Download trition - American Entomologist

nity Ecology
trition
Michael G. Kaufman, Edward D. Walker,
David A. Odelson, and Michael J. Klug
This overview shows that microbial partnerships between
insects and microorganisms are widespread, complex,
and meet a variety of insect needs.
s can be said fondly of many human relationships, associations between insects and
microorganisms go back a long way. Basic
nutritIonal deficiencies, biochemical needs, and even
sexual compatibility requirements in insects are met
through microbial "partnerships" (Bourtzis and
O'Neill 1998, Moran and Telang 1998). These relationships, falling into the original broad definition of symbioses as coined by de Bary (1879), are
widespread, varied, and, in many cases, reflect a
pronounced history of co-evolution (Buchner
1965, Moran and Telang 1998). Although there is
also an impressive record of what might be considered "unhealthy" insect-microbe relationships (i.e.,
microorganisms that are pathogenic or parasitic
to insects and other hosts [see Steinhaus 1967,
Lysenko 1985, Federici and Maddox 1996]), our
focus will be on relationships benefiting insects in
some fundamental way, particularly nutritional
mutualisms and the exploitation of microorganisms as food items. The term 'microorganism' and
its derivatives used here will refer mainly to bacteria, fungi, and protozoa, with the acknowledgment
that many groups of viruses and algae also interact
closely with a variety of insect species. Bacterial,
fungal, and protozoal associations with insects have
been included in previous reviews (e.g., Buchner
1965, Steinhaus 1967, Boush and Coppel 1974,
Breznak 1984a, Jones 1984, Campbell 1989, Prins
and Kreulen 1991, Kane 1997, Douglas 1998). It
is our intent to highlight several examples from a
slightly different perspective, stressing the complexity and dynamic nature of many insect-microbe
relationships as revealed through current techniques used in microbial ecology.
There are two aspects of insect-microbial associations that warrant reconsideration, particularly
from the viewpoint of entomologists. First, it is
important to recognize that many relationships are
formed between insects and diverse microbial communities, not the simple one-on-one interactions
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so ingrained in the concept of mutualisms. The
wide range of insect taxa known or presumed to
interact nutritionally with microbial communities
is extensive and includes representatives from all
major insect groups (Table 1). Second, the microbial communities (or their components) are not
necessarily constant in composition or functionality through the course of the interaction. The idea
that particular microbial taxa always are associated with particular insect taxa is compelling, and,
in the cases of most intracellular endosymbionts
and pathogens, this specificity has been verified
(Federici and Maddox 1996, Douglas 1998,
Moran and Telang 1998). However, in the cases of
insect-microbial community interactions the microbial end of the relationship may be composed of
scores of unknown microbial species, and verification of taxa specificity has been impractical. Even
so, the constancy of these microbial assemblages
often is assumed because stability is implicit in the
concept of an evolved community. Yet, currently,
there is little direct evidence to support this assumption in the vast majority of insect-microbial systems and increasing evidence to suggest fluctuation
in both microbial composition and function.
In insect-microbial community interactions,
overall functionality has been examined more often than individual microbial components. It can
be argued that this functionality is the only important level of interaction for the insect partner in a
mutualism. However, Harris (1993), in a review of
aquatic invertebrate-microorganism associations,
emphasized that these communities should not be
treated as "single functional entities," but that "individual populations require examination" because
those particular populations clearly are instrumental in any efficient functionality. An understanding
of the microbial details is, therefore, a fundamental
prerequisite for understanding insect-microbial
community interactions. Further, much of the literature-base concerning microorganisms and in173
Table 1. Insect taxa known to or likely to have strong nutritional interactions with microbial communities.
Order
Families or subtaxa
Interaction type
Selected literature sources or reviews
Thimm et at. 1998
Collembola
all?
detritus/biofilm microbes', gut microbes, fungivores
Protura
all?
detritus/biofilm microbes
Diplura
all?
detritus/biofilm microbes
Microcoryphia
all?
detritus/biofilm microbes
Thysanura
all?
detrituslbiofilm microbes
Ephemeroptera
Ephemeridae,
Heptageniidae,
Leptophlebiidae,
(plus other
detritivorous taxa)
detritus/biofilm microbes, gut microbes
Grylloblatidae
?
detritus/biofilm microbes
Phasmida
Orthoptera
Blattaria
Gryllidae,
Gryllotalpidae,
Acrididae
all
Isoptera
all
Meitz 1975, Cummins and Klug 1979,
Austin and Baker 1988, Lawson and Klug 1989
Embiidina
phylloplane microbesb gut microbes (?)
Cazemier et at. 1997
detritus/biofilm microbes, gut microbes,
phylloplane microbes
Martoja 1966; Hunt and Charnley 1981;
Mead et al. 1988; Kaufman et al. 1989;
Santo Domingo et at. 1998a, b
detritus/biofilm microbes, gut microbes
Bignell 1977; Cmden and Markovetz 1987;
Zurek and Keddie 1996, 1998; Kane 1997
detritus/biofilm microbes, gut microbes,
fungal cultivarslfungivores
O'Brien and Slaytor 1982, Breznak and Brune
1994, Kane 1997
detritus/biofilm microbes?
Plecoptera
Pteronarcidae,
Peltoperlidae,
Taeniopterygidae,
Nemouridae,
Leuctridae,
Capniidae
detritus/biofilm microbes, gut microbes
Meitz 1975, Sinsabaugh et at. 1985,
Lawson and Klug 1989
Zoraptera
?
gut microbes, detritus/biofilm microbes?
Choe 1992
Psocoptera
detritus/biofilm microbes
detritus/biofilm microbes?
Phthiraptera
Heteroptera
Reduviidae,
Lygaeidae,
(plus other
herbivorous taxa)
gut microbes, phylloplane microbes
Homoptera
?
phylloplane microbes
Thysanoptera
phylloplane microbes, fungivores
sects has dealt with obligatory functional associations (i.e., those in which at least one partner could
not survive or reproduce independently). However,
many relationships between organisms are more
facultative and variable in nature. Mutualisms often are conditional, with cost/benefits to each partner varying spatially and temporally and with
environmental and nutritional status (Keeler 1981,
Bronstein 1994, Herre et a!. 1999). Currently, the
prevalence of conditionality in insect-microorganism associations is unknown, but the common occurrence of facultative mutualisms in nature
suggests a need to explore this issue.
Because of technical limitations in characterizing microbial communities, their study has lagged
behind that of "macro" community ecology. For
entire generations of ecologists, microbial communities have been represented as a "black box" of
populations and/or processes. The individual components and detailed mechanisms within these conceptual black boxes still are poorly understood,
largely due to inadequate methods, but the lid is
174
Jones 1984, Beard et at. 1998, Benedict et al.
1991, Hackstein and Stumm 1994
being pried off with newly developed technologies.
Traditional methods of culturing and identifying
microorganisms from the environment (including
insects and insect habitats) fail to account for more
than 85% of the organisms present (Amman et a!.
1994, Tiedje 1995). Fortunately, tools have become available that allow closer examination of
the microbial players in situ. Molecular methods
such as ribosomal RNA-targeted probes, 165 RNA
gene sequencing, and polymerase chain reaction
(PCR) have been added to the traditional arsenals
of microbial cultivation techniques, advanced microscopy, and phenotypically based identification
systems (Pace 1996). Indeed, many of the specific,
intracellular microorganisms found in insects are
resistant to cultivation outside of the insect and
must be examined with these novel methods
(Amman et a!. 1994, Baumann and Moran 1997).
We will illustrate with several examples that looking at microbial communities from the entomological point of view as a constant-function black
box containing a few "loyal" microbial species is
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Order
Families or subtaxa
Interaction type
Megaloptera
Corydalidae
Coleoptera
Scara baeidae,
detritus/biofilm microbes, gut microbes,
Scolytidae,
phylloplane microbes, fungivores
Silphidae,
Bostrichidae,
Lagriidae,
Ipidae,
Elmidae,
Cerambycidae,
Chrysomelidae,
(plus other herbivorous,
wood-boring, and
detritivorous taxa)
gut microbes
Selected literature sources or reviews
Meitz 1975
Cummins and Klug 1979, Bayon and Mathelin
1980, Jones 1984, Benedict et al. 1991,
Hacksteinand Stumm 1994, Vasquez-Arista et
al. 1997, Cazeimer et al. 1997
Siphonaptera
?
Diptera
Tipulidae,
detritus/biofilm microbes, gut microbes,
Chironomidae,
phylloplane microbes, fungivores
Tephritidae,
Drosophilidae,
Anthomyiidae,
Culicidae,
Simuliidae,
Ceraropogonidae,
(plus other herbivorous
and detritivorous taxa)
Cummins and Klug 1979, Klug and Kotarski
1980, Starmer and Fogleman 1983, Sinsabaugh
et al. 1985, Kaufman et al. 1986, Lawson and
Klug 1989, Drew and Lloyd 1991, Merritt et al.
1992, Clyde 1996
Trichoptera
Limnephilidae,
Lepidosromatidae
(plus other
detritivorous taxa)
detritus/biofilm microbes, gut microbes
Meitz 1975, Cummins and Klug 1979,
Sinsabaugh et al. 1985, Lawson and Klug 1989
Lepidoptera
Galleriinae
(plus herbivorous and
wood-boring taxa)
phylloplane microbes, gut microbes
Jones 1984, Campbell 1989, Hackstein and
Stumm 1994, Benedict et al. 1991, McKillip
et al. 1997
Hymenoptera
Apidae,
fungal cultivars/fungivores, detritus/biofilm microbes Martin 1987, Caetano 1989, Gilliam 1997
? (epidermal microbes?)
Siricidae,
Formicidae
Only representative
"Microorganisms
layer on more inert
bMicroorganisms
famlies are listed in the cases of orders where a high percentage of taxa are likely to have strong interactions.
(principally bacteria, fungi, and heterotrophic protists) associated with decaying organic matter, or those forming an organic
substrates (e.g., rock surface).
(principally bacteria and fungi) inhabiting the surfaces of leaves on living plants.
antiquated, and, further, that opening the black
box is essential to understanding how many insect-microbe interactions work. The ability of new
techniques to enhance our resolution of viewing
microbial community structure can aid our understanding of the subtle and dramatic dynamics in
insect-microbe relationships that ultimately impact
the dynamics of insect populations.
Microbial Communities in Insect
Digestive Tracts
The primary insect habitat for microorganisms
is the digestive tract (Bignell 1984, Hackstein and
Stumm 1994, Cazemier et al. 1997), and the termite gut is the most intensively studied example.
The fascinating mutualism illustrated by the interdependence of these insects and a complex assemblage of microorganisms (mainly protozoans and
bacteria) in their digestive tracts is now a classic
illustration of animal-microbial interaction. The
relationship is built around the metabolism of refractory diet components (i.e., wood) by a diverse
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microbial community in the case of lower termites,
and a more balanced attack of insect enzymes and
microbial activity in the higher termites (O'Brien
and Slaytor 1982, Breznak 1984a, Breznak and
Brune 1994). Microorganisms (presumably different groups) are involved intimately in carbon acquisition
(cellulose breakdown),
nitrogen
acquisition (fixation of atmospheric nitrogen), and
nitrogen recycling (uric acid metabolism) in the termite gut (Breznak 1984a). Aside from the key feature of celluloytic enzymes, the system is driven by
anaerobic metabolism of wood carbohydrates by
protozoa (lower termites) and bacteria (lower and
higher termites). The mutualism works because
wood fiber components are oxidized incompletely
to acetate that subsequently is assimilated as an
energy source and lipid precursor in termite tissue
(Breznak 1984a).
Theoretically, a single species of microorganism
would be adequate for transforming cellulose to
acetate in a termite gut. Anaerobic protozoans in
lower termites do just that. Why then, does the
175
termite gut contain hundreds of microbial species,
most of which are yet to be described fully (Okhuma
and Kudo 1996)? Part of the answer may lie in the
multiple functions of the gut community. In lower
termites, for example, nitrogen fixation and recycling are accomplished by various bacterial groups,
whereas protozoans handle the bulk of cellulose
degradation. Oxygen-scavenging activities of facultatively anaerobic bacteria are important in maintaining low redox conditions that ultimately make
the mutualism work (Tholen et al. 1997). Additionally, different microbial groups play different
roles in the flow of carbon compounds as they are
degraded. A consequence of anaerobic degradation of carbohydrates by microorganisms is that
hydrogen gas (Hz) is produced during regeneration of electron acceptors such as oxidized nicotinamide adenine dinucleotide (NAD+). If hydrogen
gas accumulates, even in low concentrations, the
reactions regenerating electron acceptors back up,
and the entire decomposition process slows down.
However, if Hz is kept low by hydrogen consumers, the overall process can proceed more efficiently
(Fenchel and Finlay 1994).
In the termite gut and other anaerobic environments, hydrogen consumers fall into two major
categories: methanogenic bacteria and acetogenic
bacteria. Both groups appear to co-exist in many
termite species, but their relative proportions and!
or activities vary greatly across taxonomic and feeding categories (Brauman et al. 1992, Wheeler et al.
1996, Sugimoto et al. 1998). Which group predominates, however, has important implications
for termite nutrition. Methanogens use COz and
Hz to make methane, whereas acetogens, among
other varied metabolic capabilities, use COz and
Hz to produce more acetate. Both aid in speeding
the processing of wood sugars, but the acetogens
supply additional acetate (up to one-third of respiratory demand) for the termite. Clearly, production of acetate over methane is advantageous to
termites and, not incidentally, represents much less
of a concern to humans than a potent greenhouse
gas. Recent work has detailed some of the dominant methanogens and acetogens within the gut of
Reticulitermes flavipes (Leadbetter and Breznak
1996, Leadbetter et al. 1999), but the relative distribution of various hydrogen-consuming bacteria continues to be investigated.
The preceding only touches on part of the diversity found in the termite/microbe symbiosis.
Within the termite gut, in a sort of Chinese puzzle
box of murualisms, protist mutualists harbor their
own endosymbiotic bacteria (e.g., methanogensLee et al. 1987, Fenchel and Finlay 1994) and external symbionts that aid in their motility (Breznak
1984b). Much of the overall diversity has yet to be
described, but modern molecular methods are revealing the phylogenetic relationships of the spirochetes and novel bacteria found in termites
(Okhuma and Kudo 1996, Paster et al. 1996,
Leadbetter et al. 1999) and the potential contributions of these microorganisms to the functionality
of the system.
176
In contrast to termites, several omnivorous
groups of insects have developed associations with
gut microbial communities in which the interdependency is less pronounced and could be described
best as supplemental. Well-studied examples include
the cockroaches (Cruden and Markovetz 1987),
whose gut microorganisms are incompletely described but are dynamically responsive to diet and
growth stages in the animal (Kane and Breznak
1991, Zurek and Keddie 1998). As in termites,
part of the nutritional relationship between cockroaches and their gut microbial communities appears to involve anaerobic degradation of plant
polymers and the activities of hydrogen-consuming bacteria, particularly methanogens (Hackstein
and Stumm 1994, Zurek and Keddie 1998). Relative proportions
of bacterial
groups and
methanogenic activity change dramatically when
.he insects change diets (Kane and Breznak 1991,
Gijzen et al. 1994, Zurek and Keddie 1998), suggesting that an adaptable and responsive community of microorganisms may playa role in the
nutritional ecology of roaches.
Crickets also serve as an example of insects
whose diet breadth might be expected to affect the
dynamics of microbial symbionts in their digestive
tracts. Most species of gryllids and gryllotalpids
harbor dense populations of bacteria in the anterior regions of their hindgut (Martoja 1966, Fig.
1; Ulrich et al. 1981). These bacterial communities
metabolize dietary carbohydrates and produce
volatile fatty acids that can be used by the host.
Interestingly, only gryllotalpids appear to harbor
methanogenic bacteria (Kaufman 1988, Santo
Domingo et al. 1998a), whereas acetate-producing
hydrogen consumers are present in both field and
mole crickets (Kaufman 1988). The bacterial activity probably is not essential to the insect; however,
it aids in the overall efficiency of the animal to use
ingested material and may "buffer" the insect's use
of carbohydrates from variable dietary sources
(Kaufman et al. 1989, Kaufman and Klug 1990).
Some evidence also suggests that the bacterial community competes with the host for nitrogen when
dietary N sources are scarce (Kaufman et al. 1989).
Thus, both the community and its role may change
during the life of the insect.
We employed the amplified ribosomal DNA restriction analysis (ARDRA; see Massol-Deya et al.
1995) technique to examine changes in overall community composition of cricket hindgut microorganisms in response to diet and to determine if the
response was primarily enzymatic or involved a
shift in dominant bacterial populations. This technique essentially is an restriction fragment-length
polymorphism (RFLP) fingerprint of 16S RNA
genes amplified with PCR from whole-community
microbial DNA. It originally was used to distinguish individual bacterial species but is applicable
for comparisons of dominant populations in relatively simply communities (Martinez-Murcia et al.
1995, Massol-Deya et al. 1995). House crickets,
Acheta domesticus (L.), were reared in groups to
adults on a standard basal diet (Purina Cricket
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Gut wall surface
Cross-section
Fig. 1. Cricket digestive tract and associated bacteria in anterior hindgut (AGH) region. Bacterial cells fill the
entire space between the gut wall (gw) and peritrophic membrane (pm) surrounding the food bolus (fb) in the
cross-sectional view, and coat the interior gut wall and associated structures. CR, crop; PV, proventriculus;
GC, gastric cecum; V, ventriculus (midgut); MT, Malphigian tubules; PHG, posterior hindgut; R, rectum.
(drawing from R. Martoja [1966] with permission of publisher).
Chow, Purina Mills, St. Louis, MO) and then
switched to one of three new diets. The diets, sugar
beet pulp (high carbohydrate, low protein content),
alfalfa (intermediate carbohydrate and protein content), and a casein-based artificial diet (high protein, low carbohydrate content) differed from each
other and the basal diet in major digestible components (Santo Domingo et a!. 1998a, b). A set of
control animals was reared individually for the
same period but kept on the original chow diet.
Individual anterior hindguts were removed, homogenized, and subjected to freeze/thaw extraction procedures to release microbial DNA. Primers
for highly conserved regions of the 165 ribosomal
gene (Massol-Deya et a!. 1995) were used to generate a PCR product that was digested subsequently
with restriction endonucleases. Gel electrophoresis
yielded a pattern of DNA fragments that was quantified with Gelcompar image analysis and software
(Applied Maths BVBA, Kortrijk, Belgium). Band
position and intensity values were analyzed with
multivariate statistical methods (principal component analysis [PCA]) to compare gene fragment
patterns of hindgut microbial communities from
each diet treatment quantitatively. PCA reduces
complex variable sets and groups samples along
axes that describe the overall variability in the data.
In this case, samples with similar gel patterns will
group, whereas samples that differ in band position and intensity will separate. Distances between
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samples in the PCA plots are, therefore, indicators
of the degree of genetic similarity in the microbial
communities from different cricket individuals.
Principal component analysis groupings of
cricket ARDRA patterns (Fig. 2) showed a separation of samples from three of the four diet treatments. Diet changes from cricket chow to beet pulp
or alfalfa caused similar changes in microbial composition, whereas a switch to a casein-based diet
elicited a much different response in the dominant
bacterial groups. The general trends in samples
suggest that genetic patterns of dominant hindgut
community members can be related to diet composition along a carbohydrate gradient. The beet pulp
and alfalfa both had relatively low protein and
higher carbohydrate content than either the basal
chow or casein-based diets. The basal chow diet
was intermediate in that it was mainly carbohydrate based, but contained a higher proportion of
protein than pulp or alfalfa. Interestingly, the pattern from a freshly collected field cricket, Gryllus
pennsylvanicus Burmeister, specimen grouped between the chow and alfalfa/pulp diet treatments of
A. domesticus. This suggests an overall similarity
in microbial community composition in crickets
that cuts across species lines yet may be altered
fundamentally by diet habits of individuals.
In a related study, the microbial community
structure of the cricket hindgut was investigated
by using fluorescently labeled, rRNA-targeted oli177
•
1.2
1
0.1
•
•
o
)(
pulp dlOl
.I'alf.
dlOl
.•.
•
chow diet
proteln dlOl
lIeld crickOl
.•.
.•.
0••
...11
.•.
0.1
1.2
1
0••
0.1
•
0
0••
0.2 PC 3
0
0.2
-0.2
-0••
-0.2
-0.1
-0••
, -0.1
Fig. 2. Principal component analysis of ARDRA
patterns generated from microbial communities in
hindgut of A. domesticus adults fed different diets.
Diets are as follows: pulp = sugar beet pulp; alfalfa =
alfalfa hay: chow = Purina cricket chow; protein =
casein-based artifical diet. See text for more details.
The percent total variance explained by PC 1, 2, and
3 was 30.4, 14.6, and 11.8, respectively.
gonucleotide probes. This technique involves the
construction of molecular probes from variable
regions of the 16S rRNA genes designed to detect
specific taxonomic groups of bacteria and allows
enumeration of these groups using epifluorescence
microscopy. Using this technique, Santo Domingo
et a!. (1998a) were able to assess the number of
Bacteroides-Prevotella type organisms in the hind-
Fig. 3. Epifluorescent
micrograph
of bacteria in
hindgut of A. domesticus. Bacteria stained with a
"universal" DNA-binding fluor (DAPI) (A), and the
same field viewed after probing with 168 rRNAtargeted oligonucleotide
with Bacteroides-type
of
178
J.
probe designed to hybridize
bacteria (B). Photos courtesy
Santo Domingo (Santo Domingo 1994).
gut of A. domesticus (Fig. 3). By using probes targeting archae bacteria (e.g., methanogenic bacteria)
and several major groups of eubacteria ("true"
bacteria), they found that both diet and species
could affect the composition of the hindgut communities, that differences were most-pronounced
between the families Gryllidae (house and field crickets) and Gryllotalpidae (mole crickets), and that
the probes used still accounted for less than 50%
of bacteria observed in direct counts. Archeabacteria were found only in species of gryllotalpids, an
observation consistent with reports that these animals are methanogenic, whereas house and field
crickets are not (Kaufman 1988, Hackstein and
Stumm 1994, Santo Domingo et a!. 1998a). Interestingly, both diet and gut structure differences in
mole crickets may account for the microbial community differences
(Kaufman 1988, Santo
Domingo et a!. 1998a).
The dynamic nature of microbial populations
in omnivores may be one key to explaining the
adaptability of these animals. For crickets, cockroaches, and similar insects, having a flexible packet
of microbial diversity may be one way to deal with
inconsistent and unpredictable dietary resources.
The idea that microbes serve to mediate diet variability has been put forth more eloquently elsewhere (e.g., Jones 1984) and has been discussed in
terms of plant-herbivore interactions (Barbosa et
a!. 1991). For crickets, gut bacteria serve to spare
some dietary carbohydrate needs (Kaufman et a!.
1989) and handle dietary changes (Kaufman and
Klug 1991). The primary metabolites (i.e., volatile
fatty acids) provided by microorganisms to the insect partner do not change markedly with diets
(Kaufman and Klug 1990). Therefore, the insect is
assured a more or less constant supply of these
compounds across a range of conditions. This consistent supply is maintained almost certainly because the microbial populations involved are not
constant.
Although a flexible microbial community would
be advantageous to a mobile omnivore or generalist herbivore, important questions arise about the
extent of the flexibility (e.g., how much is the community a function of local environmental or diet
conditions?). Do gut microbial populations change
from within, or does each different diet introduce
novel members that then become established because they are pre-adapted to the new food item?
To date, these questions remain open, but diet effects on gut microorganisms are well known from
studies of mammalian systems (Lee 1985). Even
so, the specifics of microbial population responses
often are unclear and the degree of flexibility in the
communities is debatable.
Insects That Feed upon Microbial
Communities
Microorganisms often comprise or modify the
primary food source for insects. For example, there
is evidence to suggest that a wide range of herbivorous insects are influenced by phylloplane (leaf surface) microbial communities and associated
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actIvItieS that alter host plant characteristics
(Barbosa et al. 1991, Benedict et al. 1991). Fungal
cultivation by some ant and termite groups is another classic example of animal-microbe nutritional
interactions. These insects exhibit an almost total
reliance upon the tissues and enzymes of specific
fungi (Martin 1987).
Detritivores are a widespread functional group
of insects and other invertebrates that are less particular in their consumption of particular microbial species but are nonetheless dependent upon
microbial biomass and/or activities for growth and
development. Detritivores are well represented in
many insect taxa (Table 1). An entire guild of
aquatic detritivores known as "shredders" are tied
fundamentally to the microbial community on decaying plant material (Cummins and Klug 1979).
The presence of this mainly bacterial and fungal
community is apparently essential to shredders,
and components of the community can strongly
influence feeding and developmental rates. It is wellestablished that for several of these detritivores,
some species of aquatic hyphomycetes (a fungal
group) are palatable and serve as nutritional resources, whereas others are unacceptable and deter growth (Barlocher 1985). The constituent fungal
species relevant to detritivore growth are known in
a few cases, but the remaining microbial components (bacteria and protozoans) are almost entirely
undescribed.
The larva of the cranefly Tipula abdominalis
Say (Diptera: Tipulidae) is a particularly interesting example of an insect detritivore. This shredder
not only requires microorganisms and microbial
conditioning of decaying leaf material for development, it also receives internal assistance from gut
bacteria (Klug and Kotarski 1980, Lawson and
Klug 1989). A complex assemblage of bacteria and
fungi on leaf material stimulates normal feeding
and growth, whereas fungi alone appear to be inadequate (Lawson et al. 1984). This complex microbial assemblage can change substantially during
leaf decay and presents somewhat of a "moving
target" for a fastidious detritivore like Tipula
(Lawson et al. 1984). Interestingly, recent preliminary investigations (Clyde 1996) suggest that the
gut bacterial community in T. abdominalis changes
in response to stream habitat and/or diet. Therefore, the microbial communities in the food source
may be directly or indirectly influencing microbial
communities internally-Tipula's nutritional physiology is sandwiched almost literally between the
dynamics of diverse microbial groups.
In most cases, the quantitative role of microbial
communities in the nutrition of insect detritivores
is unknown because these insects ingest large portions of the leaf substrate along with the associated
microbial biomass. However, there is one group of
insects that unquestionably relies primarily upon
decay-associated microorganisms for food. Larval
mosquitoes, particularly those in the genus Aedes,
are known to filter microorganisms from the water column and to browse them from surfaces in a
variety of small aquatic habitats (Merritt et al.
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1992). They feed indiscriminately on microorganisms within a size range of particles from 1 to 50
11M (Merritt 1987). The quantity and composition of these microbial food sources are likely the
limiting factors for mosquito development in a wide
variety of natural habitats.
We have examined microbial communities in
the larval habitats of Aedes triseriatus Say. This
mosquito breeds in small containers formed by
buttressing roots at the base of trees, rot holes
formed when limbs break off, and in discarded
tires and other containers inadvertently provided
by humans. A. triseriatus is a vector of La Crosse
encephalitis virus, and its larval feeding ecology
and habitat are similar to the more notorious Aedes
aegypti L., carrier of dengue and yellow fever viruses, and the Asian tiger mosquito, Aedes
albopictus (Skuse), a competent vector of several
human diseases and recent invader to the Americas. Larval habitat energetics are driven by leafmaterial input and other allochthonous organic
debris. Larval development can be influenced
strongly by the amounts and type of this material
(Carpenter 1983, Walker et al. 1997), but this influence is indirect because the organic particulates
are subjected first to microbial decay and incorporation into microbial biomass. In treehole systems,
the principal microbial components appear to be
fungi (aquatic hyphomycetes, yeasts), protozoans
(ciliates and small flagellates), and heterotrophic
bacteria. All of these groups, to some extent, are
consumed by mosquito larvae to the point where
their populations are noticeably or measurably
Fig. 4. Scanning electron
exposed to feeding by A.
excluded from larvae (8).
each panel. Photos by J.
micrograph of leaf surfaces
triseriatus larvae (A), or
Magnification is identical in
Schuette.
179
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larval effect
70
Larvae present
larvae absent
p = 0.016
Larval effect
p
= 0.006
60
t/)
Q)
50
as
(5
t/)
40
as
30
0
•••
~
20
0
10
0
Pseudomonaceae
Enterobacteriaceae
BACTERIAL GROUP
Fig. 5. Composition
presence
of cultivable
water column bacterial groups in response to the
of larval A. triseriatus.
Results
of ANOVA showing
significant
larval effects
are
listed in the figure.
16
«
14
u.
::J
0..
•
Larvae present
•
Larvae absent
Larval effect
Habitat effect
Interaction
12
Z
«X~
10
(.)...J
.....
"Cl
Zc::
O~
...J
...J
8
6
«
I-
4
I-
2
0
0
Microcosms
Fig. 6. Concentrations
Treeholes
Tires
of long chain (>20 carbons)
polyunsaturated
in larval A. triseriatus habitats. Results of ANOVA comparison
show highly significant
differences
fatty acids (PUFAs)
are presented
in box and
in habitat and larval main effects.
depressed either in treeholes or similar systems
(Fish and Carpenter 1982, Walker et al. 1991,
Cochran-Stafira and von Ende 1998).
As part of a long-term investigation of larval
mosquito interactions with the microorganisms in
their habitats, we are examining microbial community dynamics in the container habitats of A.
triseriatus. In general, larval feeding depresses total abundance of bacteria in both the filtered zone
of the water column and on leaf surfaces (Walker
et al. 1991, Kaufman et al. 1999; Fig. 4). Larval
feeding
also decreases
protozoan
and
microeukaryote abundance in larval feeding zones
(Kaufman et al. 1999; E.D.W., unpublished data).
Effects on fungal populations are less consistent,
probably because much fungal tissue is located
within the detrital matrix and is not consumed directly by larvae (M.G.K., unpublished data).
Apart from overall effects on abundance, larval
feeding can alter microbial community structure in
the environment such that the nutritional composition of food sources changes and microbial ac180
tivities associated with inorganic nutrient dynamics in the systems are affected. In the case of bacterial communities, we have shown that larval
presence can increase the relative proportion of
facultative anaerobes in general and enteric bacteria in particular (Kaufman et al. 1999; Fig. 5). Presumably, these groups are more resistant to
ingestion/digestion than other bacterial groups, illustrating that as larvae feed they are changing their
critical nutritional link to the particulate organic
matter that drives the system. This impact on microbial community structure also can alter microbial metabolism of nitrogen compounds, thereby
affecting the dynamics of ammonium (a substance
potentially toxic to larvae) and supplies of nitrogen for growth of decay organisms at the base of
the food web (Kaufman et al. 1999).
Aedes triseriatus also can reduce protozoan
populations substantially and may overconsume
this source of essential lipid nutrients. Dietary polyunsaturated fatty acids are necessary for development of mosquitoes and probably all other
arthropods (Dadd 1981, Brett and Muller-Navarra
1997). Polyunsaturated fatty acids are characteristic of eukaryotic organisms and in small heterotrophic habitats, such as treeholes, are most
likely biochemical indicators of protozoans (Vestal
and White 1989, Kaufman et al. 1999). In recent
studies using fatty acid profiles as indicators of
microbial community composition, we found that
larval feeding significantly reduced concentrations
of polyunsaturated fatty acids in various habitats
(Kaufman et al. 1999; Fig. 6). This reduction corresponded well with our observations of drastic
declines in protozoan abundance when larvae were
present (E.D.W., unpublished data) and may indicate a key factor controlling mosquito production
in these habitats.
Larval consumption of protists not only serves
to diminish a highly valuable nutritional resource
but can feedback on bacterial community composition as well. An unidentified bacterial type appears in treeholes when larvae are not present
initially or when they have been removed experimentally (Fig. 7). The large size and filamentous
nature of these bacteria render them resistant to
grazing by protists (Jurgens and Gude 1994) that
have proliferated in the absence of mosquito larvae. We can only speculate on how these bacteria
might affect processes within container habitats or
whether they represent a potentially valuable food
resource for mosquito larvae that disappears along
with protozoans when larvae feed. From a broader
perspective, the interaction of invertebrates in the
microbial food webs of aquatic systems is an active
area of research that may benefit from studies of
mosquito larvae feeding upon their microbial resources (Bott 1995, Cochran-Stafira and von Ende
1998).
Concluding Remarks
The preceding not only is relevant toward an
understanding of what leads to mosquito growth
in various habitats but, also, has some serious im&"1ERlCAN
ENTOMOLOGIST
•
Fa/l2000
ments from several anonymous reviewers greatly
improved the manuscript. Portions of this research
were funded by NSF (BSR-8706480) and the National Institutes of Health (AI21884).
Fig. 7. Large bacterial cells seen in treehole habitats
from which larval A. triseriatus had been removed.
Note representatives
(arrows) of more typically
sized bacteria seen in these habitats. Photo by W.
Morgan.
plications for the successful engineering and release of novel microbial pesticides developed for
mosquito control. Several such organisms have been
proposed as transgenic vectors for the Eti toxin
(Porter et al. 1993). For these novel organisms to
provide effective long-term control, they must be
able to compete with existing microorganisms,
be
ingested and digested by mosquito larvae, and be
resistant to predation by protists and other organisms when larval abundance is low. All require a
working knowledge of microbial community dynamics and composition within natural larval habitats.
The importance of understanding
the structure
and changes in microbial communities associated
with insects is becoming increasingly clear. These
dynamics often are critical in interactions ranging
from complex mutualisms to simple food acquisition and encompass a wide range of insects and life
stages. Beyond the examples discussed here, microbial community structure plays an important
role in how insects deal with ingested toxins (e.g.,
Jones 1984, Campbell 1989), survive overwintering (Lee et al. 1996), survive pathogens (Dillon and
Charnley 1995), communicate
with conspecifics
(Leufven 1991), and transmit pathogens to other
species (Jones 1984, Beard et al. 1998, Watanabe
et al. 1998). Microbial community ecology has been
an important factor in shaping insect evolution,
and it is evident that entomologists'
conceptions
also must evolve to recognize and understand this
interaction.
~
Acknowledgments
We gratefully acknowledge the technical assistance of Bob Murray and Ann Bagdley (Central
Michigan University, Mt. Pleasant), and Tracy Smith
(Michigan State University, East Lansing). We also
thank the National Science Foundation's
(NSF)
Center for Microbial Ecology at Michigan State
University for facilities and support and are indebted to the Annales Entomologie de France for
permission to reproduce a published figure. ComA~lERICAN E1\'TOMOLOGIST
•
Volume 46, Number 3
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Zurek, 1., and B. A. Keddie. 1998. Significance of
methanogenic symbionts for development of the
American cockroach Periplaneta americana. J. Insect Physiol. 44: 645-651.
Michael Kaufman is visiting assistant professor at
Michigan State University's W. K. Kellogg Biological Station (Hickory Corners, MI 49060) and department of Entomology (E. Lansing, MI48824).
In addition to his interests in insect-microbe interactions, he teaches and does research in the field of
stream ecology. Edward Walker is an associate professor of entomology at Michigan State and is the
resident medical entomologist (East Lansing, MI
48824). He has broad interests in the ecology of
vector-borne
diseases
and in insect ecology
and
behavior. David Odelson is a molecular microbial
ecologist who originally worked on biochemical
aspects of the termite-gut microbe symbiosis and is
now directing research and application of microbial treatments of soil problems at Eco Soil Systems, Inc. (11890 Thornmint Road, San Diego,
CA 92127). Michael Klug is a professor of microbiology at Michigan State and director of the W. K.
Kellogg Biological Station. He has wide-ranging
research interests in microbial ecology including
insect-microbe interactions, soil and sediment microbiology, and bioremediation.
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184
AMERICAN
ENTOMOLOGIST
•
Pal/2000
Know
-
KnOWledge
r
is your best weapon in the battle
against urban and structural pests; and the best
source of that knowledge
is the Handbook of Urban
The handbook
than 100 color photographs,
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Edited by Roger E. Gold and Susan C. Jones, this
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• Urban Entomologists
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• Human Dangers
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A~IERICAN ENTOMOl.OGIST
•
Volume 46, Number 3
185