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First published online May 2, 2008
Journal of Experimental Biology 211, 1594-1602 (2008)
Published by The Company of Biologists 2008
doi: 10.1242/jeb.017244
Synergy and specificity of two Na+–aromatic amino acid symporters in the model alimentary canal of mosquito larvae
,
The Whitney Laboratory for Marine Bioscience, University of Florida, 9505 Ocean Shore Boulevard, St Augustine, FL 3208, USA
Author for correspondence (e-mail:
dmitri.boudko{at}rosalindfranklin.edu)
Accepted 17 March 2008
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Summary |
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 TOP Summary INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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Key words: insect, mosquito, essential amino acid, nutrient amino acid transporter, NAT, co-transporter, phenylalanine, tryptophan, monoamine neurotransmitter, malaria, Anopheles gambiae
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INTRODUCTION |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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). Accordingly,
interruption of an essential amino acid absorption mechanism in mosquito
larvae could be employed to reduce the vector component of disease
transmission in endemic areas. However, safe and effective application of this
attractive strategy requires comprehensive understanding of specific and
universal properties of essential amino acid transport in target and key
risk-of-exposure metazoan organisms.
Earlier studies documented the active absorption of essential amino acids
in mosquitoes (Uchida et al.,
2003; Uchida et al.,
2001; Uchida et al.,
1990) and other insects
(Caccia et al., 2005;
Castagna et al., 1997;
Giordana et al., 1989;
Nedergaard, 1972;
Wolfersberger, 2000). Using
genome data mining in combination with comparative phylogenetic analysis of
transporters in selected organisms with published genomes, we identified and
compared key families of secondary transporters that contribute to the amino
acid traffic network in metazoans (Boudko
et al., 2005c). Each identified family provides a unique but
complementary part in the contiguous traffic and balance of amino acids;
however, the molecular identity and phylogeny of a core mechanism for active
absorption of essential amino acids remained uncertain. An intriguing
paralogous expansion of `orphan' transporters was identified among insect
members of the sodium neurotransmitter symporter family (SNF; also known as,
solute carrier family 6; SLC6) (Boudko et
al., 2005a; Boudko et al.,
2005b; Boudko et al.,
2005c). Based on their phylogenetic closeness with characterized
neutral amino acid transporters – two from the tobacco hornworm larva,
Manduca sexta, msKAAT1 (Castagna
et al., 1998) and msCAATCH1
(Feldman et al., 2000); and
one from the yellow fever mosquito Aedes aegypti, aeAAT1 – we
anticipated a homologous physiological role for other members of the
identified group (Boudko et al.,
2005a). All of the identified genes appear to encode moderately
conserved, monovalent cation–amino acid symporters, which mediate the
uptake of a set of neutral amino acids, almost all of which are essential for
insects and other metazoans. Moreover, this insect-specific cluster neighbors
a cluster of B0 system transporters that mediates the absorption of
a broad spectrum of neutral amino acids in mammals
(Broer et al., 2006a); it also
neighbors a set of orphan Caenorhabditis elegans transporters,
several of which are extensively transcribed in the worm alimentary canal
(www.wormbase.org).
The entire group, designated as nutrient amino acid transporters (NATs),
represents a functionally segregated subfamily of SNF (SLC6)
(Boudko et al., 2005a;
Boudko et al., 2005b;
Boudko et al., 2005c).
Drosophila melanogaster, An. gambiae and Ae. aegypti have 7,
6 and 9 (possibly +1) NAT members, respectively, demonstrating strong
paralogous diversification and variation in gene numbers. This observation
suggests that a rapid duplication and functional specialization of NAT members
occurs (Boudko et al., 2005a).
The retention and consistent expansion of paralogous NATs seen from bacteria
to metazoans imply the conservation of a fundamental role of these
transporters during metazoan evolution. The NAT-SLC6 population appears to
have evolved as an integrated system that performs high-throughput absorption
of essential amino acids and their derivatives
(Boudko et al., 2005c).
Recently, we have cloned and characterized two NATs with unique transport
properties from An. gambiae larval midgut
(Assis et al., 2004;
Boudko et al., 2005b;
Meleshkevitch et al., 2006).
Both transporters mediate Na+- or K+-coupled
voltage-gradient-driven absorption of specific aromatic substrates. However,
AgNAT6 (AAT07965) preferably absorbs tryptophan and indole-branched
substrates, whereas AgNAT8 (AAN40409) preferably absorbs phenyl-branched
substrates. To determine the physiological significance of such an
extraordinary specialization we examined the relative distribution of these
aromatic NATs in the model system of the alimentary canal from mosquito
larvae. In addition, we analyzed the assembly and docking of the NATs in a
heterologous expression system, Xenopus laevis oocytes.
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MATERIALS AND METHODS |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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Antibody preparation
Antigen motif selection was based on antigenicity plots and the putative 3D
structure of AgNAT6 and AgNAT8 that were cloned and sequenced in our
laboratory (accessions AAT07965 and AAN40409). Selected motifs were unique
relative to each other, to the rest of AgSLC6 and to the Anopheles
protein database. Two synthetic oligopeptides, EQSLPRDRSLVRRMFDNVFS and
GPIDPATHYEYKKFIDED, corresponding to the C-terminal 20 and 18 amino acids of
AgNAT6 and AgNAT8, respectively, were synthesized at a 20 micromolar scale,
emulsified in Freunds complete adjuvant and used according to a 90-day
immunization protocol (Open Biosystems, Huntsville Alabama, USA; operating in
accordance with animal use regulations). Antibodies were prepared by
immunizing rabbits with these synthetic oligopeptides conjugated to keyhole
lympet hemocyanin (KLH). A separate rabbit antibody with low binding of
pre-bleed sera as tested by ELISA was selected for each immunization. The
immunization series included a primary injection, 14th day first boost and
28th day second boost injections of oligopeptides. Bleeds were collected every
28 days after the primary immunizations. The crude sera were affinity purified
against the respective immunizing peptides.
Heterologous expression of AgNATs in Xenopus oocytes
Oocytes were collected from live Xenopus laevis Daudin (Xenopus
Express Inc., Brooksville, FL, USA) under sterile aseptic conditions as
approved by the University of Florida (IACUC 6032). Frogs were anesthetized by
immersion in ice-cold 0.1% tricaine solution and 1 cm cuts were made on the
lower ventral abdomen from which eggs were harvested; operated frogs were
euthanized according to the approved protocol. Apparent stage V–VI
oocytes were isolated from egg clusters after treatment with a 2% collagenase
solution in Ca2+-free ND96 medium (96 mmol l–1
NaCl, 2 mmol l–1 KCl, 1 mmol l–1
MgCl2, 10 mmol l–1 Hepes, pH 7.4). Separated
oocytes were conditioned for a few hours in Ca2+ trace ND96 before
being used in experiments. cRNA was prepared from pXOOM plasmids of NAT clones
using a high yield capped RNA transcription kit, mMESSAGEmMACHINE®
(Ambion, Foster City, CA, USA) as described previously
(Boudko et al., 2005a;
Meleshkevitch et al., 2006).
Approximately 20 ng cRNA was injected into each oocyte using a capillary glass
micropipette attached to a Nanoliter 2000 injector (WPI, Sarasota, FL, USA).
Oocytes were incubated at 16°C for 4 days after which they were evaluated
for amino acid transport activity using a standard two-electrode voltage clamp
technique (Meleshkevitch et al.,
2006) followed by immunolabeling.
Isolation of membranes from larvae and oocytes
Cell membrane fractions were isolated from An. gambiae larvae and
X. laevis oocytes that had been injected with either distilled water
(control), AgNAT6 or AgNAT8 cRNA according to a modified Hill protocol
(Hill et al., 2005). Briefly,
tissues were suspended in homogenization buffer (HB; 250 mmol
l–1 sucrose, 5 mmol l–1 MgCl2, 10
mmol l–1 Hepes, pH 7.4) containing a 1:1000 dilution of
protease inhibitor cocktail (Sigma-Aldrich, St Louis, MO, USA). Larvae and
oocytes were homogenized using a glass Dounce or plastic pestle with a
disposable 1.5 ml tube, respectively, followed by ultrasound agitation.
Homogenates were centrifuged at 500 g for 5 min and the
supernatant was recovered. Pellets were resuspended in the same volume of HB
and processed as before. The second supernatant was combined with the first
one and the crude membrane suspension was overlaid on a discontinuous sucrose
gradient consisting of 20% sucrose–HB and 50% sucrose–HB, and
centrifuged at average Relative Centrifugal Field (RCFav) 111 000
g at 4°C for 30 min using a Beckman SW-41 rotor. The layer
visible at the gradient interface, representing the membrane fraction, was
collected using a micro-syringe, diluted threefold in HB, and recovered by
centrifugation at RCFav 111 000 g at 4°C for 30
min. The resulting membrane pellet was resuspended in HB with added protease
inhibitors and quantified for protein content using the Bio-Rad protocol
(Bradford, 1976).
SDS-PAGE and western blot analysis
Protein samples were treated with NuPAGE® LDS sample buffer
(Invitrogen, Carlsbad, CA, USA) and β-mercaptoethanol and heated at
70°C for 10 min. Samples (4 µg per lane) were separated under reducing
conditions on a denaturing 4–12% Bis–Tris polyacrylamide gradient
gel. Separated proteins were blotted onto nitrocellulose membranes (0.22
µm; Millipore, Billerica, MA, USA) using a tank transfer system. Blots were
stained with Fast Green to confirm equal loading and transfer and to visualize
lane boundaries; then they were cut into strips for probing with various serum
samples. Before being probed, blots were incubated in blocking buffer
containing 5% non-fat dry milk powder (Carnation®) in Tris-buffered saline
(TBS) for 1 h at room temperature. Blots were then incubated with AgNAT6 or
AgNAT8 antibody at a dilution of 1:500 in 2% dry milk–TBS with 0.05%
Tween-20 (TBST) overnight at 4°C. As a control, lanes with identical
membrane samples were incubated with pre-immunization serum. Nitrocellulose
membranes were washed in TBST and incubated with alkaline
phosphatase-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch, West
Grove, PA, USA) diluted at 1:1000 in 2% dry milk–TBST for 1 h at room
temperature and then washed in three changes of TBST for 10 min each, followed
by one 10 min wash in TBS. Bound antibodies were detected by an alkaline
phosphatase color precipitation reaction (Bio-Rad, Hercules, CA, USA).
Whole-mount immunolabeling
Oocytes were fixed in 4% paraformaldehyde (PFA) in 0.1 mol
l–1 PBS for approximately 3 h and rinsed with three changes
of ice-cold PBS for 5 min each. Fourth instar larvae were immobilized in
ice-cold PBS, injected with 4% PFA and dissected as described previously
(Meleshkevitch et al., 2006).
The protocols for immunolabeling of oocytes and larvae were similar. Fixed
oocytes and larvae samples were permeabilized in 0.1% Triton X-100 in PBS
(PBT) for 6–12 h at 4°C on a shaker. Next, the samples were
incubated in blocking solution (BS; 2% normal goat serum, 1% bovine serum
albumin in PBT) for 12–24 h at 4°C. The primary antibodies against
AgNAT6 or AgNAT8 were then added at a dilution of 1:100 in BS and left at
4°C on a shaker for a further 24 h. The preparations were rinsed four
times in BS for 30 min each, and incubated with Alexa Fluor 488-conjugated
goat anti-rabbit (GAR) secondary antibody (Invitrogen) at a dilution of 1:800
in BS overnight at 4°C. Preparations were subsequently labeled with
phalloidin–Rhodamine (Invitrogen; 1:250 in PBS) for 15 min at room
temperature to visualize muscle actin, and with DRAQ-5 (Biostatus Limited,
Shepshed, UK; 1:1000 in PBS) for 5 min to visualize cell nuclei. Then they
were washed and mounted in 3:1 glycerol:PBS on microscope slides. To prepare a
relative-fluorescence-intensity graph two selected images of whole-mount
labeled preparations of AgNATs from identical fourth instar larval stages were
scanned using the Plot Profile tool of the ImageJ 1.38 software package
(http://rsb.info.nih.gov/).
Apparent intensity values were copied to a SigmaPlot spreadsheet, normalized
and used to generate a graph.
Frozen section immunolabeling
A surgically isolated and paraformaldehyde-fixed alimentary canal was
incubated in a 30% sucrose–PBS solution for 12 h, saturated in
TissueTek® embedding medium and mounted at –20°C on a cryostat
base. Sections (15 µm) were prepared using a cryostat microtome. For
immunolabeling, the sections were warmed at room temperature, resaturated and
permeabilized in BS for 2 h. The slides were rinsed in PBT and incubated in a
humidified chamber with primary antibodies (1:50 in PBT) for 4 h at room
temperature. Sections were washed in PBT and incubated with Alexa Fluor
488-GAR secondary antibodies (Invitrogen; 1:500 in PBS) for 1 h at room
temperature. Subsequently, preparations were washed and incubated with
phalloidin–Rhodamine and DRAQ-5 as described above. Images of
immunolabeling were acquired using a Leica laser scanning confocal microscope
with laser excitation at 495 nm, 550 nm and 678 nm for fluorescein, Rhodamine
and DRAQ-5, respectively. Scanned frame stacks were reconstructed as
three-dimensional projections and selected planes were converted to BMP images
and assembled into final plates using the Corel Draw X3 software package
(Corel Corporation, Ottawa, ON, Canada).
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RESULTS |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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3). Strong bands of smaller molecular mass were present in the
membrane fractions from the heterologous expression system. These bands may
reflect labeling of incomplete NAT fragments, which could form upon
heterologous overexpression. Intense high molecular mass bands were also
observed between putative monomer and homodimers bands in heterologously
expressed but not in tissue samples. These bands may suggest accumulation of
abnormal peptide associations, e.g. oligomerization of mature NATs with small
molecular mass peptides such as the immature NAT fragments proposed above. The
differences in electrophoretic migration of heterologous and tissue fractions
were similar for AgNAT6 and AgNAT8 expression, suggesting generalization of
the protein maturation process and escalation of maturation problems upon
heterologous overexpression. The upper band of a putative dimer represents the
dominant form of AgNATs in epithelial tissue samples, whereas putative
monomers and incomplete dimers were prevalent in the heterologously expressed
samples.
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Localization of heterologously expressed AgNATs in Xenopus oocytes
No notable staining was observed in control, water-injected oocytes
(Fig. 2A,B) or in control
preparations treated with pre-immune sera (data not shown). By contrast,
Xenopus oocytes expressing AgNAT6 and AgNAT8 proteins for 3–4
days were specifically labeled with the transporter-specific antibodies. This
labeling corresponds to efficient expression and predominant incorporation of
the proteins in the oocyte plasma membranes
(Fig. 2D,E,G,H). DW-injected
control oocytes produced only minor responses on application of tryptophan or
phenylalanine (Fig. 2C). By
contrast, the immunolabeling of both AgNATs in Xenopus oocytes
unequivocally correlates with functional expression, as determined by robust
substrate-induced Na+ currents (100% tests, N20 for
each transporter; Fig. 2F,I)
and previously reported radiolabel uptake of substrates
(Meleshkevitch et al., 2006;
Boudko et al., 2005b) (and data
not shown). The aromatic amino-acid-induced responses revealed very specific,
consistent Na+ dependency, repeatable response kinetics and
maximums of substrate-induced currents
(Fig. 2F,I; summarized in
Fig. 3). A diagram constructed
by multi parameter sorting and Gaussian pick fitting of apparent transport
efficiencies showed a very clear gain of specificity for indole- and
phenyl-branched substrates in the AgNAT6 and AgNAT8 transporters
(Fig. 3; right row; blue and
red data sets, respectively).
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Localization of AgNATs in whole mounts of the larval alimentary canal
Simple whole-mount preparations of mosquito larval alimentary canal provide
unique opportunities to compare global patterns of spatial expression of the
transporters. Steady-state patterns of immunolabeling in the larval alimentary
canal were observed for both aromatic transporters (N=22;
Fig. 4A,B). Fluorescence
intensities were highest in the anterior and posterior regions of the
alimentary canal, suggesting a synergetic contribution of both NATs to
specific functions attributed to these epithelial regions
(Fig. 4C). Nevertheless, upon
closer comparison there were differences in the spatial distribution of the
two transporters (Fig. 4).
AgNAT6 was concentrated in the gastric caeca (GC) and central midgut (CM) as
well as the anterior part of the posterior midgut (PMG;
Fig. 4A). Less intense labeling
was observed in the anterior midgut (AMG) and Malpighian tubules (MT). AgNAT8
antibodies produced intensive labeling in the salivary glands (SG), GC and PMG
(Fig. 4B). Labeling of AgNAT8
was more intense in GC than in PMG, the opposite of AgNAT6 labeling in the
same areas. In addition, labeling for AgNAT8 in the PMG was notably more
posterior than that of AgNAT6 (Fig.
4C; N=8). AgNAT8 antibodies also labeled MT more
intensely than those against AgNAT6. AgNAT8 antibody produced very weak
labeling in the AMG and the large transitional region (CM) between AMG and PMG
(N=6) in contrast to AgNAT6 antibodies which labeled this region very
well. Specific labeling of AgNAT8 but not AgNAT6 was observed in the proximal
portion of the MTs, which may reflect the distinct specialization of
individual epithelial cells in this area
(Fig. 4).
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Localization of AgNATs in frozen sections of larval AC
The general pattern of immunolabeling in frozen sections
(Fig. 5) corresponded well to
the distribution of the two NATs in whole-mount preparations
(Fig. 4) suggesting that the
antibodies had permeated the tissues well in the latter. The higher resolution
analysis of frozen sections was sufficient to determine intracellular
distribution and plasma membrane localization of the aromatic AgNATs within
epithelial tissues. Specific polar sorting of the transporters was found in
both posterior and anterior regions of the larval alimentary canal. In PMG
both transporters were most prevalent in columnar cell apical membranes. In
AMG and CM labeling was distinct in both apical and basal membranes of the
epithelial cells. The labeling intensity was higher at the apical pole of AMG
and CM, with each transporter differing with respect to intensities and
spatial profiles (Fig. 5). The
SG cells showed strong expression and uniform basal localization of both
transporters (N=5). By contrast, in GC AgNAT6 was predominantly
associated with the apical membrane whereas AgNAT8 was associated with both
apical and basal membranes, with higher intensity in the latter. AgNAT8 was
also strongly expressed in basal membranes of the cardia and in apical
membranes of the principal cells of the proximal MT.
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DISCUSSION |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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;
Linser et al., 2007). The
alimentary canal epithelium in mosquito larvae consists of several thousands
of large cells that mediate mineral ion balance, digestion, nutrient
collection and metabolic waste emission. Thus it provides distinctive
advantages for the study of the integrative and cellular basis of mineral ion
and pH balances (Boudko et al.,
2001a; Boudko et al.,
2001b; Clark et al.,
2007; Okech et al.,
2008) as well as absorption of essential amino acids
(Boudko et al., 2005a;
Boudko et al., 2005b;
Boudko et al., 2005c). It has
an explicit regional pattern of functional specialization and provides an
outstanding opportunity for parallel studies of molecular, cellular and
systemic aspects of various membrane transporters
(Boudko et al., 2005a;
Meleshkevitch et al., 2006;
Rheault et al., 2007).
Specificity of aromatic NATs in the amino acids traffic network
Each cell in metazoan organisms depends on the enduring supply of essential
amino acids. Central to this supply network are the active initial absorption
of these nutrients through apical membranes of the alimentary canal epithelial
cells and their active re-absorption via plasma membranes of other
cells (Boudko et al., 2005c;
Broer, 2002). NAT-SLC6 members
(a.k.a. B0 system transporters) mediate this absorption by coupling
uphill transport of the essential amino acids to downhill co-transport
(symport) of alkali metal cations that is driven by ionic transmembrane
electrochemical gradients (Boudko et al.,
2005a; Broer et al.,
2006a). This secondary active absorption of essential amino acids
in insects is conducted by B0 system-like transporters with
specific substrate profiles. For example, AgNAT6 and AgNAT8, display little
overlap in substrate spectra (Fig.
3). Both transport aromatic substrates but AgNAT6 translocates
indole-branched substrates whereas AgNAT8 transports phenyl-branched
substrates (Fig. 2).
Duplication and specialization of aromatic NATs along with balanced relative
expression can enhance primary absorption from midgut lumen and can provide
for more precise cellular distribution of aromatic amino acids. Similar
duplication and specialization of aromatic NATs in parallel with the
extinction of complex, energy-consuming pathways for the synthesis of the
aromatic substrates appears to be a key trade-off in the evolution of
heterotrophy; however, it raises the question: exactly how do organisms
provide and regulate NATs in response to metabolic demands in specific organs,
tissues and cells?
Synergy of aromatic NATs in aromatic substrate absorption
The indole- and phenyl-branched amino acid transporters, AgNAT6 and AgNAT8,
respectively, share high expression and apical location in the PMG. In this
area the final stage of protein digestion produces elevated concentrations of
free amino acids. The apical membranes of PMG cells have very long, densely
packed microvilli which appear in light micrographs as a brush border
(Clements, 1992;
Zhang and Nichols, 1994) and
which greatly expands the absorptive surface area. Although significantly
higher transcript concentrations of AgNAT6 and AgNAT8 were detected in PMG by
in situ hybridization (Boudko et
al., 2005a; Meleshkevitch et
al., 2006), there was no direct evidence that the active nutrient
amino acid transport mechanism is restricted to the apical membranes there. We
provide here the first solid proof that two NAT members of the SLC6 family are
expressed in the apical membranes of PMG in a pattern which suggests strongly
that NATs act in synergy to ensure complete and effective apical absorption of
nutrient aromatic amino acids. The results also suggest that the PMG is a
primary site for the initial absorption of essential aromatic amino acids. In
a broader sense, the synergy between AgNATs 6 and 8 suggests that they play
universal, complementary role as essential substrate providers
(Boudko et al., 2005a). This
increased transcription and expression of aromatic NAT transporters in the PMG
most likely corresponds to a gain of function and/or increased turnover of the
proteins. Similar, elevated PMG transcriptions were found for other AgNATs and
a few NATs from different dipteran species, e.g. Ae. aegypti and
D. melanogaster (Boudko et al.,
2005b; Meleshkevitch et al.,
2006; Thimgan et al.,
2006). Several members of the mammalian NAT subfamily
(Boudko et al., 2005a;
Broer, 2006;
Hoglund et al., 2005) that is
adjacent to the insect NAT cluster (Boudko
et al., 2005a) are highly expressed in posterior parts of the
mammalian alimentary canal (Bohmer et al.,
2005; Broer et al.,
2006b) and have been shown to dock in apical membranes of
absorptive epithelia of kidney and small intestine
(Romeo et al., 2006;
Verrey et al., 2005).
Collectively, these data support the hypothesis that the NAT-SLC6 group is
evolving universally across the animal kingdom and providing a mechanism for
the active absorption of essential amino acids from the digestive tract lumen
across the plasma membranes of its epithelial cells. Based on the variety of
substrates for individual, characterized transporters (e.g. aromatic AgNATs
presented here) we propose that different members of the NAT SLC6 subfamily
evolved distinct patterns of selectivity and suggest that the entire group
appears to be evolving and working in synergy to ensure comprehensive
absorption of a required set of essential amino acid substrates.
Benefits from NATs duplication and morphological segregation
The results presented here raise two questions inherent to NAT regulation.
(1) Why do mosquitoes possess separate, selective transporters for indole- and
phenyl-branched substrates whereas mammals appear to absorb both of these
substrate groups via more universal transport mechanisms such as the
broad substrate spectra NATs of system B0 transporters? The
apparent benefit of such duplication is a reduction of substrate competition
for a single transporter. Mammalian transporters may operate in a contiguous
mode, clearing essential amino acids one by one in successive regions of the
relatively long intestine. By contrast, absorptive regions in the larval
alimentary canal are only 2–3 mm long and the absorption window is open
but a brief few minutes. Considering that concentrations of essential
substrates may be quite disproportionate, the bulk of minor substrates, such
as aromatic amino acids, could be lost to excretion while dominant substrates
occupy a universal transport mechanism. Thus, duplication followed by strong
specialization for the transport of indole- and phenyl-branched substrates is
the key adaptation, which leads to a more efficient profile of nutrient
absorption consistent with survival and rapid development of mosquito larvae
under the pressure of limited nutrients.
(2) Why do mosquitoes possess spatial differences in the relative expression of aromatic AgNATs in the posterior midgut rather than expressing both transporters in identical epithelial loci or cells? The answer appears to be that the separation may reduce AgNAT competition for electrochemical energy and inorganic cation driving forces. Perhaps for the same reason, the membrane sector through which AgNAT6 absorbs the typically scarcer tryptophan is remarkably larger than the sector through which AgNAT8 absorbs the three to four times more concentrated phenylalanine and tyrosine substrates (based on the frequency of these amino acids in protein sequences). More generally, modulation of the relative expression of NATs with distinct substrate profiles in the alimentary canal may be an important facet of physiological and genetic adaptation of an organism to a nutrient chain with a limited source of essential amino acids. It is therefore reasonable to propose that duplication, specialization, and morphological segregation are major steps in NATs adaptation and evolution.
Role of the AgNAT6–AgNAT8 duet in secretory epithelia
In addition to their role in housekeeping protein synthesis, insects use
large amounts of aromatic substrates during ecdysis
(Sugumaran, 2000), cuticle
hardening and tanning (Sugumaran and
Nelson, 1998; Vincent,
2002), egg chorion formation
(Li, 1994); wound healing
(Galko and Krasnow, 2004;
Shi et al., 2006), immunity
(Siva-Jothy and Thompson,
2002), melanotic encapsulation of pathogens
(Hillyer et al., 2003;
Koella and Sorensen, 2002),
neurotransmission (Caveney and Donly,
2002; Osborne,
1996) and hormonal signaling
(Gade and Goldsworthy, 2003;
Kelly et al., 1994). Hence the
demand for aromatic substrates is expected to be accompanied by an elevated
expression of aromatic NATs. High densities of both AgNATs were identified in
anterior domains of the larval alimentary canal, including salivary glands,
cardia, gastric caeca and some areas of AMG
(Fig. 4). Earlier we proposed
that the gain of aromatic NAT transcription in the anterior region of the
larval alimentary canal may support secretory functions of this region, e.g.
secretion of peritrophic polymer precursors and salivary enzymes
(Meleshkevitch et al., 2006).
The location of aromatic NATs on the basal membrane of epithelial cell in the
secretory region provides them with access to the large pool of free amino
acids in the hemolymph. Strong support for this hypothesis is the unequivocal
basal localization of both aromatic transporters in the salivary glands (Figs
5 and
6). Intriguing differences in
the expression of AgNAT6 and AgNAT8 are found in other anterior structures.
Thus AgNAT8, but not AgNAT6, is highly expressed in the cardia (Figs
5 and
6). This difference may reflect
the high consumption of phenyl-branched substrates associated with phenol
oxidase-mediated polymerization of peritrophic membrane peptides
(Tellam et al., 1999). The
opposite polarity of AgNAT6 and AgNAT8 suggests sorting of aromatic substrates
in the gastric caeca. Perhaps it favors the absorption of indole substrates in
parallel with the secretion of phenyl-branched components. The bipolar
expression of AgNAT6 in the anterior midgut may correspond to an extensive
turnover of alkalinization-maintaining components in this area. However, it
may also imply the lack of a protein polar docking mechanism in this area of
transient functional specialization. The variety of NATs expression has been
demonstrated in different mammalian tissues using PCR
(Broer et al., 2004;
Broer et al., 2006b;
Kowalczuk et al., 2005;
Takanaga et al., 2005a;
Takanaga et al., 2005b) and
antibodies (Romeo et al.,
2006). However, the alternative polarizations and spatial
expression patterns of NATs in the functionally different parts of the
metazoan alimentary canal are shown here for the first time. Although the
mechanisms of relative expression and alternative polar docking of NATs
remains to be clarified, the facts reported here establish the principal role
of two NATs in the redistribution of essential aromatic substrates. The
spatial tuning of expression and trafficking of individual NATs in the
epithelial tissue may be a particular example of a more fundamental mechanism
for functional pattern generation in the absorption of essential amino acid in
different metazoan tissues and cells.
|
10–20 kDa but not 50 kDa. The
upper bands are predominant in the gut tissues but not in the heterologous
expression system (Fig. 1)
suggesting that the putative homodimers are the prevalent structure for both
aromatic transporters in the gut tissues. The facts that symmetrical dimers
were identified in other member of SLC6 by similar analysis
(Hastrup et al., 2001), by
FRET (Bossi et al., 2007) and
upon crystallization of bacterial NAT AaLeuT
(Yamashita et al., 2005)
supports our hypothesis and suggests that dimer formation is conserved and
essential for the physiological role of SLC6 members. A detailed analysis
would be required to determine the exact nature and functional roles of
quaternary modifications of NATs.
Integration of AgNATs in ion recycling pathways
Our previous electrophysiological analysis showed that both aromatic AgNATs
use membrane potentials to drive Na+- or K+-coupled
amino acid symport (Boudko et al.,
2005a; Meleshkevitch et al.,
2006). Both of these ions are at low concentrations in the
freshwater habitat of An. gambiae larvae. The alkalinization to pH
>10 of larval AMG involves the luminal resorption of chloride anions and
secretion of strong cations, such as Na+ or K+
(Boudko et al., 2001a;
Corena et al., 2001). The high
pH has long been associated with digestion of nutrients and protection from
pathogens in mosquito larvae (Clements,
1992). It is reasonable to propose that AMG alkalinization is also
part of Na+ and K+ recycling, which provides NATs in the
PMG with an essential pool of alkali metal ions
(Fig. 6). As key users of
cation gradients in the PMG, NATs complete a systemic loop for cation
recycling between hemolymph and AMG. MT and the rectal gland are important
components of a mineral cation recycling framework in freshwater mosquitoes
(Smith et al., 2007) and may
support cation recycling under low nutrient conditions, when NAT activity
would be reduced. Alkali metal cations are replaced in the midgut lumen
through the alkalinization that is mediated by basal H+ V-ATPase in
synergy with putative cation secretion and anion absorption mechanisms
(Linser et al., 2007;
Zhuang et al., 1999). Upon
saturation of a luminal cation pool the NAT function can be energized by
apical H+ V-ATPase in the larval PMG. The H+ V-ATPase
hydrolyses ATP to move protons (H+) outwardly across the cell
membrane thereby hyperpolarizing the apical membranes; the resulting
transmembrane voltage generates electrophoretic forces sufficient for inward
movement of Na+ and perhaps K+ (considering that the
apical K+ reversal potential is relatively high) through apical
membranes of epithelial cells. NATs couple the translocation of alkali metal
cations to that of neutral amino acids. Thus, the H+ V-ATPase is
probably the most important source of electromotive force for the
electrochemical coupling of NATs to metabolic energy in mosquitoes, as it is
in caterpillars (Gluck, 1992;
Harvey, 1992;
Zhuang et al., 1999). However,
this primary membrane energization via proton efflux would be
incomplete without electrical coupling to secondary alkali metal translocation
that in turn is coupled by NATs to tertiary amino acid translocation. For
instance, the action of an electrophoretic cation/nH+ antiporter
coupled to a H+ V-ATPase activity would be essential to support
luminal secretion of sodium and potassium ions. Potentially
Na+,K+-ATPase can be involved as well in the localized
energization of NAT functions. Being located apically in the AMG and basally
in the PMG (Okech et al.,
2008) it would sustain an inward Na+ gradient that
would provide ions for Na+-coupled amino acid absorption in the
epithelial cells by exchanging intracellular Na+ for K+
(Castagna et al., 1998).
Indeed, studies on other insect species have demonstrated the interaction of
amino acid absorption with a basally located
Na+,K+-ATPase
(Castagna et al., 1998;
Goberdhan et al., 2005;
Mbungu et al.,
1995).Therefore, it seems reasonable that both,
Na+,K+-ATPase-energized and (Na+ or
K+), H+ antiporter-H+ V-ATPase-energized
mechanisms support NAT function.
LIST OF ABBREVIATIONS
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Acknowledgments |
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Footnotes |
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Present address: Department of Physiology and Biophysics, Rosalind Franklin
University, School of Medicine, North Chicago, IL 60064, USA
Present address: A. N. Belozersky Institute, Moscow State University,
Moscow 119899, Russia
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References |
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TOPSummaryINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONReferences |
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