naive 3% PGG 12.8 ± 0.3b 3.2 ± 0.4b

a Data given are the mean ± SE of the 3-d averages for each group of five rats. Numberswithin a column with different subscript letters are significantly different (P < 0.05) asdetermined by Tukey’s HSD.

cages to prevent contamination of the 24-hr fecal collections with spilled food.The 5-g meal containing the 14C-PGG was fed within 30 min of the start of thedark cycle, and rats were allowed to feed for 1 hr. Before the rats were placedback in their respective metabolic cages, they were thoroughly wiped with wetpaper towels to remove any diet particles present on their fur, tails, or whiskers.The nonradiolabeled 3% PGG diet was then provided to the rats in the metaboliccages. Food intake and body weight were measured daily, and feces were collectedas described above.

Fecal Analysis. Fecal samples were lyophilized and reweighed to determinedry matter content. Dried fecal samples were ground using a mortar and pestle andpartitioned for liquid scintillation counting (LSC), proline analysis, and nitrogenanalysis.

Radioactivity in the feces was used as a measure of the amount of 14C-PGG that was excreted in the feces. The labeled PGG was synthesized using[14C]-D-glucopyranose, and so hydrolysis of the 14C-PGG in the stomach or smallintestine of the rats would release [14C]-D-glucose that would be readily absorbedand metabolized, not excreted in the feces. However, chemical degradation studiesin artificial gastric or intestinal fluid demonstrate stability of PGG to hydrolysisfor at least 24 hr (Hagerman and Carando, unpublished data). Similarly, Bravoet al. (1994) found that tannic acid is a poor substrate for fermentation by ratceacal microflora, and that 75–95% of the initial tannic acid added was recoveredafter fermentation. Since PGG is one of the polyphenolic compounds presentin tannic acid it was assumed that the 14C-PGG that reached the rat’s hindgutwould not be readily fermented and, therefore, would be excreted intact in thefeces. For LSC, 0.02 g of feces were rehydrated in 8-ml scintillation vials with0.1 ml of deionized water. The rehydrated feces were solubilized by adding 1 mlof Soluene 350 (Sigma-Aldrich, St. Louis, MO) and placing the vials in a 50˚Coven until all particulate matter had been dissolved. Five microliter of scintillationfluid (Ecolume, ICN Costa Mesa, CA) containing 5% acetic acid (to decreasechemiluminesence) were added to the sample vials, and the samples were counted


to obtain dpm 14C with a Wallac WinSpectral 1414 liquid scintillation counter(Perkin–Elmer, Wellesley, MA).

Ground-dried feces were also analyzed for proline content to determine ifthe rats were excreting PRPs. Fecal proline excretion was used as an index for theinduction and secretion of salivary PRPs (Jansman et al., 1994) in order to avoidthe stress caused by saliva collection. Fecal samples were analyzed for prolineusing a 24-hr acid hydrolysis in 6 N HCl at 100◦C under nitrogen followed by acolorimetric ninhydrin reaction (Magne and Larher, 1992). Total proline (free andprotein) is detected by this method.

Ground-dried feces were also analyzed for nitrogen content to determine theeffect of PRPs on apparent nitrogen digestibility. The nitrogen content of the feceswas measured via a micro-Kjedhal digestion followed by a phenol-hypochloritecolorimetric determination of ammonia (Weatherburn, 1967).

Diet Analysis. The diets were analyzed for dry matter content by heatingthem in 50◦C drying oven. The nitrogen and proline contents of the diet wereanalyzed as described for fecal samples. Subsamples of the 14C-PGG diet fed eachday were analyzed via LSC.

Dry Matter Calculations. Apparent dry matter (DM) digestibility was calcu-lated on a 24 hr basis. DM digestibility = [(gram DM intake) – (gram dry weightof feces)]/[gram DM intake].

To calculate the amount of fecal DM excreted as PRPs it was assumed thatrat PRPs are 40% proline (Mehansho et al., 1983). PRP excretion = (gram prolinein feces)/0.40.

To calculate the mass of PGG in the feces of animals consuming 3% PGGin the diet it was assumed that 14C fecal excretion was proportional to total PGGexcretion. PGG excreted = [(gram food intake) × 0.03] × percent 14C in feces.

Nitrogen Calculations. Apparent nitrogen (N) digestibility was calculatedon a 24 hr basis. Apparent N digestibility = [(N intake) – (fecal N excretion)]/[Nintake]

To calculate the amount of fecal nitrogen excreted as PRPs it was assumedthat rat PRPs are 40% proline (Mehansho et al., 1983) and 16% nitrogen. Nexcreted as PRPs = [(gram proline in feces)/0.40] × 0.16.

Calculations of Reductions in Digestibility. Reductions in apparent N andDM digestibility were calculated as the difference in the 3-d average between d 1a–3a and 8–10. Reduction in digestibility = [(digestibility d 1a–3a) – (digestibilityd 8–10)]/[digestibility d 1a–3a].

14C-PGG Dose Calculation. The dosage of 14C-PGG consumed was cal-culated based on the concentration of 14C in the diet and food intake. 14C-PGGdose = [dpm 14C/gram diet] × [gram diet eaten].

Statistical Analysis. Food intake, weight gain, fecal proline, and apparentdry matter and nitrogen digestibility were analyzed as 3-d averages. Apparent


dry matter and nitrogen digestibility, % 14C excreted in feces and % reductionsin apparent dry matter and nitrogen digestibility were normalized by an arcsinesquare root transformation. Data were analyzed by repeated measure analysis ofvariance (ANOVA) with treatment and time as factors using SYSTAT (Wilkinsonand Coward, 2000). Differences between individual means were determined byTukey’s honestly significant difference (HSD). All data are expressed as mean ±SE unless stated otherwise and P values of 0.05 or less were used to determinesignificance.

RESULTS

Food Intake. During the habituation period, food intake by the PGG-consuming group diminished to almost half that of the PGG-naı̈ve group (comparebetween groups on d 1a–3a in Table 2, P < 0.05). During the experimental period,the rats in the PGG-habituated group were pair fed to rats in the PGG-naı̈ve groupto eliminate differences in food intake (compare between groups on d 1–3 and8–10 in Table 2). As the rats became habituated to the 3% PGG diet, their foodintake increased (compare within groups on d 1–3 and 8–10 in Table 2, P < 0.05).

Weight Gain. As expected based on their low food intake, the rats initiallylost weight on the 3% PGG diet (Table 2, compare between groups on d 1a–3a andwithin the tannin naı̈ve-group on d 1a–3a and 1–3, P < 0.05). As the rats becamehabituated to the dietary PGG, their weight gain increased but did not reach that ofthe animals not consuming PGG (Table 2, compare d 8–10 to PGG-naı̈ve d 1a–3a,P < 0.05).

Proline Excretion. The total proline defecated (mg/d) increased as the ratsbecame habituated to the 3% PGG diet (Figure 2) with maximal PRP excretion8–10 d after initiating the tannin diet (Mehansho et al., 1983). Rats that had beenon 3% PGG diets for 10–11 d had almost 10 times more proline in their fecesthan those on either the basal diet or on the 3% PGG diet for only 3 d (Figure 2,42.2 mg/d, d 1–3 vs. 4.6 mg/d, d 1a–3a in the PGG-habituated group, P < 0.05,and 32.5 mg/d, d 8–10 vs. 4.1 mg/d, d 1a–3a in the PGG-naı̈ve group, P < 0.05).The increase in fecal proline is coincident with the induction of salivary PRPs(Mehansho et al., 1983), suggesting that the proline is derived from PRPs. How-ever, other endogenous proteins such as mucins and sloughed cells, or bacterialproteins, could also contribute to fecal protein and consequently fecal proline.

14C in Feces. PGG habituation increased fecal excretion of 14C over threefold(Figure 3, d 2, PGG-habituated excreted 11.4% of ingested 14C vs. naı̈ve excretionof 3.5%, P < 0.05). Fecal excretion of 14C increased as the PGG-naı̈ve groupbecame habituated to PGG (Figure 3, d 2 vs. d 8, P < 0.05). The PGG-habituatedgroup also excreted significantly more 14C as the experiment progressed (Figure 3,


FIG. 2. Amount of proline defecated as rats habituated to PGG consumption. The PGG-habituated group was placed on the 3% PGG diet starting d 1a and had been eating the3% PGG diet for 9 d by d 1. The PGG-naı̈ve group was on the basal diet on d 1a–3a andwas placed on the 3% PGG diet on d 1. Data are represented as the mean ±SE. Bars withdifferent letters (a, b, c, d, and e) denote means significantly different from each other(P < 0.05) as determined by Tukey’s HSD.

FIG. 3. Percent of 14C consumed that was defecated within 48 hr in PGG-habituated ratsvs. PGG-naı̈ve rats. Rats were dosed with pentagalloyl [14C]-D-glucose on d 1, 3, 5, and 7and feces were analyzed on a 48 hr basis. The PGG-habituated group had been on the 3%PGG diet for 8 d prior to d 1 while the PGG-naı̈ve group was first exposed to the 3% PGGdiet on d 1. Data are represented as the mean ± SE. Bars with different letters (a, b, c, d,and e) denote means significantly different from each other (P < 0.05) as determined byTukey’s HSD.


11.4% on d 2 to 20.0% on d 8, P < 0.05). There were no significant differencesbetween the two groups after 8–10 d of tannin consumption (Figure 3, d 8 PGG-naı̈ve vs. d 2 PGG-habituated, P > 0.05).

Apparent Dry Matter and Nitrogen Digestibility. During the first 3 d on the3% PGG diet, there was no significant change in either apparent DM or N di-gestibility (Figure 4, compare between groups on d 1a–3a, P > 0.05). However,as the rats became habituated to the 3% PGG diet, both apparent DM and N di-gestibility significantly decreased, with N digestibility affected more strongly. Inthe PGG-habituated group, there was a 7% decrease in the apparent DM digestibil-ity from d 1a–3a to 8–10 (P < 0.001). For the PGG-naı̈ve group, the decreasein apparent DM digestibility was 5% from d 1a–3a to 8–10 (P < 0.05). ThePGG-habituated group had a 25% decrease in the apparent N digestibility fromd 1a–3a to 8–10 (P < 0.001), while the PGG-naı̈ve group had a 21% decrease inthe apparent N digestibility from d 1–3 to 8–10 (P < 0.05).

On the basis of fecal proline and 14C excretion, we calculated PRP andPGG fecal excretion. By our calculations, fecal PRP and PGG excretion accountsfor a maximum of 24% of the reduction in apparent DM digestibility, and fecalPRP excretion accounts for a maximum of 38% of the reduction in apparent Ndigestibility that we measured in PGG-habituated rats (Table 3).

DISCUSSION

The ability of tannins to bind proteins has led to the widespread notionthat dietary tannins reduce protein digestibility (Feeney, 1968; Robbins et al.,1987; Horigome et al., 1988; Hanley et al., 1992; Ortiz et al., 1994; Jansmanet al., 1995). However we found that when rats consume PGG, a hydrolyzabletannin, for a few days, there is no significant effect on apparent dry matter andnitrogen digestibility. Only after consuming PGG for 8 d or more was there asignificant decline in apparent dry matter and nitrogen digestibility (7 and 25%,respectively). We speculate that production of salivary PRPs, which in the ratare induced during the first 8–10 d of ingesting dietary tannin (Mehansho et al.,1983), could be responsible for observed reductions in digestibility seen duringconsumption of tannin-containing diets in some animals. Our idea contradicts thewidely held hypothesis that PRP production protects animals from digestibilityreduction (Mole et al., 1990; Robbins et al., 1991; McArthur et al., 1995).

We are not the first to report that the reduction in nitrogen digestibilityobserved when animals consume tannin is due to endogenous nitrogen loss andnot due to direct decreases in the digestibility of dietary nitrogen (Glick and Joslyn,1969; Mitjavila et al., 1977; Jansman et al., 1995). However, our study is the firstto show a reduction in apparent nitrogen digestibility within the same animalsacross three different dietary situations: tannin-free diets, 3% PGG diet fed to


FIG. 4. PGG habituation vs. apparent diet digestibility. (A) Shows the 3-d average of ap-parent dry matter digestibility. (B) Shows the 3-d average of apparent nitrogen digestibility.The PGG-habituated group was placed on the 3% PGG diet starting d 1a, and had beeneating the 3% PGG diet for 9 d by d 1. The PGG-naı̈ve group was on the basal diet ond 1a–3a and was placed on the 3% PGG diet on d 1. Data are represented as the mean ±SE. Bars with different letters (a, b, c, d, and e) denote means significantly different fromeach other (P < 0.05) as determined by Tukey’s HSD.

animals not producing PRP, and 3% PGG diet fed to animals that were producingPRP. We found that there was a significant reduction in apparent nitrogen anddry matter digestibility only when the rats were fed the 3% PGG diet for along enough period to induce PRP secretion. PRP secretion did not mitigate thepresumed digestibility reducing effects of PGG, because PGG itself had no effect


TABLE 3. THE PERCENT OF THE REDUCTION IN APPARENT

DIET DIGESTIBILITY EXPLAINED BY CALCULATED FECAL

EXCRETION OF PGG AND PRP

Apparent digestibility (mean ± SD)

Group Nitrogena Dry matterb

PGG-naı̈ve (n = 5) 21.6 ± 3.6 12.0 ± 3.1PGG-habituated (n = 5) 38.2 ± 3.5 23.7 ± 5.7

aBased on fecal PRP excretion.bBased on fecal PRP and PGG excretion.

on apparent digestibility. Instead PRP secretion may have increased the fecalloss of endogenous materials, thus reducing apparent digestibility, especially fornitrogen.

We calculated that fecal PRP excretion accounts for up to a maximum of38% of the decrease in apparent nitrogen digestibility while fecal PRP and PGGexcretion account for a maximum of only 24% of the decrease in apparent drymatter digestibility seen in PGG-habituated animals. Gastrointestinal mucous maycomprise at least part of the remaining nitrogen and dry matter losses in thePGG-habituated animals, as suggested by the elevated fecal glucosamine andsialic acid noted in rats on a 1% tannic acid diet (Mitjavila et al., 1977). Anincrease in mucous production and secretion can occur after the integrity of thedigestive epithelium is impaired (Deplancke and Gaskins, 2001). The mucousmay protect the gastrointestinal tract from oxidative damage incurred due to thepro-oxidative activities of some polyphenolics (Mitjavila et al., 1977; Ortiz et al.,1994; Long et al., 2000). The PAR-2 pathway controls both mucous production ingastrointestinal cells and secretion by the salivary glands (Kawabata et al., 2001),leading us to speculate that dietary tannin may use a single pathway to induce bothPRP and mucous secretion.

We believe that PRPs do play an important role in protecting animals fromthe effects of hydrolyzable tannins that are not related to digestibility. This hypoth-esis is supported by the increase in food consumption and weight gain observedafter habituation to dietary tannin compared to the low intakes and weight lossesrecorded for tannin-naı̈ve animals when first introduced to dietary tannin. Lowintakes and consequent weight loss may be due to the astringent taste of PGG-containing diets, which may be alleviated by complexation of the polyphenolsto PRPs (McArthur et al., 1995). By modifying astringent tastes, salivary PRPsmay provide an important mechanism for animals to ingest sufficient amounts ofotherwise unpalatable tannin-rich food and, thus, utilize these foods as part oftheir diets.

PRPs may also minimize toxicity of PGG by minimizing absorption of thepolyphenolic or its hydrolysis products. Our data did not indicate whether the


unexcreted PGG was absorbed or was hydrolyzed in the GI tract. Hydrolysis wouldrelease glucose that could be metabolized by either the rat or its gut microflora toproduce energy, but absorption of either gallic acid or intact PGG would imposethe metabolic costs of detoxification and excretion (Booth et al., 1959; Zong et al.,1999; Niho et al., 2001). That metabolic cost could be minimized by excretionof intact PRP–PGG complexes in the feces. Although only a maximum of 20%of the ingested PGG was excreted in the feces of PGG-habituated animals, this isa three- to fourfold increase in excretion over the PGG-naı̈ve animals, and couldreflect significant reduction in metabolic costs.

In laboratory rats, induction of PRPs conferred the benefit of a three- tofourfold increase in the amount of PGG excreted in the feces but it came withthe cost of increasing the loss of endogenous mass and nitrogen measured as adecline in apparent dry matter digestibility by up to 7% and apparent nitrogendigestibility by up to 25%. Unbound hydrolyzable tannins could cause tissuedamage within the intestinal tract (Mitjavila et al., 1977; Ortiz et al., 1994),target site toxicity post absorption (Murdiati et al., 1991; Niho et al., 2001), andcreate a metabolic cost for the detoxification and excretion of absorbed tannincomponents (Lindroth and Batzli, 1983; Thomas et al., 1988; Iason and Murray,1996). In contrast, a reduction in apparent nitrogen and dry matter digestibilitymay only be detrimental in situations where either nitrogen or overall energy islimiting. By increasing endogenous losses of nitrogen through the secretion ofPRPs, animals on a nitrogen-limited diet may alter their overall nitrogen balance,reallocating nitrogen from growth and reproduction to PRP production (Meyerand Richardson, 1993). Wild animals could counteract the loss of nitrogen byeating more, which may come with the cost of an increased risk of predation. Therats in this study were fed a diet containing 20% protein. The minimum proteinrequirement for laboratory rats for growth is 15% (NRC, 1995) and, therefore,nitrogen was not limiting. Thus, the cost of an increase in endogenous nitrogenloss would have been easily paid by the rats, well worth the increased protectionthe PRP may have provided from potential tissue damage and target site toxicitypost absorption.

Acknowledgments—We thank Kathleen Lacci, Sarah Pabien, and Bret Valentine for their adepttechnical assistance. Support for this research came from USDA (Hatch) WISO4322, NSF IBN-9723793, and IBN-0216709.

REFERENCES

AMERICAN INSTITUTE OF NUTRITION. 1977. Report of the American Institute of Nutrition ad hoccommittee on standards for nutritional studies. J. Nutr. 107:1340–1348.

AUSTIN, P. J., SUCHAR, L. A., ROBBINS, C. T., and HAGERMAN, A. E. 1989. Tannin-binding pro-teins in saliva of deer and their absence in saliva of sheep and cattle. J. Chem. Ecol. 15:1335–1347.


BERNAYS, E. A., DRIVER, G. C., and BILGENER, M. 1989. Herbivores and Plant Tannins, pp. 263–302,in M. Begon, A. H. Fitter, and E. D. Ford (eds.). Advances in Ecological Research, Vol. 19.Academic Press, New York.

BOOTH, A. N., MASRI, M. S., ROBBINS, D. J., EMERSON, O. H., JONES, F. T., and DEEDS, F.1959. The metabolic fate of gallic acid and related compounds. J. Biol. Chem. 234:3014–3016.

BRAVO, L., ABIA, R., EASTWOOD, M. A., and SAURA-CALIXTO, F. 1994. Degradation of polyphenols(catechin and tannic acid) in the rat intestinal tract. Effect on colonic fermentation and faecaloutput. Br. J. Nutr. 71:933–946.

CHEN, Y., HAGERMAN, A. E., and MINTO, R. E. 2003. Preparation of 1,2,3,4,6-penta-O-galloyl-[U-14C-D-glucopyranose. J. Labelled Compd. Radiopharm. 46:99–105.

DEPLANCKE, B. and GASKINS, H. R. 2001. Microbial modulation of innate defense: Goblet cells andthe intestinal mucus layer. Am. J. Clin. Nutr. 73:1131S–1141S.

FEENEY, P. 1968. Effect of oak leaf tannins on larval growth of the winter moth Operophtera brumata.J. Insect Physiol. 14:805–817.

GLICK, Z. and JOSLYN, M. A. 1969. Effect of tannic acid and related compounds on the absorption andutilization of proteins in the rat. J. Nutr. 100:516–520.

HAGERMAN, A., ZHAO, Y., and JOHANSON, Y. 1997. Methods for determination of condensed andhydrolysable tannins, pp. 209–222, in F. Shahadi (ed.). Phytochemicals in Foods. AmericanChemical Society, Washington, DC.

HAGERMAN, A. E. and BUTLER, L. G. 1981. The specificity of proanthocyanidin-protein interactions.J. Biol. Chem. 256:4494–4497.

HAGERMAN, A. E. and BUTLER, L. G. 1989. Choosing appropriate methods and standards for assayingtannin. J. Chem. Ecol. 15:1795–1810.

HAGERMAN, A. E., RICE, M. E., and RITCHARD, N. T. 1998. Mechanisms of protein precipitation fortwo tannins, pentagalloyl glucose and epicatechin16(4→8)catechin (procyanidin). J. Agric. FoodChem. 46:2590–2595.

HAGERMAN, A. E., ROBBINS, C. T., WEERASURIYA, Y., WILSON, T. C., and MCARTHUR, C. 1992.Tannin chemistry in relation to digestion. J. Range Manage. 45:57–62.

HANLEY, T. A., ROBBINS, C. T., HAGERMAN, A. E., and MCARTHUR, C. 1992. Predicting digestibleprotein and digestible dry matter in tannin-containing forages consumed by ruminants. Ecology73:537–541.

HASLAM, E. 1989. Plant Polyphenols. Cambridge University Press, Cambridge, UK.HORIGOME, T., KUMAR, R., and OKAMOTO, K. 1988. Effects of condensed tannins prepared from

leaves of fodder plants on digestive enzymes in vitro and in the intestine of rats. Br. J. Nutr.60:275–285.

IASON, G. R. and MURRAY, A. H. 1996. The energy costs of ingestion of naturally occurring nontanninplant phenolics by sheep. Physiol. Zool. 69:532–546.

ILLIUS, A. W. and JESSOP, N. S. 1995. Modeling metabolic costs of allelochemical ingestion by foragingherbivores. J. Chem. Ecol. 21:693–719.

JANSMAN, A. J., FROHLICH, A. A., and MARQUARDT, R. R. 1994. Production of proline-rich proteinsby the parotid glands of rats is enhanced by feeding diets containing tannins from faba beans(Vicia faba L.). J. Nutr. 124:249–258.

JANSMAN, A. J., VERSTEGEN, M. W., HUISMAN, J., and VAN DEN BERG, J. W. 1995. Effects of hulls offaba beans (Vicia faba L.) with a low or high content of condensed tannins on the apparent ilealand fecal digestibility of nutrients and the excretion of endogenous protein in ileal digesta andfeces of pigs. J. Anim. Sci. 73:118–127.

KAWABATA, A., KINOSHITA, M., NISHIKAWA, H., KURODA, R., NISHIDA, M., ARAKI, H., ARIZONO, N.,ODA, Y., and KAKEHI, K. 2001. The protease-activated receptor-2 agonist induces gastric mucussecretion and mucosal cytoprotection. J. Clin. Invest. 107:1443–1450.


LINDROTH, R. L. and BATZLI, G. O. 1983. Detoxification of some naturally occurring phenolics byprairie voles: A rapid assay of glucuronidation metabolism. Biochem. Syst. Ecol. 11:405–409.

LONG, L. H., CLEMENT, M. V., and HALLIWELL, B. 2000. Artifacts in cell culture: Rapid generation ofhydrogen peroxide on addition of (–)-epigallocatechin, (–)-epigallocatechin gallate, (+)-catechin,and quercetin to commonly used cell culture media. Biochem. Biophys. Res. Commun. 273:50–53.

LOWRY, J. B., MCSWEENEY, C. S., and PALMER, B. 1996. Changing perceptions of the effect of plantphenolics on nutrient supply in the ruminant. Aust. J. Agric. Res. 47:829–842.

MAGNE, C. and LARHER, F. 1992. High sugar content of extracts interferes with colorimetric determi-nation of amino acids and free proline. Anal. Biochem. 200:115–118.

MCARTHUR, C., SANSON, G. D., and BEAL, A. M. 1995. Salivary proline-rich proteins in mammals:Roles in oral homeostasis and counteracting dietary tannin. J. Chem. Ecol. 21:663–689.

MEHANSHO, H., ASQUITH, T. M., BUTLER, L. G., ROGLER, J., and CARLSON, D. M. 1992. Tannin-mediated induction of proline-rich protein synthesis. J. Agric. Food Chem. 40:93–97.

MEHANSHO, H., HAGERMAN, A., CLEMENTS, S., BUTLER, L., ROGLER, J., and CARLSON, D. M. 1983.Modulation of proline-rich protein biosynthesis in rat parotid glands by sorghums with high tanninlevels. Proc. Nat. Acad. Sci. U. S. A. 80:3948–3952.

MEYER, M. W. and RICHARDSON, C. 1993. The effects of chronic tannic acid intake on prairie vole(Microtus ochrogaster) reproduction. J. Chem. Ecol. 19:1577–1585.

MITJAVILA, S., LACOMBE, C., CARRERA, G., and DERACHE, R. 1977. Tannic acid and oxidized tannicacid on the functional state of rat intestinal epithelium. J. Nutr. 107:2113–2121.

MOLE, S., ROGLER, J., and BUTLER, L. 1993. Growth reduction by dietary tannins: Different effectsdue to different tannins. Biochem. Syst. Ecol. 21:667–677.

MOLE, S., ROGLER, J., MORELL, C. J., and BUTLER, L. G. 1990. Herbivore growth reduction by tannins:Use of Waldbauer ratio techniques and manipulation of salivary protein production to elucidatemechanisms of action. Biochem. Syst. Ecol. 18:183–197.

MUELLER-HARVEY, I. 2001. Analysis of hydrolysable tannins. Anim. Feed Sci. Technol. 91:3–20.MURDIATI, T. B., McSWEENEY, C. S., and LOWRY, J. B. 1991. Complexing of toxic hydrolysable

tannins of yellow-wood (Terminalia oblongata) and harendong (Clidemia hirta) with reactivesubstances: An approach to preventing toxicity. J. Appl. Toxicol. 11:333–338.

NATIONAL RESEARCH COUNCIL. 1995. Nutrient Requirements of Laboratory Animals, 4th edn. rev.National Academy Press, Washington, DC.

NIHO, N., SHIBUTANI, M., TAMURA, T., TOYODA, K., UNEYAMA, C., TAKAHASHI, N., and HIROSE, M.2001. Subchronic toxicity study of gallic acid by oral administration in F344 rats. Food Chem.Toxicol. 39:1063–1070.

ORTIZ, L. T., ALZUETA, C., TREVINO, J., and CASTANO, M. 1994. Effects of faba bean tannins on thegrowth and histological structure of the intestinal tract and liver of chicks and rats. Br. Poult. Sci.35:743–754.

ROBBINS, C. T., HAGERMAN, A. E., AUSTIN, P. J., MCARTHUR, C., and HANLEY, T. A. 1991. Variationin mammalian physiological responses to a condensed tannin and its ecological implications. J.Mammal. 72:480–486.

ROBBINS, C. T., HANLEY, T. A., and HAGERMAN, A. 1987. Role of tannins in defending plants againstruminants: Reduction in dry matter digestion? Ecology 68:1606–1615.

THOMAS, D. W., SAMSON, C., and BERGERON, J. 1988. Metabolic costs associated with the ingestionof plant phenolics by Microtus Pennsylvanicus. J. Mammal. 69:512–515.

WEATHERBURN, M. W. 1967. Phenol-hypochlorite reaction for determination of ammonia. Anal. Chem.39:971–974.

WILKINSON, L. and COWARD, M. 2000. SYSTAT 10 Statistics 1. SPSS, Chicago, IL.ZONG, L., INOUE, M., NOSE, M., KOJIMA, K., SAKAGUCHI, N., ISUZUGAWA, K., TAKEDA, T., and

OGIHARA, Y. 1999. Metabolic fate of gallic acid orally administered to rats. Biol. Pharm. Bull.22:326–329.

Do Salivary Proline-Rich Proteins Counteract Dietary Hydrolyzable Tannin in Laboratory Rats?

Documents

Transcript of Do Salivary Proline-Rich Proteins Counteract Dietary Hydrolyzable Tannin in Laboratory Rats?