Physiological and morphological responses to simultaneous cold exposure and parasite infection by wild-derived house mice


  • Deborah M. Kristan,

    Corresponding author
      *Present address and correspondence. Department of Biological Sciences, PO Box 3051, University of Idaho, Moscow, ID 83844–3051, USA. Tel. +1 208 885-6905. Fax +1 208 8857905. E-mail
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  • Kimberly A. Hammond

    1. Department of Biology, University of California, Riverside, CA 92521, USA
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*Present address and correspondence. Department of Biological Sciences, PO Box 3051, University of Idaho, Moscow, ID 83844–3051, USA. Tel. +1 208 885-6905. Fax +1 208 8857905. E-mail


  • 1Many animals respond to environmental demands with phenotypic plasticity of morphology and physiology. We examined the effects of ambient temperature and parasitism on morphology and physiology of wild-derived house mice (Mus musculus) that were exposed to cold and/or experimentally infected with a naturally occurring intestinal nematode (Heligmosomoides polygyrus).
  • 2Parasitized mice had changes in some organ masses, decreased ability to digest food, and lower rates of glucose transport but similar total glucose transport capacity as unparasitized mice. Wild-derived house mice did not use fat stores to respond to parasitism but did increase mucosal mass in the small intestine enough to maintain glucose acquisition at a similar level to unparasitized mice.
  • 3Cold-exposed mice showed increased masses of some organs, lower rates of glucose transport but similar total capacity to transport glucose as warm acclimated mice.
  • 4The effects of cold exposure and parasite infection were largely independent of each other for the morphological and physiological parameters we measured.
  • 5The more recent exposure of wild-derived house mice to fluctuating temperatures and to parasite infection may help to explain the subtle differences that we observed in how wild-derived mice respond to environmental demands compared to their laboratory mouse counterparts.


In most ecosystems wild animals are subjected to variations in temperature, moisture, food availability, and other biotic and abiotic demands that can constrain an animal's time, energy, movements or other components of its daily existence. When one demand occurs at a single time, an animal may respond differently than if several demands occurred simultaneously. Response to one demand may either be enhanced or diminished in the presence of another. For example, Monroy and colleagues have shown that cold stress in mice alters their immune response and subsequent response to infection with Toxoplasma gondii (Banerjee et al. 1999; Monroy et al. 1999; Aviles & Monroy 2001). We are interested in how animals respond to the complicated array of demands they may encounter in nature.

Phenotypic plasticity of morphology and physiology often occurs with seasonal or daily fluctuations in the environment (Piersma & Lindström 1997). For example, there are increases in preflight masses of muscle and heart with concomitant decreases in mass of digestive organs for some migratory birds (Piersma, Koolhaas & Dekinga 1993; Weber & Piersma 1996; Battley & Piersma 1997; Piersma 1998; Piersma & Gill 1998; Piersma, Gudmundsson & Lilliendahl 1999). Futhermore, snakes with intermittent eating habits quickly and drastically change the morphology and physiology of their digestive system when a meal is consumed (cf. Secor & Diamond 1995; Starck & Beese 2002). These examples illustrate the importance of both detection of and response to the naturally complex environment.

A fluctuating, but common, demand in the environment is parasites. Many wild animals encounter parasites (Behnke et al. 2001) that can affect numerous aspects of their biology, including behaviour (Kavaliers & Colwell 1995; Poulin 1995; Moore & Gotelli 1996), reproduction (Feore et al. 1997; Richner 1998), food consumption (Crompton et al. 1985; Arneberg, Folstad & Karter 1996), and morphology (Kristan & Hammond 2000, 2001) even if a host's survival is not directly affected. We examined the simultaneous effects of two common demands, low temperature and an intestinal nematode parasite (Heligmosomoides polygyrus), on the morphology and physiology of wild-derived house mice (Mus musculus).

We hypothesized that cold exposure and H. polygyrus infection would affect body composition, organ masses and intestinal glucose transport of wild-derived house mice. We predicted that cold-exposed mice would have less body fat but greater organ masses and glucose transport capacity (Kristan & Hammond 2000), H. polygyrus infection would increase lean mass and decrease glucose transport by the small intestine (Kristan & Hammond 2000, 2001), and, based on our findings from laboratory mice (Kristan & Hammond 2000), that simultaneous parasite infection and cold exposure would elicit independent morphological and physiological responses. Importantly, other researchers have found that cold exposure and sublethal parasite infection interact with each other (Banerjee et al. 1999; Monroy et al. 1999; Aviles & Monroy 2001; Meagher & O’Connor 2001), so it is possible that we would find an interaction between cold and H. polygyrus infection for wild-derived house mice. We compare our findings from this study to our previous work with laboratory mice.

Materials and methods

Our experiment had two independent variables with two levels each: parasite infection (parasitized, unparasitized) and cold exposure (cold [5 °C], warm [23 °C]). We used 40 individually housed virgin female house mice, 10 per treatment group. Because parasite infection with H. polygyrus can differentially affect males and females (Dobson 1961; Dobson & Owen 1978), we used only females. We obtained first generation captive born house mice, whose parents were trapped near Flagstaff, from Dr Lee Drickamer (Northern Arizona University). Mice for this experiment were second-generation captive born and were rendered free of infectious disease (as confirmed by sentinel mouse protocol; University of California, Riverside) except for pinworm (Syphacia obvelata) which occurred in our vivarium. Pinworm infection was determined for each mouse and the effects of pinworm occurrence were statistically tested.

parasite maintenance and mouse infection procedures

Heligmosomoides polygyrus infective stage larvae (L3) were cultivated from non-experimental animals (see protocol in Kristan & Hammond 2001). Mice were given 300 ± 9 to 300 ± 11 L3 (number of worms ± 1 SD; range of values indicates more than one parasite culture used for infections) by using a 200-µL pipette tip (Fisherbrand Redi-Tip on a Pipetman) that was placed at the back of the mouse's throat. The suspension containing the L3 was dispensed into the mouse's throat and the mouse swallowed to complete the infection procedure. All mice in the unparasitized group were given an equal volume of tap water. Infection status of all mice was checked 14 days post-infection (PI) using a modified McMaster technique (Bowman 1995).

cold exposure and digestive efficiency

For the first 6 days PI, mice were maintained at 14 L: 10 D, 23 °C and fed Rodent Diet 5001 (Purina Mills, Inc.) ad libitum. On day seven 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; Diamond & Karasov 1984) 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 faeces), cold exposed mice were moved to an environmental chamber (5 °C; 14 L : 10 D) for 10 days while control mice stayed at 23 °C. For all mice, body mass, food intake rate (I, g day−1), and faecal output rate (O, g day−1) were measured on days 21–23 PI. Digestive efficiency (average percentage apparent dry matter digestibility) was calculated as [(I − O)/I] × 100 for days 21–23 PI. We then calculated digestible food intake by multiplying absolute food intake by digestive efficiency to determine the amount of food (in grams) absorbed.

organ morphology and body fat

On day 24 PI, we anaesthetied mice between 08:30 and 11:30 hours by intraperitoneal injection of a lethal dose of sodium pentobarbital (0·04 mL at 65 mg ml−1). We removed the small intestine, stomach, caecum, large intestine, heart, liver, spleen, kidneys and lungs. Excess fat and connective tissue were removed from each organ and returned to the mouse carcass. After the small intestine was removed (see below) the mice were killed by cutting the diaphragm. For stomach, caecum and large intestine, we removed contents by rinsing with mammalian Ringer's solution. We measured dry mass of organs and the carcass after drying at 55–60 °C for two or 14 days, respectively.

We ground the dried carcass then extracted lipids using petroleum ether (Goldfische apparatus; Labconco). Extracted samples were re-dried and the difference in mass before and after extraction was taken as the mass (g) of fat in the carcass. We measured fat content of all body organs combined, except the small intestine. 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 re-dried organs and the mass difference before and after extraction was the amount of fat (g) in organs. Total fat mass was the sum of carcass and organ fat masses and lean mass was the initial whole body mass minus total fat mass.

small intestine morphology and glucose uptake

While the mouse was anaesthetized, we rinsed the small intestine in situ with cold mammalian Ringer's solution. We then removed the small intestine and placed it in cold oxygenated Ringer's solution (bubbled with 5% CO2 : 95% O2 at 2–3 L min−1). We divided the small intestine into three regions of equal length (proximal, mid and distal) and measured the wet mass of each region. Total small intestine mass was the sum of the three regions, corrected for mass of the parasites (described below). We cut two 1·5-cm sleeves per region, which were weighed and then used to separate mucosal/submucosal tissue (hereafter called ‘mucosa’) from muscularis/serosal tissue (hereafter called ‘serosa’; Diamond & Karasov 1984). The dry mass : wet mass ratio was calculated for each sleeve and we used the average of these ratios to calculate ‘mucosal’ and ‘serosal’ dry mass for the entire small intestine (Diamond & Karasov 1984; Hammond & Diamond 1992).

We measured the maximal transport velocity of the brush-border d-glucose transporter (SGLT1) in vitro using the everted sleeve technique (validated for laboratory strains of Mus musculus by Karasov & Diamond 1983; Diamond & Karasov 1984). In brief, we everted each region of the small intestine and then cut four 1·5-cm long sleeves, immediately adjacent to each other: two sleeves for measuring ‘mucosal’ and ‘serosal’ mass as described above, and two sleeves for measuring glucose uptake. We mounted everted sleeves on stainless steel rods, incubated them for 2 min in 36 °C Ringer's solution containing 50 mm d-glucose and trace amounts of 14C-d-glucose and 3H-l-glucose. Glucose uptake rate of each sleeve (mmoles day−1 g wet mucosal tissue−1) was measured using liquid scintillation (LS 6500 scintillation system, Beckman). We then averaged uptake rate of the two sleeves from each region and multiplied this by the wet mucosal mass (g) of the entire region to calculate regional glucose uptake capacity. The products of each region were summed to determine total glucose uptake capacity for the entire small intestine.

parasite infection intensity and adjusted small intestine mass

Parasite infection intensity was determined by counting H. polygyrus from small intestine tissue and adding this to the estimate of worm number from sleeves used for the glucose uptake measurement (for detailed methods see Kristan & Hammond 2001). Total parasite wet mass (number of worms multiplied by average worm mass, Kristan & Hammond 2001) was subtracted from small intestine wet mass prior to calculation of small intestine dry mass. When we examined each intestinal region separately, parasite mass was subtracted only from mass of the proximal region where adult parasites occur (Bansemir & Sukhdeo 1996).


Our data consist of two independent (parasite infection, cold exposure) and numerous dependent (food intake, digestive efficiency, body mass, small intestine morphological variables, organ masses, body fat, glucose uptake rate and capacity) variables. We first used a multivariate analysis of variance (manova), to test for effects of pinworm infection (factor: pinworm occurrence). The presence of pinworm was not significant (P > 0·05) so we used manova (factors: temperature, parasite infection, temperature × parasite infection) to test for significant differences between treatments. Because this manova was significant (P < 0·05), we used independent anovas (factors: as described for manova) to determine which treatments and which dependent variables were statistically significant. We regressed each dependent variable against body mass and if the regression was significant we used body mass as a covariate in the anova and present least squares means ± 1 standard error. If the regression between the dependent variable and body mass was not significant, we present the arithmetic mean ± 1 SE. For each analysis, we examined Levene's test for equality of variances to determine if there were treatment effects on the variability of the dependent variables.


sample size and infection intensity

Three parasitized mice (one in cold, two in warm) were excluded from analyses due to technical difficulties in parasite inoculation. Of the remaining 37 mice, all mice given parasites developed mature infections and no control mice became infected. Regressions of body mass with each dependent variable were significant for all variables except spleen mass and digestive efficiency. Data for some parasitized mice were not useable for the glucose uptake measurements because of problems with scintillation chemicals therefore the final sample size for the glucose uptake data was three parasitized mice for each temperature treatment group.

Infection intensity (number of worms) was similar for mice in cold and warm treatments (cold: 200 ± 20, n = 9; warm: 196 ± 21, n = 8; P > 0·05) with a minimum infection intensity of 79 worms and a maximum of 306 worms. Overall infection success ([no. adult worms recovered/no. L3 administered] × 100) averaged 66 ± 5%.

body mass, body composition and organ morphology

Whole body mass, lean body mass, and total body fat were similar for all mice regardless of treatment (P > 0·05, Fig. 1). We examined effects of each treatment on total organ mass. Parasitized mice had larger kidneys (by 6%; F1,32 = 50·78, P < 0·0001) and spleens (by 87%; F1,32 = 39·40, P < 0·0001) and marginally larger lungs (by 5%; F1,32 = 3·47, P = 0·072) than unparasitized mice (Table 1). Parasitized mice had smaller stomachs (by 14%; F1,32 = 24·79, P < 0·0001) and large intestines (by 10%; F1,32 = 50·78, P < 0·0001) than unparasitized mice. Cold exposed mice had larger kidneys (by 12%; F1,32 = 24·73, P < 0·0001), spleens (by 30%; F1,32 = 7·52, P = 0·01), hearts (by 24%; F1,32= 32·34, P < 0·0001), and large intestines (by 12%; F1,32 = 6·11, P = 0·019) than warm temperature mice (Table 1). There was a significant interaction between treatments for kidney mass (F1,32 = 6·98, P = 0·013) because the increase in kidney mass of parasitized mice only occurred when mice were also cold-exposed. Variance was greater for parasitized than unparasitized mice for spleen mass (Levene's test: F3,33 = 3·91, P = 0·017) but did not differ for other organs.

Figure 1.

Body mass and composition (lean vs. fat mass) for wild-derived house mice given cold exposure or parasite infection. Percent body fat is written on each bar. Values are mean + 1 standard error of the mean.

Table 1.  Least squares means adjusted for whole body mass ± 1 standard error for organ dry masses (g) from parasitized or unparasitized mice exposed to cold or room temperature. Sample size is shown in parentheses
Cold (n = 9)Warm (n = 8)Cold (n = 10)Warm (n = 10)
  • *

    Arithmetic means.

Liver0·335 ± 0·0120·323 ± 0·0130·329 ± 0·0110·320 ± 0·011
Kidney0·080 ± 0·0020·068 ± 0·0020·071 ± 0·0020·068 ± 0·002
Spleen*0·016 ± 0·0020·012 ± 0·0010·008 ± 0·0010·007 ± 0·001
Heart0·034 ± 0·0010·026 ± 0·0010·033 ± 0·0010·028 ± 0·001
Lung0·029 ± 0·0010·028 ± 0·0010·028 ± 0·0010·027 ± 0·001
Stomach0·030 ± 0·0010·029 ± 0·0010·035 ± 0·0010·034 ± 0·001
Caecum0·013 ± 0·0010·012 ± 0·0010·014 ± 0·0010·012 ± 0·001
Large intestine0·036 ± 0·0020·035 ± 0·0020·040 ± 0·0020·033 ± 0·002

Parasitized mice had a 15% longer small intestine than unparasitized mice (F1,32 = 41·37, P < 0·0001) but small intestine length did not differ with cold exposure (Fig. 2). Because qualitative results are the same for absolute small intestine dry mass and dry mass per unit length, we report results only for absolute dry mass. Small intestine dry mass was 46% greater for parasitized than unparasitized mice (F1,32 = 50·67, P < 0·0001) and was 11% greater for cold exposed than warm temperature mice (F1,32 = 4·42, P = 0·044; Fig. 2). Across the entire small intestine, parasitized mice had 38% more mucosal and 72% more serosal tissue than unparasitized mice (F1,31 = 32·35, P < 0·0001 and F1,32 = 40·55, P < 0·0001, respectively; Fig. 2) and cold exposed mice had 15% more mucosal tissue (F1,31= 6·44, P = 0·016) but similar mass of serosal tissue as warm temperature mice.

Figure 2.

Small intestine dry mass (showing ‘serosal’ and ‘mucosa’ mass) and length for wild-derived house mice given cold exposure or parasite infection. Values are mean + 1 standard error of the mean.

The effects of parasites on small intestine mass occurred for all regions (proximal: F1,32 = 25·62, P < 0·0001, mid: F1,32 = 37·36, P < 0·0001, distal: F1,32= 10·34, P = 0·003), but the effects of cold exposure only occurred for the mid region (F1,32 = 7·11, P = 0·012; Fig. 3). When we examined each region separately, mucosal mass was greater with parasite infection for all regions (proximal: F1,31 = 14·04, P = 0·001, mid: F1,32 = 27·22, P < 0·0001, distal: F1,32 = 10·56, P = 0·003) and with cold exposure for the mid region only (F1,32= 6·29, P = 0·017; Fig. 3). Serosal mass was greater for parasitized than unparasitized mice only for the proximal and mid regions (F1,32 = 36·06, P < 0·0001 and F1,33 = 14·33, P = 0·001, respectively) and there were no effects of cold exposure on serosal mass for any region (Fig. 3).

Figure 3.

Dry mass of three small intestine regions (proximal, mid and distal) showing ‘serosal’ and ‘mucosal’ masses for wild-derived house mice given cold exposure or parasite infection. Values are mean + 1 standard error of the mean.

food intake and digestive efficiency

Food intake did not differ with parasite infection but cold-exposed mice ate 52% more food than mice at room temperature (F1,32 = 124·84, P < 0·0001; Fig. 4). Digestive efficiency was 2% less for parasitized than unparasitized mice (F1,32 = 16·63, P < 0·0001) but did not differ with cold exposure (Fig. 4). Because of the significant effect of parasite infection on digestive efficiency, we examined digestible food intake. Although digestible food intake was 7% less for parasitized than unparasitized mice, this difference was not significant (F1,32 = 3·47, P = 0·072; Fig. 4). Similar to absolute food intake, digestible food intake was 51% greater for cold exposed compared to room temperature mice (F1,32 = 115·08, P < 0·0001; Fig. 4).

Figure 4.

Absolute food intake, digestible food intake, and digestive efficiency for wild-derived house mice given cold exposure or parasite infection. Values are mean + 1 standard error of the mean.

glucose transport

Average glucose uptake rate (normalized to wet mucosal mass) for the entire small intestine (mmole g−1 day−1) was 41% less for parasitized than unparasitized mice (F1,22 = 23·85, P < 0·0001) and 25% less for cold exposed compare to warm temperature mice which was marginally non-significant (F1,22 = 3·09, P = 0·093; Fig. 5). Glucose uptake rate for parasitized mice was 62% less (F1,22 = 32·70, P < 0·0001) in the proximal region and 24% less (marginally non-significant) in the mid region (F1,22 = 3·56, P = 0·072) compared to unparasitized mice but was greater by 20% (non-significant) in the distal region (F1,22 = 2·32, P = 0·142; Fig. 5). Cold exposed mice had lower glucose transport rates than warm temperature mice by 8–24%, which was not significant, although the mid region was only marginally non-significant (F1,22 = 3·48, P = 0·075; Fig. 5).

Figure 5.

Rate of glucose uptake normalized to wet ‘mucosal’ mass for three regions of the small intestine (proximal, mid and distal) and the average for all regions for wild-derived house mice given cold exposure or parasite infection. Values are mean ± 1 standard error of the mean.

Because of the increase in mucosal tissue available for glucose transport in parasitized and cold-exposed mice (see above), total glucose uptake capacity (mmoles day−1) summed for the entire small intestine did not differ with either treatment (Fig. 6). Parasitized mice had 41% lower glucose uptake capacity in the proximal region (F1,21 = 19·53, P < 0·0001), similar glucose uptake capacity in the mid region (P > 0·05), and 70% greater uptake capacity in the distal region (F1,21 = 18·70, P < 0·0001) compared to unparasitized mice (Fig. 6). There were no effects of cold exposure on glucose uptake capacity of any region.

Figure 6.

Total glucose uptake capacity for three regions of the small intestine (proximal, mid and distal) and summed for the entire small intestine for wild-derived house mice given cold exposure or parasite infection. Values are mean ± 1 standard error of the mean.


The most telling measures of intestinal accommodation to any nutrient challenge are the digestible food intake (which indicates the combined effects of digestive efficiency and food consumption) and the glucose uptake capacity of the small intestine. In this study, parasitized, and to a lesser extent cold-exposed, mice overcame lower digestive efficiency or increased food intake (load on the gut) by increasing small intestinal mucosal mass. Increase in small intestine mucosal mass as well as several other measures of response to challenges was observed for both house mice and laboratory mice but there were also some differences in how these two mouse strains responded to experimentally imposed demands.

energy acquisition and body composition

Despite a slight decrease in ability to digest food when parasitized, house mice did not show changes in food intake, body mass or, unlike our prediction, body composition. Similar to results found with tapeworm infection (Hymenolepis citelli) in the white-footed mouse (Peromyscus leucopus; Munger & Karasov 1989, 1991), our results indicate that the diminished digestive efficiency when parasitized was probably not directly biologically relevant to the mouse (at least under the laboratory conditions of our experiment) because of the phenotypic augmentation of small intestinal mass which resulted in similar nutrient uptake capacity of parasitized and unparasitized mice. It is possible, of course, that growing more small intestine tissue to accommodate the lower digestive efficiency could be biologically important and this concept deserves further investigation.

Despite increased thermoregulatory demands with cold exposure, wild-derived mice met this demand with increased food intake and intestinal accommodation, not by using body fat. In contrast, cold-exposed laboratory mice used body fat to help meet the demands of both cold exposure and parasite infection (Kristan & Hammond 2000). Even though the wild-derived house mice in our study were born and raised in captivity under ad libitum food and water availability, they had relatively little body fat (approximately 14%) compared to Swiss-webster laboratory mice of similar age (approximately 20%; Kristan & Hammond 2001). Using fat to meet demands may not be a viable option for wild-derived house mice or, by extrapolation, for house mice in nature. Indeed, Bronson, Heideman & Kerbeshian (1991) also found that fat resources were not a viable option to meet energy needs during cold exposure even over a few days for wild-derived peripubertal house mice. Therefore changes in other physiological, behavioural or morphological parameters may be a more viable solution to meet thermoregulatory demands by parasitized wild house mice that can not rely on body fat as can the typical laboratory mouse.

Wild-derived house mice did not eat more, lose body mass or use fat when parasitized with H. polygyrus, which implies that the cost of being infected (e.g. immune response, tissue repair) for these mice may be less than for Swiss-Webster laboratory mice which mobilized fat resources and substantially increased lean mass (by approximately 10%) and resting metabolism. Wild-derived house mice showed similar trends in body composition changes with parasite infection (14% decrease in fat mass; by 1·5% increase in lean mass) to laboratory mice, but these changes in resource allocation were not statistically significant. There is variation in susceptibility to H. polygyrus among laboratory mouse strains (Brindley & Dobson 1983; Dobson & Jian-Ming 1991) and it is therefore plausible that our wild-derived house mice differ in their response to H. polygyrus in part due to genetic differences from Swiss-Webster mice.

phenotypic plasticity of organ masses

Despite the differences noted above, wild-derived house mice showed many similarities in plasticity of organ masses to laboratory mice during both H. polygyrus infection and cold exposure (Kristan & Hammond 2000). During H. polygyrus infection, both mouse strains increased spleen (presumably related to an immune response) and small intestine (in regions occupied and unoccupied by parasites) masses but unlike laboratory mice wild-derived mice increased kidney mass when parasitized. Cold-exposed mice of both strains showed some similar organ mass increases but also increased masses of organs that were different between the strains (Kristan & Hammond 2000).

Simultaneous demands can elicit either independent or interactive responses in an animal. As predicted, responses of wild-derived mice to cold exposure and H. polygyrus infection were independent of each other (with the exception of kidney mass). Given that both experimental demands increased some of the same organ masses and similarly affected glucose transport by the small intestine, it is possible that response to one demand allowed mice to accommodate the second demand without much additional change in morphology and physiology.

phenotypic plasticity of small intestine structure and function

The total amount of mucosal tissue available can affect the functional capacity of the small intestine because it is cells in the mucosal tissue that are responsible for glucose and amino acid transport (Ferraris 1994; Wright et al. 1994). In contrast to laboratory mice (Kristan & Hammond 2000), wild-derived house mice did not have greater intestinal glucose transport capacity after cold exposure. Interestingly, cold-exposed house mice increased their total mucosal mass by 15% whereas laboratory mice increased their total amount of mucosa by only 6% (Kristan & Hammond 2000). Therefore, while laboratory mice enhanced their total glucose transport capacity with cold exposure by increasing the amount of tissue available, wild-derived house mice were unable to increase total glucose transport capacity to the levels of warm-acclimated mice despite a more than two-fold greater increase in small intestine tissue than was seen in laboratory mice. It may be relevant that at room temperature, laboratory mice have 23% more mucosal tissue than wild-derived mice (after accounting for body mass effects; Kristan & Hammond 2000), so the laboratory mouse maintains a baseline of mucosal tissue that exceeds the wild-derived house mouse and thereby enables the laboratory mouse to respond differently to demands requiring increased energy acquisition at the level of the small intestine.

Despite diminished glucose transport rate with parasite infection, the total glucose transport capacity of the small intestine was the same for parasitized and unparasitized house mice. This is due in part to a 38% increase in mucosal tissue with parasite infection. Therefore, although small intestine tissue was less functional on a per unit basis, the small intestine as a whole maintained similar function in parasitized and unparasitized mice. It should be noted that small sample size of infected mice for this part of the study may be a problem and indeed our statistical power is only 19% to detect effects of H. polygyrus and 7% to detect effects of cold exposure on total glucose transport capacity. Given the current level of variation in our data, we would have needed a sample size of 28 to detect effects of parasite infection and of 287 to detect effects of cold exposure (at an alpha of 0·05) which is approximately two (for parasite infection) to 200 (for cold exposure) times greater than the sample sizes used in our previous study with laboratory mice where we detected effects of both H. polygyrus and cold exposure on glucose transport capacity (Kristan & Hammond 2000).

Of interest is that, although H. polygyrus occurs only in the proximal portion of the small intestine, the effects of parasite infection in house mice are detected in more distant regions. A similar situation is found in laboratory mice, and we have previously interpreted this in terms of changes in nutrient density of digesta (Kristan & Hammond 2000). We propose that diminished function of proximal small intestine with parasite infection results in digesta with a greater nutrient density reaching the mid and distal regions of the small intestine compared to an unparasitized mouse. Changes in nutrient density of ingesta can increase nutrient absorption by the small intestine (Ferraris, Villenas & Diamond 1992; Ferraris 1994) such that diminished function in the proximal region may result in enhanced function in the distal region (Reville et al. 1991). An examination of the effects of H. polygyrus on nutrient density of digesta throughout the small intestine mice would provide valuable insight toward understanding the relationship between changes in small intestine structure (‘mucosal’ mass) and function (glucose transport).

In addition to many of the similarities noted above, wild-derived house mice showed some potentially important differences from laboratory mice. Specifically, unlike laboratory mice, parasitized wild-derived mice had a slight decrease in digestive efficiency, did not use fat stores, and did not have diminished total capacity to transport glucose, and cold-exposed wild-derived mice did not increase mass of mucosal tissue enough to increase total glucose transport capacity so had to rely only on increased food consumption to meet the thermoregulatory demand. Differences in morphological and physiological plasticity of laboratory mice and wild-derived house mice may reflect the more recent evolutionary history that wild-derived mice have with a variable and demanding environment. Phenotypically plastic responses to environmental demands may have long-term consequences (e.g. to life history and survival) that warrant further investigation using animals only recently removed from the wild.


Stacey Leder and Joanne Narksompong assisted with glucose uptake experiments and organ fat extractions. We thank two anonymous referees for their helpful comments. Experiments were approved by the Animal Care and Ethics Committee at the University of California, Riverside. This research was supported by a NSF Doctoral Dissertation Improvement Grant (IBN-0073229) to D.M.K. and by UCR Academic Senate Grant to K.A.H.