Journal of Insect Physiology 59 (2013) 525–531
Contents lists available at SciVerse ScienceDirect
Journal of Insect Physiology
journal homepage: www.elsevier.com/locate/jinsphys
A metabolic pathway assembled by enzyme selection may support
herbivory of leaf-cutter ants on plant starch
Maurício Bacci Jr. a,⇑, Odair Correa Bueno b, André Rodrigues a, Fernando Carlos Pagnocca a,
Alexandre Favarin Somera a, Aline Silva c
a
b
c
Universidade Estadual Paulista, Instituto de Biociências de Rio Claro, Centro de Estudos de Insetos Sociais/Departamento de Bioquímica e Microbiologia, Brazil
Universidade Estadual Paulista, Instituto de Biociências de Rio Claro, Centro de Estudos de Insetos Sociais/Departamento de Biologia, Brazil
Departamento de Ciências Biológicas, Universidade Estadual de Santa Cruz, Brazil
a r t i c l e
i n f o
Article history:
Received 22 October 2012
Received in revised form 13 February 2013
Accepted 15 February 2013
Available online 14 March 2013
Keywords:
Amylase
Attini
Maltase
Mutualism
a b s t r a c t
Mutualistic associations shape the evolution in different organism groups. The association between the
leaf-cutter ant Atta sexdens and the basidiomycete fungus Leucoagaricus gongylophorus has enabled them
to degrade starch from plant material generating glucose, which is a major food source for both mutualists. Starch degradation is promoted by enzymes contained in the fecal fluid that ants deposit on the fungus culture in cut leaves inside the nests. To understand the dynamics of starch degradation in ant nests,
we purified and characterized starch degrading enzymes from the ant fecal fluid and from laboratory cultures of L. gongylophorus and found that the ants intestine positively selects fungal a-amylase and a maltase likely produced by the ants, as a negative selection is imposed to fungal maltase and ant a-amylases.
Selected enzymes are more resistant to catabolic repression by glucose and proposed to structure a metabolic pathway in which the fungal a-amylase initiates starch catalysis to generate byproducts which are
sequentially degraded by the maltase to produce glucose. The pathway is responsible for effective degradation of starch and proposed to represent a major evolutionary innovation enabling efficient starch
assimilation from plant material by leaf-cutters.
Ó 2013 Elsevier Ltd. All rights reserved.
1. Introduction
Mutualistic association is recognized as an important force
shaping biodiversity and co-evolution between organisms. It is
thought to be related to structure, phenotype and abundance of
species populations in many organism groups, such as plants (El
et al., 2009), insects (Nobre et al., 2010) and reptiles (Sinervo
et al., 2006).
The mutualism between ants in the tribe Attini and basidiomycete fungi has been subjected to an intense investigation regarding
evolution (Silva-Pinhati et al., 2004; Bacci et al., 2009), metabolism
(Semenova et al., 2011) and behavior (Mueller et al., 2004). Special
attention has been given to the most evolved lineage of this ant
tribe, the leaf-cutter ants which include some species with crop
pest status (Fowler and Forti, 1986).
Leaf-cutter ants cultivate the fungus Leucoagaricus gongylophorus (Silva-Pinhati et al., 2004) inside their nests on the foliar material that they cut and collect. Biochemical dependence in this
interaction is one the most preponderant factors in maintaining
⇑ Corresponding author. Address: UNESP-IB-CEIS, Avenida 24A, 1515 Bela Vista,
13506-900 Rio Claro, SP, Brazil.
E-mail address: [email protected] (M. Bacci Jr.).
0022-1910/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved.
http://dx.doi.org/10.1016/j.jinsphys.2013.02.007
the alliance between these organisms (Martin, 1970). One example
of this interaction is the fungal origin of proteinases (Boyd and
Martin, 1975) and pectinases (Ronhede et al., 2004; Schiøtt et al.,
2010) found in the fecal fluid of workers of the leaf-cutters ants
in different species.
The excretion of digestive enzymes in the fecal material is proposed as an adaptation evolved in species of insects that cultivate
fungi (Mishra, 1991). Such digestive enzymes are deposited by
Attini ants in order to stimulate the mutualistic fungus growth
on the substrate harvested by the insects (Boyd and Martin,
1975), generating mycelial mass and soluble sugars which ants
use for food (Silva et al., 2003). Martin (1974) argued that the
established portions of cultures of the mutualistic fungus produce
enzymes which are ingested and then expelled in the ants fecal
fluid on the newly introduced plant substrate inside the nest, so
the recently planted portions of the fungus culture finds the substrate previously degraded when it is inoculated on the foliar mass
by the ants. Consequently, the mutualistic fungus would be able to
grow more quickly and overcome other competitor microorganisms that live inside the nest. This was also recognized by Ronhede
et al. (2004) that demonstrated the capacity of workers to concentrate fungal enzymes in their intestines and to deposit them in regions where fast fungal growth was required. Thus, enzymes
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M. Bacci Jr. et al. / Journal of Insect Physiology 59 (2013) 525–531
circulation from the fungus to the ant intestine and then to the
plant material inside the nests is a significant factor in the metabolic integration between these mutualists.
The fecal fluid of leaf-cutter ants also contains appreciable
amounts of amylase activity digesting starch (Martin et al.,
1973). Degradation of starch from plant material inside the ant
nest is proposed as the major metabolic pathway generating glucose, which in turn is a key carbon source for the nutrition of ants
(Silva et al., 2003) and the mutualistic fungus (Siqueira et al.,
1998).
Amylases are enzymes which are widely distributed in insects
(Terra and Ferreira, 1994). Among Attini ants, amylases have been
detected in species of the genera Atta (Richard et al., 2005), Acromyrmex (Febvay and Kermarrec, 1983; Erthal et al., 2004), Cyphomyrmex, Apterostigma, Myrmicocrypta and Sericomyrmex (Martin
et al., 1973). Amylases are produced in glands of the ant digestive
apparatus, as well as in the salivary and mandibular glands and in
the medium intestine (Ayre, 1967; Febvay and Kermarrec, 1983).
Starch digestion by amylases also seems to occur in initial parts
of the intestine of hemipterans (Khan and Ford, 1967) and in the
midgut of coleopterans, dipterans and lepidopterans (Bolognesi
et al., 2008). In leaf-cutter ants there is evidence of starch digestion
by enzymes produced by the workers (Febvay and Kermarrec,
1983; Silva et al., 2003). However, despite of the major importance
of amylases for mutualists nutrition, its origin in fecal fluid remains
unknown.
The present investigation aimed to shed some light on the origin of a-amylase and maltase activity detected in the fecal fluid
of A. sexdens. To accomplish this, enzymes were purified from the
fecal fluid of the ants and from pure laboratory cultures of the
mutualistic fungus L. gongylophorus, and then characterized
biochemically.
Our results suggest that the fecal fluid contains a fungal a-amylase and a maltase acting synergistically in a selected metabolic
pathway to degrade starch, depicting an important evolutionary
innovation supporting the biochemical alliance between leaf-cutters and its mutualistic fungus.
2. Material and methods
2.1. Samples
The strain B1-97 of the symbiotic fungus L. gongylophorus, isolated from a laboratory nest of A. sexdens rubropilosa was kept at
25 °C for 20 days in culture medium with 0.075 M citrate-phosphate buffer (pH 5.0), 0.67 g 100 ml 1 Yeast Nitrogen Base (Difco,
United States, catalogue number 100690), 0.25 g 100 ml 1 starch
(Sigma, Germany, catalogue number S9765) and 0.25 g 100 ml 1
maltose (Sigma, Germany, catalogue number M9171). The culture
medium containing the enzymes produced was filtered through a
0.45 lm filter, dialyzed for 12 h at 4 °C, in order to eliminate soluble sugars, and concentrated 10 times by lyophilization, giving
rise to the crude sample which was used in the purification
process.
The fecal fluid of A. sexdens was taken from workers collected
from a laboratory nest. The abdomens of worker ants were compressed and the expelled fecal fluid droplets were collected with
a micropipette and transferred to an ice-cold micro assay tube.
Each worker released 0.5 lL fluid in average and approximately
300 lL (600 workers) were collected per day and stored at
80 °C. The collection was repeated in the next 2 days, the nest
was allowed to recover for 3 weeks and a new set of collections
was carried out. Around 5400 workers were processed, samples
were pooled, dialyzed for 12 h at 4 °C and concentrated and submitted to the purification process described in Tables 1 and 2.
2.2. Enzyme purification
Alpha-amylase and maltase purification was carried out as described in Tables 1 and 2, respectively. Chromatographic procedures were carried out in the AKTA-FPLC™ system (GE
HealthCare). Each step of enzyme purification was followed by
electrophoresis in polyacrylamide gel in denaturing conditions,
as described by Laemmli (1970).
2.3. Denaturing electrophoresis in polyacrylamide gel
Polyacrylamide gels were used at a final concentration of 15%
(v/v) and electrophoresis conditions were the same as those previously established by Laemmli (1970). After the run, the gel was
submitted to a silver stain (Heukeshoven and Dernick, 1985). All
the reagents for electrophoresis were obtained from GE HealthCare, including molecular weight standard proteins serum bovine
albumin (66 kDa), ovalbumin (45 kDa), carbonic anhydrase
(29 kDa), trypsin inhibitor (20 kDa) and a-lactalbumin (14 kDa).
Migration (mm) of each standard protein in SDS–PAGE was plotted
graphically as a function of molecular weight and the curve was
used to determine the molecular weights of the purified proteins.
2.4. Enzymes activity (U) assay
For a-amylase determination, a 100 ll sample (chromatographed sample, 1/10 fungal crude sample, 1/100 fecal fluid) was
mixed with 100 ll of solution containing 20 mg ml 1 starch,
0.002 M calcium chloride, and 0.075 M citrate-phosphate buffer
(pH 5.0). The reaction mixture was kept at 30 °C during zero, 20,
40 and 60 min, after which 500 ll iodine reagent
(0.25 g 100 ml 1 iodine (Merck, Germany, catalogue number
4761) and 0.16 g 100 ml 1 potassium iodide (Mallinckrodt,
Mexico, catalogue number 1127)), was added to the reaction
according to Bernfeld (1955). Then, the values of absorbance at
620 nm were determined, compared to a starch standard curve
and plotted to calculate the enzymatic activity, which was
expressed in a-amylase units (UA), one UA corresponding to the
consumption of 1 lg in soluble starch per minute per ml of culture
medium or fecal fluid or chromatographed sample.
For maltase determination, 100 ll of sample (chromatographed
sample, 1/10 fungal crude sample, 1/100 fecal fluid) was mixed
with 100 ll of 40 mg ml 1 maltose dissolved in 0.075 M citratephosphate buffer (pH 5.0). The reaction mixture was kept at
30 °C during zero, 20, 40 or 60 min and immediately boiled for
3 min. Then, 300 ll of water and 500 ll of glucose oxidase reagent
(Labtest, Brazil, catalogue number 34-E) were added to the reaction mixture and, after 15 min at 37 °C, the absorbance at
505 nm was determined. Glucose concentration was estimated
based on a glucose standard curve and maltase units (UM) were
calculated, one UM corresponding to the amount of maltase which
generated 1 lMol of glucose per minute per ml of culture medium,
fecal fluid or chromatographed sample.
2.5. Characterization of purified enzymes
Alpha-amylase and maltase purified from the mutualistic fungus or from the fecal fluid of ant workers were characterized by
determining the influence of temperature or pH on their activity
and stability, as well as by determining their kinetic constant Km
and molecular weight.
2.6. Influence of the temperature on purified enzymes
The temperature effect on enzyme activity was evaluated by
determining a-amylase or maltase activity in 1h-incubations at
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M. Bacci Jr. et al. / Journal of Insect Physiology 59 (2013) 525–531
Table 1
Purification steps for a-amylase from A. sexdens or L. gongylophorus.
Steps
Crude samples
1
2
3
4
Total protein (mg)
Total activity (UA)
Specific activity (UA mg
Ant
Fungus
Ant
Fungus
Ant
72
31
14
3.1
0.8
280
180
22
8.0
2.1
1098
980
730
309
176
3775
3250
1250
482
364
15
32
52
100
220
1
protein)
Fold of purification
Yield (%)
Fungus
Ant
Fungus
Ant
Fungus
14
18
57
60
173
1.0
2.1
3.5
6.7
15
1.0
1.3
4.0
4.3
12
100
89
67
28
16
100
86
33
13
10
Step 1: DEAE Sepharose equilibrated with 0.02 M sodium phosphate buffer (pH 8.0); fungal or ant a-amylase was eluted in the equilibrium buffer.
Step 2: Butyl Sepharose equilibrated with 0.02 M sodium phosphate buffer (pH 8.0) and 1.3 M ammonium sulfate; fungal or ant a-amylase was eluted in 0.02 M phosphate
buffer.
Step 3: Q Sepharose equilibrated with 0.02 M sodium phosphate buffer (pH 8.0); fungal or ant a-amylase was eluted in 0.25 M NaCl.
Step 4: Sephacryl S-100 equilibrated with 0.02 M sodium phosphate buffer (pH 8.0) at 1.0 ml min 1; fungal or ant a-amylase was eluted from 85 to130 ml buffer.
Table 2
Purification steps for maltases from A. sexdens or L. gongylophorus.
Steps
Crude samples
1
2
3
1
Total protein (mg)
Total activity (UM)
Specific activity (UM mg
Ant
Fungus
Ant
Fungus
Ant
Fungus
protein)
Fold of purification
Ant
Fungus
Yield (%)
Ant
Fungus
72
22
8.0
1.0
280
43
22
1.3
7.0
3.0
2.0
1.0
5.0
2.0
2.0
0.5
0.1
0.14
0.25
1.0
0.02
0.05
0.09
0.39
1.0
1.4
2.5
10
1.0
2.5
4.5
20
100
43
29
14
100
40
40
10
Step 1: DEAE Sepharose equilibrated with 0.02 M sodium phosphate buffer (pH 8.0); fungal maltase was eluted in 0.3 M NaCl, and ant maltase was eluted in 0.3–0.5 M NaCl.
Step 2: Butyl Sepharose equilibrated with 0.02 M sodium phosphate buffer (pH 8.0) and 1.3 M ammonium sulfate; maltases were eluted in 0.02 M phosphate buffer.
Step 3: Sephacryl S-100 equilibrated with 0.02 M sodium phosphate buffer (pH 8.0) at 1.0 ml min 1; fungal maltase was eluted from 30 to 70 ml buffer, and ant maltase was
eluted from 45 to 75 ml buffer.
zero, 20, 25, 30, 35, 40, 45, 50, 55 or 60 °C. The thermal stability of
these enzymes was evaluated by incubating them for 1 h at the
temperatures cited above, followed by determining their activity
at 30 °C. The results of both temperature effect and stability experiments were expressed in a U% (residual activity) versus temperature plot. Based on the temperature assays, it was possible to
calculate activation energy.
Thermal inactivation was evaluated by incubating each of the
purified amylases at 40 °C for 5, 10, 15, 20 or 30 min, after which
the enzyme residual activity (UA) was determined at 30 °C. The
plotting of the log of residual UA as a function of the incubation
time at 40 °C resulted a straight line that is characteristic for each
enzyme.
2.7. Influence of the pH
The influence of pH on enzyme activity was determined by
assaying the purified enzymes at 30 °C in 0.075 M citrate-phosphate buffer at pH values 4.0, 5.0, 6.0, 7.0 or 8.0. To evaluate the
influence of pH on the stability of enzymes, each purified enzyme
was kept for one hour at 8 °C in 0.075 M citrate-phosphate buffer
at pH values of 4.0, 5.0, 6.0, 7.0 or 8.0. Then, residual activities were
determined at 30 °C in the optimum pH for each enzyme: pH 5.0
for fungal or fecal a-amylase and fungal maltase; and pH 6.0 for fecal maltase.
3. Results
3.1. Purification of a-amylases and maltases from the ants or the
fungus
The crude samples containing a-amylase or maltase activity
from the ants or fungus were purified to electrophoretic homogeneity by chromatographic procedure as shown in Tables 1 and 2,
respectively.
3.2. Molecular weight determination
A single and predominant molecular species of each enzyme
was found and purified to homogeneity. The values of molecular
weight, determined with denaturing electrophoresis in polyacrylamide gel were 27 and 29 kDa for ant and fungal a-amylases,
2.8. Michaellis–Menten kinetic constant (Km)
Enzyme activity was determined with nine substrate concentrations ranging from 1.0 to 10 mg ml 1 of soluble starch (Sigma, Germany, catalogue number S-9765) or 0.2 to 10 mM of maltose
(Sigma, Germany, catalogue number M-9171) and Km values were
calculated using the Lineweaver–Burk plot. Testing of differences
in Km values was carried out by an independent t test at 0.05
significance.
Fig. 1. Denaturing electrophoresis polyacrylamide gel of molecular markers (1),
fungal 29 kDa a-amylase (2), ant 27 kDa a-amylase (3), ant 63 kDa maltase (4) and
fungal 74 kDa maltase (5).
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M. Bacci Jr. et al. / Journal of Insect Physiology 59 (2013) 525–531
respectively, and 63 and 74 kDa for ant and fungal maltases,
respectively (Fig. 1).
Log Residual UA (%)
2.2
3.3. Influence of the temperature
The optimal temperature was 35 °C for both fungal and ant aamylases (Fig. 2A), 55 °C for fungal maltase and 40 °C for ant maltase (Fig. 2B).
The thermal stability test showed loss of activity at any of the
temperatures utilized, with fungal or ant a-amylase presenting a
similar pattern of activity loss and total inactivation occurring at
50 or 55 °C, respectively (Fig. 2C). However, fungal or ant maltases
differed from each other in their pattern of activity loss, with ant
maltase being less resistant to temperature increase and reaching
complete inactivation at 45 °C as fungal maltase conserved 9% of
its initial activity at 60 °C (Fig. 2D).
Activation energy values were 21.4 and 21.2 cal mol 1 for ant or
fungal a-amylases and 16.9 and 13.0 cal mol 1 for ant or fungal
maltases, respectively.
Since fungal and ant a-amylases have been quite similar to each
other, an additional test, the kinetic inactivation at 40 °C, was carried out aiming to differentiate these enzymes. The two linear
curves obtained from this test (Fig. 3) had nearly identical inclination, so no significant difference was found at thermal inactivation
between these enzymes.
3.4. Influence of pH
Optimum pH value was 5.0 for the ant or fungal a-amylase
activity (Fig. 4A), as well as for the fungal maltase activity
(Fig. 4B), and it was 6.0 for the ant maltase activity (Fig. 4B). Fungal
L. gongylophorus
A. sexdens
2.0
1.8
1.6
1.4
1.2
0
15
20
25
30
Fig. 3. Thermal inactivation at 40 °C of a-amylase from A. sexdens or L. gongylophorus, showing two curves with nearly identical angular coefficients.
or ant a-amylases had also similar stability, which was higher at
pH values 4.0–6.0 (Fig. 4C). However, fungal or ant maltase had
distinct pH stability profiles: ant maltase was more stable at pH
6.0 as fungal maltase was instable at the tested pH values (Fig. 4D).
3.5. Kinetic constant Km
Fig. 5 shows the Lineweaver–Burk linear graphs used to calculate values for the Michaelis–Menten kinetic constant (Km) of aamylases and maltases. The Km value for purified amylases from
workers was 5.86 ± 0.14 mg ml 1, which was not significantly different (t = 2.75; p = 0.052) from the 6.23 ± 0.19 mg ml 1 found for
(B)
L. gongylophorus
A. sexdens
80
60
40
L. gongylophorus
A. sexdens
100
Relative UM(%)
Relative UA(%)
10
Time (min)
(A)
100
5
20
80
60
40
20
0
0
20
30
40
50
20
60
Temperature ( °C)
30
40
50
60
Temperature (°C)
(C)
(D)
70
60
L. gongylophorus
A. sexdens
40
30
20
10
L. gongylophorus
A. sexdens
60
Residual UM (%)
Residual UA (%)
50
50
40
30
20
10
0
0
20
30
40
50
Temperature ( °C)
60
20
30
40
50
60
Temperature ( °C)
Fig. 2. Effect of temperature on enzyme activity (A and B) and thermal stability (C and D) of purified a-amylase (A and C) or maltase (B and D).
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M. Bacci Jr. et al. / Journal of Insect Physiology 59 (2013) 525–531
(B)
100
100
90
80
Relative UM (%)
Relative UA (%)
(A)
80
70
60
L. gongylophorus
A. sexdens
50
60
40
20
L. gongylophorus
A. sexdens
0
4
5
6
7
8
4
5
pH
100
80
80
Residual UM (%)
Residual UA (%)
7
8
7
8
(D)
(C)
100
60
40
L. gongylophorus
A. sexdens
20
6
pH
60
40
L. gongylophorus
A. sexdens
20
0
0
4
5
6
7
8
pH
4
5
6
pH
Fig. 4. Effect of pH on activity (A and B) and stability (C and D) of the purified a-amylase (A and C) and maltase (B and D) from the A. sexdens or L. gongylophorus.
the fungus amylase. However, a significant difference (t = 7.29;
p = 0.0019) was found in the Km values found for maltase purified
from
workers
(2.63 ± 0.29 mM)
or
from
the
fungus
(1.24 ± 0.17 mM) (Fig. 5).
4. Discussion
Martin (1983) stated that many insects compensate their incapacity to degrade complex nutrients by ingesting fungal enzymes
from microorganisms. The fungal origin of cellulases, xylanases
and cellobiases has been demonstrated in beetles (Martin et al.,
1981; Kukor and Martin, 1986), wasps (Kukor and Martin, 1983)
and in termites, the latter living in the Tropical Region of the African and Asian continents and cultivating a mutualistic basidiomycete (Abo-Khatwa, 1978; Martin and Martin, 1979; Rouland et al.,
1986; Veivers et al., 1991; Matoub and Rouland, 1995). Pectinases
produced by the mutualistic fungus was also found in leaf-cutter
ants (Ronhede et al., 2004; Schiøtt et al., 2010).
Therefore, the investigation on the origin of enzymes in the
intestine of insects is essential from a nutritional and ecological
point of view, since it reveals the enzymatic interactions between
insects and microorganisms.
A biochemical study of the relevant enzymes present in the fecal fluid of the leaf-cutter ants is important because these enzymes
are responsible for the process of degrading polysaccharides from
the foliar material which is cut and carried by the ants to their nest.
This degradation generates major food resources for the ant (Silva
et al., 2003) and its mutualistic fungus (Siqueira et al., 1998).
In the present investigation, our results allowed the understanding of the participation of fungal and ant enzymes in starch
degradation, providing an excellent opportunity for detailing the
metabolic integration maintaining Attini ant and fungus
mutualism.
4.1. The origin of a-amylases in the intestine of the A. sexdens workers
By comparing the biochemical characteristics from the a-amylases obtained in our work, we found identical or nearly identical
responses to temperature or pH, as well as activation energy, substrate affinity and molecular weight. This is compatible with the
proposition that fungal and ant a-amylases are in fact a single enzyme. In other words, the a-amylase secreted by the mutualistic
fungus is likely the one found in the ant intestine. On the other
hand, maltases from ants or from the fungus differed from each
other in most of the biochemical characteristics assessed, indicating these enzymes are distinct from each other.
4.2. Selection of fungal and ant enzymes in the intestine
Our findings indicate that the ant intestine environment selects
some of the enzymes which are produced by the mutualists. The
maltase we found to be produced by L. gongylophorus or the aamylases produced by the ants in the salivary and mandibular
glands and in the medium intestine (Ayre, 1967; Febvay and Kermarrec, 1983) were not found in the fecal fluid. These enzymes
are probably inactivated in the ant intestine. On the other hand,
the a-amylase produced by the mutualistic fungus seems to be ingested by the ants, pass through the intestine and remain active in
the fecal fluid. The fecal fluid also contains an active maltase likely
produced by the workers. Therefore, the ant intestine seems to
positively select the fungal a-amylase and workers maltase which
are found in the fecal fluid as negative selection seems to be imposed to fungal maltase and ant a-amylases.
Negative selection on fungal maltase is somehow unexpected
because fungal maltase is better than ant maltase regarding temperature resistance (Fig. 2D) and substrate affinity expressed by
Km values.
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M. Bacci Jr. et al. / Journal of Insect Physiology 59 (2013) 525–531
(A)
L. gongylophorus
A. sexdens
2.5
2.0
1/v
1.5
1.0
0.5
-0.4
-0.2
0.0
0.2
0.4
1/S
0.6
0.8
1.0
(B)
5. Conclusions
L. gongylophorus
A. sexdens
6
3
The intestine of the leaf-cutter ant Atta sexdens positively selects the Leucoagaricus gongylophorus fungus a-amylase and a maltase likely produced by the ants. These enzymes compose a
catabolic pathway to assimilate starch from vegetal matter. The
pathway represents a major evolutionary innovation supporting
ant herbivory.
2
Acknowledgement
5
1/v
4
1
-2
-1
It could be argued that the fungus would not deeply benefit
from starch degradation promoted by the fecal fluid, because the
fungus is able to generate glucose from starch independently of
the ants and because fungal maltase has higher substrate affinity
than fecal fluid maltase. However, fungal maltase (and also fungal
amylase to a less extent) is highly sensitive to catabolic repression
(Silva et al., 2006) at glucose concentrations prevailing in the ant
nest (Silva et al., 2003). Thus, the structuring of the starch catabolic
pathway in the fecal fluid seems an evolutionary advantage which
resulted more efficient than the starch degradation carried out by
each of the mutualists individually.
Glucose generation from plant starch occurs when ants deposit
fecal droplets on the surface of cut leaves inside the nests. Glucose
is an essential carbon source for the survival of both the ant (Silva
et al., 2003) and its mutualistic fungus (Siqueira et al., 1998).
Therefore, the generation of glucose through the starch degradation pathway proposed here is a major evolutionary innovation
which plays a central part in ant and fungus nutrition on plant
material.
We thank FAPESP 2011/50226-0 for funding this research.
0
1
2
3
4
5
1/S
Fig. 5. Lineweaver–Burk plot for A. sexdens or L. gongylophorus purified a-amylases
showing Km values not significantly distinct from each other (A) and similar plot (B)
for purified maltases for A. sexdens or L. gongylophorus which had Km values
significantly distinct from each other.
It is possible that enzyme selection occurs by enzyme inactivation during its passage through the ant intestine. A potential reason for enzyme inactivation is degradation by proteases which
are present in the fecal fluid, albeit proteases are thought less active in the rectal region of the intestine of some the leaf-cutter ants
(Erthal et al., 2004).
4.3. Starch degradation in leaf-cutter ant-fungus system
Enzyme selection in the ant intestine seems to have structured
a metabolic pathway to degrade starch by the fecal fluid. In this
proposed pathway, fungal a-amylase is the first enzyme acting to
hydrolyze starch, producing reducing sugars, such as the disaccharide maltose, which is then hydrolyzed by a maltase to result glucose. The functionality of this pathway is reinforced by our finding
that these have been the unique starch degrading enzymes present
in the fecal fluid and that the fecal fluid is able to efficiently degrade starch (Martin et al., 1973).
This conclusion is in agreement with the observations by Richard et al. (2005), which showed that leaf-cutter ants and their
mutualistic fungus have complementary enzymatic activities, the
fungus displaying polysaccharidase activity and the ants being specialized in the degradation of low molecular weight substrates.
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