Gastric function and its contribution to the postprandial metabolic response of the Burmese python Python molurus
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
(e-mail: ssecor{at}biology.as.ua.edu)
Accepted 19 February 2003
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Summary |
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Key words: reptile, snake, Python molurus, digestion, stomach, gastric pH, specific dynamic action
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Introduction |
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It has therefore been suggested that the python's large postprandial
metabolic response is attributed to the digestion of large intact meals (up to
100% of body mass), their relatively low standard metabolic rate (SMR) above
which SDA is quantified, and the added cost of upregulating their quiescent
guts immediately following feeding (Secor
and Diamond, 1995). Understandably, the digestion of large intact
meals would significantly contribute to the python's large SDA response. The
digesta exiting the stomach consists of a soup-like chyme for practically all
vertebrates, whereas the physical state of food entering the stomach differs
dramatically among species. Humans swallow small macerated pieces of highly
digestible food, most carnivores swallow small intact prey that have been
crushed or pieces of meat torn from larger prey, and many reptiles (lizards,
turtles, and crocodilians) commonly ingest crushed or fragmented animal or
plant material. In stark contrast, snakes swallow only intact prey and must
delegate to the stomach the whole job of breaking down that prey before its
passage into the small intestine. Therefore, for snakes, the digestion of a
meal requires a relatively larger effort by their stomachs. Given that gastric
function (i.e. acid and enzyme secretion) is energetically demanding
(Reenstra and Forte, 1981
;
Helander and Keeling, 1993
),
the gastric breakdown of the python's meal may occur at a relatively high cost
(compared with that of other carnivores) and thus explain their comparatively
larger SDA.
Studies that have attempted to elucidate the relative contribution of the
various components of digestion to SDA have divided SDA into `mechanical SDA'
and `biochemical SDA' (Tandler and Beamish,
1979,
1980
;
Jobling and Davies, 1980
;
Carefoot, 1990
). Mechanical SDA
represents the cost of physically processing the food (i.e. chewing,
swallowing and peristalsis), whereas biochemical SDA is the postabsorptive
cost of assimilation (including nutrient transport and protein and tissue
synthesis). Finding that meal passage represents only 812% of SDA,
whereas protein synthesis contributes as much as 44% to SDA, researchers have
concluded that postabsorptive costs (biochemical SDA) dominate SDA
(Jobling, 1981
;
Brown and Cameron, 1991
;
Lyndon et al., 1992
). Contrary
to this conclusion, python SDA may be largely dominated by preabsorptive
costs.
The goal of the present study was to ascertain information on gastric digestion and its energetic cost for the Burmese python. I hypothesized that gastric breakdown of the large intact meals consumed by Burmese pythons would incur a substantial energetic expense and thus be an important contributor to their SDA. The aims of this study were to: (1) document the gastric breakdown of an intact meal; (2) profile the postprandial pattern of intragastric pH; (3) assess changes in the postprandial metabolic response to decreasing gastric workloads and (4) estimate the relative contributions of gastric performance, protein synthesis and gastrointestinal upregulation to the python's SDA. As shall be shown, pythons maintain a highly acidic environment within their stomach during digestion, the cost of which may dominate their SDA.
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Materials and methods |
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Rate of gastric digestion
I used two methods to examine the rate at which ingested rat meals were
broken down within the python's stomach and passed into the small intestine.
First, four pythons (449±60 g) were fed one or two intact rats equaling
28.6±0.7% of their body mass. These snakes were maintained at
2730°C and were x-rayed each day for 6 days following feeding in
order to visualize the breakdown of the skeleton of the ingested rat(s) within
the python's stomach. Second, 36 pythons (882±37 g) were fed rat meals
(13 rats) equaling 25.7±0.6% of their body mass. Following
feeding, snakes were maintained at 2730°C and sacrificed at 12 h
and at 1, 2, 3, 4, 6 and 14 days postfeeding (38 snakes per time). For
each snake, stomach contents were weighed and compared with the original wet
mass of the ingested meal in order to quantify the percentage of the ingested
meal still remaining within the stomach.
Gastric pH
I monitored gastric pH of seven pythons (1250±130 g) following their
ingestion of a single intact rat equaling 24.7±1.7% of the snake's body
mass. Stomach pH was measured using an infant gastric pH electrode (model
91-0011; Synectics Medical Inc., Irving, TX, USA) sutured to the head of the
ingested rat. The electrode cable extended from the snake's mouth and was
sutured to the side of the snake immediately posterior to its head. A
reference electrode (model 4011; Synectics Medical Inc.) was attached to the
dorsum of each snake, and both electrodes were connected to a pH monitor
(Digitrapper Mk II, Synectics Medical Inc.). Soon after feeding and for the
remainder of the measurements, snakes exhibited no observable discomfort to
the cable extending from their mouth or to the electrode attached to their
back. Position of each pH electrode was checked by x-ray. Prior to use, each
pH electrode was calibrated using commercial pH buffers of pH 1.07 and 7.01
(Synectics Medical Inc.). Following feeding, pH was recorded sporadically at
24 h intervals for up to 14 days while snakes were maintained at
2328°C. Measurements ended when either the snake was able to
dislodge the probe from its mouth (one case) or when stomach pH returned to
its initial level (approximately 7.5). Data from all snakes were grouped into
3 h intervals (5.6±0.2 snakes per interval) beginning at the time of
feeding.
Feeding treatments to assess gastric workload
I assessed the effects of different gastric workloads on SDA by feeding
pythons one of five meal treatments (46 snakes per treatment) and
measuring their postprandial metabolic responses. Meal treatments included
normal intact rats and the following four meals designed to reduce the
workload of the stomach while still inducing full postgastric responses:
steak, ground rat, liquid meal and ground rat infused directly into the
proximal small intestine. Five snakes (2390±340 g) consumed 23
intact rats whose combined mass equaled 25.0±0.1% of their body mass.
Steak meals (25.3±0.2% of body mass) were made from slabs of lean
sirloin steak (3.8% fat) rolled into a tube with either a rat or chicken head
attached to one end to entice pythons (2320±430 g; N=6) to
consume them. Six snakes (2430±190 g) were fed meals (25% of body mass)
of finely ground pre-killed rats by forcing the meal through a tube inserted
into the snake's esophagus. The liquid meal (25% of body mass) was formulated
to match the nutrient composition of an intact rat and consisted (by mass) of
70% mammalian Ringer's solution (composition of mammalian Ringer's solution in
mmol l1: NaCl 128, KCl 4.7, CaCl2 2.5,
KH2PO4 1.2, MgSO4 1.2, NaHCO3 20,
pH 7.4, 290 mosmol l1), 15% casein (C-7078; Sigma, St Louis,
MO, USA), 12% homogenized chicken fat and 3% D-glucose (G-8270; Sigma). The
liquid meal was similarly gavaged into the esophagus of four pythons
(5240±510 g).
Ground rat was infused through rubber catheters surgically inserted into the proximal end of the small intestine of four pythons (1670±280 g). To implant catheters, snakes were anesthetized with halothane (Halocarbon Laboratories, River Edge, NJ, USA), their ventral midsection scrubbed with a topical antiseptic (Betadine solution; Purdue Frederick Co., Norwalk, CT, USA), and a 6-cm incision was made between the ventral scales and the first set of lateral scales at a site approximately 65% of the distance from the snout to the cloaca. The incision was retracted open and a small hole was made in the proximal end of the small intestine just distal to the pyloric sphincter and the junction with the pancreaticobiliary duct. A 10-cm rubber catheter (8 mm diameter) was inserted through the hole, extended 2 cm downstream into the intestinal lumen and attached to the intestinal wall by a series of `purse-string' 4-0 silk sutures. The other end of the catheter was exteriorized through a small incision in the snake's body wall and sutured to lateral scales. The incision through the body wall was closed with an inner (muscular layer) and outer (scales) set of interrupted sutures (3-0 Vicryl; Ethicon Inc., Somerville, NJ, USA), followed by an application of New-Skin® (Medtech Laboratories Inc., Jackson, WY, USA). Immediately following surgery, each snake was given a single injection of antibiotic (1 ml kg1 enrofloxacin; Baytril, Bayer Co., Shawnee Mission, KS, USA) and analgesic (0.5 mg kg1 flunixin meglamine; Phoenix Pharmaceutical Inc., St Joseph, MO, USA). Snakes recovered from anesthesia within 1 h and were allowed one month of recovery before the start of the experiment. The ground rat meal was infused at 6-h intervals over a period of 5 days, such that the combined mass of the infusate was equivalent to 25% of the snake's body mass.
Measurements of oxygen consumption and quantification of SDA
I measured rates of oxygen consumption
(O2) of pythons
using closed-system respirometry as described by Vleck
(1987
) and Secor and Diamond
(1997a
). Snakes were placed
into individual respirometry chambers (939 liters) and maintained
within an environmental chamber at 30°C. Each respirometry chamber was
constructed with an incurrent and excurrent air port, each attached to a
three-way stopcock. For each metabolic trial, a 50-ml gas sample was withdrawn
from each chamber, the chambers were then sealed (closing the incurrent and
excurrent stopcocks), and a second gas sample withdrawn 0.51 h later
from the reopened excurrent air port. Gas samples were pumped through a column
of H2O 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 h1) and
mass-specific (ml g1 h1) rates of
O2 consumption corrected for standard temperature and pressure.
Each SDA trial began by measuring the
O2 of fasted
snakes once or twice a day for 3 days. For each snake, I assigned the lowest
measure of its
O2 during those
days as its standard metabolic rate (SMR). In this and previous studies
(Secor and Diamond, 1997a
),
the lowest measures of
O2 were commonly
recorded during the morning (06.0008.00 h), a time when pythons were
least active. Following SMR measurements, snakes were fed (or infused) and
metabolic measurements were continued at 12-h intervals for 2 days and
thereafter at 1-day intervals for up to 7 days. For each trial, I quantified
the following six variables as described and illustrated by Secor and Diamond
(1997a
): `SMR', as described
above; `peak
O2', the highest
recorded
O2
during digestion; `factorial scope of peak
O2', calculated
as peak
O2
divided by SMR; `duration', time from feeding when
O2 was no longer
significantly greater than SMR; `SDA', the total energy expended above SMR
during the duration of significantly elevated
O2, quantified
as kJ and kJ kg1; and `SDA coefficient', SDA quantified as a
percentage of the energy content of the meal. I calculated SDA (kJ) from the
extra oxygen consumed above SMR over the duration of significantly elevated
O2, assuming
that 19.8 J are expended per ml of O2 consumed (Gessaman and Nagy,
1988). Energy content of each meal was calculated as the product of meal wet
mass and the energy equivalent of that meal (kJ g1 wet
mass). Energy equivalent of the intact rat, steak and ground rat meals were
8.0 kJ g1 wet mass, 6.2 kJ g1 wet mass and
8.0 kJ g1 wet mass, respectively, as determined by bomb
calorimetry, and 8.0 kJ g1 wet mass for the liquid diet,
assuming 17.6 kJ g1 of casein (protein), 39.3 kJ
g1 of fat and 17.6 kJ g1 of glucose
(Schmidt-Nielsen, 1997
).
Statistical analyses
Postfeeding changes in stomach content (analyzed as calculated percentages
and actual mass of stomach content) were evaluated using analysis of
covariance (ANCOVA) with body mass as a covariate. A repeated-design analysis
of variance (ANOVA) was applied to test for significant effects of sampling
time on gastric pH and
O2 for each meal
treatment. To test the effects of meal treatment on metabolic variables, I
used ANCOVA (body mass as a covariate) for whole-animal measures and ANOVA for
mass-specific measures. Because of the significant variation in body mass
among the five meal treatments, I recalculated whole-animal measures of SMR,
peak
O2 and SDA
of each snake assuming a body mass of 2400 g. Adjusted values were calculated
from allometric equations presented in table 2 from Secor and Diamond
(1997a
), assuming mass
exponents of 0.7, 0.9 and 1.01, respectively, for SMR, peak
O2 and SDA. In
conjunction with ANOVA and ANCOVA, post-hoc pairwise mean comparisons
(TukeyKramer procedure) were used to compare treatments (sampling times
or meal type). I present the P value results of ANOVA and ANCOVA and
significant pairwise mean comparisons. The level of 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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Gastric pH
Feeding triggered a rapid decrease in gastric pH
(Fig. 3). During the first 12 h
after feeding, gastric pH dropped (P<0.0001) from 7.5 to 2.9,
representing, on average, more than a 10-fold increase in intragastric
[H+] every 3 h. By 24 h postfeeding, gastric pH had declined to
approximately 2, and for the next 57 days held steady between 1.1 and
1.8 (mean pH during that time was 1.52±0.05). After the meal had passed
from the stomach (usually 68 days after feeding), gastric pH increased
at a rate that mirrored the rapid postfeeding decrease, such that over a span
of only 1824 h gastric pH had returned to 7.5. The individual variation
in the time it took gastric pH to return to initial levels (712 days
postfeeding) is explained by the differences among snakes in relative meal
size (19.732.9% of body mass), as larger meals took more time to digest
and thus induced longer episodes of acid production
(Fig. 4).
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Varying gastric workload and SDA
For each meal type,
O2 (ml
g1 h1 or ml h1) varied
significantly (all P<0.0001) among pre- and postfeeding samples
(Fig. 5). Whereas mass-specific
SMR (ml g1 h1) varied significantly
(P=0.008) among meals, whole-animal measures of SMR (ml
h1; recorded or adjusted to a body mass of 2400 g) did not
differ (Table 1). Pythons
reached peaks in
O2 at 12 h
(infused ground rat), 36 h (intact rat, steak and ground rat) or 48 h (liquid
meal) postfeeding. Peak
O2 varied
significantly (all P<0.0001) among meal treatments as peak values
during the digestion of intact rats were significantly (all
P<0.007) greater than during the digestion of ground rat, which
was greater (all P<0.004) than that during the digestion of the
infused ground rat (Table 1).
The factorial scope of peak
O2 (peak
O2/SMR) was
highest for the digestion of intact rats, significantly (P=0.005)
lower for the digestion of the ground rat and even lower (both
P<0.05) for the digestion of the liquid meal and
intestinal-infused ground rat (Table
1). The duration of significantly elevated
O2 was 8 days
for the digestion of the intact rat meals and 6 days for the other four meal
treatments (Table 1).
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SDA differed significantly (all P<0.0001) among meal treatments (Table 1). Although not differing between the intact rat and steak meals, SDA (adjusted to 2400 g) decreased significantly (P<0.013) with the digestion of ground rat and decreased even further (both P<0.003) during the digestion of the liquid meal and intestinal-infused ground rat. By infusing ground rat directly into the small intestine, thereby bypassing the workload of the stomach, SDA was reduced to one-third of that generated by the intact rat meals. Whereas SDA coefficients (SDA expressed as a percentage of the ingested meal energy) likewise did not differ between the intact rat and steak meals, values were significantly (both P<0.001) lower with the ground rat meal, decreasing again (P=0.022) with the liquid meal and declining even further (P=0.021) with the intestinal-infused ground rat meal (Table 1).
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Discussion |
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Gastric breakdown
Following gastric upregulation during the first 12 h after feeding,
secreted HCl and the protease pepsin begin to digest away the rat's head and
anterior thorax. Concurrently, gas begins to build up within the body cavity
of the ingested rats, a phenomenon that was also observed by Blain and
Campbell (1942) in digesting
boa constrictor (Boa constrictor) and indigo snake (Drymarchon
corais couperi). Once HCl and pepsin have breached the rat's body cavity,
the gas is released and gastric distention is relaxed. Breakdown of each rat
proceeded from its head to its tail (rats were all swallowed head first), as
it was continuously pushed towards the more distal portion of the stomach. The
last materials to exit the stomachs were mats of hair, suggesting that the
indigestible hair is either selectively held back, giving priority to the
passage of more nutritional and digestible material, or is simply more
difficult to pass through the pylorus.
In an x-ray study evaluating body temperature effects on gastric digestion,
Skoczylas (1970a) observed
rapid decomposition of ingested frogs within the stomachs of grass snakes
(Natrix natrix) maintained at 25°C. Following their consumption
of frog meals equaling 20% of body mass, N. natrix had cleared their
stomachs within 3 days, which is considerably faster than the 57 days
it took pythons to empty their stomachs. Plausible explanations for these
differences are that N. natrix had consumed relatively smaller meals,
their frog meals were digested more rapidly compared with rat meals due to a
thinner integument, they had consumed a single frog whereas two-thirds of the
pythons had consumed at least two rats (a single large prey item may pass
faster than multiple smaller prey items) and, as a frequent feeder, N.
natrix are able to initiate gastric digestion faster than the
infrequently feeding P. molurus, which must first upregulate gastric
function before digestion can commence. In support of this last point, 24 h
following consumption of a common meal size (25% of body mass), frequently
feeding snake species pass twice the percentage of ingested prey (30%) from
their stomachs than do infrequently feeding species (15%;
Secor and Diamond, 2000
).
Gastric pH
During fasting, the quiescent stomach of pythons apparently does not
secrete acid, as is evident by the slight alkalinty of the gastric lumen
immediately after swallowing the rat meal. The presence of the meal within the
stomach (if not sooner) triggers the secretion of H+,
Cl and enzymes from the oxyntic cells of the gastric
epithelium. This causes a rapid decrease in luminal pH to a stable level of
approximately 1.5. Once the meal passes from the stomach, acid production
ceases and luminal pH returns to around 7.5. Similarly, following ingestion of
a frog meal, gastric pH of N. natrix decreases from 7.2 to 3.3 within
4 h, a response noticeably faster than that of P. molurus
(Skoczylas, 1970b). For
turtles, lizards, alligators and other snake species, gastric pH of fasting
individuals ranges between 7 and 8 and declines to a range of 1.54
during digestion (Blain and Campbell,
1942
; Coulson et al.,
1950
; Wright et al.,
1957
; Fox and Musacchia,
1959
).
The pre- and postprandial profile of gastric pH of pythons (and other
reptiles) is markedly different from that of mammals. In contrast to the
slight alkalinity of the fasting python's stomach, mammals maintain a highly
acidic environment (pH 1.13) within their stomachs between bouts of
digestion (Youngberg et al.,
1985; Evans et al.,
1988
; Cilluffo et al.,
1990
; Viani et al.,
2002
). Whereas pythons experience a dramatic postprandial decrease
in gastric pH, the luminal pH of mammal stomachs increases rapidly after
feeding to range between 3 and 6, presumably as the ingested meal buffers the
gastric acid (Savarino et al.,
1988
; McLauchlan et al.,
1989
; Cilluffo et al.,
1990
). Within a few hours after feeding, intragastric pH of
mammals drops as acid production, which has increased 20-fold, overwhelms the
buffering capacity of the food, which is then being passed through the pyloric
sphincter into the small intestine
(Fordtran and Walsh,
1973
).
The duration of gastric acid and enzyme production is a function of meal
size, meal composition and body temperature. For pythons, increasing meal size
by 65% (from 19.7% to 32.9% of body mass) resulted in a 50% increase in the
duration that gastric pH was maintained at 1.5
(Fig. 4). Decreasing the
structural composition of the python's meal results in an apparent decrease in
gastric workload and acid and enzyme secretion
(Fig. 5;
Table 1). Intuitively, intact
meals possessing a hard exo- or endoskeleton would require more time and
effort to digest and pass than fragmented and/or soft-bodied food items. The
turtle Kinixys spekii and the toad Bufo marinus required
more time and energy, respectively, to digest millipedes and superworms
(Zophobas larva), both possessing a chitinous exoskeleton, than to
digest soft fungi and earthworms (Hailey,
1998; Secor and Faulkner,
2002
). There is a direct relationship between body temperature and
rates of chemical reactions; therefore, decreasing body temperature decreases
rates of acid and enzyme secretions and thus increases the duration of
digestion (Skoczylas, 1970a
;
Stevenson et al., 1985
). The
snakes used to monitor gastric pH were maintained at a lower body temperature
(2328°C) than those used to evaluate gastric digestion
(2730°C) and consequently experienced longer bouts of
digestion.
Cost of gastric digestion
The results of this study suggest that as the workload of the python's
stomach is reduced, so is the cost of digestion
(Fig. 5; Table 1). One means to reduce
gastric workload is to reduce meal size; for pythons, smaller meals are
digested faster and incur a lower SDA and SDA coefficient
(Secor and Diamond, 1997a).
Another means (employed in this study) is to reduce the structural integrity
of the meal while maintaining a constant meal size, thereby accelerating its
passage into the small intestine. The rolled steaks lacked the outer
integument and bones of an intact rat. The ground rat was finely pureed,
thereby greatly increasing its surface area for acid and enzymatic
degradation, and the liquid diet was already equivalent to or beyond the state
of particle reduction characteristic of the chyme exiting the stomach. Whereas
the steak meal passed from the stomach faster than intact rats but did not
produce a lower SDA, both the ground rat and liquid diet reduced gastric
resident time and SDA. In bypassing the stomach and infusing ground rat
directly into the small intestine, the generated SDA was 33% of that resulting
from the digestion of intact rat meals.
The large reduction in SDA when the functions of the stomach are bypassed
suggests that gastric digestion is apparently an expensive component of the
python's SDA. Contributing to this cost are three well-cited activities of the
stomach; motility, enzyme production and acid secretion. Contractions of the
stomach's smooth muscles serve to churn and grind the ingested meal, thereby
facilitating contact of the food with enzymes and HCl, and to drive chyme
through the pyloric sphincter. For non-mammalian species, a single cell type,
the oxyntic cell, is responsible for the secretion of pepsinogen and HCl
(handled by the chief and parietal cells, respectively, in mammals;
Helander and Keeling, 1993).
Secreted pepsinogen, when exposed to a luminal pH of 23.5, is cleaved
to the active proteolytic enzyme pepsin. Pepsin begins the process of protein
digestion by acting on collagen and hydrolyzing proteins. HCl is formed from
Cl and H+; Cl is passively
released from oxyntic cells whereas H+ is actively pumped from
cells by the ATP-driven H+/K+-exchanger
(H+/K+-ATPase or proton pump;
Forte et al., 1980
).
While motility and pepsinogen production undoubtedly both contribute to the
cost of gastric digestion, at least five lines of evidence emphasize the cost
of acid production: (1) pythons maintain an intragastric pH of 1.5 in spite of
the large buffering capacity of the rat meals for 57 days; (2) the
production of such a quantity of HCl requires the proton pumps of the oxyntic
cells to move H+ from the cytosol into the gastric lumen against a
concentration gradient in excess of a million-fold
(Helander and Keeling, 1993);
(3) the proton pumps operate via the hydrolysis of ATP with a
stoichiometry of one H+ pumped per ATP hydrolyzed
(Reenstra and Forte, 1981
;
Norberg and Mårdh,
1990
); (4) the gastric parietal cells of mammals contain the
highest concentration of mitochondria (3444% by volume) compared with
any other mammalian cell type (Helander
and Hirschowitz, 1972
;
Helander et al., 1986
), and,
in a preliminary study, I found python oxyntic cells to be 40% mitochondria by
volume; and (5) acid secretion is absolutely dependent upon oxygen delivery
(Forte et al., 1975
;
Berglindh, 1984
). Collectively,
these findings indicate that pythons expend considerable amounts of cellular
energy via aerobic metabolic pathways to generate the vast quantity
of HCl necessary to digest their large intact meals.
Components of SDA
One goal of this project was to ascertain the relative contribution of pre-
and postabsorptive activities to the python's SDA. To begin, I calculated that
a 1 kg python digesting a 250 g rat (25% of snake body mass) would experience
an SDA of 600 kJ based on the published regression equation: log SDA = log
body mass x 1.01 0.25 (table 2 in
Secor and Diamond, 1997a). I
next estimated the cost of gastric performance as 330 kJ (55% of SDA) based on
the differences between the SDA resulting from digesting intact rat meals (528
kJ kg1) and that generated by the intestinally infused
ground rat meals (175 kJ kg1) and considering that the
infused ground rat meals may not have fully generated all postgastric
activities (therefore decreasing the assumed cost of gastric function).
Lacking, at least from the infused ground rat response, is the production and
secretion of bicarbonate solution by the pancreas and small intestine in
response to the introduction of the acidic chyme from an ingested meal.
Following feeding, pythons upregulate the performance of their dormant guts
in order to digest their meals (Secor and Diamond,
1995,
1997b
). To quantify the cost
of gastrointestinal (GI) upregulation, I first estimated that the difference
in SMR over a 24-h period (time taken to upregulate the gut) between a 1 kg
python with a quiescent gut (from table 2 in
Secor and Diamond, 1997a
) and
a 1 kg python with an upregulated gut (calculated from the SMR of frequently
feeding snakes that maintain an upregulated gut;
Secor and Diamond, 2000
) was
equivalent to 12 kJ. Next, I calculated that the cost of the postprandial
increase in stomach and intestinal mass for a 1 kg python was 17.5 kJ. I
assumed that these organs gained 2.5 g in protein with an energetic value of
44 kJ (17.5 kJ g1 protein) and that the cost of protein
synthesis is 0.4 kJ expended per kJ of protein synthesized
(Aoyagi et al., 1988
).
Combining metabolic and growth costs, the estimated cost of GI upregulation is
29.5 kJ, which is 4.9% of SDA (Fig.
6).
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To calculate the cost of post-absorptive protein synthesis, I estimated
that a 1 kg python would gain 103 g in body mass from the digestion of a 250 g
rat, assuming a growth efficiency (mass gained/mass consumed) of 41.2% (S. M.
Secor, personal observations; based on 40 pythons each consuming 10 meals,
each equaling 25% of body mass). Assuming that 22.5 g of the mass gained is
protein (the rest being water, bone and fat), with an energy value of 394 kJ
(17.5 kJ g1 protein), the cost of protein synthesis is
therefore 158 kJ (0.4 kJ g1 protein), equivalent to 26.3% of
SDA (Fig. 6). The combined cost
of gastric performance, GI upregulation and protein synthesis is 517.5 kJ,
86.2% of the projected SDA. The remaining component, 82.5 kJ, 13.8% of SDA,
would include the activity costs of the pancreas, gallbladder, liver, kidneys
and small and large intestines (Fig.
6). Also included are the costs stemming from the postprandial
increase in pulmonary and cardiovascular performance
(Secor et al., 2000).
A universal phenomenon following feeding is activation of tissues of the GI
tract to propel food through the oesophagus, stomach and intestines, to
secrete H+, enzymes, bicarbonate solution and bile, and to
hydrolyze and transport nutrients. Given that these activities are energy
consuming and are initiated prior to assimilating the meal, all digesting
organisms must first expend energy before harvesting and metabolizing any of
the ingested nutrients. Secor and Diamond
(1995) in discussing this
physiological phenomenon used the analogy `pay before pumping' in reference to
self-service fuel stations. Burmese pythons, like other organisms, must expend
energy (pay) to generate the HCl and pepsin necessary to initiate gastric
digestion, propel food into the small intestine, produce and release bile,
enzymes and bicarbonate, upregulate and operate intestinal hydrolases and
nutrient transporters, and lengthen intestinal microvilli before they can use
any of the ingested nutrients in metabolic pathways (pump). Assuming that none
of the ingested nutrients has crossed the intestinal wall by 18 h after
consuming an intact rat (25% of body mass;
Fig. 5), the minimal cost that
is paid upfront is 62 kJ, which is 5.5% of SDA. This start-up cost must be met
by endogenous energy stores, most likely by lipids mobilized from fat bodies.
Support for this response is the rapid postprandial 50-fold increase in plasma
triglycerides observed for Burmese pythons
(Secor and Nagy, 2000
).
The Burmese python's impressive upregulation of their GI tract following
feeding was earlier suggested as one of several important contributors to
their relatively large SDA (Secor and
Diamond, 1995). The combination of 5- to 20-fold increases in
small intestinal nutrient transport rates, up to a 3-fold increase in
pancreatic and intestinal enzyme activities, a doubling of small intestinal
mass, and a 5-fold increase in microvillus length was reasoned to have an
impact on their SDA (Secor and Diamond,
1995
,
1998
). As previously
calculated, the cost of GI upregulation may represent approximately 5% of the
python's SDA. Similarly it was concluded from metabolic measurements taken
during overlapping digestive bouts (thereby the gut was not allowed to down-
and upregulate performance) that the cost of postprandial GI upregulation for
the turtle Kinixys spekii and Burmese python is not large
(Hailey, 1998
;
Overgaard et al., 2002
).
Granted that the postprandial upregulation of the python's GI tract occurs at
some cost, it apparently does not dominate SDA.
Outlook on python gastric physiology
The findings of this project raise several interesting points warranting
future investigation of the python's gastric physiology. First, the python's
ability to activate and deactivate gastric function, well beyond that of
mammals, avails them as an excellent model to investigate the underlying
mechanisms involved in the regulation of gastric performance. For the python,
the intragastric presence of the meal undoubtedly triggers neural and
endocrine signals that stimulate the production of HCl and pepsin, and induces
gastric hypertrophy and motility. It is well known that the hormone gastrin
stimulates HCl production and gastric hypertrophy in mammals
(Walsh, 1994), although recent
attempts to assay gastrin in pythons using mammalian probes have been
unsuccessful (Secor et al.,
2001
). Of interest is determining whether pythons possess a
gastrin structurally distinct from that of mammals or employ a different
regulatory peptide to control gastric function.
Second, mammals characteristically maintain an acidic environment within their stomachs between digestive bouts, whereas for pythons intragastric pH is kept slightly alkaline between meals. It has been suggested that mammals maintain an acidic gastric lumen as a protective means against ingested bacteria and other pathogens. Therefore, are pythons susceptible to being colonized by pathogenic microorganisms or do they possess some alternative protective mechanism against them?
And third, given that their large postprandial metabolic response is dominated by gastric function, do pythons possess an oxidative capacity of their stomach and oxyntic cells of unprecedented magnitude for an ectothermic vertebrate? This is suggested from the effort they expend to generate the HCl that can maintain an intragastric pH of 1.5 for a week against the constant buffering actions of their large meal, to constantly produce pepsin and perhaps other enzymes during that interval, to propel a large amount of material through the pyloric sphincter, and to produce new mucosal cells. Thus, the stomach's capacity for high aerobic performance may be necessary for pythons to digest their large intact meals.
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Aoyagi, Y., Tasaki, I., Okumura, J.-I. and Muramatsu, T. (1988). Energy cost of whole-body protein synthesis measured in vivo in chicks. Comp. Biochem. Physiol. A 91,765 -768.[CrossRef][Medline]
Benedict, F. G. (1932). The Physiology of Large Reptiles with Special Reference to the Heat Production of Snakes, Tortoises, Lizards, and Alligators. Carnegie Inst. Wash. Publ. No. 425. Washington, DC: Carnegie Institution of Washington.
Berglindh, T. (1984). The mammalian gastric parietal cell in vitro. Annu. Rev. Physiol. 46,377 -392.[CrossRef][Medline]
Blain, A. W. and Campbell, K. N. (1942). A study of digestive phenomena in snakes with the aid of the roentgen ray. Am. J. Roentgenol. Radium Ther. 48,229 -239.
Brody, S. (1945). Bioenergetics and Growth. New York: Reinhold Publishing Company.
Brown, C. R. and Cameron, J. N. (1991). The relationship between specific dynamic action (SDA) and protein synthesis rates in the channel catfish. Physiol. Zool. 64,298 -309.
Carefoot, T. H. (1990). Specific dynamic action (SDA) in the supralittoral isopod, Ligia pallasii: identification of components of apparent SDA and effects of dietary amino acid quality and content on SDA. Comp. Biochem. Physiol. A 95,309 -316.
Cilluffo, T., Armstrong, D., Castiglione, F., Emde, C., Galeazzi, R., Gonvers, J.-J. and Blum, A. L. (1990). Reproducibility of ambulatory gastric pH recordings in the corpus and antrum. Scand. J. Gastroenterol. 25,1076 -1083.[Medline]
Coulson, R. A., Hernandez, T. and Dessauer, H. C. (1950). Alkaline tide in the alligator. Soc. Exp. Biol. Med. 74,866 -869.
Evans, D. F., Pye, G., Bramley, R., Clark, A. G., Dyson, T. J. and Hardcastle, J. D. (1988). Measurement of gastrointestinal pH profiles in normal ambulant human subjects. Gut 29,1035 -1041.[Abstract]
Fordtran, J. S. and Walsh, J. H. (1973). Gastric acid secretion rate and buffer content of the stomach after eating. J. Clin. Invest. 52,645 -657.[Medline]
Forte, J. G., Machen, T. E. and Öbrink, K. J. (1980). Mechanisms of gastric H+ and Cl transport. Annu. Rev. Physiol. 42,111 -126.[Medline]
Forte, T. M., Mache, T. E. and Forte, J. G. (1975). Ultrastructural and physiological changes in piglet oxyntic cells during histamine stimulation and metabolic inhibition. Gastroenterology 69,1208 -1222.[Medline]
Fox, A. M. and Musacchia, X. J. (1959). Notes on the pH of the digestive tract of Chrysemys picta.Copeia 1959,337 -339.
Geesaman, J. A. and Nagy, K. A. (1988). Energy metabolism: errors in gas-exchange conversion factors. Physiol. Zool. 61,507 -513.
Hailey, A. (1998). The specific dynamic action of the omnivorous tortoise Kinixys spekii in relation to diet, feeding patterns, and gut passage. Physiol. Zool. 71, 57-66.[Medline]
Helander, H. F. and Hirschowitz, B. I. (1972). Quantitative ultrastructure studies on gastric parietal cells. Gastroenterology 63,951 -961.[Medline]
Helander, H. F. and Keeling, D. J. (1993). Cell biology of gastric acid secretion. Baillière's Clin. Gastroenterol. 7,1 -21.[Medline]
Helander, H. F., Leth, R. and Olbe, L. (1986). Stereological investigations on human gastric mucosa: I. normal oxyntic mucosa. Anat. Rec. 216,373 -380.[Medline]
Janes, D. N. and Chappell, M. A. (1995). The effect of ration size and body size on specific dynamic action in Adélie penguin chicks, Pygoscelis adeliae. Physiol. Zool. 68,1029 -1044.
Jobling, M. (1981). The influences of feeding on the metabolic rate of fishes: a short review. J. Fish Biol. 18,385 -400.
Jobling, M. and Davies, P. S. (1980). Effects of feeding on metabolic rate, and the specific dynamic action in plaice, Pleuronectes platessa L. J. Fish Biol. 16,629 -638.
Kleiber, M. (1975). The Fire of Life. New York: Krieger.
Lyndon, A. R., Houlihan, D. F. and Hall, S. J. (1992). The effect of short-term fasting and a single meal on protein synthesis and oxygen consumption in cod, Gadus morhua. J. Comp. Physiol. B 162,209 -215.[Medline]
McLauchlan, C., Fullarton, G. M., Crean, G. P. and McColl, K. E. L. (1989). Comparison of gastric body and antral pH: a 24 hour ambulatory study in healthy volunteers. Gut 30,573 -578.[Abstract]
Norberg, L. and Mårdh, S. (1990). A continuous technique for analysis of stoichiometry and transport kinetics of gastric H, K-ATPase. Acta Physiol. Scand. 140,567 -573.[Medline]
Overgaard, J., Busk, M., Hicks, J. W., Jensen, F. B. and Wang, T. (1999). Respiratory consequences of feeding in the snake Python molurus. Comp. Biochem. Physiol. A 124,359 -365.
Overgaard, J., Andersen, J. B. and Wang, T. (2002). The effects of fasting duration on the metabolic response to feeding in Python molurus: an evaluation of the energetic costs associated with gastrointestinal growth and upregulation. Physiol. Zool. 75,360 -368.[CrossRef]
Pope, C. H. (1961). The Giant Snakes. New York: Knopf.
Reenstra, W. W. and Forte, J. G. (1981). H+/ATP stoichiometry for the gastric (K++H+)-ATPase. J. Membr. Biol. 61, 55-60.[Medline]
Savarino, V., Mela, G. S., Scalabrini, P., Sumberaz, A., Fera, G. and Celle, G. (1988). 24-hour study of intragastric acidity in duodenal ulcer patients and normal subjects using continuous intraluminal pH-metry. Dig. Dis. Sci. 33,1077 -1080.[Medline]
Schmidt-Nielsen, K. (1997). Animal Physiology, Adaptation and Environment. Cambridge: Cambridge University Press.
Secor, S. M. (2001). Regulation of digestive performance: a proposed adaptive response. Comp. Biochem. Physiol. A 128,565 -577.
Secor, S. M. and Diamond, J. (1995). Adaptive responses to feeding in Burmese pythons: pay before pumping. J. Exp. Biol. 198,1313 -1325.[Medline]
Secor, S. M. and Diamond, J. (1997a). Determinants of post-feeding metabolic response in Burmese pythons, Python molurus. Physiol. Zool. 70,202 -212.[Medline]
Secor, S. M. and Diamond J. (1997b). Effects of meal size on the postprandial responses in juvenile Burmese python (Python molurus). Am. J. Physiol. 272,R902 -R912.[Medline]
Secor, S. M. and Diamond J. (1998). A vertebrate model of extreme physiological regulation. Nature 395,659 -662.[CrossRef][Medline]
Secor, S. M. and Diamond, J. (2000). Evolution of regulatory responses to feeding in snakes. Physiol. Biochem. Zool. 73,123 -141.[CrossRef][Medline]
Secor, S. M. and Faulkner, A. C. (2002). Effects of meal size, meal type, body temperature, and body size on the specific dynamic action of the marine toad, Bufo marinus. Physiol. Biochem. Zool. 75,557 -571.[Medline]
Secor, S. M. and Nagy, T. (2000). Postprandial response of plasma lipids and the hormone leptin in pythons. Am. Zool. 40,1205 .
Secor, S. M., Hicks, J. W. and Bennett, A. F.
(2000). Ventilatory and cardiovascular responses of a python
(Python molurus) to exercise and digestion. J. Exp.
Biol. 203,2447
-2454.
Secor, S. M., Fehsenfeld, D., Diamond, J. and Adrian, T. E.
(2001). Responses of python gastrointestinal regulatory peptides
to feeding. Proc Natl. Acad Sci. USA
98,13637
-13642.
Skoczylas, R. (1970a). Influence of temperature on gastric digestion in the grass snake, Natrix natrix L. Comp. Biochem Physiol. 33,793 -804.[CrossRef]
Skoczylas, R. (1970b). Salivary and gastric juice secretion in the grass snake, Natrix natrix, L. Comp. Biochem. Physiol. 35,885 -903.[CrossRef]
Soofiani, N. M. and Hawkins, A. D. (1982). Energetic costs at different levels of feeding in juvenile cod, Gadus morhua L. J. Fish Biol. 21,577 -592.
Stevenson, R. D., Peterson, C. R. and Tsuji, J. S. (1985). The thermal dependence of locomotion, tongue flicking, digestion, and oxygen consumption in the wandering garter snake. Physiol. Zool. 58,46 -57.
Tandler, A. and Beamish, F. W. H. (1979). Mechanical and biochemical components of apparent specific dynamic action in largemouth bass, Micropterus salmoides Lacépède. J. Fish Biol. 14,343 -350.
Tandler, A. and Beamish, F. W. H. (1980). Specific dynamic action and diet in largemouth bass, Micropterus salmoides (Lacépède). J. Nutrit. 110,750 -764.[Medline]
Tandler, A. and Beamish, F. W. H. (1981). Apparent specific dynamic action (SDA), fish weight and level of caloric intake in largemouth bass, Micropterus salmoides Lacepede. Aquaculture 23,231 -242.
Viani, F., Verdú, E. F., Idström, J. P., Cederberg, C., Fraser, R., Fried, M., Blum, A. L. and Armstrong, D. (2002). Effect of omeprazole on regional and temporal variations in intragastric acidity. Digestion 65, 2-10.[CrossRef][Medline]
Vleck, D. (1987). Measurement of O2
consumption, CO2 production, and water vapor production in a closed
system. J. Appl. Physiol.
62,2103
-2106.
Walsh, J. H. (1994). Gastrointestinal hormones. In Physiology of the Gastrointestinal Tract (ed. L. R. Johnson), pp. 1-128. New York: Raven.
Wang, T., Morten, Z., Arvedsen, S., Vedel-Smith, C. and Overgaard, J. (2003). Effects of temperature on the metabolic response to feeding in Python molurus. Comp. Biochem. Physiol. A 133,519 -527.
Wright, R. D., Florey, H. W. and Sanders, A. G. (1957). Observations on the gastric mucosa of reptilia. Quart. J. Exp. Physiol. 42, 1-14.
Youngberg, C. A., Wlodyga, J., Schmaltz, S. and Dressman, J. B. (1985). Radiotelemetric determination of gastrointestinal pH in four healthy beagles. Am. J. Vet. Res. 46,1516 -1521.[Medline]
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