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THE ROLE OF AN AEROBIC BACTERIAL METABOLISM

IN THE NUTRITION OF CRICKETS (ORTHOPTERA: GRYLLIDAE)

by

Michael G. Kaufman

A DISSERTATION

submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCI‘OR OF PHILOSOPHY

Department of Entomology

1988

ABSTRACT

THE ROLE OF AN AEROBIC BACTERIAL METABOLISM
IN THE NUTRITION OF CRICKETS (ORTHOPTERA; GRYLLIDAE)

by

Michael G. Kaufman

The relationship between members of the family Gryllidae and
the bacterial community in their anterior hindgut was investigated
from two perspectives. In one approach, A. domesticus larvae were
reared with (conventional) and without (germfree) microorganisms
under different conditions of diet quality. In general, growth rates
and measures of adult fitness were affected more by diet quality
than by the presence of bacteria. In all cases, however, conventional
larvae digested more, consumed less, and converted a greater portion
of available food into biomass than germfree counterparts. Based on
measures of excretion rates of uric acid and total N, the mechanism
of increased food utilization efficiency was not related to nitrogen
metabolism by gut bacteria. Digestibility values of major nutrient
classes indicated that the gut bacteria bacteria acted principally on
soluble carbohydrates. Enzymatic profiles also revealed that the
bacterial community enhanced carbohydrase activity in general and
produced at least one enzyme class not found in germfree crickets.

In the second approach, the overall bacterial community
metabolism was investigated with a focus on carbon-based
fermentation products. Volatile fatty acids (VFA) were found to be

produced in quantities estimated to account for 75% of the observed

food utilization advantage observed in conventional crickets (ca 400
nmol/ cricket/ hr.). VFAs were consistently found in the anterior
hindguts of a number of gryllid species, and over a range of diets
and rearing conditions. The compounds were produced far in excess

of their excretion rates, and were shown to readily transverse the
hindgut wall. Acetate was the dominant acid formed and evidence
indicated that a substantial portion of acetogenesis (ca 30%) could be
derived from C02 reduction with H2. The bacterial community/cricket
symbiosis, therefore, appears to be based upon carbon conservation
mechanisms via anaerobic bacterial metabolism. Gut microbes
increase the metabolic diversity of crickets and allow a more

efficient use of dietary carbohydrate resources.

ACKNOWLEDGMENTS

I've been exceptionally fortunate to have had the opportunity to
interact with many outstanding scientists and teachers during my
graduate program. I gratefully acknowledge the advice and
encouragement from committee members Drs. J. Breznak, M. Martin,
W. Mattson, and J. Miller. I'm particularly grateful to Dr. Rich Merritt
for his unwavering support, and especially to Dr. Mike Klug who
provided the means and inspiration to continue. I also wish to thank
Dr. Dan Lawson for invaluable discussions, computer consultation,
and friendship. I greatly appreciated the technical assistance of C.
Fulmer, G. Walker, and L. Jorissen, and thank G. Eldred and Dr. T.
Walker for providing me with the insects. Finally, I wish to thank my
wife, Suzanne, whose support and patience were the mainstay of my

efforts.

iv

TABLE OF CONTENTS

List of Tables ........................................................................................ vi
List of Figures ...................................................................................... vii
Introduction .......................................................................................... 1

Chapter 1 Growth and food utilization parameters of germfree Acheta
domesticus (Orthoptera: Gryllidae) ........................................... 24

Chapter 2 The production of volatile fatty acids by the bacterial
community in crickets (Orthoptera: Gryllidae): potential

significance to the host .......................................................... 66
Synopsis ........................................................................................... 110
AppendixI Early growth studies ........................................................... 116
AppendixII Whole insect respiration ....................................................... 118
List of References ................................................................................ 120

LIST OF TABLES

Table1.l Adult size, maturation time, mortality, and fecundity values
for germfree and conventional A. domesticus on three diets.
Adult weights were taken 5 days after the final molt. Values are
mean + S. E. (n). ..................................................................... 36

Table 1.2 Overall digestibility and biomass gain per 100 mg of food
consumed by germfree and conventional A. domesticus during
the larval stage. Values are combined estimates from all growth
periods. All values are mg dry weight. ............................................ 40

Table 1.3 Nitrogen content of combined body tissues of last instar A.
domesticus larvae. Values are mean %N (S.E.), n = 6. ........................ 41

Table 1.4 Composition of crude nutrient fractions in diets and fecal material,
and approximate digestibility estimates for each fraction in
germfree and conventional A. domesticus during growth period

4. Values are mean + S. E., n = 6. ................................................. 45
Table 1.5 Percent solubilization of 14c-1abened cellulose in excised cricket

digestive tracts. Mean (S.E.) n = 3 for all cases. ................................. 48
Table 2.1 VFA excretion rates in last instar larvae of conventional A.

domesticus . ........................................................................... 84
Table 2.2 Hydrogen and methane in headspace of gut incubation vials after

4 hrs. Values are mean i SE. (n). ................................................ 85
Table 2.3 Percentage of added 14002 fixed under various conditions of gut

incubation. Mean (S.E.), n = 4-6. ................................................. 85
Table 2.4 Comparison of gut VFA in various animal groups. .............................. 96

Table AI.1 Adult size (10 days beyond molt) and maturation
times of first generation germfree and conventional A.
domesticus . Values are mean 1'. SE. (n). ....................................... 117

Table AI.2 Adult size (10 days ), fecundity, mortality, and maturation

times of second generation germfree and conventional A.
domesticus .Values are mean 3; S.E.(n). ......................................... 117

vi

Figure 1.1
Figure 1.2

Figure 1.3

Figure 1.1

Figure 1.2

Figure 1.3

Figure 1.4

Figure 1.5

Figure 2.1

LIST OF FIGURES

Gryllid digestive tract. From Martoja (1966). ................................ 14

Scanning electron micrographs of the luminal side of the

anterior hindgut wall of A. domesticus. (A) Conventional

adult 3 weeks after molt; 200x. (B) Germfree adult 3

weeks after molt; 200x. (C) Close -up of setacaceous

papillae in conventional adult; low mag 540X, inset

4500X. (D) Corresponding view of (C) in germfree

version; low mag 540X, inset 3000X. (E) Close-up of wall

between papillae in conventional adult; low mag 540X,

inset 2700K. (F) Corresponding view of (E) in germfree

version; low mag 180X, inset 2000K. ........................................ 15

Transverse section through anterior hindgut of germfree
and conventional A. domesticus adults (SEM, freeze-
fracture prep.). Material between peritrophic membrane
(pm) and ileum wall (il) in conventional animal is
bacteria and associated extracellular matrix. Bar =

lOOuM. ........................................................................... 17
Growth of germfree and conventional A. domesticus
larvae on three diets. Bars are 8.13., n = 25-30. ............................. 35

Digestibility, food utilization efficiencies, and relative

consumption rates for germfree and conventional A.

domesticus larvae on three diets. Periods are growth

intervals illustrated in Figure 1.1. Bars are S. E., n = 6-10. ............... 38

Total nitrogen (N) and uric acid (U A) excreted by

germfree and conventional A. domesticus larvae on

three diets. Periods are growth intervals illustrated in

Figure 1.1. Bars are S. E., n = 6-10. ........................................ 42

Activity levels of major enzyme classes in the digestive
tract of germfree and conventional A. domesticus . ......................... 47

Percent survival of germfree and conventional A.

domesticus reared on diets with alfalfa hay. Top -

survivorship when hay is alternated with basal diet

(diet one). Bottom - Survivorship on 100% alfalfa with

and without growth supplements. ............................................. 63

VFA concentration in the anterior hind gut of various
gryllid species. Bars are S. E., n = 6-14. ..................................... 75

vii

LIST OF FIGURES (cont)

Figure 2.2 Production rates and % composition of VFA in the
anterior hindgut of three gryllid species. Each point in
the production cm've is a single gut replicate. Error bars
are SE, n = 7-14. .............................................................. 76

Figure 2.3 Comparison of VFA production rates and composition in
anterior hindguts of lab-reared and field-collected G.
rubens. Each point in the production curve is a single gut
replicate. Error bars are S. E., n = 7-8. ....................................... 78

Figure 2.4 Percent composition of VFA in the ectoperitrophic fluid
of the anterior hindgut of A. domesticus.. Error bars are
SE, n = 5. ....................................................................... 79

Figure 2.5 Production rates and composition of VFA in the anterior
hindgut of young (20-25 days) A. domesticus larvae.
Each point in the production curve is a single gut
replicate. Error bars are S. E., n = 8. ......................................... 81

Figure 2.6 Production rates and composition of VFA in the anterior
hindgut of last instar larvae of A. domesticus reared on
four diets. Each point in production curve represents a
single gut replicate. Error bars are S. E., n = 4-20. ......................... 82

Figure 2.7 Distribution of fixed 14C02, specific activity of
partitioned label, and total VFA formed per gut during
"suspension" method incubations with and without
CHCL3 block. H2 = hydrogen atmosphere, BLOCK =
hydrogen atmosphere and CHCL3 added to the
suspension. ....................................................................... 86

Figure 2.8 Distribution of fixed 14002. specific activity, and total
VFA produced per gut in "whole gut " incubations. N2 =
nitrogen atmosphere, H2 = hydrogen atmosphere. .......................... 88

Figure 2.9 Rate of movement of 14C-labelled VFA across the
anterior hindgut wall of A. domesticus in virro. Each
point represents the percentage of added label found in
the saline incubation medium. .................................................. 91

Figure 2.10 Comparison of linear best fit model with computer
interpolation curve of VFA production in A. domesticus
reared on a diet of 100% alfalfa hay. ......................................... 102

Figure AI.1 Larval growth curves of germfree and conventional A.
domesticus reared on diets one and two, for two
successive generations. ........................................................ 1 16

viii

INTRODUCTION

Insect-microbe symbioses1 are widespread and almost as
diverse as the Class Insecta itself. These associations have been
cataloged since the late 1800's, and several books and literature
reviews attest to the multitude of insect species that serve as "hosts"
to various types of microorganisms (Buchner 1965, Koch 1967,
Richards and Brooks 1958, Breznak 1982, Houk and Griffiths 1980).
In spite of nearly a century of active research in the area, and the
detailed work with several particular groups of insects, most of our
present knowledge of insect-microbe associations is descriptive.
Given the estimate that less than 20% of insect species are thought to
be known to man, and that only a fraction of these have been
examined for specific microbial symbionts, it is safe to conclude that
our ignorance in the area far exceeds our understanding.
Investigations of these symbioses are fascinating and valuable from
the pure science perspective, and have been instrumental in
addressing questions of nutritional ecology, co-evolution, and
microbial ecology. With the advent of pesticides derived directly

from microorganisms and the need for non-chemical control agents,

 

1 Within the context of this discussion, symbiosis and subcategories will follow
the broad definitions of Starr (1971) and Buchner (1965) referring to the close
physical association of two different species of organisms throughout all or
part of their life cycles.

1

2

microbial symbionts of insects present a potential opportunity for
manipulations leading to control of particular insect populations; an
idea that has intrigued insect pathologists for decades (Lysenko
1985). At the very least, our knowledge of insect physiology and
ecology - an invaluable prerequisite to any applied control
measures - must include an awareness of the potential contributions
of microorganisms.

The types of insect-microbe symbioses range from tightly-linked
mutualisms and parasitisms; where the relationship between
microorganism and insect is species-specific and obligate, to
facultative associations in which neither member demonstrably
benefits nor suffers from the relationship. In most cases the
association has not yet been classified in terms of its degree of
interdependence. Microorganisms can be categorized as
ectosymbionts (living outside the body) or endosymbionts (living
within the body of the animal). Endosymbionts can be further
subdivided into extracellular or intracellular forms. The type of
association is thought to indicate the degree of
specialization/interdependence of the symbioses; with intracellular
forms representing the extreme of mutualistic specialization and
presumably a more highly co-evolved situation (Buchner 1965, Koch
1967). The evolution of intracellular endosymbionts is generally
thought to have progressed from a condition of extracellular
microbes living in the digestive tract to microbes becoming an
integral component of specialized cells or groups of cells
(mycetocytes). Microbes housed in specialized regions of the gut or

contained in specialized organs within the hemoceol (mycetomes)

3

represent intermediate forms. Microorganisms housed in
mycetocytes are transferred from generation to generation of insect
via elaborate mechanisms, and in many ways must be considered
inseparable components of the insect tissue. Examples include those
found in several families of Homoptera( Houk and Griffiths 1980),
Coleoptera (Buchner 1965), and Orthoptera (Brooks 1970, Richards
and Brooks 1958). Although intracellular endosymbionts clearly
represent a high degree of specialization, several other types of
associations suggesting highly evolved symbioses occur in those
relationships classified as ectosymbioses or extracellular
endosymbioses. The relationship between bark beetles (Scotylidae)
and certain fungi instrumental in the beetles' utilization of wood is
one example (Graham 1967), and recent studies have demonstrated
the importance of acquired enzymes from particular fungal
ectosymbionts in the nutritional ecology of several insect groups
(Martin 1984). Some extracellular endosymbioses are also thought to
be highly specific relationships; an example being the association of
unique protozoans with lower termites (Breznak 1982).

Although the insect-microbe associations which attract the most
attention are those in which some specific interdependence or
pathology is clearly suggested, the vast majority of associations
cannot yet be classified in terms of any functional interaction. Even
among the specific intracellular endosymbioses there is little
empirical evidence in many instances that demonstrates the "role" of
the microorganism. There is also some controversy regarding the
validity of evidence reported by researchers investigating the

functional roles of microbial symbionts. Buchner's (1965) treatise on

4

endosymbionts points out several sources of misinformation and
erroneous interpretations from earlier work. More recently, Brooks
(1970) has questioned much of the work with bacteroid symbionts in
cockroaches, and Campbell and Nes (1983) have indicated that
certain functional roles of aphid endosymbionts in cholesterol
metabolism may have been misrepresented. Hill et al. (1976) have
presented evidence that does not support the reported role of
Rhodnius symbionts in B-vitamin production. Additionally, several
assumed mutualistic associations in dipterans have been shown to be
less obligatory than previously thought (Schneider et. al., 1983,
Howard et al .1985). This is not to dismiss the aforementioned
examples as inconsequential symbioses; clearly further research is
required to examine additional hypotheses about the relationship.
It is rather to underscore the point that our understanding of even
the best-known associations is fragmentary and sometimes based
upon untested assumptions.

Microorganisms are ubiquitously found as residents of insect
digestive tracts. It is currently unknown how many insect-gut
microbe associations represent any type of symbiosis within the
definitions employed here, but in general they are more apt to be
facultative and non-vital associations. Many gut microbes are
merely transients associated with ingested material; eventually
ending up as nutritional sources themselves or being passed out with
undigested material. In several insect groups, however, microbes
proliferate in the gut tract and become permanent or semi-
permanent symbionts. Ironically, gut microbes in insects are

probably both the best and least—understood of non-pathological

5

insect/microbe associations. Research on termites has elucidated a
story of nutritional interdependence that has become a classic
example of mutualism. Termites and their intestinal protozoa and
bacteria represent a highly-specialized, extracellular endosymbiosis.
However, in most other insects that possess a permanent gut flora,
mutualism has yet to be conclusively demonstrated. Further, in most
other insect groups, the constancy and function of alimentary tract
microorganisms is completely unknown.

Bignell (1984) discusses the insect digestive tract as an
environment for microorganisms and points out that the highest
densities of microbes and the more permanent populations are
usually found in the hindgut region. The hindgut in insects is well-
suited to microbes in many cases since it presents a relatively stable
environment, presents a source of nutrients, and is osmotically
favorable to most groups. The region is often differentiated
morphologically and houses dense, multispecies communities of
microbes. These hindgut modifications are variously termed
fermentation chambers, diverticuli, paunches, or cecae (McBee 1977,
Buchner 1965). The relative size and extent of development of
hindgut modifications in insects is strongly correlated with diet
(Buchner 1965). Known examples of insects with modified hindgut
regions include all families of Isoptera, Scaraboidea, and the
Tipulidae, Blattidae, and Gryllidae.

One of the principal roles which has been demonstrated and
often postulated for gut microbes in insects is the degradation of
refractory polymers that are components of the insects' diet. The

digestion of native (crystalline) cellulose is a complex process

6

involving the action of at least three enzymes, and, with the possible
exception of silverfish (Lasker and Giese 1956), it has not been
conclusively demonstrated that any insect possesses the capability to
completely digest the compound without the aid of one or more
microbial enzymes (Martin 1983). In the case of lower termites and
woodroaches, digestion of native cellulose in their wood-based diets
is accomplished principally by flagellated protozoans inhabiting the
hindgut (Breznak 1982, O'Brien and Slaytor 1982, Hogan et al. 1988).
The lower temite, Coptotermes lacteus, produces enzymes active
against soluble forms of cellulose in addition to those supplied by the
gut protozoa (Hogan et al. 1988). Higher termite groups, which lack
the protozoan component in their hindgut microbial community, are
thought to synthesize some cellulose-degrading enzymes in their own
tissue, and the activities of hindgut bacteria do not appear to be
involved (O'Brien and Slaytor 1982). Bacteria in the hindguts of
cockroaches (Periplaneta americana) are thought to be responsible
for the utilization of carboxymethylcellulose by the insect (Bignell
1977, Cruden and Markovetz 1979). Cellulase activity was not
specifically demonstrated in Oryctes nasicornis (Scarabaeidae)
larvae by Bayon (1980); however, she indicates that hindgut
microbial fermentation activity in the larvae is unaffected when 3-
cellulose is supplied as the only diet constituent. Hindgut bacteria in
the field cricket, Gryllus bimaculatus , and house cricket, Acheta
domesticus , have also been implicated in the degradation of cellulose
(Martoja 1966, McFarlane and Distler 1984).

Although cellulose metabolism is the most widely-investigated

of gut microbe activity, the roles of bacteria in the hindguts of insect

7

groups may include the utilization of other polymers. Hemicelluloses
are apparently degraded by hindgut bacteria in termites (Breznak
1982) and by bacterial isolates from the grass grub, Costelytra
zealandica (Coleoptera: Scarabaeidae) (Bauchop and Clarke 1975),
but little is known about the importance of these activities in viva.
Chitin can be utilized by some isolates from cockroach hindguts
(Cruden and Markovetz 1987), but there is no evidence to suggest
that such a capability is quantitatively important in the insect, since
roaches produce their own chitinase (Bignell 1981, House 1974).
Less-resistant polysaccharides such as starch, and most
oligosaccharides, fall within the metabolic capabilities of many gut
bacteria isolated from insects, and these substrates are also normally
degraded by insect enzymes. In general, gut microbes in insects are
likely to contribute to the diversity of carbohydrases present (House
1974 and references therein), but research thus far has been limited
in its focus.

Microorganisms in insect hindguts also are known to contribute
to the nitrogen economy of the host. Again, the termite literature
provides much of the evidence. Nz-fixation, an activity restricted to
prokaryotic organisms, has been demonstrated in several species of
termites from both higher and lower families (Prestwich et al. 1980,
Breznak 1982, O'Brien and Slaytor 1982). The capability to fix
atmospheric nitrogen in these insects is clearly related to the low
nitrogen content of their wood diet and the activity can be reduced
by supplementing the diet with additional nitrogen sources (Breznak
1982). The resulting available nitrogen could account for substantial

portions of termite growth requirements, depending upon the

8
species, caste, and diet involved (Breznak 1982 Bentley 1984). N2-
fixing bacteria have been isolated from cockroaches (Cruden and
Markovetz 1987), however its significance to the host is doubtful.

Recycling of uric acid nitrogen via gut microbes has also been
demonstrated in termites by Potrikus and Breznak (1981). The insect
is itself incapable of degrading the compound and hindgut bacteria
are responsible for making a portion of the carbon and nitrogen from
uric acid available to the termite in the form of acetate and
ammonia (Potrikus and Breznak 1981). A similar mechanism has
been postulated for cockroaches which, like termites, store uric acid
internally (Cochran 1985), however mycetocyte symbionts are
usually invoked as the main uricolytic sites in the insect. Uricolytic
bacteria have been isolated from cockroach hindguts (Cruden and
Markovetz 1987) and house cricket iliea (Ulrich et. al. 1981), but
their in situ importance remains to be demonstrated.

'The metabolism of other nitrogenous compounds by hindgut
bacteria in insects is even less understood. The synthesis of some
amino acids and the composition of termite protein and free amino
acid pools is affected by the presence of gut microbes (O'Brien and
Slaytor 1982)). It has also been demonstrated that gut microbes
from cockroaches and crickets are capable of converting cysteine to
methionine (Bignell 1981, Martoja 1966); a conversion not thought to
generally occur in insect tissue (Dadd 1985). It has also been
frequently postulated that gut bacteria supply B-vitamins to insects
(House 1974, Wigglesworth 1972, Dadd 1985). However, evidence
had come mainly from studies of Rhodnius and its gut symbionts

(Baines 1956, Lake and Friend 1968), and the role of gut microbes in

9

this insect has been subsequently questioned by Hill et al. (1976).
Although it seems quite clear that hindgut microorganisms could
offer several classes of nitrogenous compounds to the host, it has
rarely been shown that they would be available to the insect. Studies
with mammals indicate that nitrogenous compounds from hindgut
microbes are normally unavailable to the animal without coprophagy
or some other means of harvesting gut microbe cellular components
(McBee 1977, Reddy et al 1968).

Other possible nutritional roles of gut microbes are largely
speculative. Incorporation of sulfate into the amino acids methionine
and cysteine by gut bacteria has been demonstrated in a few insects
(Martoja 1966, Dadd 1985), but the quantitative significance is
unknown and the presence and availability of sulfate in normal diets
has yet to be shown. Clayton (1960) has indicated that gut bacteria in
roaches supply sterols, however, no bacteria are known to synthesis
this class of lipids. Bacterial metabolism of other lipids in insect
digestive tracts has not been investigated, nor has their possible
influence on mineral metabolism, osmolarity maintenance, and
excretion. The latter are a potentially important considerations in
view of the hindgut/Malpighian tubule interface and the normal
function of most hindgut tissue in resorption of water and soluble
material (Bradley 1985, Phillips et al 1986).

Hindgut bacteria in insects may also interact beneficially in ways
not directly related to nutrient metabolism. The production of
aggregating pheromones by bark beetles (Scotlylidae) (Borden 1982),
locusts (Nolte 1977), cockroaches (Mcfarlane and Alli 1986), and

house crickets (McFarlane er al. 1983) has been attributed to

10

metabolites produced by hindgut bacteria. However, Charnley et al.,
(1985) have recently presented evidence to contradict this
contention in the case of locusts. It has also been suggested that gut
microbes may act to detoxify ingested compounds (Jones 1984,
Guthrie and Apple 1961). This is an interesting hypothesis that
deserves investigation, however it currently remains untested.
Among other possible non-nutritional roles, the inhibition of
pathogenic microoganisms by endemic gut microbes has been of
interest to invertebrate pathologists, but little effort has been
specifically directed to this question. Dillon and Charnley (1986) have
shown the bacterial community in locust hindguts inhibits the
germination and viability of the fungus Metarhizium anisopliae
Proliferation of certain pathogenic bacteria is inhibited by members
of the "normal flora" (Goodwin 1967, Lysenko 1985). Yet as Lysenko
(1985) notes, studies of the normal flora of insects have "received
surprisingly little attention" .

The roles of gut microorganisms in vertebrate groups have been
studied more intensively, due in large part to the interest of
agricultural (livestock) scientists and the medical research
community. The subject is beyond the scope of this discussion and
has been reviewed elsewhere (Luckey 1969, Gordon and Pesti 1971,
Savage 1977, 1984, Hungate 1966, 1975, Wolin 1979). All of the
same gut microbe metabolic characters discussed above for insect
systems have been documented in vertebrate systems. Further, more
detailed and specific examples of metabolic interactions with the
host have been shown. Indeed, many hypotheses concerning the

activities of hindgut microbes in insects originated from studies of

ll

vertebrates. Research on vertebrate gut systems have also placed a
much greater emphasis on interaction between members of the gut
microbial community; particularly as it relates to pathogen resistance
(Savage 1977, Lee 1985). In spite of the relatively intense research
in the area, understanding of gut microbe/vertebrate associations,
like similar research with insects, suffers from a lack of breadth in
the type of organisms examined. Information has been heavily
skewed toward ruminants and laboratory rodents. There are no
doubt many parallels between vertebrate and invertebrate gut
microbe systems, however the basis of comparison is currently
restricted to a few examples.

The preceding discussion has emphasized the diversity of insect
microbe associations, the potential metabolic capabilities of gut
microbial communities, and the paucity of information available on
all but a few systems. Within insect groups in particular, research in
the area has focused upon obligate microbial associations in species
feeding upon incomplete diets. Even the excellent and detailed
microbiological characterization of the cockroach hindgut (Cruden
and Markovetz 1987) has not clarified the relationship between
bacteria and this omnivorous species. An intriguing association of
bacteria with the anterior hindgut of crickets (Gryllidae,
Gryllotalpidae) has been investigated thus far in only a descriptive
manner. This insect hindgut/bacteria system is provocative and
deserving of investigation because the association is non-obligate
(Martoja 1966) and the insect is not restricted to particular,
incomplete diets. Additionally, unlike roaches, nutritional

relationships between gut microbes and insect would not be clouded

12

by the presence of intracellular symbionts as no cricket species are

known to harbor such symbionts.

I! .1 I. l .

Crickets are members of the order Orthoptera and as such
represent an ancient group of hemimetabolous insects. There are
over 2400 species in the family Gryllidae distributed worldwide in
temperate and tropical regions (Alexander 1968). The most familiar
groups are the subfamilies gryllinae, nemobiinae, and gryllotalpinae
(house and field, ground, and mole crickets, resp.). Most gryllid
species are associated with grassland and field soils, and mole
crickets are almost exclusively subterranean. House crickets (Acheta
domesticus), an introduced species in the United States, inhabit
human dwellings and refuse dumps in Europe (Bates 1969,1971). As
a group, crickets are omnivorous but generally consume plant
material (Tennis 1983, VanHook 1971, Gangwere 1961, Walker and
Masaki in press). The list of potential diet items is long and varied,
and very little is known about the normal feeding and foraging
behaviour of these insects. Mole crickets are also omnivorous, with
degrees of herbivory and carnivory varying with species (Matheny
1981). Mole crickets (Scapteriscus spp.) are important pests of turf in
parts of the southern United States (Walker and Nickle 1981, Walker
and Ngo 1982) while field crickets (Teleogryllus commodus) are a
serious pastureland pest in New Zealand (Blank and Olson 1981,
Blank and Bell 1982). House crickets are the basis of a multi-million
dollar industry which supplies bait shops, pet shops, and zoos in N.

America.

13

In addition to their economic importance, crickets have served
as models in several areas of research. Several gryllid species have
been studied to address questions of speciation (Alexander 1968,
Harrison 1979), mating behaviour (e.g. Cadel981), and physiology
(e.g. Woodring et al. 1977, 1979). As such, there is a considerable
information base, particularly in the area of physiology and nutrition,
for these insects. A recent publication (see references in Walker and
Masaki in press) reviews areas of research employing gryllids. There
is limited information, however, concerning the microbial community
associated with the digestive tract of these insects.

The digestive system of all gryllids is similar and is illustrated
in Figure 1.1. The digestive tract is largely unmodified from the
presumed primitive insect gut (Chapman 1985) and reflects the
ancient lineage of the group. Unique to crickets is a section of the
hindgut located anterior to the insertion of the Malpighian tubules
(ileum) and containing setaceous structures with a dense bacterial
community. All gryllid subfamilies except for the myrmecophilinae,
and all gryllotalpids examined thus far have a similar anatomy and
association with bacteria (Martoja 1966, Nation 1983). In most
members of the Gryllidae, the bacteria occupy the space between the
peritrophic membrane and the gut wall (Fig. 1.2, 1.3): termed the
ectoperitrophic space. The peritrophic membrane usually separates
the food bolus from the bacterial community and is disrupted by
muscular activity at the junction of the anterior hindgut and the
remainder of the proctodeum. This membrane is absent in

gryllotalpinae (Thomas and Nation 1984a).

l4

   

Qastric Malpighian
cecae tubules
proventriculus ‘ rectum
buccal cavity m ureter
£5: .
‘ 3
‘\
posterior hindgut
ventriculus (colon)
crop anterior hlndgut

(ileum)

Figure 1.1 Gryllid digestive tract. From Martoja (1966)

The anterior hindgut (ileum) in insects is generally thought to
function mainly in the export of undigested material and products of
metabolism excreted via the Malpighian tubules to the rectum.
Mechanisms of water and solute resorption are associated with the
posterior hindgut (rectum) (Bradley 1985). It is now known that the
anterior hindgut is more than a passive channel to the rectum
(Phillips et al. 1986), and in the case of crickets, plays a considerable
role in digestion and absorption (Thomas and Nation 1984a, b).

The cricket hindgut as an environment for bacteria is likely to be
influenced by substrates passing from the midgut, excretory
products and activities of the surrounding tissues, and by the
activities of the bacteria themselves. The presence of a peritrophic
membrane will restrict the availability of large compounds (> 20 nM)
to the bacterial community (Richards and Richards 1970, Chapman

1985). Contributions from the Malpighian tubules are likely to

Figure 1.2

15

Scanning electron micrographs of the luminal side of the
anterior hindgut wall of A. domesticus. (A) Conventional
adult 3 weeks after molt; 200X. (B) Germfree adult 3
weeks after molt; 200X. (C) Close -up of setacaceous
papillae in conventional adult; low mag 540X, inset
4500X. (D) Corresponding view of (C) in germfree
version; low mag 540X, inset 3000X. (E) Close-up of wall
between papillae in conventional adult; low mag 540X,
inset 2700X. (F) Corresponding view of (E) in germfree
version; low mag 180X, inset 2000X.

16

 

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include a pool of soluble materials (inorganic ions, amino acids,
soluble carbohydrates) that are a primary filtrate of the insect's
hemolymph (Bradley 1985). Additionally, the tubules concentrate
uric acid for excretion as the insect's major nitrogenous waste
product. It is unclear how much if any of the Malpighian tubule fluid
contents are available to the bacteria in the cricket anterior hindgut,
since the community is located "upstream" of the ureter's insertion
point (Figure 1.1). However, anterior movement of ectoperitrophic
space fluid is known to occur in insect midguts (Dow 1986) and such
a flow might carry Malpighian tubule effluent to cricket hindgut
microbes in this case also.

The physiochemical environment of the gryllid hindgut is
incompletely known. pH conditions within the digestive tracts of
most gryllids is circumneutral; with the anterior hindgut reported to
range from 6.2 - 7.8 (Teo and Woodring 1985, Thomas and Nation
1984b, Srivastava and Baigal 1983). Oxygen concentrations and eH
values for the region have not been reported, but Ulrich et al. (1981)
reported the isolation of strict anaerobes from the anterior hindgut
of A. domesticus . Maintenence of anaerobic conditions is presumably
a consequence of the metabolism of the underlying epithelium and
the facultative bacteria present (Veivers et al 1980).

The bacterial community is found in the hindgut of very young
instars and is present throughout the life of the insect (Martoja
1966). However, the community is periodically disrupted during the
molting process since the hindgut cuticle is derived from ectoderm
and must be shed for growth. This event occurs from 6 to 12 times

during development of A. domesticus (Patton 1978) and necessitates

20

the redevelopment of the community. Martoja (1966) has observed
the molting process and reports that the hindgut cuticle is shed
separately and subsequent to the exoskeleton. He states that the
bacterial community is reestablished by the fraction of the
populations remaining after the old hindgut cuticle is defecated and
that the recolonization is quite rapid (< 2 days). The original inocula
for hatchlings is not known for certain, but clearly is initiated
through the ingestion route. There is no evidence of transovariole
(within the egg) transmission of microorganisms in crickets. Martoja
(1966) concluded that the egg surface is contaminated with gut
microbes as it is laid (some microbes possibly supplied by the male
parent through spermatophore transfer) and that these provide the
inoculum for hatchlings. There is, however, no demonstrated
evidence of this event.

The composition of the bacterial community in gryllids has been
examined in a descriptive manner in only one species, A. domesticus
(Ulrich er al 1981). Facultative anaerobes such as Citrobacter spp.,
Klebsiella spp., Yersinia spp., and unidentified strains of
Enterobacteriaceae dominated the isolates obtained. Strict anaerobes
were less numerous and included Bacteroides spp. and
F usobacterium spp. Overall, the isolates and facultative nature of the
community suggest a similarity with the bacteria isolated from
termites (Schultz and Breznak 1978), however the isolates obtained
from general media are likely to represent only a small percentage of
forms present in vivo. Other gut systems such as the rumen,

vertebrate hindguts, and the cockroach hindgut contain a higher

21

porportion of strict anaerobes (Hungate 1975, McBee 1977, Cruden
and Markovetz 1987).

The metabolic capabilities of isolates obtained by Ulrich er al
(1980) were characterized with API system techniques and were not
remarkable in degradative capabilities. Isolates did not grow on ball-
milled filter paper based media, but a high percentage were able to
degrade uric acid. Fermentation products included acetate,
propionate, and butyrate. However, in viva capabilities were not
assessed.

Although bacterial isolates from A. domesticus display no
particularly unusual degradative capabilities, it is unclear whether
members of the hindgut community supplement the cricket
digestive system by acting on substrates for which no host enzymes
are produced. Teo and Woodring (1985) found that only raffinase
activity is likely to be restricted to the hindgut of house crickets.
Interestingly, a F usobacterium sp. isolated from crickets shows
activity against the plant trisaccharide raffinose (Ulrich et al. 1981).
It also seems likely that at least part of the microbial activity in the
anterior hindgut could augment the cricket's own digestive
capabilities by duplicating the enzymatic processes of the host;
essentially providing an additional attack on solubles that escape
digestion/absorption in the midgut. Teo and Woodring (1985) and
Thomas and Nation (1984a) have shown that degradation of
substrates in the anterior hindgut segment of crickets is reflective of
enzymatic activity in the gastric cecae and midgut segments.
Presumably, this activity results from the continued action of insect-

produced enzymes. However, since activity was assayed as

22

disappearance rates of substrates, evidence of enzyme action in the
anterior hindgut would also include microbial attack on the

compounds.

We;

It is clear that basic information about the relationship between
crickets and their hindgut microbes is fragmentary. The same can be
said of omnivore/gut microbe associations in general. This study
examines the functional relationship between gryllids and the
microbial community in their digestive tract. In a broader sense, it
addresses the question of interaction between host and gut microbe
in omnivorous animals; where the association is non- obligate and
inconspicuous from the perspective of nutritional interdependency.
The approach taken here is bipartite and attempts to: a) determine
the effect of the microbial community's presence on growth and food
utilization by the insect over a range of diet conditions and b) to
characterize the microbial community's metabolic activities most
likely providing the interface between insect and microbe.

The first portion "of the investigation addresses the question
"Does the hindgut microbial community in A. domesticus contribute
beneficially to the insect across conditions of varying diet quality?" It
is hypothesized that any microbe-mediated nutritional benefit would
be more pronounced as diet conditions deteriorate, and conversely,
that microbial contributions to nutrition would be inconsequential
when an abundant and complete diet is available.

The second part of the study is directed toward describing the

bacterial community's fermentative metabolism in a range of cricket

23

species, identifying and quantifying metabolites potentially available
to the insect, and determining the constancy of these metabolites
across diets and in a range of gryllid species. Research is focused on
terminal processes of anaerobic metabolism in order to gain an
understanding of important interactions within the bacterial
community and because these end products represent the primary

pool of compounds available to the insect.

CHAPTERI

Growth and food utilization parameters

of germfree Acheta domesticus (Orthoptera: Gryllidae)

24

Introduction

The presence of symbiotic microorganisms in the digestive
tracts of insects is usually correlated with refractory and/or
incomplete diets. Most familiar are the obligate microbial associations
in xylophagous insect species (Breznak 1982, Buchner 1965). The
anterior hindgut of crickets (Gryllidae) is also characterized by high
densities of microorganisms (Martoja 1966, Ulrich et al. 1981);
however, potential nutritional roles of this bacterial community are
obscured by the fact that crickets are omnivorous, opportunistic
feeders and not restricted to particular, incomplete diets.
Additionally, the associaton of gut microbes with crickets appears to
be nonobligatory to the insect (Martoja 1966). It has been shown
that considerable digestion and transport of nutritionally valuable
compounds occurs in the gryllid anterior hindgut (Thomas and Nation
1984 a,b), but the contribution of the bacterial community to these
processes is not yet understood.

In omnivorous cockroaches, hindgut microbes are thought to
contribute to carbon and nitrogen metabolism in the insect through
the degradation of cellulose (Bignell 1981) and uric acid (Cochran
1985). There are conflicting reports regarding utilization of cellulose
by crickets. Martoja (1966) and McFarlane and Distler (1982)
reported digestion of the polymer in Gryllus bimaculatus and Acheta
domesticus , respectively. However, Teo and Woodring (1985) could
find no evidence of a cellulase in A. domesticus and Ulrich et al.

(1981) could not isolate cellulolytic bacteria from A. domesticus
25

26

digestive tracts. A potential for nitrogen recycling in crickets was
suggested by the finding that a large percentage of microbes isolated
from A. domesticus hindguts were able to degrade uric acid (Ulrich
et al. 1981), but it remains to be seen if this activity is important in
vivo.

The following report details initial investigations into the
nutritional interactions between Acheta domesticus (Orthoptera:
Gryllidae) and the bacterial community inhabiting its anterior
hindgut. The objective was to determine the overall effects of the
bacterial community's presence on the development of A. domesticus
reared under dietary conditions varying in plant carbohydrate and

nitrogen content.

Materials and Methods

Insects: Acheta domesticus individuals were obtained from
stock cultures maintained in the laboratory for several years. The
stock cultures were kept at 30°C, 60% R.H., and on a 12:12 L:D cycle.
Crickets were fed a commercial diet (described below).

Germfree mm: The terminology employed here to
describe animals reared with and without microbes is discussed by
Coates and Gustafsson (1984) . "Germfree" (=axenic) will refer to
conditions of no detectable bacteria and fungi, while "conventional"
will apply to insects with a "normal", undefined compliment of
microorganisms.

Germfree cricket hatchlings were obtained in the following

manner. Stock culture females oviposited into a dish of moist,

27

autoclaved sand for a 24 hr. period. Eggs were incubated in the
oviposition dish for five to seven days and separated from the sand
by washing with distilled water. Broken and discolored eggs were
discarded, and the remainder rinsed several times with sterile
distilled water. The eggs were then immersed in a 0.3% solution of
benzalkonium chloride and gently agitated for 10 minutes. Following
several rinses with sterile distilled water, eggs were transferred
individually to brain heart infusion (BHI Difco Co., Detroit, MI) plates
using aseptic techniques. After 24 hrs. incubation at 30°C, eggs
showing no sign of contamination were aseptically transferred to
fresh BHI plates and incubated for an additional 24 hrs at 30°C.
Groups of 20-25 eggs which showed no evidence of microbial growth
were transferred to sterile plastic petri plates (100 x 80 x 25 mm)
containing moistened filter paper and incubated at 30° until
hatching. All BHI plates used were also incubated for an additional
4-7 days to assay for slower growing contaminants. In practice, the
majority of contamination manifested itself in the first 24 hrs. and
subsequent prolonged incubations rarely indicated new growth.
Subsamples (2-3) of young larvae (5-7 days after hatch) from each
group were assayed for contaminants by crushing them with forceps
to expose and open the digestive tract, and then inoculating media
with the entire carcass. Several media were initially employed as
screens; including BHI agar and broth, blood agar, PYG (peptone-
yeast extract-glucose) plates, nutrient agar, and 2% malt extract
broth. Preliminary work indicated that BHI plates and broth were
equal to or better than other aerobic media in detection efficacy, and

utilized anaerobic BHI or PYG broth (Holdeman and Moore 1971)

28

after transferring larvae to an anaerobic glove bag (Coy Manufact.
Ann Arbor, MI). Though contaminant growth was rarely seen in
malt extract broth, the media was used as a screen for yeasts in all
initial and final tests.

All manipulations of surface sterilized eggs and subsequent
transfers of germfree crickets were done at a laminar flow work
station. Plates and growth chambers were kept in microisolater units
(Lab Products Inc., Haywood, NJ.) which provided an effective
secondary barrier during incubations in environmental chambers.

Groups of hatchlings deemed germfree with the above tests were
subdivided into smaller groups as they matured by transferring the
insects to fresh petri plates. Sterile food and water were aseptically
placed in the 100 x 80 x 25 mm plastic petri plates (henceforth
referred to as "growth chambers") prior to the introduction of insects.
All food (diets described below) was stream-sterilized (121°C) for 30
min and dried. Water was introduced in 3 dram vials with cotton
rolls as wicks. The vials, placed horizontally in the growth chambers,
were equipped with stainless steel wire collars to reduce rolling
movements. Crickets were transferred to new growth chambers at
approximately weekly intervals. Transferring the insects was
facilitated by chilling them (5°C, approx. 30 min.) prior to handling.
Routine assay for contaminants was accomplished by plating fecal
material and moist cotton swabbings of used chambers on aerobic
BHI. Critical assays for contamination performed at the end of an
experiment included aerobic and anaerobic incubation of fecal
material and swabs of each chamber, phase microscopic examination

of fecal material, placing whole, crushed insects in aerobic and

29

anaerobic broth, and examination of the anterior-hindgut intima of
several animals with Scanning Electron Microscopy (SEM) (Ulrich et
al. 1981). Efficacy of aerobic BHI plates for routine containment
checks was confirmed by failure to detect microorganisms with any
of the above methods when BHI plates showed no sign of growth.

Successive generations of germfree A. domesticus were obtained
by transferring mated females to larger chambers (Pyrex Corning
#3250 storage jars) containing oviposition dishes.

111111: A "natural" diet for crickets and other opportunistic
omnivores is unknown and variable enough to be essentially
undefinable. A commercially available "cricket feed" (Country Mark
Inc., Columbus, OH) was used as a basic diet for the maintenance of
stock cultures and as an "optimal" reference diet (Diet one) for
growth studies because it contained mixtures of unrefined plant and
animal material that would be similar to the general classes of
material encountered by "wild" crickets. The commercial diet
consisted largely of a mixture of wheat middlings and soy bean meal,
with animal protein and fat, vitamins, and minerals added as
supplements. The diet was similar to that employed by Woodring et
al. (1977) in studies of growth and food utilization by A. domesticus.
Though ill-defined, cricket feed supported good growth of A.
domesticus , contained no antibiotics, and remained palatable to the
insects after autoclaving.

Diets two and three consisted of 50:50 (wt.:wt.) dilutions of Diet
one with alfalfa hay and cellulose (ball-milled filter paper),
respectively. Diet two increased the level of plant structural

carbohydrates in the diet while only marginally decreasing total

3O

nitrogen levels. Diet three increased structural carbohydrates and
halved nitrogen content of ingested material. Diets two and three
also presumably diluted fat and vitamin pools. Basic component
percentages for each diet are presented with the results (Table 4).
All diets were ball milled for 48 hrs., sieved (#40 mesh), and stored
under desiccation. Diets were autoclaved (121°C) for 30 min. and
dried prior to use in growth studies for both germfree and
conventional animals.

W5: Ten groups consisting
of three larval crickets of each type (conventional and germfree)
were reared from hatching to 10 days beyond the adult molt on each
of the three diets. Germfree animals were third generation under
axenic conditions and conventional larvae were taken from a parallel
stock culture originating from the same batch of eggs. Conventional
animals were reared in chambers identical to those described for
germfree larvae except that the petri dish lids were perforated.
Humidity levels within the conventional chambers were
consequently 10-15% below levels within the germfree chambers.
The humidity reduction was necessary to control occasional mold
growth on uneaten food over the course of the experiment and did
not appear to affect growth rates in preliminary experiments.

The food utilization parameters: approximate digestibility (AD),
efficiency of conversion of ingested matter to biomass (ECI),
efficiency of conversion of digested matter to biomass (ECD), and
relative consumption rates (RCR) were calculated following
Waldbauer (1968) and were determined during four consecutive

periods encompassing the most linear portion of the larval growth

31

curve. Fecal material weight was corrected for uric acid content in
estimates of digestibility (Waldbauer 1968). The growth periods
were arbitrary divisions, but most larvae had reached the last instar
by the start of period #4 on all diets. Larvae were weighed after
each period and transferred to fresh chambers. Remaining food and
fecal material was dried (50°C, 72 hrs), sorted, and weighed. Fecal
material was then ground and stored under desiccation for later
chemical analysis. Larval fresh weights were converted to dry mass
with linear regression equations derived from wet-to-dry
measurements of larvae taken from parallel cultures of germfree and
conventional insects. Larvae were separated prior to the final molt
and were weighed 5 days after reaching the adult stage. Adult
females were also weighed and sacrificed 10 days after the final molt
to determine the number of eggs produced.

Chg-mica! analyses: Uric acid (UA) and nitrogen (N) content of
fecal material was determined for each replicate group of crickets at
each growth period. For UA determination, duplicate 10 mg
subsamples of material were extracted for 30 min at 60°C in a
Li2CO3/boric acid buffer (Cox et al. 1976), centrifuged, and the
supernatant analyzed with a modified version of an HPLC method
devised by Marquardt et al. (1983). A Regis ODS C18 column (25 x
4.6 mm) was used for separation of the compound eluted with a
mobile phase of 50 mM Na P04 acidified to pH2 with phosphoric acid.
The flow rate was 1.0/ml/min. at 40°C. U.V. dectection at 305 nm
was used in the assay.

Total N content was measured for triplicate subsamples of each

fecal sample using a Carlo Erba Elemental Analyzer (Model 1102).

32

Nitrogen content was also determined for six subsamples of each
autoclaved diet using the above methods. Nitrogen in whole body
tissue was determined for six larvae of each type during the final
growth period. Larvae were obtained from parallel cultures of
germfree and conventional crickets and were prepared as follows.
The crop and contents were removed and the carcass was then dried
and subsequently ground in a micro-Wiley mill (#40 mesh).
Triplicate subsamples of each larvae were then analyzed as above for
total N content.

Total lipid, soluble carbohydrate, insoluble carbohydrate, protein
and ash content were estimated for food and fecal material collected
during growth period 4. Food material analyses were done on six
subsamples of each autoclaved diet, while analysis of fecal material
was done on material from individual larvae replicates. Total lipids
were estimated by the methods of Bligh and Dyer (1959). Soluble
carbohydrates were determined as those sugars released after
extraction with 0.1N H2SO4 and quantified with the phenol-sulfuric
method (Dubois et al. 1956). Woodring et al.. (1979) reported that
virtually all carbohydrate available to the normal A. domesticus
digestive system is released upon extraction with 0.1 N H2SO4, and
the soluble carbohydrate refered to here is equivalent to their
"digestible" carbohydrate. Total residual (=insoluble) carbohydrate
was estimated by successive extraction of the material with hot
water (100°C, 30 min) and hot 1.0N H2804 (100°C, 30 min), followed
by chloroform/methanol (25°C, 30 min) to remove lipids and serve

as a final wash. The ash-free dry weight of the material remaining

33

remaining after these extractions was taken to be a relative estimate
of refractory structural carbohydrate content.

Protein content was estimated from total nitrogen values (x
6.25) after adjustment for uric acid, and percent ash was determined
by standard methods. Approximate digestibility (AD) values for each
component were calculated as described above for total dry weight
(Waldbauer 1968).

W: Comparisons of germ free and conventional
enzyme activity were made using API ZYM system (API Analytical
Products, Sherwood Medical, Plainview, NY). Digestive tracts were
excised from eight insects of each type and subdivided into two
regions. The midgut region included gastric cecae and the
ventriculus, and the hindgut segment consisted of the bacteria-rich
ileum. Gut segments from each category were pooled and
homogenized in 2 ml of deionized water with a tissue grinder. Eight
each of germ free and conventional adult A. domesticus midgut
sections were pooled for the analyses, and eight germ free and four
conventional hindgut segments were used. Fewer conventional
hindgut segments were used in order to keep the weight of
homogenized material approximately equivalent to the germ free
material. A subsample of each homogenate was inactivated by
heating at 95°C for 15 minutes and used as a blank. Subsamples of
the homogenate were added to substrate containers as per API
instructions, and incubated for 5 hrs at 30°C. Semi-quantitative
measures of enzyme activity levels were then determined by
comparisons of color development in active vs blank homogenate

treatments for each gut segment category.

34

14C-ee11u1ose (ICN) was purified by methods adapted from
Papson (1963). and added as an aqueous suspension to excised
digestive tracts of conventional and germfree A. domesticus, and the
field cricket, G. pennsylvanicus, in 2.5 ml Venoject® tubes. The
suspension and gut material were homogenized with a glass rod,
after which the tubes were flushed with N2 and incubated for 24 hrs.
at 30°C. The suspension was then diluted with 1.0 ml deionized
water, clarified by centrifugation, and filtered (0.22 nM). Filtrates
were assayed for radioactivity in aqueous solution with liquid
scintillation techniques.

Statistiss: Parameters were compared as their log transformed
values with standard ANOVA and regression techniques (Sokal and
Rohlf 1969, Kleinbaum and Kripper 1978). Experiments were initially
analyzed in a three-way factorial design. When significant interaction
was found, further analyses were performed with separate ANOVA
of parameters within each diet or growth period. Unless otherwise
indicated, probability levels less than 0.05 were considered

significant.

Results

W: Growth rates (Figure 1.1), size at maturity,

maturation time, and fecundity estimates (Table 1.1) were generally
very similar for germfree and conventional insects. Although

conventional animals tended to be slightly larger than germfree

35

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counterparts on diets one and two, the only statistically significant
difference in any of the growth parameters was found between
fecundity estimates for crickets reared on diet three; conventional
crickets produced fewer eggs during the first 10 days of maturity.
Adult male size of conventional insects reared on diet three was also
reduced, but not significantly.

W: Differences between germfree and conventional
animals were manifested clearly in the insects' abilities to digest
food. Conventional crickets digested a greater percentage of each
diet (AD, Figure 1.2 ) and converted food into biomass more
efficiently (ECI, Figure 1.2 ). The AD differences were significant
across diets and time periods, and ECI differences were significant.
Overall, crickets with hindgut microbes digested 34-50% more of
ingestia than germfree crickets and gained 21-39% more biomass per
unit of food ingested on average over the four time periods (Table
1.2). Germfree crickets tended to convert digested food into biomass
more efficiently (ECD, Figure 1.2 ), but the difference was significant
only within periods 1 and 3 of the Diet two treatments and period 2
of the Diet three treatment. Though conventional animals were more
efficient in converting food to biomass, growth rates were similar to
those in germfree animals because conventional crickets consumed
significantly less food (RCR, Figure 1.2 ) across all diets and time
periods.

Food utilization parameters changed with diet treatments for
both germfree and conventional crickets. Efficiencies decreased from
diets one to three, while consumption rates generally increased with

the exception of period 4. Changes in food utilization parameters

38

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41

across diets were the same for germfree and conventional animals;
i.e. differences between germfree and conventional animals in these
parameters remained nearly constant as diet quality was decreased.

Nitrogen excretion (Figure 1.3) paralleled overall food
utilization efficiency patterns on diets one and two; conventional
larvae excreted a smaller portion of consumed N and apparently
conserved the element. However, older instars of both larval types
on diet three showed no difference in overall N balance. Total N
percentages of combined body tissues did not differ significantly
between types on any diet treatment, but larvae reared on diet three
tended to contain a smaller percentage of N (Table 1.3). Although
conventional animals excreted less uric acid in total than did
germfree versions (Figure 1.3), the two groups produced similar
amounts of the compound per unit N consumed (Figure 1.3). On diets
one and two, however, younger instars (growth periods 1 and 2) of
conventional crickets excreted less UA and older instars excreted
more than germfree larvae. On diet three, uric acid excretion did not
increase with age and no difference was found between germfree

and conventional larvae during any period.

Table 1.3 Nitrogen content of combined body tissues of last instar
A. domesticus larvae. Values are mean %N (S.E.), n = 6.

 

Diet one Diet two Diet three
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The cricket/microbe complex was more efficient at removing
water-soluble and acid—soluble carbohydrates from food material.
Fecal material from germfree animals contained a high proportion of
soluble carbohydrate, while material from conventional crickets was
correspondingly higher in residual carbohydrates (Table 1.4) on all
diets. Differences in calculated digestibility of the acid-soluble pool
were significant between groups, while digestibility of the remaining
insoluble carbohydrate was virtually identical in germfree and
conventional animals. Lipid and ash digestibility estimates did not
differ significantly between the two groups of crickets, but
conventional animals digested protein (N) more efficiently than
germfree animals on diets one and two.

Enzyme activity levels of gut sections from germ free and
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levels than hindgut preparations. Conventional hindgut extracts,
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insects could not. Some a-galactosidase activity was also noted in the
midguts of conventional crickets. Purified cellulose was not
solubilized above blank values in the isolated guts of any cricket

examined (Table 1.5).

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carbohydrase

Figure 1.4 Activity levels of major enzyme classes in the digestive

tract of germfree and conventional A. domesticus.

48

Table 1.5. Percent solubilization of 14C-labelled cellulose in excised
cricket digestive tracts. Mean (S.E.) n = 3 for all cases.

 

 

A. domesticus (conventional) 8.3 (1.2)

A. domesticus (germfree) 8.0 (1.2)

Gryllus pennsylvanicus 7.7 (0.7)

(field cricket)

Blank 6.3 (0.3)
Discussion

Based solely on growth performance, the presence of bacteria in
the digestive tract of A. domesticus would appear to be
inconsequential, or even detrimental to reproductive allocation when
diet conditions are poor. Upon closer examination, however, the gut
symbionts enhance the extraction of soluble nutrients from the food
bolus; reducing the amount of ingested material necessary for
optimal growth across a range of diets.

Comparisons of axenic and conventional versions of insects in
nutritional studies have usually focused on differential survival,
growth, or fecundity in the presence or absence of symbionts.
Relatively few insects which normally possess large and permanent
gut microbe populations have been examined with this approach, and
none have quantified food utilization efficiency parameters. Charnley
et al. (1985) found little distinction in growth and development

between germfree and conventional locusts free of gregarine

49

parasites. Bracke et al. (1978) reported that cockroaches developed
poorly when a large portion of the normal gut bacterial community
was eliminated with antibiotics. Although not quantified, Martoja
(1966) notes that Gryllus bimaculatus seemed to develop normally
when reared without gut bacteria. Studies of vertebrate animals are
much more extensive (reviews in Savage1984, Gordon and Pesti
1971, Luckey 1969) and indicate that germfree animals generally
show better growth than conventional individuals if the diet is
”complete". Cecal microorganisms did, however, enhance the
digestibility of dry matter in rodents and rabbits (Gordon and Pesti
1971, Yoshida et al. 1968), but the degradation of crude fiber was
the principle mechansim. Findings here suggest the cricket hindgut
microbes interact with the host in a manner which may be analogous
to cecal microorganisms, but that the mechanism is not as dependent
upon structural carbohydrate degradation.

The mechanisms of increased food utilization efficiency by
conventional crickets are presently unknown, but would not seem to
involve substantial degradation of cellulose. Calculated digestibility
for the residual carbohydrate fractions do not differ with the
presence and absence of gut microbes (Table 1.4) and growth of
conventional crickets is not enhanced as this fraction increases as a
dietary constituent. It is also apparent that purified cellulose is not
degraded to any extent (Table 1.5). Additionally, ADs for both types
of crickets are almost exactly halved as the basic diet is diluted by
50% with cellulose (Figure 1.2). Previous reports of cellulose
digestion in gryllids (Martoja 1966, McFarlane and Distler 1982) may

reflect impurities in the cellulose used, or perhaps a difference in the

50

microbial communities found in the cricket populations assayed.
There are several additional lines of evidence to suggest that
substantial degradation of plant polymers would be an improbable
occurrence in the cricket digestive tract. First, gut retention times
are relatively short (ca 3 hrs) for both field and house crickets
(Chauvin 1947, Turcek 1967). Secondly, the bulk of the microbial
community in gryllids lies within the ectoperitrophic space in the
anterior hindgut (Martoja 1966). This would greatly restrict contact
of all but the smallest of cellulose fragments with the bacteria and
would necessitate a profusion of extracellular enzymes. Although
extracellular release of enzymes is not unknown in bacteria,
degradation of plant polymers in the rumen usually require close
physical association with the substrate (Hobson and Wallace 1982).
Finally, Ulrich et al. (1981) and Teo and Woodring (1985) could find
no evidence of cellulolytic bacteria or cellulase in A. domesticus
digestive tracts. None of these observations absolutely and
independently prove that cellulolytic activity does not occur in the
hindguts of crickets; some species may harbor capable microbes and
further research is warranted. The observations also do not preclude
the possibility of gut microbe attack on various soluble forms of
cellulose (e.g. cellobiose, carboxymethylcellulose). The soluble
carbohydrate fraction assayed in this study could include a wide
range of compounds including sugars, hemicelluloses, gums, and
fragments thereof. Therefore it is possible that microbial activity
includes attack of soluble fiber. A further partitioning of extracted

carbohydrates will be necessary to address the question.

51

Enzyme activity levels in germfree and conventional hindguts
(Figure 1.4) demonstrate that gut microbes enhance the ability of the
insect to attack carbohydrates , and to some degree proteins. It
should be pointed out that activity levels illustrated were measured
for equal weights of gut material, but that on a per cricket basis, the
conventional levels would increase by a factor of two. The results are
completely consistent with the calculated digestibilty values (Table
1.4) which reveal that carbohydrate utilization accounts for much of
the gut microbe activity. The results also suggest that gut microbes in
crickets do not substantially inactivate host enzymes. In some
vertebrate systems, host enzyme levels drop in the hindgut of
conventional animals; presumably because gut microbes use the
enzymes as protein sources for growth (Yoshida et al.. 1968, Coates
1984). Charnley et al.. (1985) showed, however, that activity of a-
glucosidase was increased in the locust hindgut in the presence of gut
microbes in locusts. They suggest that the finding reflects an enzyme
of microbial origin and not an enhancement of the activity of host
enzyme present.

Since the results only show activity levels of broad classifications
of enzymes, it cannot be determined how much of the microbial
activity is a duplication of insect degradative capabilities. It is likely
that the bacteria provide different isozymes within a broad substrate
category, and that this allows more efficient attack of carbohydrate
classes. It is also possible that insect enzyme activity levels are
enhanced due to microbial maintenance of more "optimal"
physiochemical conditions. Midgut pH and anterior hindgut pH are

similar in normal A. domesticus (Tea and Woodring 1985); implying

52

that insect enzymes would operate with similar efficiency throughout
most of the digestive tract in this species. Hindgut environmental
condition differences in germfree and conventional crickets have not
been examined; however, pH and osmolarity in rat cecae are
considerably altered by germfree conditions (Gordon and Bruckner
1984, Gordon 1968).

The finding here that microbes are responsible for a-
galactosidase activity confirms the suspicion of Teo and Woodring
(1985) that the raffinase activity found in the anterior hindgut of A.
domesticus was produced by microbes. Raffinose is a plant
trisaccharide composed of galactose and sucrose in an alpha linkage.
It is likely to be only a minor component of most plant tissues.
However, the presence of raffinase activity demonstrates that cricket
gut microbes not only raise enzyme activity levels in general, but
they also increase the enzymatic diversity of the insect. It is likely
that other (e.g. Gryllus spp.) species of crickets harbor a more diverse
bacterial community than lab-reared A. domesticus , and the
potential for increased metabolic diversity in the hindgut is even
greater in field species..

Whether gut microbes in crickets act chiefly on common
substrates or on those that are normally unavailable to host
enzymes, it is clear that the bacteria process "leftovers" and make a
portion of the energy available to the insect. The fact that fecal
material from germfree insects contains a substantial portion of
soluble carbohydrate indicates that the house cricket digestive
system is relatively inefficient to begin with, and that

microorganisms are not competing with the animal under normal

53

conditions. It remains to be seen how microbial trapping of
"leftover" energy is transferred to the insect.

There are two major routes of transfer possible. The first would
involve a utilization of material by the bacteria and the subsequent
release of metabolites which are absorbed by the insect. This
scenario is thought to characterize the transfer of energy in
carbohydrates in most gut/microbe systems studied to date (Oldeson
and Breznak, 1983, Hungate 1975). Anaerobic production of organic
acids like acetate provides useful compounds to the host with a
minimal loss of potential ATP. Such compounds are rapidly and
preferentially transported by the hindgut epithelium (Phillips et al.
1986). The hindgut community in gryllids is anaerobic and produces
several short-chain organic acids (Martoja 1966, Chapter 2), and the
quantitative significance of these metabolites is currently being
investigated. Additional compounds such as vitamins and cell
exudates are also conceivably made available to the insect. Vitamin
synthesis is thought to be one function of hindgut microbes in
vertebrates but there is conflicting evidence regarding the
availability of the compounds to the host animal and in some cases
microbes compete with the host for essential vitamins (Coates 1984).
The issue has not been critically examined in insects. Although
conventional crickets seemed to benefit little from microbes when
reared on the vitamin-diluted diets two and three in this study, the
question of vitamin nutrition was not specifically addressed in the
experiment.

A second possible method of converting excess digestia to cricket

biomass would involve the harvesting of microbial cells. Microbial

54

biomass is generally discounted as a possible nutritive source in
animals with hindgut microbes, except in cases where coprophagy is
employed (McBee 1977). There is no evidence to indicate that
crickets are coprophagic under the rearing conditions employed
(excess food), but natural populations have not been observed.
Reasons that microbial biomass harvesting have been dismissed as a
significant mechanism in hindgut systems center around the fact that
microbes are concentrated posterior to the major sites of digestion
and adsorption in the animal (McBee 1977). The unusual anatomy of
the cricket anterior hindgut, however, may allow an exception to
these generalities. The presence of a peritrophic membrane in the
hindgut presents the potential for anterior movement of fluid in the
ectperitrophic space, as has been demonstrated in the midguts of
several insect species (Dow 1986). It is not known if a similar
mechanism occurs in crickets, but bacterial cells in the anterior
hindgut conceivably could move anteriorly to digestive/absorptive
sites in the midgut. The evidence of some a-galactosidase activity in
the midgut preparations of conventional crickets hints that a portion
of the microbes or their extracellular products may move forward in
the digestive tract.

Nitrogenous excretion patterns do not clarify the issue, as they
suggest microbes aid in N conservation when dietary N is in excess,
but when dietary N becomes less available they appear to compete
with the host. This is implied because overall nitrogen economy is
not enhanced by microbes in late instar larvae on diet three (Figure
1.3), and egg production; a highly N-dependent process (Chen 1985),

was reduced in conventional animals on the same diet. A

55

scavenging/recycling of nitrogen as it becomes scarce in the diet is
often postulated as a role for gut microbes (Salter 1984, Cochran
1985), but the evidence from the cricket system suggests otherwise.
If microbial protein is being harvested, it may be highly dependent
upon the population dynamics and metabolic activities of the
bacterial community. Conceivably, population turnover rates were
greatly reduced in the gut bacterial community as low nutrient
intake by the larvae on diet three became critical. In ruminant
systems, microbial biomass production is reduced as diet quality
deteriorates (Hungate 1975). When nutrient levels were adequate
(diets one and two), population turnover rates and biomass
production would be greater; potentially making more microbial N
available to the insect. The possibility of hindgut bacterial protein
being made available to the host is supported by the findings of
Martoja (1966) who reported incorporation of gut bacteria amino
acids into the tissue of G. bimaculatus.

The low levels of soluble carbohydrates in diet three may also
have changed the composition of the bacterial community and/or
caused a shift in its metabolism. With fewer readily available
carbohydrates, the bacteria may have switched to utilizing
nitrogenous compounds (amino acids, peptides) that would have
been uncontested under conditions of excess. There is evidence to
suggest that gut microbes in other systems act mainly on
endogenous, soluble pools of nitrogenous compounds (Salter 1984)
and hindgut fluid may contain significant quantities of amino acids
from normal excretory processes (Bradley 1985). If hindgut bacteria

in crickets shift to utilizing these compounds as other sources are

56

depleted, then nitrogen reclamation routes normally employed by
the insect could be disrupted. The tendency for a decreased nitrogen
utilization efficiency with age in conventional crickets reared on diet
three suggests that the microbial use of nitrogen changes or that the
insect's own nitrogen metabolism is disrupted only gradually by
microbial competition for substrates.

The role of microbes in the nitrogen metabolism of crickets
remains unclear, but there is little evidence from this study to
suggest that uric acid degradation by gut bacteria constitutes a
meaningful nitrogen conservation mechanism. The performance of
conventional animals on the low nitrogen diet indicates that if uric
acid is degraded, then the nitrogen is unavailable to the insect or is
balanced by a loss of exported microbial nitrogen. A peculiarity in
the localization of the hindgut microbes - they are concentrated
anterior to the insertion of the Malpighian tubules - may preclude
opportunities to degrade significant quantities of the compound in
any case. Most of the urate would enter the digestive tract
"downstream" of the microbial community, and be passed out of the
system in an insoluble form (Bradley 1985). The uric acid-degrading
capabilities of the hindgut microbes in crickets may be more
important to the survival of those cells which are subsequently
exported to a different environment (e.g. soil) with egested material.

Gut bacteria have been shown to recycle uric acid in termites
(Potrikus and Breznak 1981) and have been implicated in
performing a similar function in cockroaches (Cochran 1985) and
birds (Salter 1984, Mortensen 1984). Termites and roaches are

known to store uric acid internally for mobilization under conditions

57

of N stress (Mullins and Cochran l975a,b). Such a system of "storage
excretion" (Cochran 1985) is not evident in house crickets (Cochran
1976, Ulrich et al. 1981, Nation and Patton 1961), however, and
strategies for nitrogen conservation are apparently different or less
important in this insect.

Interestingly, efficiency of food utilization in conventional larvae
exceeded that of germfree larvae even when overall nitrogen
digestion and excretion patterns were identical in late growth
periods of the diet three treatment. This would imply that the
mechanism of enhanced efficiency is not principally channeled
through nitrogenous compounds. It is not known how any weight
gain was partitioned between carbohydrate, lipid, and protein during
this study. However, it has been established that larval growth in A.
domesticus, particularly during late instars, is proportionally greater
in the lipid component (Lipitz and McFarlane 1971, Woodring et al.
1979). Microbes do not appear to affect the overall digestibility of
lipids, but they do salvage carbohydrates which may serve as lipid
precursors.

Crickets were apparently unable to totally compensate for
nutrient dilution with increased consumption as has been
demonstrated for locusts (McGinnis and Kastins 1967) and
cockroaches (Gordon 1972). In general, food was consumed at a
higher rate by young cricket larvae and individuals reared on diet
three. Patterns of consumption appear to be different in larvae
reared on diet three, but the high RCRs of larvae during growth
period 4 on diets one and two (Fig. 2) is most likely an artifact of the

growth period duration. The fourth growth period of diets one and

58

two treatments was shortened to 3 days because food became
depleted in some germfree chambers. At this time, most larvae were
in the middle of the last instar and consumption rates decline rapidly
after that point (Woodring et al. 1979). Therefore, if the period had
been extended to 7 days (length of period 4 in diet three treatment),
little additional food would have been consumed, but the calculated
RCR (mg/mg/day) would have dropped to approximately 1/2 of the
rate illustrated in Figure 1.2; effectively making the patterns of
consumption with age similar across the diet treatments.

It is difficult to reconcile the facts that crickets with microbes
used food more efficiently, but failed to translate the benefit into
better growth. The reduction in consumption of food by conventional
animals, which accounts for the discrepancy, may be related to a
change in physiology due to microbial activity or the physical
presence of the bacterial community: e.g. the mass of microbes in
the anterior hindgut acts on stretch receptors that signal a slowing of
feeding because the region is perceived to be "full". There is
evidence to suggest that food movement and feeding behavior can be
partially controlled by such feedback sensors in various regions of
insect digestive tracts, including the hindgut (Bernays 1985, Simpson
1983). Gut retention times were measured using a chromic oxide
marker in house crickets and no discernable differences between
germfree and conventional animals were found (Kaufman unpub.).
The rates were between 2 and 3 hrs., but it was impossible with the
method to estimate rates with a precision of greater than ;l-_ 15
minutes. In any case, the regulation of feeding behavior is certainly

more complex than merely controlling the passage of ingestia.

59

Internal physiological status (e.g. hemolymph titres of sugars and
amino acids) may play a role in determining how frequently and how
much an insect consumes, and osmolarity of the hemolymph has
been shown to affect feeding behavior of locusts (Bernays 1985). It
is possible that microbial activity interacts at that level by releasing
metabolites and products of digestion to the hemolymph, which
subsequently act to inhibit ingestion.

Growth rates and food utilization parameters reported here for
conventional animals reared on diet one are in close agreement with
values measured for other lab populations of A. domesticus given a
similar diet (Clifford et al. 1977, Patton 1978, Woodring et al. 1977,
Crossley and Van Hook 1970). ECI values from this study are also
comparable to those reported for ultimate instar A. domesticus
reared on a totally synthetic diet (McFarlane and Distler 1982).
Growth and food utilization for both germfree and conventional
crickets generally decreased with diet quality and most likely varied
because of diet nitrogen (available protein) content, as was
determined by Gryllus bimaculatus (Merkle 1977). Dietary nitrogen
was not controlled specifically and variation in performance could be
related to several factors. A more defined diet manipulation would
be necessary to resolve the question; however, in preliminary
experiments using only conventional larvae, the addition of
cholesterol, B-vitamins, and vitamins E and K to diet three did not
increase growth performance (Kaufman, unpub.).

Although conventional crickets digested a greater percentage of
ingested material on all diets, conversion of digested material into

body mass (ECD) was in general, less efficient than in germfree

60

insects. This also supports the idea that the flow of nutrients and
energy in the excess digested material goes through the bacterial
community prior to being incorporated into insect tissue. Some of the
potential energy and biosynthetic precursors would be lost to
bacterial metabolism and export of bacterial cell matter with the
feces. Approximately 1 x 107 bacterial cells are excreted per mg of
fecal material by crickets from stock cultures (Kaufman, unpub.).

It is somewhat perplexing that an increased efficiency was not
translated into a growth advantage by conventional crickets. The
tendency toward larger body size in conventional larvae and adults,
though not statistically significant, would seem to indicate a
beneficial interaction when sufficient nitrogen is present. Results
from several preliminary growth studies employing diets one and
two failed to demonstrate any difference in growth parameters
between conventional and germfree crickets and the combined data
merely support the observations of this study (Appendix 1.).
Conditions of the experiment, however, may have influenced the
growth performance. As indicated, relative humidity levels in
conventional animal growth chambers were necessarily lower to
inhibit fungal growth, allowing quantitation of food and fecal
material. Though our preliminary work and that of Bates (1971)
indicated no substantial difference in growth rates within a humidity
range of 60-85%, and that conventional and germfree crickets
contained identical water as percent body weight on all diets, there
may have been a subtle interaction between humidity and diet. As
Clifford et al. (1977) have pointed out, crickets will not drink to

repletion from wick sources, and conventional crickets in slightly

61

drier chambers may have paid higher metabolic costs to maintain a
water balance. The poorer egg production of conventional animals on
the low quality diet may, in particular be related to relative
humidity levels, since nutrition in many insects is a synergistic
function of nitrogen and water content of food material (Mattson
1980)

The experimental conditions also precluded selective exclusion of
parasitic or pathogenic organisms from conventional animals. Zuk
(1987) showed that parasitism by gregarine sporozoans did not
adversely affect host field crickets unless diet conditions were
stressful, and there is substantial documentation that host
susceptibility is related to nutrient status in many other insects
(Vago 1963). Though A. domesticus cultures were free of protozoan
parasites - typical orthopteran gregarine parasites do not seem to
colonize house crickets - the prevalence of other pathogens has not
been examined.

Although growth performance of house crickets was not
enhanced by the presence of microorganisms, a general increase in
food utilization efficiency across a range of diets might be of
substantial benefit to crickets in natural environments. Gryllids are
omnivores which feed sporadically; food resources are likely to be
limited in quantity and variable in quality. House crickets occupy a
variety of habitats indirectly created by humans (Bates 1969,1971)
and are probably restricted in their foraging by predation and other
factors. An ability to efficiently extract nutrients from an

unpredictable resource would certainly be adaptive.

62

In an attempt to simulate fluctuating diet conditions, a
preliminary experiment was performed using alternating diets
(Figure 1.5). Germfree and conventional larvae were reared from
hatching to an age of 15 days on diet one, and then given a diet of
100% alfalfa hay for one week. Diets (diet one and 100% alfalfa) were
then alternated weekly until the adult molt and insects were kept on
100% alfalfa for the duration. Results showed that germfree larvae
suffered substantial mortality when diets were switched to alfalfa,
while conventional larval survival was 100%. Further investigation
with the alfalfa diet showed that germfree crickets repeatedly
showed higher mortalty rates which were not reduced by the
addition of cholesterol, fat-soluble vitamins, or water soluble
vitamins. Conventional larvae also showed increased mortality
independent of vitamins and cholesterol when kept exclusively on
the alfalfa diet. Growth rates were also reduced; however, the
presence of microbes allowed most conventional larvae to reach the
adult stage, while no germfree crickets survived to that stage.
Digestibilty of the hay diet was enhanced by microbes following the
patterns seen in the study discussed above. Mean AD (approximate
digestibilty) was 26.6 _-l_-_ 1.3 (n =7) for conventional larvae, and 8.7 1;
2.5 (n = 6) for germfree larvae. It seems likely that this added
digestive capability provided the minimum nutrition necessary for
survival over the short term and may aid in growth over long term
exposure to poor diets. These results support the evidence gathered
from studies with the diluted diets. With sufficient nitrogen levels in
the diet, the microbial community's extraction of carbohydrates

allowed subsistence and minimal growth. The gut microbial activity

63

Figure 1.5 Percent survival of germfree and conventional A.
domesticus reared on diets with alfalfa hay. Top -
survivorship when hay is alternated with basal diet (diet
one). Bottom - Survivorship on 100% alfalfa with and
without growth supplements.

96 SURVIVAL

96 SURVIVAL

1201

 

64

ALTERNATING DIET

 

lg

 

 

alfalfa
added

 

 

 

 

 

TIME (days)

65

enabled the insect to utilize a food source that was apparently
inadequate and unavailable to the insect's endogenous systems.

An important concern in viewing the any of the results
presented here is that laboratory populations of insects were utilized.
Acheta domesticus is cosmopolitan in its native Europe and Asia
(Bates 1971), but it is an introduced species in the US; reared
commercially in large concentrated populations (cricket "farms") or
maintained in laboratories as a tool for insect physiologists. It
remains to be seen if lab populations of crickets contain a "normal"
bacterial community and some previous work would suggest that
rearing conditions and diet do affect gut microbe community
composition in other insects (Brooks 1963, Hunt and Charnley, 1981,
Carpentier et al 1978). Other authors, however, have emphasized the
stability in gut microbe community composition over a range of
conditions (Savage 1977, Lee 1985). Work with various gryllid
species (Chapter 2) indicates that the bacterial community in crickets
is functionally similar between lab and field-collected individuals.
Furthermore, because fermentation metabolites of microbes in the
anterior hindgut of A. domesticus are identical to those seen in
native populations of field crickets (Gryllus spp.), it appears that the
Gryllidae as a group is likely to posses a functionally similar

microbial community.

Chapter 2

The production of volatile fatty acids by the
bacterial community in crickets (Orthoptera: Gryllidae):

Potential significance to the insect.

66

Introduction

Crickets are omnivorous insects which possess a densely-
populated bacterial community in a modified region of the anterior
hindgut (Ulrich et al.. 1981, Martoja 1966). In the house cricket,
Acheta domesticus , the bacteria enhance food utilization efficiency
by larvae and appear to conserve soluble carbohydrates that escape
digestion by the insect (Chapter 1). Little is known about the
metabolic activities of the bacterial community in gryllids, but the
mechanism of increased efficiency is likely to be expressed through
these activities.

In almost all other animal/hindgut microbe systems studied to
date, fermentation metabolites in the form of volatile fatty acids
(VFA) represent the principle pool of microbially-produced
compounds which are available to the host. The importance of these
compounds to the host is thought to vary considerably with the
species of animal and its feeding habits (McBee 1977). Among
invertebrates, VFA produced by hindgut microbes in termites could
support almost all of the insects basal metabolism energy
requirements (Odelson and Breznak 1983). Although fermentation
metabolites have been measured in the gut systems of other insects
(Bracke et al. 1980, Bayon 1980, Bauchop and Clarke 1975), their

significance is unknown.

67

68

The bacterial community in gryllids is thought to function under
anaerobic conditions (Ulrich et al. 1981) and utilizes soluble
carbohydrates (Chapter 1). These observations strongly suggest that
fermentation metabolites in the form of VFA are produced;
potentially providing a link between microbial activity and food
utilization efficiency by the insect. It is the intent of this study to
examine and characterize the major end products of the fermentation
in gryllid hindguts, and to evaluate their potential contribution to the
insect. Effort has been focused on the production and distribution of

VFA over a range of cricket species and diets.

Materials and Methods

Insects: A. domesticus were obtained from stock cultures and
maintained as reported previously (Chapter 1). Field specimens of
Gryllus rubens and Scaptericus vicinus (gryllotalpinae) were
supplied by Dr. T. Walker, Univ. Fla., Gainesville and were collected
from fields adjacent to the campus. Dr. Walker also provided lab-
reared specimens of G. assimilis. Field specimens of G.
pennsylvanicus were collected from grassland fields near the Kellogg
Biological Station, Hickory Corners, MI. All field-collected specimens
were either analyzed immediately or were maintained for short
periods (1-3 days) in covered aquaria and fed autoclaved cricket
feed (see Chapter 1) prior to analysis. A stock culture of G. rubens

was maintained in the laboratory under the same conditions as A.

69

domesticus colonies. Germfree A. domesticus were obtained with
methods reported in Chapter 1.

Diets: Diets for A. domesticus have been described previously
(Chapter 1). They consisted of a commercial cricket feed (Country
Mark, Inc., Columbus, OH) as the basic diet, with dried alfalfa hay and
cellulose (ball-milled filter paper) as constituents added to decrease
protein and soluble carbohydrate content.

Won: Anterior hindgut segments were quickly
excised from live crickets and placed individually in 300 pl of
deionized water. 100 pl of 1 mM 3-methyl pentanoic acid in NaOH
was added as an internal standard. The gut was then homogenized
with a stainless steel rod and the suspension was clarified by
centrifugation. Supernatants were collected and stored at -70°C.
VFAs were quantified after distillation following methods of (Lovley
and Klug 1982). VFA pool size of fluid in the ectoperitrOphic space
was assessed by exposing the anterior hindgut segment of living A.
domesticus adults and collecting fluid in capillary pipettes. The
ventral portion of the segment was first pierced with an insect pin
and fluid was collected as it flowed from the opening. Samples from
several insects were pooled until approximately 20 .11 were obtained.
Five separate, pooled samples were each placed in 300 pl water with
100 pl internal standard, and samples were treated as above. Cell
material content of the fluid was determined by collecting
subsamples of the fluid in capillary pipettes, sealing the opening with
paraffin, and centrifuging the pipettes for 30 min. at 10,000 rpm

(Odelson and Breznak 1983). Percent cell material content was taken

70

to be the volume of the pellet as a percentage of the total volume
collected.

yEA production: Estimates of VFA production rates were
obtained following the general methods of Odelson and Breznak
(1983) using a time-zero approach. Individual anterior hindgut
segments were excised and placed in 2.5 ml Venoject® flushed with
an atmosphere of 95% N2, 5% C02. These were then incubated from
0, 2, and 4 hrs at 30°C. Freezing at -70°C terminated the incubations
and samples were subsequently assayed for VFA content as
described above.

Estimates of VFA excretion were obtained by homogenizing 10
mg of fresh (< 1hr.) fecal material collected from terminal instar
larvae in 0.5 deionized water and subsequent analysis of the extract
as described above. Excretion rates were then estimated by the
product of VFA / mg feces and total hourly fecal production during
the last larval growth period (Chapter 1).

Non-volatile fatty acids: A. domesticus anterior hindguts were
prepared as above for VFA production up to the point of termination
of the incubation. Samples were then prepared for HPLC as follows.
Guts were homogenized in 400 ul purified water and the homogenate
clarified by centrifugation, followed by acidification of the
supernatant with 2.5 N H2804 (10% acid: extract, v:v). Equipment and
column conditions were as reported by Lawson and Klug (in press).

I_I_2 and CH4= evolution: Isolated anterior hindgut segments were
incubated for four hours at 30°C in 2.5 ml Venoject® tubes flushed
with N2. Headspace gas was sampled and analyzed by gas

chromatography (3 M Porapak N column, nitrogen carrier). Living

71

crickets were examined for H2 and CH4 gas production by placing
them in 36 ml serum vials containing soda lime to remove C02. 02
was added hourly in 0.5 ml quantities to compensate for respiratory
consumption (see Appendix II.). Headspace samples were taken
after incubation at 30°C for 4 hrs. and analyzed by gas
chromatography (Dacey and Klug 1982).
@W The potential for VFA production
through reduction of C02 with H2 was examined in A. domesticus
anterior hindguts following the methods of Breznak and Switzer
(1986). This method will subsequently be referred to as the
"suspension" method to distinguish it from a separate experiment.
Ten gut segments were homogenized in 2.0 ml BSS (Breznak and
Switzer 1986) in an anaerobic glove box (atmosphere 5% H2, 10%
C02, 85% N2). 200 pl of the suspension was added to each of nine 2.5
m1 Venoject® tubes which were then removed from the glovebox and
flushed with H2. Chloroform, which blocks the methylation of
formate to acetate in the acetogenic reaction (Prins and Langhurst
1977), was added to each of six tubes in a final concentration of 10,
50, and 100 uM (two replicates per concentration). The remaining
three tubes served as controls and received deionized water in
volumes equal to those added to the CHCl3 block treatments. 5 pl of
36mM Na14CO3 (8.3 uCi/ nM) was then injected into the tubes as an

aqueous solution (pH 9.5). The tubes were then incubated with

agitation at 30°C for 3 hrs. The incubation was terminated by the
addition of 50 ul of 1.0 N HCL and the tubes wereflushed under CO2

for 10 min. 60 ul of 1.0 N NaOH was added to neutralize the

suspension, followed by 180 pl of deionized water to bring the

72

volume up to 500 pl. Following centrifugation, 50 ul subsamples were
counted by liquid scintillation to determine the total counts fixed in
the soluble fraction. The remaining supernatant was prepared for
HPLC as described above for nonvolatile organic acids. Fractionation
of the HPLC eluent was accomplished by methods reported in Lawson
and Klug (in press). Under conditions of the analysis, formate and
fumarate co-eluted and fractions reported here as formate are those
collected from the single formate/fumarate peak. As a consequence,
concentrations of formate could not be determined.

The acetogenic reaction was also examined in a slightly different
manner; henceforth referred to as the "whole gut" method. Two gut
segments were placed in 2.5 ml Venoject® tubes under a flow of N2
or H2. One ul of Na14CO3 (aqueous, pH = 9.5) was injected through
the wall of each segment and the guts were then macerated with a
stainless steel rod to form a more homogenous slurry and to
distribute the label more uniformly. It was thought that this
procedure would be an effective compromise between the disruption
of the spatial distribution of the bacterial community likely to occur
with the suspension method, and the problems associated with an
uneven distribution of label as it is introduced by injection into the
intact gut system. The tubes were incubated for 3 hours at 30°C
without agitation. Incubations were terminated and homogenates
analyzed as described above for the suspension method.

W: The permeability of the hindgut cuticle
and epithelium to major VFAs was assessed by injecting 14C-labelled
acetate, propionate, and butyrate into isolated hindgut segments

from A. domesticus. guts were removed from living insects and

73

quickly immersed in a saline solution isosmotic with cricket
hemolymph (J. P. Woodring, Louisiana State University, person.
com.). The composition of the saline was as follows: NaCl-160mM,
KCL-6mM, CaC12.2H20-8mM, Na-citrate-SmM, NaCO3-5mM, glycine-
10mM. The gut segments were ligated at the hindgut/colon junction
and label was introduced by inserting a 30 gauge needle with
blunted tip through the lumen of the midgut and into the anterior
hindgut segment. The label (1 ul) was slowly injected as the needle
was withdrawn and a ligature was tightened at the midgut/hindgut
junction immediately after the needle was withdrawn. The isolated
gut was then rinsed in fresh saline, blotted, and placed in 100 ill of
saline. Subsamples of the saline bath were taken after 10 minutes
and at hourly intervals for two hours. Subsample volume (50 1.11) was
replaced with fresh saline at each time point. Gut integrity was
assessed by inclusion of 3H-labelled polyethylene glycol (PEG, 4000
mw, 0.1 uCi per til solution) in the injected solution as described by
Bracke et al. (1980). Each acid was added at concentrations within
the range of those found in the ectoperitrophic space fluid of
conventional insects and were as follows: acetate, 100 mM (5.4 uCi);
propionate, 10 mM (0.5 uCi); butyrate, 5 mM (0.13 uCi).

Statistjgs: Data were compared as their log transformed values
with standard ANOVA and regression techniques (Sokal and Rohlf
1969, Kleinbaum and Kripper 1978). Probability level for significance

was 0.05 unless otherwise indicated.

74

Results

VFAs were found in the anterior hindgut segments of all cricket
species examined (Figure 2.1). Acetate dominated the pool in all
species, but propionate was prominent and characteristically
accounted for 15 to 20% of the total acids. Other VFAs generally
consisted solely of butyrate, with trace quantities of isobutyrate,
isovalerate, and valerate found inconsistently. Germfree A.
domesticus hindguts contained significantly less total VFA, with
acetate comprising over 98% of the pool. Lab-reared species and field
collected species showed strikingly similar patterns. Only S. vicinus,
the mole cricket representative, seemed to differ; however, neither
total VFA concentration nor % composition differed between
conventional species.

Production rates of VFA and the percent composition of acids
produced during the incubations (Figure 2.2) were also very similar
between field-collected G. rubens and G. pennsylvanicus, and lab-
reared A. domesticus. Estimated rates (slopes from linear regression)
were not significantly different, nor were acid distributions.
Germfree A. domesticus hindgut incubations (not illustrated) failed to
demonstrate any VFA production. In a direct comparison of lab-
reared and field-collected G. rubens (Figure 2.3), production rates
and percent composition did not differ significantly, although field
crickets' rates tended to be slightly lower and the VFA pattern
indicated slightly elevated levels of propionate.

Fluid in the ectoperitrophic space of A. domesticus (Figure 2.4)

contained similar proportions of acids to those measured for entire

75

  
 
 

 

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Figure 2.1 VFA concentration in the anterior hindgut of various
gryllid species. Bars are S. E., n = 6-14.

76

Figure 2.2 Production rates and % composition of VFA in the
anterior hindgut of three gryllid species. Each point in the

production curve is a single gut replicate. Error bars are
S.E., n = 7-14.

TOTAL VFA (nmol/mg)

77

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1001 ectoperitropmc fluid
A. domestlcus

36 TOTAL ACIDS

 

 

ACETATE PROPIONATE OTHER VFA

Figure 2.4 Percent composition of VFA in the ectoperitrophic fluid

of the anterior hindgut of A. domesticus.. Error bars are
S.E., n = 5.

8O

guts. Cell material volume of the fluid was calculated to be 48%
(S.E.=l.4, n=5) and mean concentrations for acetate, propionate, and
total other VFA were 8O (:97) mM., 13 mM (i 1.9), and 5 (10.7) mM,
resp.

The rates and acid composition of hindguts from young (20 -25
days old) A. domesticus larvae (Figure 2.5) were indistinguishable
from those of late instar and adult house crickets .

Long term exposure to varying diets did not appreciably affect
estimated production rates of VFA in A. domesticus, however, the
distribution of acids was changed significantly (Figure 2.6). Dilutions
of the basic diet with alfalfa hay or cellulose tended to increase
concentrations of propionate and butyrate in incubated hindgut
segments. Maintenance on a diet of alfalfa hay alone reduced
propionate concentrations, but elevated butyrate. Other VFAs in all
cases remained at trace levels.

Non-volatile organic acid production was not evident from
incubations of isolated hindguts and subsequent HPLC analysis.
There was some indication of lactate production; but results were
inconsistent in repeated trials. Acetate and propionate did
consistently increase with incubation time in the same analyses
(results not illustrated);however, indicating that the method was able
to detect production of fermentation metabolites.

Based upon VFA production estimates of ZOO-400 nmol/ last
instar cricket] hr., excretion rates of VFA (Table 2.1) would account

for less than one percent of production.

81

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82

Figure 2.6 Production rates and composition of VFA in the anterior
hindgut of last instar larvae of A. domesticus reared on
four diets. Each point in production curve represents a
single gut replicate. Error bars are S. E., n = 4-20.

 

83

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84

Table 2.1. VFA excretion rates in last instar larvae of conventional
A. domesticus.

 

Diet Total VFA in feces Feces produced Estimated excretion

(nmol/mg) (mg/cricket/hr.) (nmol/cricket/hr.)
1 5.7 (0.9) 0.3 (0.1) 1.7
2 2.9 (0.7) 0.5 (0.2) 1.4
3 2.3 (0.6) 0.6 (0.2) 1.4

 

Analysis of gas evolution from isolated hindgut segments of G.
rubens (field collected), G. pennsylvanicus (field collected), and A.
domesticus indicated that hydrogen was produced, but no evidence
of methane was found in any of the crickets examined (Table 2.2).
Additionally, H2 gas could be detected in the headspace of vials
containing live A. domesticus, but levels were low, not consistently
detected from individual crickets, and were therefore not quantified.

14C02 was fixed by anterior hindgut microbes in a homogenized
suspension and in whole gut preparations (Table 2.3). Label was not
fixed when added to germfree hindguts (not illustrated) . The
percentage of added label fixed in was enhanced by the presence of
hydrogen and by the more thorough homogenization (suspension
method) of the hindgut material (Figures 2.7, 2.8). Label appeared
principally in acetate. However, propionate and butyrate fractions
contained considerable counts; particularly in the slurry preparation.
Label distribution varied with the incubation method: acetate specific

activity increased disproportionately with the suspension method.

85

Table 2.2 Hydrogen and methane in headspace of gut incubation
vials after 4 hrs. Values are mean i S.E. (n).

 

H2 (141/ mg) CH4
A. domesticus 0.33 i 0.03 (14) n. d. (14)
G. rubens (lab) 0.32 1 0.03 (8) n. d. (8)
G. rubens (field) 0.25 i. 0.03 (8) n. d. (8)
G. pennsylvanicus
(field) 0.29 3:. 0.07 (8) n. d. (8)

n. d. = not detected.

 

Table 2.3 Percentage of added 14C O 2 fixed under various
conditions of gut incubation. Mean (S.E.), n = 4-6.

 

Suspension Whole gut
H2 H2 + CHCL3 H2 N2
77.5 (0.71) 47.2 (1.01) 44.5 (2.1) 25.8 (2.3)

 

86

Figure 2.7 Distribution of fixed l4C02, specific activity of partitioned
label, and total VFA formed per gut during "suspension"
method incubations with and without CHCL3 block. H2 =
hydrogen atmosphere, BLOCK = hydrogen atmosphere and
CHCL3 added to the suspension.

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88

Figure 2.8 Distribution of fixed 14c02, specific activity, and total
VFA produced per gut in "whole gut " incubations. N2 =
nitrogen atmosphere, H2 = hydrogen atmosphere.

 

 

 

 

 

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90

CHC13 effectively decreased total fixation of 14C02, and label to
accumulated in formate (Figure 2.7). The total counts fixed and
distribution of label in various fractions in the block treatments was
independent of CHC13 concentration, and the illustrated results
reflect the pooled results from 10, 50, and 100 uM CHC13 treatments.
The CHC13 block reduced the specific activity of acetate and butyrate,
but propionate remained unchanged. The block also significantly
reduced acetate concentrations in the suspension, while propionate
and butyrate were unaffected (Figure 2.7).

Labelled acetate, propionate, and butyrate easily transversed the
hindgut cuticle and epithelium; most of the added label could be
recovered in the saline bath after the first hour (Figure 2.9).
Butyrate appeared to move at a slightly reduced rate compared to
the other two compounds, however, high permeability is still evident
and would indicate that all compounds could rapidly diffuse into the

hemolymph.

91

Figure 2.9 Rate of movement of l4C-labelled VFA across the
anterior hindgut wall of A. domesticus in vitro. Each
point represents the percentage of added label found in
the saline incubation medium.

92

 

 

 

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93

Discussion

VFA pool sizes and production rates found in gryllid hindguts
indicate an active fermentation that could explain a portion of the
enhanced food utilization found to be provided by microorganisms in
house crickets (Chapter 1). Excretion rates of VFA are almost
negligible in relation to production estimates and the hindgut wall is
readily permeable to the principle acids produced. Assuming
produced VFA are absorbed and oxidized completely to C02, the
acids could provide 4-5% of basal respiratory demand in late instar
A. domesticus , based on an oxygen consumption figure of 0.5 ml 02/
cricket / hr (see Appendix III). This figure is comparable to that
estimated for pigs, but generally falls below estimates for other
hindgut fermenters (McBee 1977). VFA production is calculated to
provide nearly 100% of respiration needs in termites (Odelson and
Breznak 1983 and between 5 and 16% of the demand in Tipula
abdominalis (Lawson and Klug, in press).

The production rates in crickets are not trivial, however, since
they represent a potential conservation of 0.6 mg glucose
equivalents/ cricket/ day. Using a reported growth efficiency (dry
weight gained/ dry weight food consumed) of 0.15 for late instar
conventional insects and an efficiency of 0.11 for germ free versions
(Chapter 1) the difference due to microbes is 0.04. Given a
consumption rate of approximately 20 mg/cricket/day for
conventional insects (Chapter 1, Woodring et al. 1977) the efficiency
difference would account for 0.8 mg of dry weight gain per day: a

figure remarkably similar to potential savings calculated from VFA

94

production estimates. Clearly, calculations from estimates include
many possible sources of error, and conversion of VFA into biomass
would not be 100% efficient. Nonetheless, the figures suggest that
VFA production is well within the range necessary to account for a
substantial portion of microbe-enhanced food utilization efficiency in
A. domesticus.

VFA production rates from in vitro measurements may also
be underestimates of in situ rates (Hume 1984) since removal of
products by active gut tissue would be inhibited. There is evidence
of fermentation product build up in the gut incubations reported
here because longer chain volatile fatty acids tend to increase in
proportion compared to time-zero points and ectoperitrophic fluid
(Figure 2.4). Additionally, total VFA production in the entire cricket
digestive tract was not quantified; only the anterior hindgut segment.
Conceivably, microbial activity in the posterior hindgut and rectum
contribute to the overall digestion and similar metabolites could be
formed. This has not yet been investigated.

VFA contribution to food utilization may also be
underestimated by simply calculating potential oxidation rates. VFA
may enhance overall efficiency indirectly; by interacting with gut
epithelial cells. There is evidence from vertebrate systems that
VFAs stimulate development of gut cells (Sakata and Yajuma 1984),
provide a large portion of gut cell energy requirements (Roediger
1980), and enhance fluid movement and transport activity (Demigne
et al. 1980). Insect systems are not as well studied, but locust rectal
tissue absorption of fluid and ions from the lumen is sustained by

acetate transport, and acetate also supplies a portion of the tissue's

95

energy demands (Baumeister et al. 1981). Part of the overall
digestive efficiency in crickets may include a similar mechanism.
The characteristics of fermentative metabolism in the gryllid
hindgut suggest an unusual system compared to previously-reported
hindgut fermentations. In insect systems investigated thus far,
acetate dominates the pool of detected VFAs in termites, grass grubs,
and craneflies (Table 2.4). The scarab, Orcytes nasicornis, however,
has a digestive tract fermentation that produces a suite of VFA
similar to crickets (Bayon 1980), but the standing pool of VFA in the
proctodeal dilation of this beetle was reported (by the same author)
to contain over 90% acetate (Bayon and Mathelin 1980). In
vertebrates, fermentation products include a higher percentage of
propionate and butyrate than those reported for insects; with
changes in diet causing considerable variation. Odelson and Breznak
(1983) suggest that the dominance of acetate in termites reflects
both the insoluble nature of the substrate consumed and the high
relative numbers of protozoa in the system. Aquatic detritivores
consuming a diet of decomposing leaf material also have hindgut
fermentations characterized by high levels of acetate and few longer
chain VFA (Lawson and Klug in press), but the community is
composed entirely of bacteria (Klug and Kotarski 1980). The pattern
in crickets appears to be somewhat intermediate between the few
insects studied and vertebrates. Propionate values consistently
comprise 15-20% of VFA, while butyrate values are much lower than
in vertebrate systems. In another anomaly, methane has not yet
been detected in any cricket species, whereas its production has been

measured in termites (Breznak 1982), scarab beetles (Bayon 1980)

96

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and numerous vertebrates (Prins 1977, McBee 1977). Finally, the
microbial community in crickets does not appear to rely as much
upon refractory structural polymers as substrates (Chapter 1), as is
thought to drive much of the metabolism in other gut systems.
Evidence presented thus far does not preclude the bacterial
metabolism of soluble fiber (e.g. some hemicellulose, pectins,
starches), however, and this may be of significance in certain dietary
regimes.

The constancy of the fermentation pattern seen across species of
crickets (Figures 2.1, 2.2), age (Figure 2.5), and rearing conditions
(Figure 2.4) indicates an overall functional similarity in the bacterial
communities that is maintained over a range of habitats. Hungate
(1975) has noted that although the community composition of
microbes in the rumen may vary across geographical region, the
overall pattern of VFA remains remarkably consistent. It would in
fact be surprising to find identical species of bacteria in different
cricket groups, however, if metabolite production forms the basis of
the relationship, the bacterial community composition may be
irrelevant. The results also show that lab-rearing does not appear to
alter the overall community metabolism in the hindgut. Again,
nothing is known of the bacterial species present in crickets other
than A domesticus, but a sufficiently "normal" compliment of
microbes is retained even after several generations of artificial
conditions. VFA represent only the final products of the
fermentation, however, and the initial degradative pathways in
hindguts of lab and field groups of crickets have not been critically

compared. Lab-reared crickets may be deficient in certain members

99

of the bacterial community that have more relevance (in terms of
substrate utilization capabilities) to wild cricket populations.

The shift in VFA patterns that is seen when A. domesticus is
reared on different diets (Figure 2.6) suggests a change in substrates
available to the bacteria and/or a shift in bacterial populations. As
acid-soluble carbohydrate content drops, propionate and butyrate
percentages increase. The addition of alfalfa hay to the diet raised
butyrate levels in the fermentation, while dilution with cellulose
raised levels of both propionate and butyrate. The shift to longer
chain VFA is still evident when crickets are maintained on Diet three:
a diet diluted with insoluble cellulose but containing no "new"
carbohydrates compared to the basic diet. It may be that the insect's
midgut scavenges a greater percentage of the pool of readily
utilizable carbohydrates in the diluted diet; leaving a quantitatively
different pool for the hindgut microbes. The leftovers reaching the
hindgut from diet three would be poorer in easily-assimilable
compounds, and the bacteria may have been forced to utilize other
energy sources that were ignored in the richer diet. Increases in the
relative proportions of propionate and butyrate in the rodent cecum
are associated with increasing fiber content , however, the response
is variable and dependent on the type of fiber available. In general,
it appears from studies of vertebrate gut systems, that increasing
cellulose and hemicellulose content in a mixed diet. favors more
butyrate formation (Demigne et al. 1980, Sudo and Duke 1980,
Hungate 1975). However, xylophagous insects utilizing strictly
cellulose and hemicellulose have hindgut fermentations characterized

by acetate proportions of 95% or greater (Odelson and Breznak

100

1983). The soluble pectins, guar gum and gum arabic, were also
shown to elevate butyrate and propionate concentrations in the rat
cecum (Tulung et al. 1987, Topping et al. 1985). Interestingly, a VFA
pattern similar to that reported here for crickets was shown to
develop in the rat cecum when the animal was fed wheat starch or
pectin diets (Demigne et al. 19,80). The basal diet employed here
contains a high proportion of wheat middlings and the gut microbial
community in crickets may be functioning in a similar manner to the
rat cecum community. It is currently unknown if pectins,
hemicelluloses, and soluble cellulose fragments are important
fermentation substrates for cricket hindgut bacteria, but their
increased utilization may explain the shifts in VFA patterns as diets
changed.

A point that emerges from the preceding discussion is that the
pattern of the hindgut microbial fermentation is not predictable
when diet changes are considered independently of the particular
gut system. The resulting change (or lack of) is also a response of the
endemic microbial community present. It remains to be seen how
much of the VFA pattern shift is due to metabolic changes or
population shifts within the bacterial community; or both.

From the view point of the cricket, the changes in
fermentation patterns with diet may be meaningful from a
nutritional standpoint. If production rates for total VFA remain
constant, a shift to longer chain acids represents an increase in
available carbon. Assuming equal assimilation costs, propionate and

butyrate would provide more oxidative/biosynthetic precursor per

101

mole than acetate. Details of this study are insufficient to quantify
the answer to the question, but further investigation is warranted.

The production rates of VFA do not appear to differ markedly in
response to diet, however, variation in values around time points is
considerable, and critical comparisons of slopes would necessitate
larger sample sizes in most cases. In the case of lower soluble
carbohydrate diets, estimates of rates from 4 hour incubations may
even be understated because VFA content in individual gut segments
tends to begin to level off between 2 and 4 hrs and a linear model
may not be the best fit of the data (Figure 2.10). Rates calculated
between 0 and 2 hrs in that case would be higher, and perhaps more
relevant, since transport of VFA may occur rapidly (discussed
below). The tapering-off of VFA production in some incubations may
indicate that substrates are being depleted. Production rates of
acetate in termites vary with diet and conditions of the insect
(Odelson and Breznak 1983) and VFA production rates in vertebrate
gut systems are known to increase as more readily-digestible
carbohydrates increase in the diet (Hungate 1975, Hume 1984,
McBee 1977).

Although a characteristic feature of the cricket hindgut
fermentation is the production of propionate, acetate clearly
dominates the pool in all cases and represents the principle product
available to the insect. Acetate can be derived directly through
carbohydrate fermentation , but it can also be formed through the
terminal process of COz-reduction with hydrogen (Ljungdahl 1986).
The latter may be of particular significance in the cricket system

since it represents a conservation of carbon and reducing equivalents

102

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103

The formation of acetate from C02 and H2 is strongly suggested by
the l4CO2-fixation reported here. Conclusive evidence would
necessitate the confirmation by NMR techniques of both carbon
atoms of acetate being labeled . However, the results of the
experiments here leave little doubt that the reaction occurs and is of
considerable importance to the overall fermentation in A. domesticus
hindguts. Based upon the reduction in acetate quantities that
followed the addition of chloroform, H2/CO2 acetogenesis could
account for approximately 35% of the acetate produced. This figure
is nearly identical to that estimated for termite hindguts (Breznak
and Switzer 1986). Although the incubation method (suspension)
would limit extrapolation to intact gut systems, % composition of VFA
produced in the suspension ( not illustrated) are in agreement with
values obtained for whole gut preparations. This would indicate that
the overall fermentation was not greatly disrupted by the addition of
CHCL3 to the system.

Results from the "whole gut" incubations also demonstrated an
active CO2 fixation into VFA. The activity was enhanced under
hydrogen, but considerable label was fixed under N2; indicating that
hydrogen evolution in the microbial community is sufficient to drive
the reaction. Overall, less fixation of added label occurred in the
whole gut method compared to the suspension incubations; indicating
that distribution of the label was not adequate in the former. Even
though label may not have been introduced uniformly in the whole
gut method, its distribution in VFA after incubation is puzzling.

Specific activities in acetate, propionate, and butyrate are

104

approximately equal in the whole gut incubatons, (Figure 2.8) while
acetate specific activities in the suspension are disproportionately
greater (Figure 2.7). The discrepancy requires further investigation,
but may indicate that disruption of spatial relationships of bacteria
in the community affects the transfer of fixed product between
different groups.

The substantial incorporation of label into propionate raises
questions about mechanisms involved in propionate formation. Of
those known to involve CO2 fixation are a Bacteroides -type pathway
(Macy and Probst 1979) and a less well known reductive
carboxylation of acetate (Schink 1984). C02 incorporation into
propionate occurs via succinate in several species of Bacteroides
(Macy and Probst 1979). Since this genus has been isolated from
crickets (Ulrich et al. 1981) a potential explanation for the high
specific activity found in propionate here may be due to Bacteroides
metabolism of CO2. There was little evidence of labeled succinate in
any of the incubations, but turnover to propionate may have been
too rapid for detection. Results from the chloroform block (Figure
2.7) support a Bacteroides-type fixation. l“CO3 incorporation into
propionate occurs independent of the reaction leading to acetate:
specific activity of propionate does not change if acetate production
from H2 and CO2 is inhibited (Figure 2.7). The evidence from the
CHC13 treatment, then, is contrary to expectations, if propionate was
formed from a reductive carboxylation of acetate. Incorporation of
substantial amounts of 1"'CO3 (up to 79%) into propionate have been
reported by researchers primarily concerned with CO2 fixation into

acetate, but few have discussed the issue (Prins and Lankhorst 1977,

105
Goldberg and Cooney 1981, Schink 1984). The formation of C3

compounds from C2 compounds was called "enigmatic" by Schink
(1984) and deserves more attention; particularly in systems where
propionate is a significant end product. The high specific activity of
propionate relative to acetate in the slurry incubations remains
unexplained, but suggests that reductive carboxylation of acetate to
propionate should not be dismissed as a possible pathway of CO2
fixation in vivo.

The CHCl3 block does indicate that incorporation of 14CO3 into
butyrate is dependent upon the acetogenic reaction. Condensation of
two acetate molecules to form butyrate has been demonstrated
previously (Kirby et al. 1983). The reaction may take on greater
importance in crickets fed an alfalfa-based diet, as the butyrate
proportion increases in the hindgut fermentation on that diet (Figure
2.6)

The significance of acetogenesis to terminal metabolism in
anaerobic systems is only beginning to be understood. The process is
generally thought to be less important than methanogenesis in most
anaerobic systems, including the rumen (Breznak and Switzer 1986,
Hungate 1975). Interest is currently focused on why methanogens
seem to be more prevalent or active as terminal anaerobic
metabolism. Since methanogenesis results in the net loss of carbon
and reducing equivalents, the formation of acetate from H2 and CO2
would be advantageous in systems where fermentation products are
linked to animal metabolism. In the case of termites, acetogenesis in
the hindgut is relatively more important than methanogenesis, and

represents a benefit to the insect (Breznak and Switzer 1986).

106

Although crickets are not as dependent upon their gut microbes, the
presence of acetogenic bacteria as terminal process organisms fits in
well with the apparent strategy of carbohydrate conservation in the
insect. Methanogenesis apparently outcompetes acetogenesis in the
hindgut of another omnivore; the American cockroach (Periplaneta
americana) (Breznak and Switzer 1986). The reasons for dominance
of acetate formation over methane formation in cricket and termite
gut systems is an intriguing question: in the case of termites,
acetogens appear to outcompete methanogens in the system.
However in crickets, there is no evidence that methanogens ever
establish in the system. Further research will hopefully clarify the
issue.

The ability of injected VFA to move across the hindgut wall of
crickets adds support to several other studies that reveal the insect
hindgut permeable to a wide variety of small soluble organic
compounds (Maddrell and Gardiner 1980, Hogan et al. 1985, Bracke
et al. 1980, Phillips et al. 1986, Lawson and Klug, in press, Thomas
and Nation 1984, Bayon and Mathelin 1980). As Phillips et al. (1986)
point out, most studies involving ligated, in vitro gut incubations do
not really measure transport rates. The research reported here for
crickets is more accurately a measure of potential transport rates
which would ultimately have to be determined in vivo. VFA
movement across the cricket anterior hindgut wall was rapid. Rates
were comparable to rates reported for termites (Hogan et al. 1985),
but generally was higher than in viva estimates for cockroaches
(Bracke et al. 1980), cranefly larvae (Lawson and Klug, in press), and

scarab beetles (Bayon 1980). The incubation temperature was

107

higher in this study (30°C) than in the aforementioned works,
however, and may account partially for the high rates reported here.
Additionally, the incubation bath used in this study did not contain
VFA at physiological concentrations.

The pattern of label accumulation into the saline medium for all
three acids suggests a passive transport of VFA (diffusion) in the
anterior hindgut. Much of the absorption of compounds from insect
midgut tissue is passive (Dow 1986), and the cricket anterior hindgut
is in some ways a hybrid of midgut and proctodeum (Nation 1983).
Thus, a passive transport is consistent with known mechanisms of
absorption. Concentration gradients between the lumen and
hemolymph would favor movement of VFA from the hindgut.
Acetate concentrations in the hemolymph are approximately 10 mM
(Kaufman unpub), while ectoperitrophic fluid concentrations are 80
mM (see above). Butyrate and propionate concentrations in
hemolymph are below the mM range, and these acids would also
have a favorable concentration gradient for leaving the hindgut.

It has been demonstrated previously that the anterior hindgut of
crickets is permeable to products of digestion. Martoja (1966) found
that 14C from labelled cellulose in the diet could be found in the
hemolymph of G. bimaculatus and that microorganisms were
responsible for the activity. He did not identify the form of the
transported label, but speculated that it likely was in the form of
organic acid. Thomas and Nation (1984b) demonstrated that glucose,
glycine, and palmitate could move across the gut wall of G. rubens, G.
assimilis and Scaptericus acletus, but that palmitate tended to remain

in the gut tissue, while glycine and glucose passed into the bathing

108

medium. These studies and the data presented here leave little
doubt that the cricket anterior hindgut represents an absorptive site
for products of microbial metabolism and possibly, cellular
components released upon cell lysis (e.g. longer chain fatty acids).
Although rates of movement in in vitro incubations are merely
estimates of potential transport, they suggest that the hindgut can
allow movement of produced VFA. Additionally, rectal tissue in
some Orthoptera has been shown to actively transport acetate at
rates that would greatly exceed production estimates reported here
(Baumeister et al. 1981). Acetate transport mechanisms are
reportedly unsaturable in the hindgut of Panesthia cribrata (Hogan et
al. 1985). This has yet to be demonstrated in crickets, but similar
salvaging of acetate and other VFA is implied by the low excretion
rates of these compounds.

Since VFA are produced and absorbed in crickets, it is
assumed they are utilized. Acetate is readily detectable in the
hemolymph of A. domesticus in concentrations approaching lOmM
(Kaufman, unpub.). Propionate and butyrate have also been found in
hemolymph in trace amounts (< 100 nM), but detection limits were
approached and no reliable quantitation is available. Acetate has
been shown to be incorporated into lipids (Blomquist et al. 1982) in
A. domesticus and serves as precursor to hormones, chitin and
several other components of insect tissue. Additionally, there is little
doubt it is oxidized as a source of energy in insect tissue (Odelson
and Breznak 1983). Propionate and butyrate have received little
attention, but presumably could serve as lipid precursors or oxidized

substrates also. In vertebrates, propionate is particularly important

109

since it is one of the few lipids capable of being converted to
carbohydrate via gluconeogenesis. Propionate is also incorporated
into cuticular hydrocarbons in insects (Blomquist et al. 1979 and
references therein). In view of the prominence of propionate in the
cricket hindgut fermentation, and the role of gut microbes in food
utilization efficiency (Chapter 1), the incorporation of propionate into
insect carbohydrate deserves further research attention.

Although many aspects of the relationship between hindgut
microbes and gryllids remain unclear, the results of this and
previous studies (Chapter 1) strongly suggest a role in carbon
conservation. Approximate digestibility of food components and
enzymology comparisons between germ free and conventional A.
domesticus reveal that soluble carbohydrates are the principle
substrates of the microbiota. The hindgut bacteria add to the
diversity of carbohydrases present and provide an increased
degradative capability to the insect through a duplication of enzyme
activity. VFA patterns also indicate that microbes ferment
carbohydrates and that these compounds are made available to the
insect in the form of products of anaerobic metabolism. The
evidence for acetate formation from H2 and C02 also suggests that
microbial recycling of compounds that would normally escape the
system is a significant piece of the carbon efficiency story. In many
ways, the cricket hindgut/microbe system is unusual in comparison
to the insect-microbe symbioses previously examined. Further
research is needed to determine if similar mechanisms exist in other

omnivorous species.

SYNOPSIS

The preceding chapters have examined the relationship between
crickets and their hindgut symbionts from two slightly different
perspectives. In one investigation, the effects of the microbial
community on larval growth and digestive capabilities were
examined. In another, the microbial fermentative metabolism was
characterized and related to observed improvement of digestion
efficiency in conventional animals. A synthesis of the two appears to
explain the relationship between insect and microbe in A. domesticus
and, possibly, other members of the Gryllidae.

The absence of a microbial community in the hindgut of A.
domesticus had little obvious outward effect on growth, appearance,
and behavior. The principle and consistent difference between
conventional and germfree insects was the relative inefficiency of
the latter in the utilization of food. It was apparent that acid-soluble
carbohydrates were the microbes' principle substrates, and that
carbohydrase activity in general was enhanced by the presence of
hindgut bacteria. Nitrogenous compounds may also have been
utilized more effectively by conventional crickets under some
conditions. Unexpectedly, however, results from rearing the insects
under low dietary N conditions did not indicate that a nitrogen
conservation mechanism was operating, and in fact suggested that
microbes interfere with nitrogen metabolism in adult females. The
increased food utilization efficiency of conventional larvae was still

maintained on a low nitrogen diet, however, and is likely explained
110

111

by the disproportionate increase in lipid content as cricket larvae
mature.

The germfree/conventional comparison, therefore, strongly
suggested a role for the hindgut bacteria in carbon metabolism and
conservation. The cricket hindgut bacterial community was found to
produce volatile fatty acids in quantities greatly exceeding excretion
rates. The types of acids produced were entirely consistent with
those expected of a carbohydrate fermentation, with acetate and
propionate dominating the pool. In addition, a terminal aaerobic
process and mechanism. of carbon and reducing equivalent
conservation was found to contribute substantially to the
fermentation: acetogenesis from carbon dioxide and hydrogen. Since
the bacterial metabolism is anaerobic and a mechanism for recycling
C 02 exists within the bacterial community, much of the energy of the
carbohydrate precursors would still be available to the host. The
potential savings amount to approximately 0.6 mg dry
weight/cricket/day and agree well with estimated microbial
enhancement of weight gain per unit of food consumed. These acids
are highly permeable to the hindgut cuticle and epithelium, and have
been shown elsewhere to be incorporated into lipid and
carbohydrate components of insect tissue.

Through microorganisms, crickets increase their metabolic
diversity. Although the study presented here uncovered only two
specific examples (a—galactosidase activity and C02 fixation into
acetate), there are undoubtedly many more. This would potentially
allow the insects to utilize a wider range of diets, and the

experimental evidence indicates that microbial-enhanced food

112

utilization is maintained across a range of diets. In natural
situations, diet is likely to be even more variable.

Kinnear et al. (1983) have postulated that the chief benefit of
gut microbes is to allow "niche expansion". They hypothesize that the
success of ruminants and ruminant-like animals is not principally
based upon the microbes abilities to degrade specific compounds (i.e.
cellulose), but rather that microbes allow the utilization of a wider
array of plant compounds which allows them to outcompete other
herbivores in low nutrient situations. In other words, ruminants are
not specialists on high fiber diets, but they are generalists with a
large diet breadth. Certainly gryllids and other omnivores have not
evolved as tight a dependence on microbes as ruminants, however, a
portion of their success may also lie with expanded food utilization
capabilities due to microbial metabolism. Jones (1984) has also
forwarded several hypotheses concerning microbial associations with
insect herbivores. Among others, he hypothesized that microbial
mutualists should allow a utilization of marginal resources and
function to "even out resource heterogeneity". His ideas are similar to
those presented by Kinnear et al. (1983) as he emphasizes the
increased metabolic diversity supplied by microbe mutualists and
suggests that they are beneficial in low nutrient conditions, but
detrimental in situations of better resources.. Buchner (1965) has
postulated that microbial associations in general are more specific
and pronounced as diets become specialized and deficient in one or
more nutrients. How then, is the microbial symbiosis in crickets and

other omnivores compatible with the above hypotheses?

113

The role of microbes in the nutrition of omnivores is likely to be
more subtle and supplemental than in herbivores; but similarly not
linked exclusively to particular diets. It may be a more general
mechanism likely to be valuable throughout a range of fluctuating
conditions. The idea is still compatible with the discussed
hypotheses because increased food utilization efficiency in
conventional crickets does become {112111211 more important as
dietary carbohydrate is diluted. Conventional cricket food utilization
efficiency was 40% greater than germfree values on the poorest diet,
and 20% greater than germfree values on the highest quality diet
(Table 1.2). In contrast to the predictions of Jones' (1984)
hypotheses, however, the symbionts do not appear to be detrimental
on diets of higher nutrient levels. Further, in terms of nitrogen
metabolism, symbionts in crickets may be beneficial on high quality
diets, and detrimental on nitrogen poor diets. The preceding study
demonstrates that definitions of mutualism between microbes and
animals must qualified in many cases because of interactions with
nutrient type and availability, and growth stage of the animal. Diets
employed in this study may not have been sufficiently different to
test the above hypotheses, however, because the suite of available
nutrients did not differ greatly in quality of compounds, but were
more accurately a gradient of quantity. Support for the above
hypotheses is provided by the differential survival of conventional
and germfree crickets on the alfalfa diet (Figure 1.5) and indicate
that microbes do indeed allow exploitation of marginal resources
through increased metabolic diversity. Crickets as a group are

diverse in habitat and feeding strategies. Examination of the

114

microbial symbioses in other species of crickets over a range of
natural food items might provide a more rigorous test of the ideas.
Although microbial metabolites in the form of VFA may account
for a substantial portion of the food utilization story in A. domesticus,
there are several other tenable postulates concerning contributions
from the bacteria. As discussed in previous chapters, a primary
target for further investigation should be the question of cricket
utilization of bacterial biomass and cellular constituents from the
hindgut. 0f most insect hindgut/bacteria systems studied thus far,
the gryllid system has one of the highest potentials for such a
mechanism independent of coprophagy because of its peculiar
anatomy: the bulk of the bacterial biomass is not associated with the
posterior moving food bolus. The unexpected observation that
nitrogen may be conserved by conventional crickets on a high
nitrogen diet, but that microbes may have a detrimental effect when
nitrogen is scarce, is still unexplained. The examination of this will
question require an understanding of the bacterial population
dynamics within the hindgut under various nutrient conditions.
There also remain the possibilities of essential amino acid
metabolism, supply of vitamins, and inhibition of pathogens as
important roles. These questions were not directly addressed in this
study, however, the germfree methodology employed will allow a
means to investigate the possibilities. Additionally, the methods
should allow an assessment of the microbe function in other species
of crickets. The cricket hindgut/bacterial system remains a viable
and intriguing opportunity for further research. The results

presented in this dissertation represent only an initial, broad-based

115

look at the symbiosis, and it is hoped that future investigations will

provide details.

APPENDICES

APPENDIX I

Early growth studies

 

 

 

 

 

 

 

 

 

 

DIETONE DIETTWO
400‘ «tout. ‘1“900- —-— Gumtree

3 —a-— 00m
5

a:

m

m

a:

II-

‘6

5. ..

O

a mod -

3 , .

g

m 1”: d

u:

u. d i

0 r I ' F ' I ' ' I ' I T r '
0 10 20 30 40 10 20 30 40
‘95 (day!) AGE (days)

Figure AI.1 Larval growth curves of germfree and conventional A.
domesticus reared on diets on and two, for two
successive generations.

116

1 17
APPENDIX I. (cont.)

Table AI.1 Adult size (10 days beyond molt) and maturation
times of first generation germfree and conventional A.
domesticus. Values are mean .t SE. (n).

 

Diet one Diet two
Males Convent. 328.1 i 4.4 (16) 296.9: 6.0(15)
(mg) Germfree 337.6 i. 6.8 (14) 275.5 1'. 10.4 (13)
Female Convent. 415.3 i 9.8 (12) 356.61; 7.5 (15)
(mg) Germfree 411.1 i 4.5 (13) 357.01; 7.6 (12)
Mat. time Convent. 39 i 0.5 (28) 40 _-I_-_ 0.6 (30)
days Germfree 38 i 0.4 (27) 40 :1; 0.6 (25)
Total % Convent. 0 6.7
mortality Germfree 0 10.0

 

Table AI.2 Adult size (10 days ), fecundity, mortality, and
maturation times of second generation germfree and
conventional A. domesticus. Values are mean_ +.S E. (n).

 

Diet one Diet two

Males Convent. 344.2 i 7.7(16) 307.8 i 7.9 (15)
(mg) Germfree 323.4 1; 6.5 (14) 276.9 1 10.4 (13)
Females Convent. 414.1 i 7. 8 (15) 349.3 i 11.3 (15)
(mg) Germfree 411.1 .1: 8.4 (12) 339.8 1; 7.5 (15)
Mat. time Convent. 38 i 0.4 (27) 39 i 0.3 (29)
(days) Germfree 38 :L' 0.4 (27) 39 i 0.3 (30)

# eggs per Convent. 428 i 11.2 (13) 318 i 12.5 (14)
female Germfree 434 + 9.8 (12) 309 + 10.8 (15)
Total % Convent. 0 6.7
mortality Germfree 0 6 7

 

APPENDIX II

Whole Insect Respiration

02 consumption and C02 production of germ free and
conventional A. domesticus was measured by placing insects in wire
mesh tubes (6 x 30 mm) plugged with cotton at both ends to limit
movement by the animal. The tubes were then placed in 36 ml
serum vials and incubated at 30°C. For 02 consumption, soda lime
was placed in the serum vial prior to introduction of the insect in
order to eliminate C02 accumulation during the incubation.
Headspace was subsampled at 3 minute intervals for 2 hrs and
analysed for 02 and N2 concenu'ations by GC (Dacey and Klug 1982).
C 02 production was assessed in a similar manner (soda lime was
eliminated frc the incubation vessel). 02/N2 and C02/N2 ratios
were used in place of absolute concentration values to correct for
subsampling without replacement of headspace gas. Rates are
reported as the slopes from linear regression analysis of computed
ratios vs incubation time. R2 values for all regressions were 0.98 or

higher.

118

119

Table AII.1 Oxygen consumption and carbon dioxide production by
last instar,germfree and conventional larvae of A.
domesticus. Values are mean (S. E.), n = 5.

 

02 Consumed C02 produced R. Q.

(ml/g lhr-) (ml/g lhr)
Conv. female 1.5 (0.1 1.9 (0.1) 0.8 (0.1)
Conv. male 1.9 (0.2) 2.2 (0.3) 0.9 (0.1)
GF female 1.4 (0.1) 1.8 (0.1) 0.8 (0.0)

GF male 1.8 (0.1) 2.2 (0.2) 0.8 (0.1)

 

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Bate, J. 1969. On some aspects of the biology of Acheta domesticus
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_. 1971. Life history of Acheta domesticus (Insecta, Orthoptera,
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Bauchop, T. 1977. Foregut fermentation. In R. Clark and T. Bauchop,
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Bayon, C. 1980. Volatile fatty acids and methane production in
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Bentley, B. L. 1984. Nitrogen fixation in termites: Fate of newly fixed
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Bemays, E. A. 1985. Regulation of feeding behaviour. 3;. G. A. Kerkut
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