To investigate the effects of parasitism and caloric restriction on morphology (body composition, organ mass) and physiology (resting metabolism, intestinal glucose transport capacity), we gave laboratory mice intestinal parasites (Heligmosomoides polygyrus, Nematoda), 30% caloric restriction, or both. Calorically restricted mice had smaller body mass, enhanced glucose transport capacity, and lower resting metabolism than ad libitum-fed mice. Parasitized mice maintained body mass, had diminished intestinal glucose transport capacity, and greater resting metabolism than unparasitized mice. Parasitized, calorically restricted mice had smaller organ masses than parasitized, ad libitum-fed mice and did not increase their glucose uptake rate as much as unparasitized, calorically restricted mice. There was a significant interaction between caloric restriction and parasite status for morphological variables but not for physiological variables. Knowing the types of phenotypic changes that occur with simultaneous parasitism and caloric restriction will provide insight into understanding human helminthiasis in food-restricted communities and also how wild animals cope with environments where parasitism and seasonal food restriction are common.
- intestinal parasites
- Mus musculus
- Heligmosomoides polygyrus
physiological and morphological effects of moderate caloric restriction vary with the duration and degree of restriction. In laboratory rodents, long-term caloric restriction (months to years) is beneficial in many respects. It increases small intestinal transport rates of some amino acids and monosaccharides (6), increases tissue glucose metabolism (51), reduces age-related decline of organ function (19, 35, 44), decreases the occurrence of carcinogenesis (24, 33), improves thermal stress tolerance (15), and increases longevity (28, 29). In contrast, short-term caloric restriction (days to weeks) induces body mass loss for the first 3–4 wk (6), decreases metabolic rate (31) and reproductive output (22), does not increase intestinal nutrient transport (6), impairs immune response (39), and can increase parasite pathology (5). Therefore, to realize the benefits of long-term caloric restriction, animals must first support the predominantly negative effects of short-term restriction.
Although effects of caloric restriction in laboratory rodents are well studied, relevance of this research to humans has been questioned (34, 47, 52). Recent studies show that responses of non-human primates to long-term caloric restriction are similar to responses of laboratory rodents (17, 25, 47, 50) and that humans show similar results as other vertebrates when given short-term caloric restriction (41, 49). However, with few exceptions (2, 3, 21, 49), studies of caloric restriction in humans involve obese subjects (48), and relevance of these studies to non-obese humans and humans with parasite infection is not well established. In many developing countries, obesity is uncommon, whereas parasite infection is relatively common, and reliable access to a balanced diet is often not available although diet composition affects how people respond to caloric restriction (40,46).
We examined the effects of short-term caloric restriction and sublethal parasite infection to determine how laboratory mice (Mus musculus) respond to simultaneous demands that individually produce mainly opposing responses. For example, food shortage can decrease resting metabolic rate (31) and body mass (6), whereas some parasite infections can increase resting metabolic rate and organ masses (23). When conflicting demands occur simultaneously, demands may interact with each other or demands may remain independent of each other. We hypothesized that both parasitism and short-term caloric restriction would affect body composition, intestinal transport capacity, and resting metabolism. As a result of this hypothesis, we predicted that mice given short-term caloric restriction would have smaller whole body mass (6), lower metabolic rate (31), and similar intestinal glucose transport as ad libitum-fed mice (6). We also predicted that parasitism by itself would have no effects on body mass, would increase resting metabolic rate, and decrease intestinal glucose transport (23). Finally, we predicted that when mice were simultaneously parasitized and food restricted they would have diminished changes in intestinal glucose transport, organ masses, and resting metabolic rate compared with unparasitized mice fed ad libitum, such that both morphological and physiological responses would interact.
MATERIALS AND METHODS
Our design had two main variables with two levels each: parasite infection (parasitized, unparasitized) and caloric restriction (caloric restricted, ad libitum fed). We used 40 individually housed virgin female Swiss-Webster mice (50–90 days old; Hilltop, Scottdale, PA), 10 per treatment group. Because parasites can affect males and females differently (56), we used only females for this study.
Parasite maintenance and mouse infection procedures.
H. polygyrus infective stage larvae (L3) were cultivated by collecting infected feces from nonexperimental mice. Feces were mixed with tap water, and the mixture was strained through cheesecloth and centrifuged. The resulting pellets were spread on filter paper and incubated for 9 days. There were 26 ± 1 L3/20 μl (mean ± SD) as determined by counting the number of L3 in three 20-μl samples with a compound microscope. All mice were anesthetized via intraperitoneal injection of pentobarbital sodium (50 mg/kg) mixed with sterile saline, and recovered from anesthesia in 2–3 h. While anesthetized, mice in the parasitized group were given ∼300 L3 suspended in tap water via an 18-gauge intragastric stainless steel curved feeding tube. Unparasitized mice were gavaged with an approximately equal volume of tap water only (n = 10) or were anesthetized but not gavaged (n = 10). There is no effect of the gavage procedure on any measured variable (23).
Caloric restriction and digestive efficiency measures.
For the first 6 days postcontrol gavage or infection (PI), mice were maintained at 14:10-h light-dark cycle, 23°C, and fed standard laboratory chow (Purina Mills) ad libitum. On day 7 PI, mice were switched to a high-carbohydrate diet (Custom Karasov Diet, ICN Nutritional Biochemicals: 55% sucrose, 15% casein, 7% cottonseed oil, 2% brewer's yeast, 4% salt mix, 1% vitamin, and 16% non-nutritive bulk; Ref. 10) necessary to determine maximal glucose transport capacity. On day 14 PI, when mature adult H. polygyrus occupy the small intestine (as determined by parasite eggs in mouse feces; Ref. 4), calorically restricted mice began the 10-day restriction treatment. For each caloric restriction day, mice were weighed and fed 3.48 ± 0.01 g custom Karasov diet (70% of ad libitum intake as similarly aged mice, determined from a previous experiment). For all mice, body mass, food intake rate (I, g/day), and fecal output rate (O, g/day) were measured on days 21–23 PI. Average percent dry matter digestibility (DMD) was calculated as [(I − O)/I] × 100 fordays 21–23 PI. Daily food intake was converted to digestible energy intake by multiplying food intake by dietary caloric content (15.1 kJ/g) by average DMD measured over 3 days.
Resting metabolic rate.
On day 23 PI, we measured resting metabolic rate (RMR) of nonpostabsorptive mice as oxygen consumption (V˙o 2, ml/min) using open-flow respirometry starting between 0800 and 0930. Dried (Drierite) air entered a 600-ml Plexiglas chamber housed in a dark cabinet (30 ± 1°C; within Mus thermoneutral zone; Ref. 18) at 650–700 ml/min from mass flow controllers. Air leaving the chamber was dried, scrubbed of CO2 (soda lime), and redried before entering an Applied Electrochemistry S-3A/II oxygen analyzer that was connected to a Macintosh computer. We measuredV˙o 2 [±0.002%; calculated usingequation 4a of Withers (53)] for 3 h, recorded every 5 s, and analyzed data with customized software (WartHog Systems). We sampled two mice simultaneously, whereby reference air was sampled for 2.5 min followed byV˙o 2 for 17.5 min. This cycle was repeated until each mouse was sampled for ∼2.5 h. We calculated RMR as the mean of the three lowest 2-min intervals ofV˙o 2.
Organ morphology and body fat measures.
On day 24 PI, we anesthetized mice between 0830 and 1130 by intraperitoneal injection of 0.07 ml pentobarbital sodium (65 mg/ml). We removed the small intestine (see below) and then removed the stomach, cecum, large intestine, heart, liver, spleen, kidneys, and lungs. Excess fat and connective tissue were removed from each organ and returned to the mouse carcass. For stomach, cecum, and large intestine, we weighed each organ with and without contents (flushed with mammalian Ringer solution). We measured dry mass of organs and the carcass after drying to a constant mass at 55–60°C for 2 and 14 days, respectively.
We ground the dried carcass and then extracted lipids using petroleum ether (Goldfische apparatus; Labconco). Extracted samples were redried, and the difference in mass before and after extraction was used to calculate percent fat. Percent fat from extraction was multiplied by the dry mass of the empty carcass to determine total amount of body fat (g) for each mouse. Percent whole body fat was grams of fat divided by whole body mass multiplied by 100. We measured fat content of all body organs, except the small intestine, together. Organ fat was removed by soaking organs in 10-ml aliquots of petroleum ether for six 24-h periods (pouring off ether at the end of 24 h and replacing it with fresh ether). After fat extraction, we redried organs, subtracted extracted organ dry mass from initial organ dry mass to determine percent of fat (g) in organs, and multiplied percent fat by total organ dry mass to calculate absolute fat content of organs. Percent organ fat was grams of fat divided by wet organ mass multiplied by 100. Total fat mass of each mouse was the sum of absolute body and organ fat mass. Therefore, lean mass was calculated as initial whole body mass minus total fat mass. Lean mass was partitioned into organ and nonorgan lean mass.
Small intestine morphology and glucose uptake measurements.
While the mouse was anesthetized, we rinsed the small intestine in situ with cold Ringer solution. It was then excised and placed in cold oxygenated Ringer solution (bubbled with 5% CO2-95% O2 at 2–3 l/min). We divided the small intestine into three regions of equal length (proximal, mid, and distal), measured the wet mass of each region after lightly blotting to remove adherent Ringer solution, and summed the three masses to determine total intestinal mass (corrected for mass of the parasites as described below). We separated mucosal/submucosal tissue (hereafter called “mucosa”) from muscularis/serosal tissue (hereafter called “serosa”) for two 1.5-cm sleeves per region (10). The dry mass-to-wet mass ratio was calculated for each sleeve, and we used the average of these ratios to calculate mucosal and serosal wet mass and dry mass of the entire small intestine (10, 16).
To determine glucose uptake by the small intestine, we measured the maximal transport velocity of the brush-border d-glucose transporter (SGLT1) in vitro using the everted sleeve technique (validated for this species by Refs. 10, 20). Briefly, we everted each region of the small intestine so that the mucosa faced outward. From each region we cut four 1.5-cm-long sleeves immediately adjacent to each other: two sleeves for measuring relative mucosal and serosal mass as described above and two sleeves for measuring glucose uptake. To measure carrier-mediated glucose uptake, we mounted everted sleeves on stainless steel rods and incubated them for 2 min in 36°C Ringer solution containing 50 mMd-glucose and trace amounts ofd-[14C]glucose. The incubating solution also contained trace amounts of l-[3H]glucose that was used to correct for glucose in the adherent mucosal fluid and for passive uptake of d-glucose. The amount of isotope taken up by the tissue was measured using liquid scintillation (LS 6500 scintillation system, Beckman) to determine glucose uptake rate of each sleeve (mmol · day−1 · g wet mucosal tissue−1). We then calculated the average uptake rate of the two sleeves from each region. The glucose uptake capacity for each region of the small intestine was calculated by multiplying the average glucose uptake rate by the wet mucosal mass (g) of the region. The products of each region were summed to determine glucose uptake capacity for the entire small intestine.
Parasite intensity and mass.
During rinsing of the small intestine, some adult H. polygyrus were flushed from the intestine and collected in a petri dish. After the small intestine was everted and sleeves were removed for mucosal scraping and glucose uptake measurements, the remaining small intestine tissue was added to the petri dish. Parasites were removed from the mucosal scraped sleeves and placed in the petri dish before the sleeves were weighted and mucosa was scraped away. Parasites were not removed from sleeves used for the glucose uptake experiment so as not to damage the tissue before glucose uptake measurements. Rather, the number of parasites counted from the mucosal scraped sleeves was doubled (assuming the number of worms on the sleeves used in the glucose uptake experiment was the same as the number of worms on the adjacent mucosal scraped sleeves). Therefore, final infection intensity equaled the number of parasites in the petri dish that were either flushed from the intestine or were attached to unused tissue plus two times the number of worms from the mucosal scraped sleeves. We determined the wet mass of H. polygyrus by measuring four samples of 50 H. polygyrus (approximately equal numbers of male and female worms) that had been lightly blotted on filter paper collected from nonexperimental mice. Total parasite wet mass was subtracted from small intestine wet mass before calculation of small intestine dry mass used in analyses. When we examined each intestinal region separately, parasite mass was subtracted only from mass of the proximal region, because adult parasites only occupy this portion of the small intestine (1).
Our data consist of two independent (parasite infection, caloric restriction) and numerous dependent variables (food intake, DMD, body mass, small intestine morphological variables, organ masses, body fat, glucose uptake rate and capacity, rate of body mass loss for calorically restricted mice, and RMR). We first used a multivariate ANOVA (MANOVA) that tested for significant differences between treatments for all dependent variables together. Because this MANOVA was significant (F = 7.48, P < 0.0001), we used independent ANOVAs to determine which treatments and which dependent variables were statistically significant. After we examined effects of decreased body mass associated with caloric restriction, we regressed each dependent variable against body mass, to remove body mass effects, and used residuals of significant regressions in further analyses. For RMR data, because calorically restricted mice had smaller body mass than ad libitum-fed mice, we used analysis of covariance (ANCOVA) with whole body mass as a covariate in our analysis. Also, because parasitized mice had greater lean body mass than unparasitized mice and ad libitum-fed mice had a greater lean mass than caloric-restricted mice, we also used ANCOVA with lean mass as the covariate for RMR data. Repeated-measures ANOVA (Wilk's Lambda) was used to examine effects of different small intestine regions on glucose transport rate and capacity, on masses of each region, and for changes in body mass during the 10-day restriction period. Infection intensity of parasitized ad libitum-fed and calorically restricted mice was tested with a t-test. Throughout, we used P< 0.05 as the level of significance, and all data are presented as the mean ± SE unless otherwise stated.
Infection intensity and parasite mass.
All mice gavaged with L3 developed mature infections, and no control mice became infected. Infection intensity was similar for ad libitum-fed and calorically restricted mice (ad libitum: 191 ± 11, caloric restricted: 206 ± 8; P > 0.05). Average wet mass of 50 H. polygyrus was 0.0123 ± 0.0012 g (mean ± SD) so the wet mass of individual H. polygyrus for each infected mouse for this study was calculated as the infection intensity times 0.000246 g. Parasite wet mass was ∼1.5% of total small intestine wet mass and 3.0% of the proximal region wet mass for both calorically restricted and ad libitum-fed mice.
Body mass and composition.
Body mass was 22% less for calorically restricted than ad libitum-fed mice (F 1,36 = 113.8, P < 0.0001), but parasite infection had no effect on body mass (Fig.1). Absolute lean mass was 12% greater in parasitized than unparasitized mice (F 1,36 = 23.6, P < 0.0001) but was 13% less for calorically restricted than ad libitum-fed mice (F 1,36 = 21.3, P < 0.0001), producing a significant interaction between treatments (F 1,36 = 4.9, P = 0.033). Total fat mass and percentage of body fat were ∼20% less for parasitized than unparasitized mice (fat mass:F 1,36 = 7.2, P = 0.011; percent fat: F 1,36 = 6.7, P= 0.014). Total fat mass also was less for calorically restricted than ad libitum-fed mice (fat mass: F 1,36 = 9.7,P = 0.004, 24% less), but percent fat did not differ.
Organ mass and composition.
Total organ fat was <3% of total body fat mass for all mice (ad libitum, parasitized: 2.3 ± 0.1%; ad libitum, unparasitized: 2.1 ± 0.2%; calorically restricted, parasitized: 2.3 ± 0.1%, calorically restricted, unparasitized: 2.5 ± 0.1%), and the contribution of organ fat mass to total body fat mass did not differ with either treatment. When comparing only organs, absolute organ fat mass was 15% less for calorically restricted than ad libitum-fed mice (F 1,36 = 5.9,P = 0.02) but did not differ with parasite treatment. Percent organ fat did not differ with parasite or caloric treatment (Fig. 1). Lean organ mass was 27% less for calorically restricted than ad libitum-fed mice (F 1,36 = 63.4,P < 0.0001) but was 17% greater in parasitized than unparasitized mice (F 1,36 = 35.4,P < 0.0001).
Because we did not extract fat of each organ separately, we used data of organ dry masses (including fat content) to examine effects of each treatment on total organ mass. Parasitized mice had larger cecae (by 30%; F 1,36 = 10.9, P = 0.002), lungs (by 5%; F 1,36 = 5.5,P = 0.025), and spleens (by 45%;F 1,36 = 95.3, P < 0.0001) than unparasitized mice (Table 1). Calorically restricted mice had smaller cecae (by 19%;F 1,36 = 12.4, P = 0.001), hearts (by 18%; F 1,36 = 62.7,P < 0.0001), kidneys (by 8%;F 1,36 = 9.7, P = 0.004), large intestines (by 8%; F 1,36 = 4.6,P = 0.038), livers (by 35%;F 1,36 = 98.0, P < 0.0001), lungs (by 12%; F 1,36 = 27.8,P < 0.0001), spleens (by 30%;F 1,36 = 28.5, P < 0.0001), and stomachs (by 12%; F 1,36 = 13.9,P = 0.001) than ad libitum-fed mice (Table 1). There were significant interactions between treatments for cecae (F 1,36 = 10.8, P = 0.002), hearts (F 1,36 = 8.6, P = 0.006), large intestines (F 1,36 = 15.0,P < 0.0001), and spleens (F 1,36 = 5.5, P = 0.024); in each case, the smaller organ masses of calorically restricted mice were more pronounced or only occurred in parasitized compared with unparasitized mice. After adjusting for body mass (because calorically restricted mice had smaller body mass than ad libitum-fed mice) there were no effects of caloric restriction on any organ mass but significant interactions between caloric restriction and parasitism remained for cecae (F 1,36 = 10.0,P = 0.003), large intestines (F 1,36 = 15.0, P < 0.0001), and spleens (F 1,36 = 4.4,P = 0.043).
Parasitized mice had an 8% longer small intestine than unparasitized mice (F 1,36 = 22.2, P < 0.0001), but small intestine length did not differ between calorically restricted and ad libitum-fed mice (Fig.2). Because qualitative results are the same for absolute small intestine dry mass and dry mass per unit length of small intestine (P > 0.05), we report only absolute dry mass results for all small intestine variables. Small intestine dry mass was 38% greater for parasitized than unparasitized mice (F 1,36 = 150.5, P < 0.0001) but was 13% less for calorically restricted than ad libitum-fed mice (F 1,36 = 13.9,P = 0.001; Fig. 2). A significant interaction between treatments showed that the increased small intestine mass of parasitized mice was not as great for calorically restricted mice as it was for ad libitum-fed mice (F 1,36 = 4.5,P = 0.041).
Calorically restricted mice had mucosal and serosal dry masses that were less than those of ad libitum-fed mice (F 1,36 = 9.4, P = 0.004 andF 1,36 = 7.5, P = 0.009, respectively), but parasitized mice had more mucosal and serosal tissue than unparasitized mice (F 1,36 = 51.4,P < 0.0001 and F 1,36 = 203.5, P < 0.0001, respectively; Fig. 2). There was a greater increase in serosal dry mass for parasitized mice when they were fed ad libitum than when calorically restricted (F 1,36 = 5.0, P = 0.032). Because parasitized mice had a disproportionate increase in serosal compared with mucosal tissue, the percent mucosa was less in parasitized than unparasitized mice. We also found that both serosal and mucosal dry masses varied among intestinal regions (F 2,35 = 94.0, P < 0.0001 and F 2,35 = 103.2, P < 0.0001, respectively; Fig. 3) and that serosal and mucosal masses decreased from proximal to mid to distal regions. Regardless of caloric treatment, mass differences among regions for unparasitized mice were due primarily to changes in mucosal mass, whereas mass differences among regions for parasitized mice were due to changes in both serosal and mucosal masses (Fig. 3). There was a significant interaction between parasite treatment and intestinal region for serosal dry mass (F 2,35 = 90.0,P < 0.0001), because serosal dry mass of parasitized mice was greater in the proximal region than the mid and distal regions regardless of caloric restriction treatment. Calorically restricted mice had smaller mucosal and serosal dry masses than ad libitum-fed mice for the proximal (F 1,36 = 4.99,P = 0.032 and F 1,36 = 6.03,P = 0.019, respectively) and mid (F 1,36 = 6.54, P = 0.015 and F 1,36 = 5.88, P = 0.02, respectively) regions, but had similar masses in the distal region (Fig. 3).
Because lean body mass increased with parasite infection but decreased with caloric restriction and because whole body mass decreased with caloric restriction, we examined the relationship between both mass variables and RMR. The relationship between body mass (whole and lean) and RMR was the same for all treatment groups, and there was a significant relationship when we regressed whole or lean body mass with RMR (r 2 = 0.33, P < 0.0001 and r 2 = 0.68, P < 0.0001, respectively). RMR, corrected for whole body mass (ANCOVA), was greater for parasitized mice than unparasitized mice (F 1,35 = 7.1, P = 0.011) by 9% (least square adjusted mean ± SE, kJ/day: ad libitum, parasitized = 25.1 ± 1.0; ad libitum, unparasitized = 23.0 ± 0.9; calorically restricted, parasitized = 21.7 ± 1.1; calorically restricted, unparasitized = 19.7 ± 0.9), and RMR was 14% less for calorically restricted mice than ad libitum-fed mice (F 1,35 = 5.2, P = 0.029). When lean body mass was used as a covariate, there was no effect of parasite infection, but calorically restricted mice had a 30% lower RMR than ad libitum-fed mice (F 1,35 = 46.1,P = 0.0001; least square adjusted mean ± SE, kJ/day: ad libitum, parasitized = 27.2 ± 1.3; ad libitum, unparasitized = 25.3 ± 0.9; calorically restricted, parasitized = 18.8 ± 0.9, calorically restricted, unparasitized = 18.1 ± 1.1).
Mass loss for calorically restricted mice and digestive efficiency.
In ad libitum-fed mice, food intake was not different between parasitized and unparasitized mice and, after accounting for orts lost in bedding, calorically restricted mice actually ate 68% less than ad libitum-fed mice. Regardless of caloric or parasite treatment, DMD was 78.8 ± 0.2%. We regressed whole body mass against day of caloric restriction and found that average total mass loss during 10 days of restriction was 16 ± 2% for parasitized mice (r 2 = 0.40, P < 0.0001) and 14 ± 1% for unparasitized mice (r 2 = 0.37, P < 0.0001). There were no differences between parasitized and unparasitized mice for rate of mass loss (g/day) or total mass loss (g) during caloric restriction.
Qualitative results were the same for intestinal glucose uptake rate normalized either to dry or wet mucosal mass; we present data using wet mucosal mass. Total glucose uptake capacity (mmol/day) summed for the entire small intestine was 28% less for parasitized than unparasitized mice (F 1,36 = 42.1, P < 0.0001) and was 14% greater for calorically restricted than ad libitum-fed mice (F 1,36 = 7.0,P < 0.012; Fig.4 A). Similarly, glucose uptake rate (mmol · g−1 · day−1) was 37% lower for parasitized than unparasitized mice (F 1,36 = 19.2, P < 0.0001) and 25% greater for calorically restricted than ad libitum-fed mice (F 1,36 = 15.0, P < 0.0001; Fig. 4 B).
We next examined each region separately to determine if parasites had a localized effect in the proximal region where they occurred. There was a significant difference in glucose uptake capacity among regions (F 2,35 = 105.2, P < 0.0001; Fig. 4 A). Parasitized mice had lower uptake capacity in the proximal (F 1,36 = 68.3,P < 0.0001) and mid (F 1,36= 4.7, P = 0.037) regions but greater uptake capacity in the distal region (F 1,36 = 4.5,P = 0.041) than unparasitized mice. There was a significant interaction between parasite treatment and intestinal region (F 2,35 = 33.4, P < 0.0001), because glucose uptake capacity of parasitized mice decreased more in the proximal than the mid region and increased in the distal region compared with unparasitized mice. Glucose uptake capacity of the distal region was greater in calorically restricted than ad libitum-fed mice (F 1,36 = 6.8, P = 0.013) but did not differ in other regions (Fig. 4 A).
Glucose uptake rate also varied among intestinal regions (F 2,35 = 8.2, P = 0.001; Fig. 4 B). There was a significant interaction between parasite treatment and intestinal region (F 2,35 = 8.7, P = 0.001) because parasitized mice had a lower uptake rate in the proximal and mid regions (F 1,36 = 15.9,P < 0.0001 and F 1,36 = 17.3, P < 0.0001, respectively), but not in the distal region, compared with unparasitized mice. Glucose uptake rate was greater for all regions of the small intestine in calorically restricted than ad libitum-fed mice (proximal:F 1,36 = 5.5, P = 0.024; mid: F 1,36 = 4.4, P = 0.043; distal: F 1,36 = 7.4,P = 0.01; Fig. 4 B).
Calorically restricted mice had lower fat and lean components of both organs and nonorgan tissue, resulting in decreased whole body mass. Parasitism resulted in no overall changes in body mass, because decreased body fat mass was compensated for with increased lean mass of vital organs. Thus, as predicted, caloric restriction and parasite infection had opposing and interacting effects on body mass and composition.
Parasitism resulted in larger spleen, lung, small intestine, and cecum dry masses. Although we did not measure passage rates of digesta or cecal fermentation rates, we speculate that nutrients spilled from the small intestine (which had a lowered glucose uptake capacity) into the cecum and were either retained longer or fermented in the cecum and therefore may be responsible for increased cecum masses of parasitized mice. Increased spleen mass of parasitized mice results, at least in part, from the immune response of mice to H. polygyrus(26, 30). Parasitized mice also had increased resting metabolic rates compared with unparasitized mice, which may have resulted in larger lung masses via a greater use of the cardiovascular system. Calorically restricted mice had smaller body masses and thus smaller heart, lung, spleen, liver, kidney, stomach, small and large intestines, and cecal masses than ad libitum-fed mice. There were no additional effects of caloric restriction on organ masses after accounting for body mass.
As predicted, RMR increased as a result of parasitism but decreased as a result of caloric restriction. The lower RMR of calorically restricted mice may have resulted from a lower specific dynamic effect (cost of digestion), because calorically restricted mice ate less food and had 87% less food in their stomachs than ad libitum-fed mice (stomach content mass at dissection: calorically restricted = 0.091 ± 0.014 g, ad libitum fed = 0.716 ± 0.107 g). So, although calorically restricted mice were not postabsorptive, digestion and assimilation processes of calorically restricted mice may have produced less heat than those of ad libitum-fed mice. Although we did not measure cost of digestion directly, we previously found that this strain of laboratory mice, when truly postabsorptive, had only a 4% lower RMR than ad libitum-fed mice (Hammond, unpublished data). We found a 14–30% effect of caloric restriction on RMR, however, so effects of caloric restriction were more than simply a result of lower costs of digestion. Mice may respond similarly to rats that had decreased body temperature and thermal conductance (36), two factors that can affect RMR, when given ∼50% caloric restriction. In a separate experiment, we found that calorically restricted laboratory mice have a 0.2 ± 0.06°C lower body temperature than ad libitum-fed mice (Kristan and Hammond, unpublished data), which, if conductance remains unchanged, may slightly lower RMR. The relationship between metabolism and caloric restriction is unclear, although decreased metabolism associated with short-term food restriction or deprivation may preserve body mass and prolong survival (31) during a time when nutritional resources are unpredictable. However, others have found the decreased metabolism with caloric restriction only occurs for the first few weeks of restriction and thereafter mass-specific metabolism is similar for ad libitum-fed and food-restricted rats (28), placing into question the relevance of short-term changes in metabolism to survival.
As predicted, glucose uptake rate and summed glucose uptake capacity of the entire small intestine was lower as a result of parasitism, regardless of caloric treatment. The reduction in glucose uptake capacity occurred in the proximal and mid small intestine regions, whereas capacity increased in the distal region primarily due to increases in mucosal mass of the distal region. Similarly, the lower rate of glucose transport in parasitized mice occurred in both the proximal and mid regions, but not the distal region. Decreased glucose transport capacity of parasitized mice may be related to increased mucosal cell-turnover rates as seen in nematode infections of rats (43), leaving more immature and undifferentiated intestinal cells that do not function in nutrient absorption (54). Because parasitized mice had diminished glucose transport in the proximal region compared with unparasitized mice, the nutrient density of ingesta that reached the mid and distal regions may be greater than usual and could thereby induce changes in nutrient transport at these regions (11, 12, 32).
Similar to our findings for glucose transport, Zarling and Mobarhan (55) found that sucrase activity increased in food-restricted laboratory rats. Rats responded to caloric restriction with a decrease in mucosal protein resulting from decreased cell height, which did not decrease disaccharidase activity (55) or nutrient absorption in the small intestine. Although we found increased rate of glucose transport for all intestinal regions in mice, rats had increased glucose transport only in the proximal and mid regions (55). In contrast to our findings with young mice, short-term caloric restriction in older mice did not alter glucose transport by the small intestine (6). Therefore, age or age-related morphological and physiological changes can affect the functional response of the small intestine to short-term caloric restriction. Despite opposing responses of glucose transport by our mice when parasitized vs. calorically restricted, there was no significant interaction between these physiological demands.
Simultaneous demands with opposing responses.
In general, the morphological responses to parasite infection that we observed were diminished in the presence of caloric restriction. Parasitized mice given caloric restriction had diminished or no increases in organ lean mass compared with parasitized mice that were fed ad libitum, regardless of whether body mass effects were removed. Therefore, when parasitism and caloric restriction were given simultaneously, they interacted with each other for morphological variables. This was true for spleen mass, for example. If the immune response, as indicated in part by splenic hypertrophy (26,30), was compromised during caloric restriction, individuals might be at greater risk for other pathologies. Shi et al. (38) showed that caloric restriction in the host can affect parasite development, survival, and fecundity when caloric restriction is given before infection, and therefore the nutritional state of the host can affect parasite infections. However, we gave caloric restriction after the immune-inducing phase of the H. polygyrus life cycle was complete (30). This may explain the similar infection intensity, and presumably host immune response, for both calorically restricted and ad libitum-fed mice.
Interestingly, unlike for morphological responses (organ masses and body composition), caloric restriction and parasitism were independent for physiological responses (metabolism and glucose transport). For example, contrary to our prediction, changes in RMR associated with parasite infection and caloric restriction remained independent when we presented both demands simultaneously. Greater RMR associated with parasite infection was partially due to greater lean mass (metabolically active tissue), whereas decreased RMR associated with caloric restriction was probably due to decreased lean mass, differences in absorptive state, and possibly lower body temperature. Although caloric restriction lowered absolute RMR, the relative increase in RMR associated with parasitism was similar for both ad libitum-fed and calorically restricted mice. So, although we have previously shown that simultaneous cold exposure and parasite infection did not interact for morphological and physiological responses (23), the results of our present study suggest that it is not simply the presence of multiple demands but the types, and likely intensities, of demands that are important determinants of how mice respond and whether demands interact.
Relevance of rodent models to humans.
There is evidence that humans respond to both helminth infections (9) and caloric restriction (41, 49) using similar morphological or physiological pathways or outcomes as laboratory rodents. Therefore, studying the effects of simultaneous parasitism and caloric restriction in rodents may help address the ongoing problems of human helminthiasis that often cooccur with chronic poverty (8) and nutritional deficiency. For example, nematodes, along with other helminths, infect an alarmingly large number of people, especially in developing countries (7, 8,42). Similar to some studies of laboratory rodents (43), people show changes in gastrointestinal morphology with nematode infection (9) that can then lead to impaired nutrition (42), especially in children (45). Childhood infections can correlate with parasite prevalence in adults (14) and may potentially affect work productivity during adulthood (13, 37). When human helminthiasis cooccurs with food restriction, responses may be different from when these demands occur alone. Therefore, understanding how demands of parasite infection and restricted caloric intake affect the morphology and physiology of rodents may provide insight into the biological basis of the perpetual and costly effects of human helminthiasis (27) in poor, undernourished communities. Future studies to determine how demands interact or remain independent under differing combinations are needed.
M. Chappell, E. Platzer, B. Kristan, M. Zuk, and two anonymous reviewers gave helpful suggestions.
This project was supported by a Newell Award, Department of Biology, and a Dissertation Research Grant, Graduate Division, UCR to D. M. Kristan and by National Institutes of Health Grant 30745 and a University of California Faculty Fellowship award to K. A. Hammond.
Address for reprint requests and other correspondence: D. M. Kristan, Dept. of Biology, University of California, Riverside, CA 92507 (E-mail:).
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- Copyright © 2001 the American Physiological Society