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Dissecting the association between a gall midge,
Asteromyia carbonifera, and its symbiotic fungus,
Botryosphaeria dothidea
Jeremy J. Heath* & John O. Stireman III
Department of Biological Sciences, Wright State University, 3640 Colonel Glenn Hwy., Dayton, OH 45435, USA
Accepted: 21 June 2010
Key words: Ambrosia gall, adaptive radiation, microbial mutualism, eclosion behaviour, sympatry,
ecological speciation, symbiosis, Diptera, Cecidomyiidae, Ascomycota, Dothideomycetes
Abstract The Ambrosia gall midge [Asteromyia carbonifera (Osten Sacken) (Diptera: Cecidomyiidae: Alycau-
lini)] consists, in part, of a complex of genetically differentiated populations that have diverged in gall
morphology on the host plant Solidago altissima L. (Asteraceae). This divergence appears to be an
incipient adaptive radiation that may be driven by parasitoid pressure. Understanding the mecha-
nisms driving this genetic and phenotypic diversification requires a close examination of the relation-
ship between the midge and its fungal associate Botryosphaeria dothidea (Moug.) Ces. & De Not.
(Ascomycota: Dothideomycetes), whose mycelia actually form the protective gall structure. We used
manipulative experiments to test the degree of interdependency of the fungus and the midge, and we
employed field and laboratory studies to gain insight into the source of fungal conidia, which our
data and observations indicate are collected by females and stored in specialized pockets (mycangia)
on the ovipositor. Manipulative experiments demonstrate that fungal proliferation on the host plant
is dependent on the midge larvae and larvae exhibit significant growth on the fungus alone. Field
observations and experiments were unable to identify the source of mycangial conidia; however, anal-
yses of conidia shape suggest a biotrophic source. We conclude that this association is an obligatory
mutualism with respect to successful gall formation. These findings corroborate recent findings that
the primary food source of the midge is the gall fungus.
Introduction
Mutualistic associations with microbes have likely played
an important role in the phenomenal evolutionary and
ecological success of the Insecta (Moran, 2002; Janson
et al., 2008). Virtually every insect species that has been
examined closely has been found to be engaged in some
form of microbial mutualism, most frequently in the form
of gut-associated bacteria (e.g., Buchner, 1965; Douglas,
1998). However, insect-fungal symbioses are also wide-
spread across many insect groups including beetles (Sco-
lytinae: Bentz & Six, 2006), ants (Formicidae; Mikheyev
et al., 2006), moths (Tortricidae: Fermaud & Lemenn,
1989), and flies (Cecidomyiidae: Borkent & Bissett, 1985;
Gagne
´, 1989; Anthomyiidae: Schiestl et al., 2006). In these
associations, the insect typically benefits from using the
fungus as a food source (leafcutter ants: Cherrett et al.,
1989; gall midges: Bissett & Borkent, 1988; Ambrosia bee-
tles: Farrell et al., 2001) in exchange for dispersing the fun-
gus or promoting fungal outcrossing (Schiestl et al., 2006).
One of the most diverse and widespread groups of
insects known to engage in symbiotic associations with
fungi are the ‘Ambrosia’ gall midges, which represent a
significant portion of the family Cecidomyiidae. The galls
that these midges induce on their host plants are typically
lined internally with fungal hyphae, which the developing
larva(e) may feed upon (Haridass, 1987; Bissett & Borkent,
1988). Borkent & Bissett (1985) have provided tantalizing
morphological evidence that at least some of these species
actively transport fungi in specialized pockets, or mycan-
gia, associated with the terminal abdominal segments.
However, the nature of the Ambrosia midge-fungus
interaction is largely speculative and there has been little
*Correspondence: E-mail: heath.22@wright.edu
2010 The Authors Entomologia Experimentalis et Applicata 137: 36–49, 2010
36 Journal compilation 2010 The Netherlands Entomological Society
DOI: 10.1111/j.1570-7458.2010.01040.x
experimental analysis of the association and the degree to
which it represents an obligate mutualism. In fact, it has
been argued that at least in some cases the fungi may
represent opportunistic colonization rather than a strict
mutualism. This controversy has been thoroughly
reviewed (Haridass, 1987; Rohfritsch, 2008; Adair et al.,
2009). Here, we employ observational and manipulative
experiments to dissect the association between the gall
midge Asteromyia carbonifera (Osten Sacken) (Diptera:
Cecidomyiidae: Alycaulini) and its associated fungus
Botryosphaeria dothidea (Moug.) Ces. & De Not. (Ascomy-
cota: Dothideomycetes) in order to classify this symbiosis
and understand its consequences for the ecology and
evolution of the gall midge.
The galls induced by A. carbonifera and its associated
fungus on host plants in the genus Solidago (goldenrod;
Asteraceae) were observed over a century ago (Trelease,
1884; Batra, 1964). However, despite the efforts of several
researchers (e.g., Batra, 1964; Gagne
´, 1968; Weis, 1982a,b;
Bissett & Borkent, 1988), the biology of A. carbonifera is
still poorly understood. Populations of A. carbonifera use
a wide range of goldenrod species as hosts (Gagne
´, 1968),
and preliminary analysis indicates that many of these
populations exhibit host associated genetic differentiation
(Stireman et al., 2010). Furthermore, within the single
host plant Solidago altissima L., at least four genetically
distinct gall morphotypes coexist, suggesting that A. carbo-
nifera is adaptively radiating on this host (Crego et al.,
1990; Stireman et al., 2008).
Understanding the details of the interaction between
A. carbonifera and its fungal associate is likely key to
understanding the adaptive diversification in this system.
Borkent & Bissett (1985) mentioned that A. carbonifera
transport fungal conidia, but only supplied photographs
of the mycangia of A. tumifica and related species. Gagne
´
(1968) attempted to initiate A. carbonifera galls in screen
cages. Galls developed in his experiment, but we suspect
that there were eggs already on the plants before they were
caged. In fact, Gagne
´(1968, p. 13) does not reject this
possibility when he states, ‘It is possible also that…some
newly laid eggs were overlooked.’ Weis (1982a,b) clearly
illustrated that larvae are variously protected from para-
sitoid attack by the fungal stroma and A. carbonifera has
never been found in plant tissue not associated with the
fungus (Trelease, 1884; Batra, 1964); nevertheless, some
still question whether this association represents a mutual-
ism. Therefore, the main goal of this study was to investi-
gate the nature of this intimate association between the
midge and the fungus. That is, is it truly a mutualism? If
so, is it an obligatory relationship?
In particular, we were interested in (1) verifying earlier
claims that adult females harbour fungal conidia in
specialized structures on their ovipositor called mycangia
and whether these conidia are present on eggs, (2) deter-
mining whether the midge is necessary for fungal growth
and vice versa, (3) initiating galls under controlled condi-
tions, (4) understanding where midges mate and obtain
fungal conidia, and (5) determining the time required for
fungus to appear on the top of the leaf after oviposition
has occurred.
Study system
Asteromyia carbonifera is involved in complex interactions
involving many interacting species. The four most promi-
nent players are goldenrod (Solidago spp.), the gall midge
A. carbonifera, the fungus B. dothidea (Bissett & Borkent,
1988; Janson et al., in press), and at least nine associated
parasitoids, predators, and inquilines. Together, the fun-
gus and midge larva produce at least four morphologically
distinct galls on the leaves of S. altissima. Crego et al.
(1990) described these four gall morphotypes on S. altis-
sima. These were named crescents, flats, irregulars, and
cushions in accordance with their overall morphology.
Stireman et al. (2008) provided photographs and descrip-
tions of these morphotypes. These galls are not deforma-
tions of plant tissue, but rather a gall formed mainly of
fungus. Aside from chlorosis, the gall causes very little
physical change at the cellular level of the plant (Camp,
1981). Other Asteromyia spp. also form similar Ambrosia
galls on related host plants (Gagne
´, 1968, 1989; Stireman
et al., 2010) and many Asphondylia spp. (Diptera: Ceci-
domyiidae) harbour the same fungal species (Bissett &
Borkent, 1988; Adair et al., 2009); therefore, understand-
ing this symbiosis in A. carbonifera may provide insight
concerning other potential adaptive radiations.
Materials and methods
Biology and life history traits
Unless otherwise specified, the biology and life history
traits were generally inferred from field observations at all
the field sites (Table 1) over the summers of 2007–2009.
To investigate the time for fungus to appear on the top of
the leaves, A. carbonifera egg clutches were located on the
most susceptible S. altissima clone in a common garden
and marked (see subsequent section for details on the
common garden). These clutches were checked daily until
at least one position within the clutch began to show fun-
gal growth on the top of the leaf (i.e., when the growth was
about 0.3 mm in diameter). The leaf the eggs were found
on was given an estimate of age in days between 0 and 3.
The plants grow about 1 cm per day and the distance
between pairs of leaves is roughly 1 cm at the top of the
plant. Age 0 leaves were on the outside of the whorl and
Asteromyia-fungus association 37
had not dropped to a horizontal position. Age 1 leaves
were still connected to the whorl, but were horizontal. Age
2 and 3 leaves were one and two nodes lower than the base
of the whorl, respectively. Eggs were only found on age 0–3
leaves, but only the new growth was searched. Females
were observed laying eggs in three instances in the field
and these eggs were tracked as above. These clutches were
followed until they formed mature, identifiable morpho-
types (clutch sample sizes per morph: cushion: 22, flat: 4,
irregular: 13, crescents: 0, and unknown: 1). A linear
model with leaf age as a covariate was used to test for dif-
ferences between the morphotypes in the time for fungus
to appear on the top of the leaf.
Nature of the midge-fungus interaction
Two types of experiments were conducted to test the inter-
dependency of the midge larva and fungus. The first exper-
iment tested whether the growth of the gall fungus
depends on the presence of the midge larva by removing
larvae from very young galls and assessing fungal growth.
The second experiment tested whether the midge larva
feeds on the fungus by isolating the larva and fungus on
growth media and tracking larval growth.
To test whether gall fungal growth is dependent on the
presence of the midge larva, the larvae from very young
galls were removed and fungal growth measured. Solidago
altissima stems with very young blister galls (<2 mm in
diameter) were collected from the Beavercreek Wildlife
Management Area (BCWMA) site (Table 1). The stems
were re-cut under water in the field and immediately
placed in a container of cut-plant solution (Aquaplus; Syn-
dicate Sales, Kokomo, IN, USA). On sunny days stems
were placed in the shade during collection. The galls on the
stems were randomly processed in the laboratory as nega-
tive controls, mocks, or removals. Controls were left
untouched and intact. Mock removals (‘mocks’) con-
trolled for possible confounding effects of the removal
process; galls were dissected from the bottom of the leaf to
the removal point and the larva was either touched with
the forceps as if it was going to be removed or not touched.
The removaltreatment was conducted in the same manner
as the mock treatment, but the larva was permanently
removed from the gall. In both mocks and removals the
disturbed fungal layer and leaf tissue was carefully placed
back in its original position. These treatments were
repeated in five independent experiments with a balanced
sample size within each experiment (n = 6–9 per treat-
ment). All galls were photographed with identical camera
and magnification settings at the time of processing and
after 10 days of incubation at room temperature. Fungal
growth before and after incubation was measured in pixels
in Photoshop (version 9.0; Adobe Systems Inc., San Jose,
CA, USA), converted to net fungal growth per day, and
standardized to the number of larvae present in the final
gall. Standardization was necessary because during the
initial stages of gall development each larva initiates a
separate tiny gall. If the initial galls are close enough they
will eventually merge leading to the appearance of a single
large gall. In some cases immature galls processed as nega-
tive controls had neighbouring galls close enough that they
eventually merged leading to the need to standardize net
fungal growth to the number of larvae present.
To test whether the midge larva feeds on the fungus,
immature galls were collected from the BCWMA site, dis-
sected, and a small portion (ca. 10 mm
2
) of black fungal
stroma was transferred to malt-extract agar plates along
with a young larva (n = 18), which was placed atop the
portion of stroma. Negative controls without fungal trans-
fer were also included (n = 17). The length of larvae was
measured with an ocular micrometre mounted on a Nikon
SMZ1000 stereomicroscope (Nikon Instruments, Melville,
NY, USA) on the day of transfer and after 2 weeks of
incubation at room temperature.
Preliminary experiments had shown that larvae grew
significantly larger when transferred with the fungus as
above, but the growth was marginal (i.e., a mean increase
over the control of only 51 lm). Furthermore, in preli-
minary experiments most of the control plates were
Tab l e 1 List of study sites with abbreviations, names, and coordinates
1
Site abbreviation Site name (all USA) Latitude (N) Longitude (W)
BCWMA Beavercreek Wildlife Management Area, OH 3945¢59.09¢¢ 8400¢15.95¢¢
HS Huffman Metropark, Dayton, OH 3948¢28.30¢¢ 8405¢33.94¢¢
KWP Koogler Wetland Preserve, Dayton, OH 3945¢57.97¢¢ 8400¢40.96¢¢
VS Varner Rd., Dayton, OH 3946¢01.45¢¢ 8400¢56.51¢¢
WSU-1 WSU Common Garden, Dayton, OH 3947¢14.96¢¢ 8403¢08.59¢¢
WSU-2 WSU Services Site, Dayton, OH 3947¢25.12¢¢ 8403¢10.85¢¢
SC-1 Millbrook Marsh, State College, PA 4049¢00.44¢¢ 7750¢04.13¢¢
SC-2 Stewart Drive, State College, PA 4049¢50.16¢¢ 7747¢44.52¢¢
1
Coordinates obtained from Google Earth (Version 4.3).
38 Heath & Stireman
also contaminated with gall fungus. Therefore, about
5 mg of medical grade Nystatin ointment (a fungicide,
100 000 USP g
)1
; E. Fougera & Co, Melville, NY, USA)
was applied to a 1-cm
2
area of both the treatment and
control plates. The larva and fungus or larva alone (in the
case of controls) was transferred atop this paste. The paste
completely prevented fungal contamination in the
controls and substantially reduced fungal growth in the
treatments. The paste did not appear to affect the behav-
iour or survival of the transferred larvae.
A linear model in R (version 2.8.1; R Development Core
Team, 2007) with ‘larval end length’ as the response and
‘start length’ as a covariate was used to determine fungal
treatment effects. The full model included the following
explanatory variables: larval start length, fungal treatment
(gall fungus added or not), and their interaction.
Asteromyia eclosion behaviour
To understand the reproductive behaviour and to gain
insight into where and how the adults obtain fungal coni-
dia, mature irregular galls on S. altissima (SC-2 site;
Table 1) and rugosa galls on Solidago rugosa P. Mill. (SC-1
site; Table 1) were marked and monitored daily in the field
from 10 July to 7 August 2007 (see Gagne
´,1968,fora
description of S. rugosa-type galls). As adults emerged
from the galls, their behaviour was recorded until they
either flew off or otherwise became unobservable. On aver-
age, they were observed for 2.5 h. Ethograms were gener-
ated from these data and the relative frequency of the
transition between behaviours calculated by dividing the
number of times a transition from behaviour x to y
occurred by the total number of behavioural transitions.
Conidia morphology
To gain insight into where fungal conidia are obtained, we
compared the shape of egg-associated conidia to conidia
obtained from other sources. Eggs of presumably different
morphotypes were collected from the BCWMA and Koo-
gler Wetland Preserve (KWP) sites (Table 1), mounted on
microscope slides in EMD
TM
lactophenol cotton blue
(Fisher Scientific, Pittsburgh, PA, USA), and photo-
graphed at 400·with a Nikon Coolpix 8800 VR camera
mounted on a Nikon Optiphot compound microscope
(Nikon Instruments). The width and length of the conidia
were measured from photographs in ImageJ (version
1.39u; National Institute of Health, Bethesda, MD, USA)
after calibration with a photograph of a stage micrometre
(2 mm, ruled to 0.01 mm; Micromaster, Fisher Scientific,
Hampton, NH, USA). The length and width of these
egg-associated conidia were compared to conidia from six
other sources: (1) gall-isolated fungus grown at room
temperature on oatmeal agar under cool white fluorescent
lights (six each, Philips, F40T12 ⁄CW Plus, 40-W bulbs,
suspended 60 cm above the plates), (2) gall-isolated fun-
gus grown onfresh-cut autoclaved goldenrod stems placed
on the surface of water agar plates, (3) egg-conidia isolates
grown on oatmeal agar, (4) egg-conidia isolates grown on
fresh-cut autoclaved goldenrod stems on water agar, (5)
conidia collected from pycnidia found on field-collected
S. altissima stems, or (6) conidia found in the mycangia of
malaise-trapped adult A. carbonifera females from the HS
site (Table 1). The gall and egg-conidia isolates (1–4
above) were all grown at the same time under the same
conditions in a randomized complete block design. Two
linear ANCOVA (analysis of covariance) models with
‘width’ as a covariate were used to test for significant
differences in conidia length (R, version 2.8.1). The
standardized residuals from these models were roughly
normal, distributed mostly between the mean ± 2 SD, and
appeared randomly associated with the fitted values. The
first model included three explanatory variables: (a) fungal
isolate source (i.e., egg-conidia isolate or gall-isolate), (b)
growth media (i.e., fresh-cut S. altissima stems on water
agar or oatmeal agar only), and (c) width as a covariate;
plus (d) all the two- and three-way interactions. This
model included only the first four treatments (1–4 above).
The second model included all seven conidia sources with
only two explanatory variables (i.e., width as a covariate,
conidia source, and their interaction).
Gall initiation in field plots
Two experiments were conducted in an attempt to initiate
galls on S. altissima accessions grown up in the greenhouse
and transplanted to a common garden (WSU-1; Table 1).
Solidago altissima rhizomes were collected from three sites
[BCWMA, KWP, and Varner Road (VS) sites; Table 1] in
early April and 10 clones of 10 source plants were started
in a greenhouse and later transplanted to the common gar-
den. Large conical tomato cages were placed over half of
the plants in the field plot (n = 50, five replications of each
accession). The cages were covered with fine-mesh sleeves
(194 holes cm
)2
) and buried about 10 cm deep. Mature
galls were collected from S. altissima from various field
sites on 9 July 2008, separated by morphotype, and placed
on the ground inside each of 40 cages (10 plants were con-
trols) in the same morphotype proportion as the total col-
lected. Each cage received 44 irregular, seven crescent, five
cushion, and four flat galls. Recently cut dried goldenrod
stems, old goldenrod stems (previous year’s growth), and
extraneous ground litter was cut into 10-cm sections and
placed in the bottom of each of the cages. It was thought
that this debris might provide a source of fungal conidia
for emerging adult females to collect. The caged plants
were checked periodically for the formation of new galls
Asteromyia-fungus association 39
and the presence of emerging adults. After 24 days all
plants were cut to 30 cm and checked thoroughly for the
presence of galls.
In a second experiment, a subset (n = 20) of the same
caged plants were allowed to re-grow for 17 days and then
mature galls were added to the cages as above (23 irregular,
16 cushion, 11 crescent, and six flat galled leaves per cage)
on 19 August 2008, but covered with freshly cut hay. To
each of these cages was added a single S. altissima stem
obviously infected with a pycnidia-forming fungus (pre-
sumably Botryosphaeria spec.) collected from the BCWMA
site. Each infected stem was placed in a bottle of cut-plant
solution, which was not allowed to go dry. The plants were
monitored periodically for the formation of galls and after
29 days all the plants were cut to 30 cm and the cut stems
checked thoroughly for galls.
Gall initiation in screen tents
Two experiments were conducted in an attempt to initiate
galls on S. altissima accessions grown up in the greenhouse
and moved to outdoor screen tents (WSU-2 site; Table 1).
Thirty goldenrod accessions (presumably different geno-
types), 14 of which were the same as those in the field plot
experiments, were divided equally and placed in each of
two 1.8 ·1.8 ·1.8-m screen tents with the floor covered
with black plastic (n = 72 potted plants per tent). The
tents were set up adjacent to one another and the plants
allowed to stand for 8 days to ensure no galls were initiated
during transport from the greenhouse to the screen tents.
One tent (treated) had the floor covered with old golden-
rod stems (previous year’s growth) collected from a field
site. The other tent was a negative control. To provide gall-
initiating adults, each tent was randomly supplied with 48
S. altissima stems infested with a mixture of mature galls
in a 20-l bucket filled with cut-plant solution on 2 July
2008. During collection of these stems, any pupal exuvia
found lodged in the galls were removed. The plants were
checked periodically for galls and finally after 23 days the
plants were cut to 30 cm and the stems and leaves checked
thoroughly for galls. During these periodic checks many
adult midges were seen alighted on the screening in both
tents. The total number of exuvia on the initiating cut-
stem galls was tallied to reveal that at least 111 irregular,
zero cushion, zero crescent, and one flat adult had
emerged in the treated tent. In the control tent at least, 107
irregular, four cushion, two crescent, and zero flat adults
emerged.
In a second experiment started on 13 August 2008, the
same set of plants and tents were used as above, but differ-
ent fungal sources were added to each tent. To one tent
were added three 40-l plastic tubs (15 cm deep) filled with
a plant-fungus-topsoil mixture. Holes were drilled in the
plastic covers and they were elevated 15 cm above the tubs
to provide shade, but allow rain water and female access to
the soil surface. The soil in each of the tubs contained a
mixture of 70 fresh S. altissima stems cut to about 5 cm
long, the agar from 30 oatmeal-agar plates (100 ·15 mm)
of gall-isolated cultured fungus, three handfuls of triple-
ground mulch, and about 10 l of old dried goldenrod
leaves. This mixture was allowed to stand for 2 weeks
before the experiment started. The second tent was sup-
plied with 54 cut S. altissima stems infested with pycnidia-
producing fungus (presumably Botryosphaeria spec.) and
each tent was also supplied with 100 gall-infested S. altis-
sima cut stems. All cut stems were kept in 20-l buckets of
cut-plant solution. Periodically and after 48 days the
plants were checked thoroughly for galls. The total number
of exuvia found on the cut stems was tallied to reveal that
at least 29 cushion, four crescent, five flat, and nine irregu-
lar adults had emerged in the plastic-tub treated tent, and
13 cushion, four crescent, four flat, and one irregular adult
had emerged in the fungus-infested-cut-stem tent.
Results
Biology and life history
The eggs of A. carbonifera are laid on the underside of the
leaf in the vicinity of the meristem (Figure 1B). Females
have up to 300 eggs at the time of emergence and fecundity
appears to differ by morphotype (JJ Heath, unpubl.). The
larvae (Figure 1C) hatch and the fungal spores germinate
(Figure 2) within a few days of oviposition and begin to
burrow ⁄grow into the leaf tissue within a few millimetres
of the ovipositional site. Once the larva has penetrated the
leaf tissue, fungal growth becomes evident on the bottom
and then the top of the leaf. The mean time for fungus to
appear on the top of the leaf was 6.7, 8.0, and 8.2 days for
cushions, flats, and irregulars, respectively. The linear
model with leaf age the eggs were found on as a covariate
provided estimates of the time for fungus to appear on the
top of the leaf, assuming oviposition occurred on zero-
aged leaves. This linear model indicated that there was a
significant interaction between the age of the leaf oviposit-
ed on and the time for fungus to appear on the top of the
leaf as well as significant main effects of leaf age and mor-
photype. The intercepts from this model were 8.6, 8.0, and
7.6 days for cushions, flats, and irregulars, respectively.
Adults (Figure 1A) emerge from the galls approximately
3 weeks after fungal growth is evident on the top of
the leaf. The entire life cycle from egg to adult is about
4–5 weeks. Galls are unisexual with only rare instances of
galls with mixed sexes; this is consistent with the findings
reported by Weis et al. (1983) for this species and for
cecidomyiids in general. In Ohio, larvae may begin to enter
40 Heath & Stireman
diapause as early as the 1st week of September, but galls
continue to be initiated as late as the 1st week of October.
Solidago rugosa galls marked in the field (SC-1 site) in July
and collected in December the same year still had late
instars within them, indicating a very early initiation of
diapause in some morphotypes. However, the physiology
of these larvae may have been altered by undetected
parasitoids. Larvae pupate in the spring and new galls can
be seen forming in late May, but larger populations are not
realized until mid to late June in Ohio. The timing and
details of certain aspects of their biology may vary with
gall morphotype. For instance, controlled experiments
indicate that crescents can oviposit and develop on mature
tissue (JJ Heath, unpubl.).
Asteromyia-fungus interdependency
In the experimental trials designed to test whether
fungal growth and gall development is dependent on
the midge larvae, we found that the galls in which
the larva was removed ceased to grow (Figure 3),
whereas the mock removals and controls were unaf-
fected and continued to develop normally (Figure 3).
However, in those trials where the larvae were
touched with forceps during the mock removals the
larva often died, causing these galls to cease develop-
ment (Figure 3). Dissection of the galls at the end of
the experiment revealed that all the galls that failed to
develop contained dead larvae, whereas developing
galls had healthy larvae.
Asteromyia larvae that were dissected from galls and
placed on agar plates with a portion of their gall fungus
(with no host-plant material) grew more than controls
with no fungal inoculation (F
1,20
= 5.80, P = 0.026; Fig-
ure 4). In preliminary experiments, some of the larvae
appeared to form gall-like structures on the agar plates
(Figure 5A–C). As expected, the difference between the
treatments was smaller when the larvae were larger at the
beginning of the experiment (i.e., a significant interaction
between start length and treatment: F
1,20
=6.60,
P = 0.018; Figure 4). Some larvae died and became com-
pletely deteriorated making a final measurement impossi-
ble; therefore, the sample sizes decreased (larva and fungus
transferred, n = 9; controls, n = 15).
Adult behaviour
Field observations of 15 S. rugosa midges, observed for on
average 2.7 h (maximum: 4.4 h), and 31 irregular midges,
observed for on average 2.3 h (max: 3.8 h), provided no
evidence of mating or conidia collection for either popula-
tion (Figure 6). However, it is clear that females collected
fungal spores somewhere in their environment, as the my-
ABC
Figure 1 Asteromyia carbonifera (A) adult, (B) eggs, and (C) larvae within a dissected gall. Adults and mature larvae are about 1–2 mm in
length. Scale bar on (B) is 200 lm and eggs are typically 180–240 lm long. See online colour version.
Figure 2 Two germinating fungal conidia on an Asteromyia car-
bonifera egg collected from the new growth of a Solidago altissima
plant. Inset shows an enlargement of the conidia. Conidia stained
with lactophenol cotton blue. See online colour version.
Asteromyia-fungus association 41
cangia of Malaise-trapped females always contain conidia
(Figure 7). This suggests that these activities occur some-
time after the period of our observations (Figure 6). A rare
behaviour consisting of touching or dragging the oviposi-
tor on the leaf surface was observed in two irregular
females, which may be associated with conidia collection
(Figure 6B). The leaves were inspected where this behav-
iour occurred and no eggs were found. Males and
females began eclosing in the early morning as the last of
the night’s dew was dried from the leaves (range of
eclosion times: irregulars = 06:05–08:25 hours Eastern
Daylight Savings Time (EDST), rugosa = 07:39–09:00
hours EDST). Males and females of both morphotypes
spent approximately 2.5 h on the bottom of the leaf from
which they eclosed before flying off (mean ± SEM, males:
2.31 ± 0.34 h, n = 6; females: 2.47 ± 0.18 h, n = 22). The
marked galls of both morphotypes were always checked
for missed eclosions before leaving the field site for the
day. In only a few cases were new exuvia or emergence
holes found the following day, indicating that the majority
of adults eclosed during the early morning hours. At most
5% of the marked galls had eclosions on any givenday, but
this was more generally 0–1%. The series of behaviours we
describe (Figure 6) are nearly identical to those provided
in the photographs of Gagne
´(1989); Plate 2, C–F) for a
different gall midge species.
Fungal acquisition and gall initiation
The examination of conidia morphology from different
sources indicated a high degree of phenotypic plasticity in
the shape of B. dothidea fungal conidia. All sources of con-
idia (e.g., gall, stem, agar cultures) produced conidia with
a range of sizes overlapping with those found on A. carbo-
nifera eggs or in their mycangia (Figures 2, 7 and 8), but
there were differences in shape between conidia obtained
from cultured fungus and those obtained directly from
midge eggs or adults. In a full ANCOVA model with all
two- and three-way interactions (first model), the only
CMR
0.0
0.2
0.4
0.6
0.8
mm2day−− 1)
A
B
0
0.012
CMR
Touched
0.002
CMR
0.010
CMR
0.001
CMR
0.003
Figure 3 (A) Mean (± SEM) fungal growth rate (mm
2
per day per larva) after 10 days of incubation; organized by treatment (bars) and
experiment(individual graphs). Bars are labelled with treatment abbreviations: negative control (C), mock removal (M), and complete
larva removal (R). Mock larvae were either ‘touched’ or ‘not touched’ with the forceps in the indicated experiments. The values of small
means are indicated above the bars. (B) Representativebefore and after photographs ordered by experiment. Photographs are scaled pro-
portionately to allow comparison within and among experiments. See online colour version.
42 Heath & Stireman
significant term was the growth media (i.e., goldenrod
stems or oatmeal agar), with conidia being slightly shorter
when grown saprophytically on goldenrod stems (Fig-
ure 8H, top four lines; F
1,403
= 14.0, P<0.001). Tests for
positional effects were not significant. With all seven
conidia sources included in the analysis (second model),
width covaried with length (F
1,552
= 155.4, P<0.001) and
the effect of conidia source was highly significant
(F
6,552
= 190.4, P<0.001). There was no significant inter-
action of conidia width and source, indicating that
the slopes were homogeneous (Figure 8). A priori ortho-
gonal decomposition comparing wild conidia sources
(Figure 8H, bottom three lines) to cultured sources
(Figure 8H, top four lines) revealed that wild conidia were
significantly shorter than cultured conidia (F
1,556
= 1 021,
P<0.001). Further decomposition of only the wild-sourced
conidia concluded that goldenrod-stem conidia were
significantly longer than those from the midge (i.e., eggs or
female mycangia) sources (F
1,145
= 23.5, P<0.001). Addi-
tional decomposition showed that mycangia conidia
obtained from adults at the HS site were also slightly
shorter than egg conidia obtained from the BCWMA and
KWP sites (F
1,56
= 11.9, P = 0.001).
Each of four experiments designed to induce the pro-
duction of galls on the leaves of S. altissima hosts failed;
not a single gall was initiated. However, galls induced by
natural populations of A. carbonifera rapidly appeared in
high numbers on a replicated set of 10 S. altissima acces-
sions in an un-caged field plot immediately adjacent to
the caged field plot (i.e., 435 irregular, 335 crescent, 91
cushion, and 42 flat morphotypes).
Discussion
Time required for appearance of fungus
The experiment to determinethe time for fungus to appear
on the top of the leaf took into account the fact that in
most cases the oviposition event was not actually observed.
This was attempted by incorporating the age of the leaf the
clutch was found on as a covariate. If the eggs were initially
laid on zero-aged leaves, then the time for fungus to appear
on the top of the leaf for a given morph should be the
intercept regardless of the age of the leaf the eggs were
found on. This linear model indicated that there was a
significant interaction between the age of the leaf oviposit-
ed on and the time for fungus to appear on the top of the
leaf as well as significant main effects of leaf age and mor-
photype. However, the intercepts and means of this AN-
0.4 0.6 0.8 1.0 1.2
0.7
0.8
0.9
1.0
1.1
1.2
1.3
Figure 4 Asteromyia carbonifera larval length (mm) after 2 weeks
of growth on malt-extract agar plates treated with Nystatin and
with (closed circles, dashed line) or without (open circles, solid
line) gall fungus added. The fine dotted line has a slope of 1 and
an intercept of 0 and denotes no change in length.
ABC
Figure 5 Asteromyia carbonifera pseudo-gall developing on (A) malt-extract agar. (B)Same as in (A), but it has been dissected. (C) The
same as (B), but at a higher magnification to show the healthy pre-pupa inside. See online colour version.
Asteromyia-fungus association 43
COVA model were only marginally different. Further-
more, one irregular female and one flat female were actu-
ally observed ovipositing on different aged leaves (i.e., ages
0 and 3); therefore, our assumption that eggs were always
initially oviposited on zero-aged leaves did not hold. These
observations make the interpretation of the biological sig-
nificance of this minor statistical interaction difficult. Nev-
ertheless, with all the morphs pooled, the overall mean
time for fungus to appear on the top of the leaf was
7.4 days, which is a good representation of all three mor-
photypes with an accuracy of ±1 day.
The nature of the Asteromyia-Botryosphaeria symbiosis
The results of these studies indicate that this insect-fungal
system is closely knit, with strong interdependence of the
fungus and the midge. This association appears to be obli-
gate for the gall midge and with respect to the successful
production of galls on S. altissima. The midge depends on
the fungus for food (Janson et al., 2009) and protection
from parasitoids (Weis, 1982b), and the fungus benefits
from dispersal and requires the midge for hyphal prolifera-
tion within the context of the gall. Although the fungus
likely persists independently in environments outside the
A
B
Figure 6 Ethograms of Asteromyia carbonifera behaviour under natural field conditions. Galls marked and observed for (A) Solidago rugosa
morphotypes and (B) Solidago altissima irregular morphotypes. Behaviour was observed from eclosion to fly-off. The thickness of the
arrows is proportional to the relative frequency of the particulartransition from one behaviour tothe next. Numbers along arrows are the
absolute number of transitions observed.
44 Heath & Stireman
context of the midge galls and could have additional
modes of dispersal, within the context of these galls, midge
dispersal is necessary. As the gall fungus only rarely pro-
duces sporulating pycnidia, the benefits (or the costs, for
that matter) to the fungus of associating with the midges
remain unclear. It is possible that the midge issimply para-
sitizing the fungus with no reciprocal benefit. However,
teasing out the costs and benefits of apparently mutualistic
associations can be tricky (e.g., Herre & West, 1997); the
true nature of this interaction will become clearer with a
better understanding of the life histories of both players.
This system is not unlike other insect-fungal associations
involving Ambrosia beetles, fungus-gardening ants, and
other flies. Recent work on other galling insects including
cecidomyiids has found that they actively manipulate plant
defensive chemistry (Tooker & De Moraes, 2007, 2008;
Tooker et al., 2008). This manipulation may be a factor in
allowing the fungus to proliferate and form the protective
gall-like structure.
Although several larvae died when transferred from
galls to agar plates, this experiment still revealed the
ability of larvae to grow on a fungus-only diet, strongly
supporting the hypothesis that the fungus is the larva’s
primary food source within the gall. Even in young galls,
we have observed that mycelium quickly envelops the
larva making direct access to the plant tissue difficult.
Furthermore, sterol analysis of fungi, plants, and midge
larvae indicates that the midge larvae obtain their sterols
mainly from the fungus (Janson et al., 2009). The obser-
vation that fungal structures resembling galls formed on
the agar plates, suggests that the larva is primarily
responsible for the general gall structure. Feeding dam-
age and ⁄or salivary-gland secretions are likely responsi-
ble for the formation of the hard, black, carbonaceous
material (stroma) that formed on agar plates in areas of
larval grazing. This material was similar to what sur-
rounds and protects the developing larva from desicca-
tion and parasitism in a natural gall. Larew et al. (1987)
describe a cecidomyiid from a lineage evolutionarily
basal to Asteromyia (Bissett & Borkent, 1988) that feeds
only on fungus and forms similar galls in the absence of
a host plant, suggesting that the ability to form fungal
galls is a plesiomorphic trait. Haridass (1987) was able
to rear Neolasioptera cephalandrae Mani larvae to adults,
solely on gall-isolated fungus growing on Petri dishes,
although they observed high mortality as well.
It is clear from our consistent observations of specific
conidia associated with female ovipositors and eggs that
A. carbonifera females intentionally obtain fungal conidia,
store them in mycangia, transport them, and deposit them
on their eggs. The fungus in these Asteromyia galls has been
identified as B. dothidea (Bissett & Borkent, 1988; Janson
et al., in press), a member of the family Botryosphaeria-
ceae. Other fungi assigned to this genus and even species
are primary and secondary plant pathogens on many
important ornamental and horticultural crops. For exam-
ple, grapes, avocados, oaks, apples, pears, olives, Prunus
species, poplars, pines, ashes, elms, and various berries are
known to harbour or present disease symptoms associated
with Botryosphaeria spp. (Bonfiglioli & McGregor, 2006).
Botryosphaeria ribis is thought to form cankers on the
stems of goldenrods (Horst, 2008). Many studies have pro-
vided evidence that ascospores of this family are primarily
wind dispersed, whereas conidia are primarily water dis-
persed (Sutton, 1981; Pusey, 1989; Ko & Sun, 1995; Ahi-
mera et al., 2004). To our knowledge, only two studies
have provided evidence for this active transportation of
fungi by gall midges (Borkent & Bissett, 1985; Adair et al.,
2009). No studies have investigated the direct role that
cecidomyiids may play in vectoring Botryosphaeria spp. to
other plants.
Gall initiation, conidia collection, and eclosion behaviour
Gall initiation. Although we were unsuccessful at initiat-
ing galls under semi-artificial conditions, we did discover
pycnidia-producing fungus on the stems of S. altissima.
However, two experiments including exposure of midges
to these infected stems did not result in the initiation of
Figure 7 Dorsal view of a malaise-trap-captured female
Asteromyia carbonifera showing mycangia filled with fungal
conidia. Conidia were stained with lactophenolcotton blue.Inset
is an enlargement of one of the mycangia (scale bar = 20 lm).
Normally, alarge proportion of the conidia would be distributed
toward the anterior as well, but pressure from the slide mount
forces the conidia toward the posterior in what appears to be
tubes or specialized folds of tissue that lead the conidia to their
point of deposition on the passing egg. See online colour version.
Asteromyia-fungus association 45
galls, suggesting that these stems are not their conidia
source.
It is possible that midge behaviour was constrained
in the enclosures. Many insects and vertebrates have
behaviours that are fixed action patterns (Matthews &
Matthews, 1978). A fixed action pattern is a sequence
of stereotypical behaviours that is relatively indivisible.
This indivisibility can be so strong as to prevent subse-
quent behaviours until the sequence is completed.
Furthermore, a number of these fixed action patterns
can be under hierarchical control, so that the occur-
rence of one may be required for initiation of another
(Matthews & Matthews, 1978). It is possible that our
field cages may have prevented behaviour, such as
medium range dispersal, that may be necessary before
conidia collection can occur. However, the field setting
and use of large cages in these experiments strongly
suggests that the failure of gall initiation is most likely
attributable to a lack of the appropriate source of
conidia.
A
B
C
D
E
F
G
H
Figure 8 Length and width (lm) of individual conidia sampled from different sources. The top four panels show conidia obtained from
either (B, F) gall or (A, E) egg-conidia fungalisolates, grown on either (A, B) oatmeal agar (oa) or (E, F) fresh-cut autoclaved Solidago
altissima stems atop water agarplates (wa-s). These four sources were grown under the same conditionsat the same time. The other sources
are (C) Asteromyia carbonifera female mycangia, (D) pycnidia on field-collected S. altissima stems, and (G) field-collected A. carbonifera
eggs. (H) The final panel is a composite overlay of all the graphsto facilitatecomparisons. The insets are representative photographs of
some of the conidia from each source. The scale bar (20 ·5lm) in (A) applies to all the insets. Conidia stained with lactophenol cotton
blue. See online colour version.
46 Heath & Stireman
Eclosion behaviour and mating. Our observations indi-
cated that females did not obtain conidia from their galls
as they eclosed. Observations of several female mycangia
from each of the four morphotypes after they had spent at
least a day in containers containing tens of galls never
revealed conidia within their mycangia (n = 35). This
appears to be the case in other Ambrosia gall-forming
cecidomyiids (Adair et al., 2009). Our field observations
also failed to reveal any evidence of behaviour associated
with conidia collection during or after eclosion. Further-
more, the exuvia effectively shields the ovipositor of the
eclosing female, making conidia collection from within
the gall unlikely. Perhaps most importantly, no pycnidia
were ever observed within or on the outside of mature galls
during eclosion.
Many cecidomyiid species have sexually dimorphic
antennae with the males being more plumose (Gagne
´,
1989). In some of these species sex-specific pheromones
have been identified (Heath et al., 2005). These species
tend to mate almost immediately after eclosion (McKay &
Hatchett, 1984; Pivnick & Labbe, 1992; van Lenteren et al.,
2002; Heath et al., 2005; Suckling et al., 2007). The anten-
nae of A. carbonifera are not sexually dimorphic and
attempts to attract males or females with virgin conspecif-
ics or actively sporulating B. dothidea fungus have failed
(JJ Heath, unpubl.). On several occasions, males and
females of the same morphotype were seen emerging from
galls in the field at the same time and very close to one
another (i.e., within 5–30 cm), but mating was never
observed. One may postulate that this is a result of
inbreeding avoidance, but females have single-sex families.
Therefore, these observations suggest that males and
females of A. carbonifera may aggregate at the conidia col-
lection site or that adults become more attractive after they
have collected fungal conidia. Although A. carbonifera are
diurnally synchronized in emergence, populations are not
strongly synchronized seasonally (B Wells, pers. comm.).
In our field eclosion observations, over 100 galls were
marked, but less than 5% of these produced eclosing adults
on any 1 day. Therefore, the probability of having a male,
female, and a sporulating fungal structure present simulta-
neously may be quite rare. Future gall initiation studies will
concentrate on one gall morphotype to increase the proba-
bility of the co-occurrence of these factors.
Conidia morphology. The morphology of conidia from
different sources demonstrates that B. dothidea produces
pleomorphic conidia and that this is affected by the growth
media. The fact that egg-conidia isolates produce long
slender conidia when grown as a saprophyte (i.e., on
oatmeal agar or autoclaved goldenrod stems) even though
the isolates originated from a population of short conidia
(i.e., egg-conidia isolates), suggests that the natural source
of conidia is not of a saprophytic nature. If the females
obtain conidia from a saprophytic source one would
expect these conidia to be long and slender, rather than the
observed ovoid shape. The morphology of the conidia
derived from pycnidia occurring on S. altissima stems in
the field was more ovoid and overlapped more with
those found in mycangia and on eggs, suggesting these as a
possible source. However, the conidia source may be from
some other plant growing in the same environment as
goldenrod, such as blackberries. Botryosphaeria dothidea is
known to attack blackberries and produce numerous
pycnidia on their stems (Maas & Uecker, 1984) that
overlap slightly in size with those found in themycangia. It
is also possible that the midges somehow select smaller
conidia mechanically or via odours; though the benefit of
this selectivity is unclear. We plan to generate isolates of
the S. altissima stem fungus and conduct genetic analysis
to determine its relationship to B. dothidea. Although
this may be a possible source, the distribution of these
pycnidia in the field is patchy and efforts to find them
early in the season, when A. carbonifera galls were preva-
lent, have failed. They are much more abundant later in
the season, when the goldenrod has started to form flower
buds.
Although, mycangia and egg conidia were slightly differ-
ent in morphology, this difference pales in comparison to
the phenotypic plasticity of the fungus indicated by com-
paring the morphology of egg-derived conidia to the mor-
phology of conidia derived from egg-conidia isolates in
culture. Furthermore, the mycangial conidia and egg coni-
dia came from different field sites, which may be responsi-
ble for the small but significant difference in morphology.
The conidia we found in the mycangia of A. carbonifera
females were very similar to those described by Bissett &
Borkent (1988) for a variety of Ambrosia cecidomyiids
that all use Botryosphaeria spp. to form their galls. This
consistency in conidia morphology across Ambrosia gall
midge taxa and a wide geographical range, suggests that
conidia morphology is key to understanding where these
midges obtain their conidia. However, the phenotypic
plasticity associated with substrate makes it imperative
that wild sources of conidia be measured. Once a source
with similar morphology is found, genetic profiling and
behavioural experiments will be necessary to verify its role
in this system. Detailed genetic profiling might also be
used to differentiate between a single fungal source and
random collection by comparing the genetic variation of
single-conidia isolates isolated from the same mycangia to
those isolated from a single fungal reproductive structure.
We can now say that A. carbonifera is one of a number
of complex Ambrosia gall midge mutualisms that hold
Asteromyia-fungus association 47
great promise to provide insight into the contribution that
mutualistic relationships make to adaptive radiation and
ecological speciation. Practically speaking, mating behav-
iour and conidia collection remain areas of future work.
Without this knowledge manipulative studies to assess
mating, host plant, and gall morph fidelity will be difficult
and direct studies involving reproductive isolation and
hybrid fitness will be nearly impossible. These issues not-
withstanding, A. carbonifera and other Ambrosia gall mid-
ges remain tantalizing model systems for studying a range
of evolutionary phenomena.
Acknowledgements
Seth Jenkins helped with the gall-area measurements.
Diego Javier Inclan Luna collected A. carbonifera adults
from the malaise trap. Both plus Hilary Devlin, Brenda
Wells, and Patrick McAfee helped transplant the golden-
rod accessions to the field. Eric M. Janson provided gall
fungal isolates. Jennifer L. Heath facilitated the field and
laboratory studies and provided helpful comments on the
manuscript. The manuscript was significantly improved
by the comments of two anonymous reviewers. This
research was supported by an NSF grant DEB-0614433 to
JOS and the Wright State University Environmental
Science PhD programme.
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