Energy metabolism in orchid bee flight muscles: carbohydrate fuels all
1 Department of Ecology, Evolution and Marine Biology, University of
California Santa Barbara, Santa Barbara, CA 93106-9610, USA
2 Department of Zoology, University of British Columbia, Vancouver, BC,
Canada V6T 1Z4
3 Biology and Wildlife Department, University of Alaska, Fairbanks, AK
99775, USA
4 Smithsonian Tropical Research Institute, Balboa, Republic of
Panama
* Author for correspondence (e-mail: suarez{at}lifesci.ucsb.edu)
Accepted 6 July 2005
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Summary |
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Key words: respiratory quotient, hovering flight, Euglossine bee, enzymes, glycolysis, flux, carbohydrate
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Introduction |
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A question that arises is whether the picture that has emerged concerning
the design and function of pathways of energy metabolism in honeybees also
applies to other species of bees. Although some insect species appear to fuel
flight with carbohydrate only, others can use both carbohydrate and fat, while
still other species rely primarily on amino acids
(Storey, 1985). The evolution
of metabolic fuel choice in insect flight muscles is not well understood;
however, much of the interspecific variation in this appears to be related to
factors such as diet, foraging ecology and dispersal. More than three decades
ago, Crabtree and Newsholme
(1972a
,b
)
conducted comparative biochemical studies that have since become part of the
foundation for our understanding of the diversity of fuel use in flying
insects. On the basis of maximum enzyme activities they reported in honeybees
and bumblebees, RQ values of 1.0 reported by other investigators (e.g.
Rothe and Nachtigall, 1989
),
and measurements of hemolymph sugars (Blatt
and Roces, 2001
; Gmeinbauer
and Crailsheim, 1993
), it is now assumed that, in general, bees
use carbohydrates to fuel flight. These, as well as the greater sensitivity of
CO2 analyzers compared with O2 analyzers, have led
researchers (including ourselves) in recent years to measure metabolic rates
only as CO2 production rates
(
CO2), assuming
that these equal O2 consumption rates
(
O2). The great
diversity in metabolic organization known amongst insects as well as the
limited number of species of bees subjected to detailed study warrant caution
in this regard.
Inspired by the pioneering work of Casey, Ellington and colleagues
(Casey and Ellington, 1990;
Casey et al., 1992
,
1985
), we embarked upon studies
of the allometric scaling of energy expenditure during flight
(Darveau et al., 2005a
) and
the biochemistry underlying such scaling
(Darveau et al., 2005b
;
Suarez et al., 2005
) in
Panamanian orchid bees. Orchid bees consist of more than 190 species belonging
to five genera and range from about 50 to >1000 mg in body mass. Given the
diversity in fuel use among flying insects, we considered it essential, as
part of our research program, to examine enzymatic capacities and rates of
fuel oxidation during flight within this clade to test the hypothesis that
carbohydrate oxidation serves as the main source of energy during flight. The
interspecific data reported herein, consisting of enzyme
Vmax values measured in vitro, as well as
metabolic rates and RQ
(
CO2/
O2)
values estimated in vivo, include nine species belonging to four of
the five extant genera. They provide support for the hypothesis that orchid
bee flight muscles have evolved to specialize in the use of carbohydrate
oxidation to supply their energetic requirements.
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Materials and methods |
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Respirometry measurements
Measurements of metabolic rate during flight were conducted immediately
after capture to minimize time-dependent changes in motivation. Smaller
species, up to 400 mg in body mass, were flown in a 0.5 l flask with sidearm,
while a similar flask of 1 l capacity was used for larger species. Air was
drawn into the flasks through perforated rubber stoppers and out through the
sidearms through Tygon® tubing at a rate of 1.5 l min-1 by a
FOX flow-through field respirometry system equipped with a Sable Systems
(Henderson, NV, USA) CA-2A CO2 analyzer. The CO2
analyzer was calibrated daily using a 5.03% CO2 span gas. Flight
durations of 2-3 min, facilitated by slight shaking and tilting of the flasks
when bees attempted to land, were sufficient to yield steady-state rates of
O2 consumption and CO2 production. Data acquisition and
analysis were performed using Datacan (Sable Systems). Because of the lower
limit to the sensitivity of our O2 analyzer, individuals of less
than 100 mg body mass often yielded
O2 values that
were difficult to reproduce. Therefore, all
O2 values from
individuals of <100 mg mass were discarded.
Enzyme activities
Other bees collected for enzyme measurements were frozen at -80°C,
shipped in dry ice to the laboratory, and stored at -80°C until
measurements were conducted. Individual thoraxes were minced with scissors and
homogenized in 19 volumes of ice-cold buffer. All further manipulations were
carried out in glass or plasticware cooled in crushed ice. The homogenization
buffer used on samples used for the measurement of hexokinase (HK),
phosphofructokinase (PFK), glycerol 3-phosphate dehydrogenase (GPDH), citrate
synthase (CS) and cytochrome oxidase (COX) consisted of 25 mmol l-1
Tris-potassium phosphate, pH 7.8 at 4°C, 2 mmol l-1 ethylene
diamine tetra-acetic acid (EDTA), 5 mmol l-1 dithiothreitol (DTT),
1 mmol l-1 fructose 6-phosphate, 3.5 mmol l-1 glucose
6-phosphate and 0.5% (v/v) Triton X-100. The use of phosphate buffer and
inclusion of sugar phosphates served to stabilize PFK activity that would
otherwise have been lost (Suarez et al.,
1996; Wegener et al.,
1986a
). The homogenization buffer used for samples designated for
measurement of glycogen phosphorylase (GP), trehalase (TR), and
phosphoglucoisomerase (PGI), consisted of 25 mmol l-1 Hepes, pH 7.3
at 4°C, 2 mmol l-1 EDTA, 5 mmol l-1 DTT and 0.5%
(v/v) Triton X-100. Minced thoraxes were homogenized three times for 10 s at
30 s intervals, using a Polytron homogenizer with a small tip (Brinkmann
Instruments, Rexdale, ON, Canada). Homogenates were then sonicated using a
Micro Ultrasonic Cell Disrupter (Kontes, Mandel Scientific, Guelph, ON,
Canada), again three times for 10 s, at 30 s intervals. Homogenates were
centrifuged (Jouan MR 1812, Winchester, VI, USA) for 5 min at 8000
g at 4°C, and the supernatants used for assays. To ensure
that these procedures resulted in complete extraction of membrane-bound
enzymes (e.g. trehalase), preliminary studies were conducted to compare enzyme
activities in uncentrifuged homogenates and supernatant fractions. Activities
obtained were equal, showing that extraction of all enzymes was complete.
Enzyme activities were measured in duplicate using a Perkin-Elmer Lambda 2
UV-Visible spectrophotometer (Norwalk, CT, USA) equipped with a Lauda
circulating water bath (Brinkman Instruments) adjusted to maintain cuvette
temperatures (monitored using a Cole-Parmer temperature probe) at 37°C.
HK, PFK, GPDH, PGI, TR, GP reactions were monitored by following the rate of
appearance or disappearance of reduced nicotinamide adenine dinucleotide
(NADH) or nicotinamide adenine dinucleotide phosphate (NADPH) at 340 nm using
a millimolar extinction coefficient () of 6.22. The CS reaction was
monitored 5,5' dithiobis-2-nitrobenzoic acid (DTNB) at 412 nm using
=13.6. The COX reaction was measured by monitoring oxidized cytochrome
c at 550 nm using
=29.5. Control (background) rates, obtained
without one substrate (indicated below), were measured and subtracted from
rates obtained with all substrates present.
Assay conditions and substrate concentrations required to elicit Vmax were as follows: HK: 50 mmol l-1 Hepes, pH 7.0, 5 mmol l-1 D-glucose (omitted from control), 4 mmol l-1 ATP, 10 mmol l-1 MgCl2, 100 mmol l-1 KCl, 0.5 mmol l-1 NADP+, 5 mmol l-1 DTT, 1 U glucose 6-phosphate dehydrogenase. PFK: 50 mmol l-1 Tris-HCl, pH 8.0, 5 mmol l-1 fructose 6-phosphate (omitted from control), 10 mmol l-1 MgCl2, 100 mmol l-1 KCl, 2 mmol l-1 ATP, 0.15 mmol l-1 NADH, 0.01 mmol l-1 fructose 2,6-bisphosphate, 5 mmol l-1 DTT, 1 U aldolase, 5 U triosephosphate isomerase, 5 U glycerol 3-phosphate dehydrogenase. GPDH: 50 mmol l-1 imidazol, pH 7.0, 1 mmol l-1 dihydroxyacetonephosphate (omitted for control), 0.15 mmol l-1 NADH. CS: 50 mmol l-1 Tris-HCl, pH 8.0, 0.5 mmol l-1 oxaloacetate (omitted for control), 0.3 mmol l-1 acetyl-CoA, 0.1 mmol l-1 DTNB. COX: 50 mmol l-1 potassium phosphate, pH 7.5, 0.05 mmol l-1 reduced cytochrome c. TR: 50 mmol l-1 potassium phosphate, pH 6.6, 1.1 mmol l-1 MgCl2, 0.5 mmol l-1 NADP, 1.1 mmol l-1 ATP, 10 mmol l-1 trehalose (omitted for control), 2.5 U of hexokinase and glucose 6-phosphate dehydrogenase. GP: 100 mmol l-1 potassium phosphate, pH 7.4, 2 mg ml-1 glycogen, 0.5 mmol l-1 NADP+, 4 µmol l-1 glucose 1,6-biphosphate, 2 mmol l-1 AMP, 10 mmol l-1 MgCl2, 10 U phosphoglucomutase and 2.5 U glucose 6-phosphate dehydrogenase. PGI: 50 mmol l-1 Tris-HCl, pH 8.0, 0.5 mmol l-1 fructose 6-phosphate, 0.5 mmol l-1 NADP+, 2.5 U glucose 6-phosphate dehydrogenase. All chemicals were from Sigma Chemical Company.
Respiration rate measurements in vitro
As individual bees do not possess sufficient flight muscle mitochondria for
isolation, we used crude homogenates of individual thoraxes to measure rates
of substrate oxidation in vitro. Bees were captured in the field and
placed in a refrigerator at 4°C until used for measurements. Before
preparing the thoraxes for homogenization, individual bees had to be warmed up
until leg movements were noticeable. For reasons that remain unknown, this
warm-up step was required prior to dissection and thoracic homogenization for
O2 consumption to occur. Preparation of homogenates from cold
thoraxes often resulted in no detectable respiration. After each thorax was
dissected from the insect, further manipulations were performed on ice.
Individual thoraxes were minced with scissors and homogenized in 19 volumes of
ice-cold 10 mmol l-1 Tris, pH 7.4, 1 mmol l-1 EGTA, 250
mmol l-1 sucrose, using a single, 10 s, low speed homogenization
using a Polytron homogenizer (Brinkman Instruments) with a small tip.
Rates of mitochondrial respiration in the crude thoracic homogenates were
measured at 37°C in a 1.6 ml water-jacketed Gilson glass chamber, equipped
with a Clark-type O2 electrode (YSI, Yellow Springs, OH, USA). The
assay buffer, consisting of 10 mmol l-1 Tris, pH 7.4, 1 mmol
l-1 EGTA, 25 mmol l-1 KH2PO4, 154
mmol l-1 KCl, was equilibrated with room air to an oxygen content
of 406 nmol O ml-1 (Reynafarje
et al., 1985) before measurements. After the addition of 50 µl
of homogenate, 10 µl of 1 mol l-1 pyruvate (or 10 µl 5 mmol
l-1 palmitoyl L-carnitine) and 10 µl 1 mol
l-1 proline were added and respiration was initiated by adding 20
µl 40 mmol l-1 ADP.
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Results and discussion |
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The biochemical data (Tables
2 and
3) reveal high enzymatic
capacities for carbohydrate oxidation. High citrate synthase (a Krebs cycle
enzyme) and cytochrome oxidase (a respiratory chain enzyme) activities are not
unexpected, given the high rates of aerobic metabolism required to support
flight and the high mitochondrial volume densities previously reported
(Casey et al., 1992). High
hexokinase activities, indicating high capacities for hexose sugar
phosphorylation, range from values of about half to up to twofold higher than
those found in honeybees (Suarez et al.,
1996
). GPDH, an enzyme catalyzing a near-equilibrium reaction
required for the maintenance of high cytosolic NAD+/NADH (as part
of the glycerol 3-phosphate shuttle;
Crabtree and Newsholme, 1975
;
Sacktor, 1976
), occurs at
higher Vmax values than all the other glycolytic enzymes
measured. Also consistent with high glycolytic capacities are the high
Vmax values for PGI, a glycolytic enzyme catalyzing a
near-equilibrium reaction (Staples and
Suarez, 1997
), and PFK, an allosteric enzyme catalyzing a
nonequilibrium reaction (Wegener et al.,
1986a
,b
).
Hydroxyacyl-CoA dehydrogenase, an enzyme involved in fatty acid oxidation
(Crabtree and Newsholme, 1975),
was not detectable in thoracic extracts of any species. Homogenates prepared
for measurement of mitochondrial O2 consumption displayed the
capacity to oxidize pyruvate, but not palmitoyl L-carnitine
(Fig. 1). Proline was included
in assays when testing for pyruvate or palmitoyl L-carnitine
oxidation, based on the results of preliminary studies (R. K. Suarez,
unpublished observations) with mitochondria isolated from honeybee flight
muscles. These revealed that, although pyruvate or proline alone are not
oxidized at significant rates, pyruvate plus proline support high rates of
coupled, state 3 (i.e. ADP-stimulated) respiration. Such results are
interpreted in terms of proline having an anaplerotic role (i.e. the
augmentation of Krebs cycle intermediates) required for high rates of Krebs
cycle activity (Sacktor and Childress,
1967
). Malate, however, is unable to serve such a role (R. K.
Suarez, unpublished observations), in contrast with mitochondria isolated from
locust flight muscles (Suarez and Moyes,
1992
), and so was not used in the studies reported here. The rates
measured in 21 individuals of eight species (Eg. sappharina, Eg.
imperialis, Ef. chrysopyga, Ef. schmidtiana, El. bombiformis, El.
nigrita, El. cingulata, El. meriana) were mass-independent and averaged
22.54±5.02 µmol O2 g-1 thorax min-1
(means ± S.D.).
|
|
The great consistency between the results presented herein and those
obtained with honeybees warrant the proposal of a hypothetical metabolic
scheme that combines features based on the results of various studies
(Fig. 2). The anaplerotic role
of proline in honeybees, also shown in Fig.
2, is further supported by results showing its depletion from
hemolymph during flight (Micheu et al.,
2000). Another reaction with such a role is that catalyzed by
pyruvate carboxylase, an enzyme found at high activities in honeybee flight
muscles (Crabtree et al.,
1972
; Tu and Hagedorn,
1992
). This intramitochondrial enzyme catalyzes the carboxylation
of pyruvate to oxaloacetate and is allosterically activated by acetylCoA. The
diagram also shows fat body as the source of trehalose
(Becker et al., 1996
), and
trehalase activity occuring in both intracellular as well as extracellular
compartments (Becker et al.,
1996
; Brandt and Huber,
1979
).
In conclusion, apart from the anaplerotic role played by proline,
carbohydrate oxidation predominates as the major source of energy for flight
in bees. We have found no evidence to indicate that fatty acid oxidation plays
a role in fueling flight. These findings provide added justification for the
measurement of
CO2 values and
the assumption that RQ=1.0 in studies of flight energetics in bees. Given the
use of fats by many other taxa of flying insects, one is led to ask why the
biochemical machinery required to use such an energy-rich substrate has
apparently been discarded by this lineage. Unlike the honeybees, orchid bees
do not store honey in hives and do not display clear eusociality. Observed
behaviour ranges from being solitary, as observed in some Eufriesea
and Euglossa, to being weakly social in Eulaema, to
`threshold eusociality' in Euglossa
(Roubik and Hanson, 2004
).
Despite this range of behaviours, reliance upon carbohydrate oxidation by
flight muscles is a feature common to all bee species studied, thus far. The
wasps from which bees evolved are represented by extant relatives that prey on
other insects. However, flight metabolism in wasps has not been examined in
detail. Clearly, the evolution of fuel use in flying insects is a subject that
warrants further investigation.
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Acknowledgments |
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Footnotes |
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