Physiological responses to feeding, fasting and estivation for anurans
Department of Physiology, University of California at Los Angeles, School of Medicine, Los Angeles, CA 90095-1751, USA and *Department of Biological Sciences, The University of Alabama, Tuscaloosa, AL 35487-0344, USA
Accepted 25 April 2005
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Summary |
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Key words: anuran, Bufo, Ceratophrys, Leptodactylus, Pyxicephalus, Rana, estivation, fasting, intestinal nutrient transport, specific dynamic action
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Introduction |
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An apparent adaptive mechanism employed by sit-and-wait foraging,
infrequently feeding snakes to reduce energy expenditure while fasting
includes the downregulation of their gastrointestinal (GI) tract upon the
completion of digestion. The link between the fasting-related reduction in gut
function and concurrent decrease in energy expenditure is suggested by the
lower standard metabolic rate (SMR) of sit-and-wait foragers compared to that
of active foraging snakes that feed more frequently
(Secor and Diamond, 2000). In
contrast to the sit-and-wait foragers, frequently feeding snakes do not
downregulate intestinal function with fasting, but rather maintain elevated
function until the next meal. For sit-and-wait foraging snakes, feeding is
consequently met with a rapid upregulation of GI function, highlighted by 10-
to 20-fold increases in intestinal nutrient uptake
(Secor and Diamond, 2000
).
Selective mechanisms to reduce tissue metabolism would predictably also be
present for both hibernating and estivating species. Evidence of such
mechanisms for hibernating amphibians, reptiles and mammals include the
overwintering reduction in intestinal mass or function
(Carey, 1992;
Csáky and Galluci, 1977
;
Qadri et al., 1970
).
Understandably, energy-conserving mechanisms would be more important for
estivating species given that their body temperatures, and consequentially
their metabolic rates, are substantially higher than hibernating species
(Pinder et al., 1992
). For
several amphibian species, metabolic rates are reduced by 5085% during
estivation, reportedly a function of decreases in organ metabolism
(Fuery et al., 1998
;
Guppy and Withers, 1999
).
Given the fasting responses of the GI tract of infrequently feeding snakes,
the benefits of gut downregulation to conserve energy, and the adaptive
plasticity of the intestine (Piersma and
Lindström, 1997
), I hypothesized that anuran species which
estivate during their dry season severely downregulate intestinal performance
with fasting and estivation, and consequently upregulate intestinal
performance rapidly after feeding.
A test of this hypothesis would be best served by first demonstrating the wide regulation of intestinal performance in response to feeding and fasting among distantly related anurans that estivate, and second the narrow regulation of intestinal performance for species closely related to the estivating species but which do not estivate. The experimental design that I implemented to test this hypothesis compares several physiological responses to fasting and feeding between an estivating and a non-estivating species for three anuran families, Bufonidae, Leptodactylidae and Ranidae. For each species, I measured metabolic rates, organ masses, and intestinal morphology and nutrient uptake of individuals fasted and digesting. In addition, I took similar sets of measurements of C. ornata and P. adspersus following 1 month of laboratory-induced estivation. My objectives in this study were to demonstrate: (1) that the wide regulation of intestinal performance has evolved independently among anuran estivators, (2) that the magnitude by which anurans regulate intestinal performance is linked to their feeding ecology, (3) that intestinal performance is further reduced during estivation, and (4) that the up- and downregulation of intestinal performance is reflected, respectively, in elevated and depressed metabolic rates.
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Materials and methods |
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Bufo marinus, C. ornata, L. pentadactylus and R. catesbeiana were purchased commercially and originated, respectively, from southern Florida, a captive-propagated colony in southern California, Suriname, and the southeastern United States. Bufo alvarius were collected under a state collecting permit in southern Arizona, and P. adspersus were captured in Zimbabwe. Individuals of each species were housed either individually in plastic storage boxes (20 liters) or together in large plastic or fiberglass containers at 2427°C under a photoperiod of 14 h:10 h L:D. Anurans had access to water in each container and were fed a diet of crickets or small rodents at 37 day intervals. Prior to metabolic and nutrient transport trials, individuals were fasted for 2 weeks to ensure that their guts were emptied and that all digestive activities had ceased. Because anurans commonly store water in their bladder, I manually emptied their bladder by inserting a glass rod into their cloaca and gently squeezing their hind section before measuring body mass. For this study, I used 10 B. alvarius [142.2±8.3 g (mean ± 1 S.E.M.)], 12 B. marinus (139.3±7.7 g), 11 C. ornata (185.9±21.6 g), 9 L. pentadactylus (157.8±14.9 g), 10 P. adspersus (227.6±10.9 g) and 11 R. catesbeiana (273.3±22.5 g). Animal care and experimentation were conducted under the approval of the UCLA Animal Research Committee and the University of Alabama Institutional Animal Care and Use Committee.
Metabolic response to feeding
I quantified metabolic rates as rates of oxygen consumption
(O2) measured
using closed-system respirometry as described by Vleck
(1987
) and Secor and Diamond
(1997
). Anurans were placed
individually into respirometry chambers (23.5 liter) and maintained at
30°C within an environmental chamber. Each respirometry chamber was fitted
with an incurrent and excurrent air port, each attached to a three-way
stopcock. Air was pumped into the chambers through the incurrent air port. A
small amount of water was placed in each chamber to prevent anurans from
desiccating from the constant air flow. For each
O2 measurement,
a 20 ml air sample was withdrawn from the excurrent air port, and both
incurrent and excurrent ports were then closed to seal the chamber.
0.51 h later, the excurrent air port was opened and a second 20 ml air
sample was withdrawn. Air samples were pumped (125 ml min-1)
through a column of water absorbent (Drierite; W. A. Hammond Drierite Co.,
Xenia, OH, USA) and CO2 absorbent (Ascarite II; Thomas Scientific,
Swedesboro, NJ, USA) into an O2 analyzer (S-3A/II; AEI
Technologies, Pittsburgh, PA, USA). I calculated whole-animal (ml
h-1) and mass-specific (ml g-1 h-1) rates of
oxygen consumption corrected for standard pressure and temperature as
described by Vleck (1987
).
Each metabolic trial began with the measurement in fasted anurans of
O2 twice a day
(at
08:00 h and
20:00 h) for 3 days. For each anuran, I assigned its
SMR as the lowest
O2 measured over
those 3 days. After SMR measurements, anurans were fed rodent (neonate rats)
meals equal in mass to 15% of anuran body mass. Anurans were then returned to
their respirometry chambers and metabolic measurements were continued at 12 h
intervals (at
08:00 h and
20:00 h) for 3 days and thereafter at 1
day intervals (at
08:00 h) for 5 more days. For each metabolic trial, I
quantified the following seven variables: SMR, peak
O2 (highest
recorded
O2
following feeding), factorial scope of peak
O2 (calculated
as peak
O2
divided by SMR), duration (time from feeding that
O2 was no longer
significantly greater than SMR), total energy expended above SMR over the
duration of significantly elevated
O2, [specific
dynamic action (SDA) quantified as kJ and kJ g-1], and SDA
coefficient (SDA quantified as a percentage of the energy content of the
meal). I quantified SDA (kJ) by summing the extra O2 consumed above
SMR during the duration of the significant metabolic response and multiplying
that value by 18.3 J ml-1 O2 consumed, assuming the
catabolism of a diet that is 70% protein, 25% fat and 5% carbohydrate, and an
RQ of 0.75 (Gessaman and Nagy, 1988). The energy content of the rodent meals
was calculated based on an energy equivalent of 5.98 kJ g-1 wet
mass, as determined from bomb calorimetry. Briefly, five neonate rats
(20.4±0.6 g) were each weighed, dried, reweighed and ground to a fine
powder which was pressed into pellets. Three pellets from each rodent were
ignited in a bomb calorimeter (1266; Parr Instrument Co., Moline, Illinois,
USA) to determine energy content (kJ g-1). Wet mass energy content
of the rodent meals (5.98 kJ g-1) represents the average energy
content of the five rodents, each calculated from the mean energy of the three
pellets (24.43±0.19 kJ g-1 dry mass) and the rodent's
relative body water content (75.6±0.4% of body mass).
Intestinal nutrient uptake
From three fasted and three fed individuals of each anuran species, I
quantified mass-specific rates of nutrient transport across the intestinal
brush border membrane using the everted sleeve technique
(Karasov and Diamond, 1983;
Secor et al., 1994
). Anurans
were killed by severing their spinal cord immediately posterior to the skull
either following a 2-week fast (`fasted') or after the 2-week fast, 1 day
following the consumption of a rodent meal (neonate rats) equaling 15% of
anuran body mass (`fed'). From each anuran, I removed and weighed the small
intestine, flushed it of its contents, reweighed it, and divided it into
thirds of equal length. Each third was weighed, everted and divided into 1 cm
segments. Sleeves were mounted on metal rods, incubated first in amphibian
Ringer's solution at 30°C for 5 min, and then incubated for 2 min at
30°C in stirred amphibian Ringer's solution containing an unlabeled and
radiolabeled nutrient and an adherent fluid marker labeled with a different
radioisotope. For each intestinal third, I measured the transport of the amino
acids L-leucine and L-proline (each at 50 mmol
l-1 and labeled with 3H) and of the sugar
D-glucose (at 20 mmol l-1 and labeled with
14C). The adherent fluid markers polyethylene glycol (labeled with
14C) and L-glucose (labeled with 3H)
corrected for the amount of radiolabeled amino acids and D-glucose,
respectively, adherent to the everted intestinal sleeve. L-glucose
measures also serve to correct for the passive diffusion of
D-glucose. With these corrections, total amino acid uptake (passive
and carrier-mediated) and carrier-mediated D-glucose uptake rates
are quantified as nanomoles of nutrient transported per minute of incubation
per milligram of sleeve wet mass. As previously observed for snakes, nutrient
uptake rates of the proximal and middle intestinal regions were consistently
similar and, in many cases, greater than those of the distal region
(Secor and Diamond, 2000
).
Therefore, as in earlier studies, I report separately the averaged uptake
rates of the proximal and middle segments (reported as the anterior
intestine), and of the distal third of the small intestine. I assigned as a
measure of overall intestinal performance the small intestine's total capacity
to transport nutrients. Intestinal nutrient uptake capacity (µmol
min-1) was calculated as the summed products of nutrient uptake
rates (nmol min-1 mg-1) times intestinal mass (mg) for
each intestinal third (Secor and Diamond,
1995
).
Two studies have demonstrated that the everted sleeve technique severely
damages the mucosa of avian intestines, thereby negating its use for
accurately measuring nutrient uptake for those species of birds
(Starck et al., 2000;
Stein and Williams, 2003
). To
assess the potential damage caused by the everted sleeve method on anuran
intestines, I compared two sets of intestinal samples removed from the
anterior portion of the small intestine of fasted and fed B. marinus
and R. catesbeiana at three stages of the procedure: prior to
eversion, immediately following eversion, and after everted tissues had been
incubated at 30°C in unstirred amphibian Ringers for 5 min and in stirred
amphibian Ringers for 2 min. Samples were prepared for light microscopy (as
explained below) and examined for damages to the mucosal layer. Following
eversion and incubation, I found no discernable damage to the intestinal
mucosa, nor any significant change in villus length (measured from the edge of
the smooth muscle layer to the tip of the villus, N=20 per treatment
per individual) for either species, fasted or fed
(Fig. 1). As observed for
reptiles (Secor, 2005
;
Tracy and Diamond, 2005
), the
everted sleeve method apparently does not damage the intestinal mucosa of
anurans, therefore the method can provide a reliable measure of intestinal
function.
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Physiological responses to estivation
To identify the metabolic and tissue responses to estivation, I measured
metabolic rate, organ masses, and intestinal function of C. ornata
and P. adspersus following 1 month of laboratory-induced estivation.
Individual frogs were induced into a state of estivation by simply removing
their access to water. Within days, frogs became stationary and assumed a
hunched-over posture as their outer layer of skin began to dry and harden
(Loveridge and Withers, 1981).
For the remaining weeks, frogs did not move within their `cocoon' of dried
skin. During this time, frogs lost on average 35% of their body mass,
presumably as water. I measured metabolic rates of C ornata
(283±44 g, N=3) and P. adspersus (235±19 g,
N=4) following 1 month of estivation as described above. Three
individuals of each species were then killed for measurements of intestinal
mass and nutrient uptake, and organ masses as previous described.
Statistical analyses
For each SDA trial, I used a repeated-measures design analysis of variance
(ANOVA) to test for significant effects of time (before and after feeding) on
O2 (as ml
h-1 and ml g-1 h-1). Each ANOVA was followed
with a post hoc pairwise mean comparison (TukeyKramer
procedure) to identify significant changes in
O2 between
sampling time points and when
O2 did not
differ from SMR. I used ANOVA for mass-specific measures and ANCOVA (log body
mass as a covariate) for whole-animal measures to test for species effects on
metabolic variables. For each anuran family, I similarly tested for
differences between the estivating and non-estivating species. Because of the
significant variation in body mass among the six anuran species
(Table 1), I recalculated
whole-animal measures of SMR, peak
O2 and SDA of
each anuran assuming a body mass of 180 g (the approximate average mass of all
36 anurans). Recalculated values were determined from allometric equations
presented in table 4 of Secor and Falkner
(2002
) for B.
marinus, assuming mass exponents of 0.69, 0.85 and 1.02, respectively,
for SMR, peak
O2, and SDA. The
postprandial responses were assessed by ANOVA for mass-specific rates of
intestinal nutrient uptake and by ANCOVA (body mass as a covariate) for
intestinal nutrient uptake capacity, intestinal mass and morphology, and the
masses of other organs. Likewise, I tested for significant changes from
fasting to estivation in metabolic rate, intestinal nutrient uptake and
morphology. ANOVA and ANCOVA results are reported in terms of their P
values, and I provide P values of selected pairwise mean comparisons.
Statistical significance is designated as P<0.05 and mean values
are reported as means ±1 S.E.M.
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Results |
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For all six species, feeding induced a significant metabolic response
(P<0.0001). Within 12 h,
O2 had risen by
100300%, and by 24 h, rates were 2001000% greater than SMR
(Fig. 2). Peaks in
O2 were attained
between 1 and 2.5 days postfeeding, with little difference in the time of peak
rates between estivating and non-estivating species. Postprandial peaks in
O2, were
significantly (P<0.0001) different among the six species, but only
for Leptodactylidae was there a significant difference between the two species
(C. ornate > L. pentadactylus;
Table 1). The factorial scope
of peak
O2
differed significantly (P<0.0001) among the six species, ranging
from 3.5 for L. pentadactylus to 11.6 for C. ornata. For
each family, the estivating species possessed significantly
(P<0.0001) greater scopes than the non-estivating species
(Table 1). The duration of
significantly elevated
O2 ranged
between 4 and 7 days, with duration of the SDA response averaging a day longer
for estivating species (Table
1).
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Even within the narrow range of body masses (100320 g) of anurans
used in this study, SMR, peak
O2 and SDA (kJ)
scaled allometrically (P<0.0006,
r2=0.540.80) with body mass for estivating and
non-estivating individuals (Fig.
3). Mass exponents of SMR differed significantly
(P<0.05) between estivating (0.71) and non-estivating individuals
(1.11), but did not differ for either peak
O2 (1.04 for
estivating and 1.24 for nonestivating species) or SDA (1.06 and 1.03,
respectively). Illustrated as a function of body mass, individuals of
estivating species possessed lower SMR, similar peak
O2, and greater
SDA than individuals of non-estivating species
(Fig. 3).
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The postprandial response of intestinal nutrient uptake capacity, a product of small intestinal mass and mass-specific nutrient uptake rates, similarly differed between the estivating and non-estivating species (Fig. 5). With feeding, all three estivating species significantly (P<0.029) upregulated their intestine's capacity to transport the three nutrients. Uptake capacity of L-leucine, L-proline, and D-glucose increased on average by 6.3-, 6.7- and 10.3-fold among the three estivating species. For the non-estivators, only B. marinus experienced a significant postprandial response in uptake capacity, D-glucose uptake capacity increasing (P=0.012) by threefold. For the non-estivating species, intestinal uptake capacity among the three nutrients averaged only 69% higher after feeding.
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Organ morphology
For all six anurans, regardless of being fasted or fed, small intestinal
mass declined distally as the proximal one-third of the intestine averaged
200% heavier than the distal one-third. For the three estivating species,
feeding triggered a dramatic increase (averaging 230%) in small intestinal wet
mass (Fig. 6). Among the
non-estivating species, only B. marinus experienced an increase
(P=0.016) in intestinal mass with feeding. For the three
non-estivating species, average mass of the small intestine was only 50%
greater after feeding. Small intestinal length also increased significantly
(P<0.024) with feeding for the three estivating species, but did
not for any of the non-estivating species
(Fig. 6). Within 24 h after
feeding, estivating species had experienced on average an 83% increase in
small intestinal length.
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For all six species, there was no significant difference in the thickness of the intestinal muscularis/serosa layer between fasted and fed individuals (Fig. 7). The thickness of the intestinal mucosa, largely a function of villus height, increased significantly (P<0.045) with feeding (by 75130%) for the three estivating species, but did not vary for any of the non-estivating anurans (Fig. 7). Feeding had a significant (P<0.04) effect on enterocyte height for both species of bufonids, increasing the height of cells by 145% and 57%, respectively for B. alvarius and B. marinus (Fig. 7). In contrast, feeding did not impact enterocyte height for either species of Leptodactylidae or Ranidae. Feeding did induce significant (P<0.04) increases in enterocyte width for all six species, as enterocytes of estivating and non-estivating species increased by an average of 90% and 40%, respectively (Fig. 7). The postprandial increase in enterocyte length and/or width generated the concurrent increase (P<0.05) in enterocyte volume for five of the six species (Fig. 7). Enterocytes increased on average 150% in volume for non-estivators, and 440% in volume for the estivators.
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Metabolic and organ responses to estivation
Ceratophrys ornata and P. adspersus experienced
significant (P<0.045) reductions in
O2 following 1
month of induced estivation within the laboratory
(Fig. 8). Compared to frogs
fasted for 1 month, but alert and hydrated, estivation reduced
O2 by 25% and
16%, respectively, for C. ornata and P. adspersus. Anterior
and distal uptake rates averaged lowered for estivating frogs compared to
fasted individuals, but the difference was only significant (P=0.012)
for D-glucose uptake by the anterior small intestine of P.
adspersus. Concurrently, both species experienced significant
estivation-induced reduction in small intestinal mass, declining on the order
of 31% and 57%, respectively, for C. ornata and P. adspersus
(Fig. 8). The loss of
intestinal mass during estivation is largely responsible for the significant
(P<0.046) lowering of intestinal uptake capacity of all three
nutrients for both species (Fig.
8). Collectively, estivating frogs experienced a 60% reduction in
intestinal uptake performance compared to fasted frogs, and a 93% reduction
compared to fed frogs (Fig. 4).
Whereas estivation resulted in a decrease in small intestinal mass, no other
organ experienced a significant change in either wet or dry mass with
estivation.
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Discussion |
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Metabolic response to feeding
Meal digestion is accompanied by an increase in metabolic rate, which
represents the specific dynamic action of meal, which in its entirety
represents the combined energy expended on meal ingestion, digestion and
assimilation (Kleiber, 1975).
The six anuran species of this study all experience significant postprandial
increases in metabolic rate by as much as 3.5- to 11.6-fold. Metabolic
increases associated with feeding have previously been documented for the
anurans B. marinus (Secor and
Faulkner, 2002
; Andersen and
Wang, 2003
), Bufo boreas, Bufo terrestris and Bufo
woodhousei (Secor and Faulkner,
2002
), Ceratophrys cranwelli
(Powell et al., 1999
) and
R. catesbeiana (Busk et al.,
2000
). Anderson and Wang (2003) fed adult B. marinus rat
pups equal in mass to 8.5% of toad body mass and observed a 2.8-fold increase
in
O2. The
larger postprandial increase in
O2 observed in
this study for B. marinus (3.9-fold) simply reflects their larger
meals (15% of body mass), and that the postprandial metabolic response
increases with meal size (Secor and
Faulkner, 2002
). For B. boreas, B. terrestris
and B. woodhousei the digestion of cricket meals equaling 10% of body
mass generated an average threefold increase in
O2, and
mass-specific SDA values (kJ kg-1) similar to those experienced by
the non-estivating species of this study
(Secor and Faulkner,
2002
).
The three- and fourfold increases in
O2 observed for
adult R. catesbeiana are well matched to meal sizes of 10%
(Busk et al., 2000
) and 15%
(this study) of body mass, respectively. Interestingly, under similar
experimental treatment (rodent meals 17% of body mass at 30°C), C.
cranwelli experienced only a fourfold increase in metabolic rate, less
than half of the magnitude of response observed for C. ornata in this
study. One explanation for this difference in the metabolic response is the
much smaller body size of C. cranwelli (mean=23.2 g) used in the
Powell et al. (1999
) study
compared to C. ornata of this study, and the findings that the scope
of postprandial peak
O2 increases
with body mass (Secor and Diamond,
1997
; Secor and Faulkner,
2002
).
The significantly greater SDA of the estivating species compared to the
non-estivators is due to the estivators' lower SMR, higher postprandial peak
in O2, and
longer duration of significant metabolic response. In combination, these
differences account for a 55% greater SDA for the estivating species compared
to the non-estivators. In a similar fashion, sit-and-wait foraging snakes that
feed infrequently possess SMR that are 50% less, postprandial peak
O2 that are 25%
greater, and twice the duration of the SDA response compared to frequently
feeding, active-foraging species (Secor
and Diamond, 2000
). Thereby, independent of meal size and body
temperature, sit-and-wait foraging snakes experience an SDA that is 80%
greater than that of active foraging species. Connected by natural long
episodes of aphagia, estivating anurans and sit-and-wait foraging snakes
appear to exhibit parallel metabolic responses to fasting and feeding.
Proximate mechanisms of intestinal regulation
As quantified, intestinal nutrient uptake capacity is the product of small
intestinal mass and mass-specific rates of nutrient uptake. For the three
estivating species, the significant postprandial increase in their intestinal
uptake capacity is a shared function of an averaged 120% increase in nutrient
uptake rates and a 160% increase in small intestinal mass. Although matched by
family, the non-estivating species exhibit much more modest postprandial
increases in both nutrient uptake (26±4% increase) and intestinal mass
(49±11% increase). Whereas intestinal function and morphology
contribute relatively equally to the upregulation of intestinal performance
for estivating anurans, the relative contribution of nutrient uptake is much
greater for infrequently feeding snakes. The five- to 30-fold increase in
nutrient uptake capacity that infrequently feeding snakes experience with
feeding is predominately due to an averaged sevenfold increase in uptake
rates, and to a lesser extent, to the doubling of small intestinal mass
(Secor and Diamond, 2000).
Hypertrophy of the intestinal mucosa is largely responsible for the
postprandial increases in small intestinal mass for both snakes and anurans.
After feeding, intestinal enterocytes of the infrequently feeding sidewinder
rattlesnake Crotalus cerastes increase in width and volume by 75% and
400%, respectively (Secor et al.,
1994
), increases that are similar to those experienced by
estivating anurans (Fig.
6).
Hypertrophy of enterocytes generates a lengthening of the intestinal villi
and a resulting increase in functional surface area. The role of increasing
luminal surface area to upregulate intestinal performance is evident for the
infrequently feeding Burmese python Python molurus. The python's
eightfold postprandial increase in nutrient uptake rates is contributed to by
a 40% increase in villus length and a 4.8-fold increase in microvillus length
(Lignot et al., 2005). From
electron micrographs of intestinal microvilli of a fasted and a fed
individual, I did not observe any significant postprandial change in
microvillus length for any of the species
(Fig. 9). Among the six
species, microvilli of fasted and fed individuals averaged 1.38±0.39
µm and 1.48±0.24 µm in length, respectively. Hence, anurans
apparently elevate intestinal performance by increase luminal surface area at
the villus level and potentially by increasing nutrient transport activity
and/or density at the cellular level.
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The adaptive advantage of gut regulation
For infrequently feeding snakes, the fasting-related downregulation of gut
performance is proposed to be an adaptive mechanism to reduce metabolic rate
and thus energy expenditure between their infrequent meals. An objective of
this study was to determine whether anurans that estivate and are aphagic for
long intervals exhibit similar integrated responses. For three anuran species
that estivate, fasting was characterized by the significant downregulation of
intestinal performance and relatively low rates of metabolism. Compared to
similar-sized members of the same family that do not estivate, the estivating
species possess intestinal uptake capacities and SMR's that are, respectively,
40% and 62% lower. In considering that whole-animal rates of metabolism
represent the summed metabolic output of all tissues, a downregulated gut
would contribute to their lower SMR. The ecological significance of a lower
SMR is apparent when calculating the longevity without feeding of P.
adspersus with R. catesbeiana. Assuming equal body size (250 g),
3% body fat, and an average body temperature of 25°C, an inactive,
non-estivating, P. adspersus would theoretically be able to survive
145 days on its fat stores alone, more than twice the duration (60 days)
predicted for an inactive R. catesbeiana. Possessing lower SMR would
similarly allow B. alvarius and C. ornata to survive longer
without food than B. marinus and L. pentadactylus.
For estivating anurans, the lowered metabolic rate and downregulation of
intestinal performance with fasting can be viewed as a precursor to the
further depression of their physiology with estivation. The depression in
metabolic rate with estivation observed in this study and others have been
attributed to reductions in cellular activity, including decreases in RNA
synthesis, protein synthesis, enzyme activity, ion pumping and protein
phosphorylation (Guppy and Withers,
1999; Storey,
2002
). The respective 45% and 74% decline in enterocyte
performance for C. ornata and P. adspersus following
laboratory-induced estivation likely contributes to the concurrent decrease in
metabolic rate. In addition to the intestine, other tissues (i.e. stomach,
liver, kidneys) are also expected to be downregulated during estivation
because of their lack of use. For the anuran, Neobatrachus centralis,
the 67% decline in liver protein synthesis during estivation contributes to
the 55% decrease in liver metabolism, which is responsible for 5% of the
reduction in whole-animal metabolism
(Fuery et al., 1998
). Organs
such as the heart and lungs may remain unregulated given that cardiac and
pulmonary performance are still maintained during estivation for P.
adspersus (Loveridge and Withers,
1981
).
The reduction in metabolic rate during estivation will serve to further
extend an anuran's survivorship. The decrease in metabolism observed in this
study for C. ornata and P. adspersus would theoretically add
another month to their dormancy. Loveridge and Withers
(1981), studying the metabolic
rate of dormant and cocooned P. adspersus, estimated that a 500 g
individual should be able to survive dormant for almost 9 months on 15 g of
body fat at 20°C. Although I did not measure the metabolic rate and
intestinal performance of estivating B. alvarius, which unlike C.
ornata and P. adspersus do not form a cocoon, I would expect
this species to exhibit similar adaptive responses to estivation.
In light of the advantages of a downregulated gut during fasting, there are
two potential setbacks. First is the added cost of upregulation, which would
contribute to the larger postprandial metabolic responses of estivating
species compared to non-estivating species. For the infrequently feeding
P. molurus it is estimated that the upregulation of the
gastrointestinal tract is equivalent in cost to approximately 5% of the
ingested energy (Secor, 2003).
For an estivating anuran, the first meal would require the upregulation in
structure and function of the GI tract; the intestine alone increases sixfold
in mass and triples in nutrient uptake rates. Second, the potential reduction
in digestive performance and efficiency with the first post-estivation meal.
Following 3 months of laboratory estivation, passage time for the first meal
doubles for Cyclorana alboguttata, although without any loss of
energy extraction efficiency (Cramp and
Franklin, 2003
). For a migratory bird (Sylvia
atricapilla) that has not fed for several days, refeeding is
characterized by an initial low rate of nutrient assimilation
(Karasov and Pinshow, 2000
).
Following 10 months of estivation, an anuran's first meal will probably
require more energy and time to process, with or without a reduction in
extraction efficiency.
Further insights in amphibian digestive physiology
Interest in amphibian physiology has had a recent reinsurgence with the
global realization that amphibian populations and species are rapidly
disappearing (Stuart et al.,
2003). While there are several proposed causes for the demise of
amphibian populations (parasites, UV radiation, pesticides, herbicides), they
are all based on a common mechanism, the inability of the amphibian's
physiology to cope with the new and in many cases unnatural environmental
perturbation. Exploring the mechanisms by which amphibians can endure natural
perturbation and regulate tissue performance would provide insights into their
ability to survive environmental challenges. The findings of this and other
studies spark many questions regarding amphibian physiology, but the following
four are relevant to their digestive physiology.
First, given the energetic advantage of gut downregulation, why don't all
anurans widely regulate intestinal performance with feeding and fasting, as
observed for the three estivating species? To borrow from the explanation on
the narrow regulation of intestinal performance by frequently feeding snakes,
it may in fact be energetically inefficient for a more frequently feeding
anuran to be constantly up- and downregulating the gut with each frequent meal
(Secor, 2001). By maintaining
the gut in a state of readiness, the frequent meals can be digested rapidly
and efficiently. Field data on feeding frequency and energetics would be
valuable in constructing energy models that demonstrate the benefits in
linking feeding ecology with gut regulation.
Second, how general is the combination of depressed metabolism and
downregulated gut performance among estivating amphibians? Among practically
all estivating amphibians studied, estivation is characterized by a depression
in metabolic rate (Pinder et al.,
1992; Guppy and Withers,
1999
). One study that has documented a change in gut structure or
function with estivation, found a 85% decline in small intestinal mass for
estivating Cyclorana alboguttata
(Cramp and Franklin, 2003
).
Further studies on other estivating amphibians (i.e. Scaphiopus,
Neobatrachus, Amphiuma, Siren) will determine the extent that metabolic
depression is aided by gut downregulation.
Third, hibernation is another life-history feature of amphibians that
entails long episodes of aphagia. To what extent does the amphibian intestine
respond to hibernation? Metabolic depression during hibernation is largely
achieved by the substantial decrease in body temperature
(Storey and Storey, 1990). In
addition, the downregulation of the gut could supplement the depressed
metabolism, allowing the animal to survive longer on endogenous energy stores.
This question could be addressed by comparing metabolic rates and intestinal
morphology and function at various times during hibernation.
Fourth, what are the mechanisms that underlie the regulation of intestinal
performance for anurans? Tissue, cellular and molecular studies could
ascertain the cascade of events that result in the enterocyte hypertrophy and
increase in nutrient uptake with feeding, and the subsequent atrophy and
downregulation with estivation. Combined with studies on the cellular
mechanisms of metabolic depression
(Storey, 2002), an integrated
picture could emerge on the estivating-induced reduction in tissue function
and metabolism.
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References |
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