Journal of Experimental Botany, Vol. 57, No. 4, pp. 747–754, 2006
doi:10.1093/jxb/erj067 Advance Access publication 31 January, 2006
FOCUS PAPER
Phloem-sap feeding by animals: problems and solutions
A. E. Douglas*
Department of Biology, University of York, PO Box 373, York YO10 5YW, UK
Received 8 July 2005; Accepted 20 November 2005
Abstract
The incidence of phloem sap feeding by animals
appears paradoxical. Although phloem sap is nutrientrich compared with many other plant products
and generally lacking in toxins and feeding deterrents,
it is consumed as the dominant or sole diet by a very
restricted range of animals, exclusively insects of the
order Hemiptera. These insects display two sets of
adaptations. First, linked to the high ratio of nonessential:essential amino acids in phloem sap, these
insects contain symbiotic micro-organisms which provide them with essential amino acids. For example,
bacteria of the genus Buchnera contribute up to 90%
of the essential amino acids required by the pea aphid
Acyrthosiphon pisum feeding on Vicia faba. Second,
the insect tolerance of the very high sugar content and
osmotic pressure of phloem sap is promoted by their
possession in the gut of sucrase-transglucosidase
activity, which transforms excess ingested sugar
into long-chain oligosaccharides voided via honeydew.
Various other animals consume phloem sap by proxy,
through feeding on the honeydew of phloem-feeding
hemipterans. Honeydew is physiologically less extreme than phloem sap, with a higher essential:
non-essential amino acid ratio and lower osmotic
pressure. Even so, ant species strongly dependent
on honeydew as food may benefit from nutrients derived from their symbiotic bacteria Blochmannia.
Key words: Essential amino acids, Hemiptera, honeydew,
phloem sap, sucrose.
Introduction
Sugar-rich diets taste good to most animals. Plants provide
three types of sugar-rich foods for animals: nectar, fleshy
fruits (berries, drupes, etc.), and phloem sap. The relationship between these plant products and animals is exploit-
ative. For nectar and fruits, the exploitation is reciprocal
and for phloem sap, the exploitation is unidirectional (animals
exploit plants). Both nectar and fruits have evolved in
response to the ‘sweet tooth’ of animals, are accessible to
animals, and are of a composition that maximizes animal
foraging: as a reward for animal-mediated pollination
(floral nectar), animal protection from herbivores (extrafloral nectar), and animal dispersal of seeds (fruits) (Herrera
and Pellmyr, 2002). Plant phloem sap has as its principal
function the long-distance transport of nutrients, especially
photosynthate, around the plant (Fisher, 2000), and the
diversion of these nutrients to animals is not in the
plant’s selective interests. In other words, the relationships of plants with nectar and fruit feeders are generally
mutualistic and those with phloem-feeders are generally
antagonistic.
Current understanding of phloem sap utilization by
animals is shaped by two facts. First, just one group of
animals, insects of the order Hemiptera, includes members
that use phloem sap as their dominant or sole food source.
This lifestyle has evolved multiple times among the Hemiptera, and is displayed by most Sternorrhyncha, including
whiteflies, aphids, mealybugs, and psyllids, many Auchenorrhyncha, for example, most planthoppers and many
leafhoppers, and most plant-feeding Heteroptera, including the lygaeids, pentatomids, and coreids (Dolling, 1991).
A few other animal groups (including some thrips and
lepidopterans among the insects, sapsuckers and hummingbirds among the birds, and some primates) consume
phloem sap occasionally, but none require it as a component
of their diet (Dailey et al., 1993; Passamani and Rylands,
2000). Second, all phloem-feeding hemipterans possess
symbiotic micro-organisms that are vertically-transmitted,
i.e. passed from mother to offspring. In the sternorrhynchans and auchenorrhynchans, the micro-organisms are
intracellular, restricted to specific cells; and in the heteropterans they are localized in specialized diverticula of the
gut (Buchner, 1965; Douglas, 1989). Several groups have
* E-mail: [email protected]
ª The Author [2006]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
748 Douglas
reverted secondarily from feeding on phloem sap to whole
plant cells. These insects (e.g. typhlocybine leafhoppers,
phylloxerid aphids) lack symbiotic micro-organisms
(Buchner, 1965), suggesting that the micro-organisms are
advantageous to the insect only in the context of phloem
sap feeding.
These considerations raise two questions: in what ways
are hemipteran insects uniquely predisposed to use phloem
sap as food; and why is symbiosis with micro-organisms
linked to this habit? In this article, as one contributory
factor, it is suggested that phloem sap poses nutritional
barriers to utilization by animals that only the hemipterans
have overcome, partly through the nutritional contribution
from their symbiotic micro-organisms. Other hemipteran
traits important to phloem feeding include the anatomy
and function of the insects mouthparts and gut, and these
are addressed by Goodchild (1966) and Douglas (2003).
Although phloem feeding is restricted to hemipterans,
other animals use phloem sap by proxy. These are the
so-called ‘cryptic herbivores’ (Hunt, 2003) that feed on
the honeydew of hemipterans. Honeydew is phloem sap
modified in composition by passage through the hemipteran
gut and released via the anus. In the latter part of this article,
the phloem sap and honeydew are compared as food.
Phloem sap as a food source
In some respects, phloem sap is an excellent diet for
animals. For most plants, it approximates to a ‘predigested’
food with high concentrations of sugars providing an
abundant source of carbon and energy, and nitrogen
predominantly in the form of free amino acids. It is also
generally free of toxins and feeding deterrents, a consequence of its being a highly specialized cytoplasm (plant
secondary compounds tend to be localized in the apoplast
and cell vacuole, and not the cytoplasmic compartment).
There are exceptions to these generalities, including the
high protein content in the phloem sap of cucurbits, at
10–40 lg llÿ1 compared with 0.1–2 lg llÿ1 in most other
plants (Thompson and Schulz, 1999), and the phloemmobile secondary compounds in various plants, for example, glucosinolates in crucifers and other Capparales
(Merritt, 1996; Brudenell et al., 1999), cardenolides in
the Asclepiadaceae (Botha et al., 1977), and pyrrolizidine
alkaloids in various groups (Hartmann, 1999). These
exceptions notwithstanding, phloem sap remains a poorlydefended, nutrient-rich food source for those animals that
can access it.
Despite this, phloem sap poses two major nutritional
problems for animals. These problems can be described as
the ‘nitrogen barrier’ and ‘sugar barrier’ that animals must
overcome to use phloem sap. The nature of these barriers
and the response of phloem-feeding insects to them will be
considered.
The nitrogen barrier to phloem sap utilization
The growth and fecundity of phytophagous insects are
generally limited by nitrogen, in two ways: the quantity of
nitrogen, i.e. the total amount of nitrogen available; and the
quality of nitrogen, or its composition. The issue of quality
arises because animals are metabolically impoverished,
lacking the ability to synthesize nine of the 20 amino acids
that make up protein. (These nine ‘essential’ amino acids
are listed on the x-axis of Fig. 1A). If the concentration of
just one of these essential amino acids is in short supply,
protein synthesis and growth of an animal are constrained.
The nitrogen barrier to phloem sap utilization is its low
nitrogen quality. Broadly speaking, the ratio of essential
amino acids:non-essential amino acids in plant phloem sap
is 1:4–1:20, considerably lower than the ratio of 1:1 in
animal protein. Data for the phloem sap of the broad bean
Vicia faba and the legume-feeding pea aphid Acyrthosiphon pisum illustrate this mismatch (Fig. 1A). The amino
acids in the phloem sap of V. faba are dominated by
asparagine, which accounts for 72% of the total amino acids
in this dataset. All the essential amino acids are detectable
in the phloem sap samples, but their combined concentration represents just 8.2% of the total and the concentration of all but one essential amino acid (histidine) is
proportionately lower in phloem sap than in aphid protein.
The essential amino acid content of phloem sap is
insufficient to support the observed growth rate of the
aphids. This can be illustrated for final instar larvae of
A. pisum that ingest, on average, 1.92 ll phloem sap over
2 d (from days 6 to 8 after birth), during which time they
increase in weight from 0.7 mg to 1.32 mg, equivalent to
protein growth of 31 lg (AE Douglas, unpublished data).
Even if the unlikely assumption is made that the aphids
assimilate the ingested essential amino acids and convert
them to protein with 100% efficiency, the phloem-derived
amino acids are inadequate to support the observed protein
growth for all essential amino acids other than histidine.
The shortfall varies from 0.06 nmol tryptophan to 21.6 nmol
leucine per aphid (Fig. 1B), and represents 5% and 60–86%
of the total aphid requirement for tryptophan and the
other essential amino acids, respectively. The shortfall in
the dietary supply of amino acids is met by an endogenous
source. As described below, there is overwhelming evidence that the endogenous source is symbiotic bacteria,
Buchnera sp., which synthesize and provide these nutrients
to the aphid. In other words, Buchnera cells enable aphids
to overcome the nitrogen barrier to phloem sap utilization.
The biology of Buchnera has been reviewed recently
in Douglas (2003). The key information required in the
present context is that Buchnera sp. is a coccoid cproteobacterium that dominates the microbiota of aphids,
accounting for >90–99% of all microbial cells in the aphid
tissues. Buchnera is obligately intracellular, restricted to
the cytoplasm of specialized insect cells, known as
Phloem-sap feeding by animals
B
80
25
20
phloem sap
60
nmol derived
from Buchnera
aphid protein
40
20
15
10
5
essential
amino acids
valine
threonine
tryptophan
phenylalanine
lysine
methionine
leucine
valine
tryptophan
threonine
phenylalanine
methionine
lysine
leucine
isoleucine
histidine
tyrosine
serine
glycine
glx
asx
arginine
alanine
non-essential
amino acids
histidine
0
0
isoleucine
mean amino acid content
(% of total amino acids)
A
749
amino acid
Fig. 1. The nitrogen barrier to phloem sap utilization: amino acid relations of the pea aphid Acyrthosiphon pisum line LL01 feeding on Vicia faba. (A)
Amino acid content of V. faba phloem sap and aphid protein (excluding the non-essential amino acids cysteine and proline, which cannot be quantified by
the method adopted) asx: aspartic acid and asparagine; glx: glutamic acid and glutamine. (B) Amino acids derived from Buchnera symbionts during the
2 d of the final larval stadium, as calculated from the difference between amino acids required for protein growth and acquired by feeding on plant phloem
sap. [Unpublished data of LB Minto, E Jones and AE Douglas and data from Wilkinson and Douglas (2003) which has been reproduced here with kind
permission of Blackwell Publishing.]
bacteriocytes, in the aphid haemocoel (body cavity) and
transferred vertically to eggs (or early embryos for
viviparous morphs) in the female reproductive organs.
The evidence that Buchnera provide aphids with essential amino acids is 3-fold: nutritional, physiological, and
genomic. The nutritional and physiological approaches
have depended on the development of two sets of techniques: to eliminate the Buchnera from the aphids using
antibiotics, generating aphids known as aposymbiotic
aphids (Wilkinson, 1998); and to rear the aphids on
chemically-defined diets that can be manipulated (Dadd,
1985). The key results of nutritional research over many
years are that aphids with their normal complement of
Buchnera can be reared on diets from which individual
essential amino acids are eliminated, but aposymbiotic
aphids have an absolute requirement for all the essential
amino acids (reviewed in Douglas, 1998). The complementary physiological experiments demonstrate that aphids
with Buchnera, but not aposymbiotic aphids, can synthesize essential amino acids from dietary precursors such
as sucrose and aspartate (Douglas, 1988; Febvay et al.,
1995, 1999; Wilkinson et al., 2001; Birkle et al., 2002).
Together, these experiments indicate that bacteria are responsible for the capacity of aphids to utilize phloem sap
poor in essential amino acids by providing the aphid tissues
with these nutrients.
The genomic evidence supporting the central role of
Buchnera in providing essential amino acids to aphids
comes from annotation of the complete genome sequences
of Buchnera, now available for isolates from A. pisum
(Shigenobu et al., 2000), Schizaphis graminum (Tamas
et al., 2002), and Baizongia pistacea (van Ham et al.,
2003). The Buchnera in all these aphid species have a
small genome (0.62–0.64 Mb) with few genes (553–630,
including pseudogenes). Most unusually for bacteria,
nearly all the genes have orthologues in other bacteria, including Escherichia coli. In other words, the Buchnera
genome approximates to a subset of the E. coli genome.
Exceptionally, Buchnera have retained genes coding for
most enzymes in the biosynthetic pathways for essential
amino acids, even though they have lost many other
metabolic capabilities, including the capacity to synthesize
most non-essential amino acids (Table 1).
These studies suggest strongly that aphids overcome
the nitrogen barrier to phloem sap utilization by their
acquisition of essential amino acid-overproducing bacterial
symbionts. As considered in the Introduction, all phloemfeeding hemipterans bear symbiotic micro-organisms,
a minority of which have been studied phylogenetically.
For example, psyllids bear c-proteobacteria, known as
Carsonella sp. (Thao et al., 2000); mealybugs have
Tremblaya sp. in the b-proteobacteria (Baumann et al.,
2002); and some planthoppers and a single tribe of aphids,
the Cerataphidini, bear ascomycete fungi of the family
Clavicipitaceae (Suh et al., 2001), misleadingly called
‘yeast-like’ symbionts in the early literature. By analogy
with the aphid–Buchnera relationship, these microorganisms can plausibly be argued to provide their insect
hosts with essential amino acids, but direct evidence is
lacking. If future research demonstrates uniformity in
the nutritional role of the various micro-organisms, it suggests that the key symbiotic trait of essential amino acid
750 Douglas
Table 1. Amino acid biosynthetic capability of Buchnera, the
symbiotic bacterium in aphids: + genes present; – genes absent:
data collated from Shigenobu et al. (2000), Tamas et al. (2002),
and van Ham et al. (2003)
Essential
amino acids
Genes for
biosynthetic
pathway
Non-essential
amino acids
Genes for
biosynthetic
pathway
Histidine
Isoleucine
Leucine
Lysine
Methionine
Phenylalanine
Threonine
Tryptophan
Valine
+
+
+
+
+a
+
+
+
+
Alanine
Arginine
Asparagine
Aspartic acid
Cysteine
Glutamic acid
Glutamine
Glycine
Proline
Serine
Tyrosine
ÿ
+b
ÿ
ÿ
6c
ÿ
ÿ
+
ÿ
ÿ
ÿ
a
The biosynthetic pathway is truncated in Buchnera relative to E. coli.
Buchnera has metH, suggesting that it can synthesize methionine from
homocysteine, but lacks metABC required for the conversion of
homoserine to homocysteine.
b
Animals including most insects can synthesize arginine from
ornithine. Arginine is a non-essential amino acid by the definition used
here, i.e. it is synthesized de novo by the animals, but it is sometimes
treated as an essential amino acid because it is synthesized at insufficient
rates to support optimal growth rates of young rats. Present in Buchnera
from Acyrthosiphon pisum and Schizaphis graminum, absent from
Buchnera from Baizongia pistacea.
c
Present in Buchnera from Acyrthosiphon pisum and Baizongia
pistacea, absent from Buchnera from Schizaphis graminum.
provisioning has evolved independently in many different
microbial lineages and is not an evolutionarily ‘difficult’
transition.
The sugar barrier to phloem sap utilization
The dominant compounds in phloem sap are sugars derived
from photosynthetic carbon fixation. In many plants, most
of the sugar is in the form of sucrose, a chemically-stable
disaccharide of low viscosity. The phloem-mobile sugars in
some plants, including many labiates, include oligosaccharides of the raffinose series, especially raffinose and
stachyose (with one or two galactose units, respectively,
transferred to the glucose moiety of sucrose); and some
plants have appreciable levels of sugar alcohols, for example, mannitol in the Apiaceae, sorbitol in the Rosaceae
(Ziegler, 1975).
The basis of the sugar barrier to phloem sap feeding is
its very high concentration in phloem sap, up to and often
exceeding 1 M sugar, and a resultant osmotic pressure 2–5
times greater than the osmotic pressure of the insect’s body
fluids. The insects ingest the phloem sap at a high rate,
partly because phloem sap has high hydrostatic pressure
and partly to ensure a sufficient supply of other phloem
nutrients at low concentrations. The predicted consequence
of the continuous flow of fluid at high osmotic pressure
into the gut is the transfer of water from the body fluids
to the gut contents and osmotic collapse of the insect.
In other words, phloem-feeding insects are expected to
shrivel as they feed.
The key evidence that aphids overcome the sugar barrier
by osmoregulation is that the osmotic pressure of the
honeydew, the egesta voided from the gut, is comparable
to that of the body fluids and is lower than the ingested
food (Fig. 2A). Equivalent data for other hemipterans are
lacking. Analysis of honeydew sugar composition provides
a clue as to how the osmotic barrier is overcome. When
aphids are reared on chemically-defined diets with sucrose
as the sole sugar, the dominant honeydew sugars are the
monosaccharides, glucose and fructose, at low dietary
sucrose concentrations (0.2–0.3 M), but oligosaccharides
comprising mostly glucose moieties at high concentrations
(0.5–1.0 M) (Fig. 2B); the sucrose-derived fructose is
assimilated with high efficiency and used principally in
respiration (Ashford et al., 2000). It is proposed that the
transformation of ingested sucrose to oligosaccharides
would tend to reduce the osmotic pressure of the gut
contents because the osmotic pressure exerted by solutes
is determined by their molality and not their weight.
These data suggest that the sugar relations of phloemfeeding insects are intimately linked with osmoregulation.
At an enzymological level, the fate of ingested sugars is
best-understood in aphids. For example, pea aphids have
very high sucrase activity localized in the gut distal to the
stomach (Ashford et al., 2000; Cristofoletti et al., 2003).
The sucrase is an a-glucosidase (i.e. it is specific for the
a-glucosyl residue of sucrose) and not a b-fructosidase
(specific for the b-fructosyl residue) (Ashford et al., 2000).
It has been proposed that the sucrase enzyme may also
mediate the synthesis of oligosaccharides by transglucosidation, i.e. by inserting glucose and not water at the
glucosidic bond (Walters and Mullin, 1988; Ashford
et al., 2000). Transglucosidation activity has also been
demonstrated in the whitefly Bemisia, generating an array
of honeydew sugars (Byrne et al., 2003), but its incidence
in other phloem-feeding insects is unknown.
An important implication of these results is that the
microbiota play no part in the capacity of insects to utilize
sugar-rich phloem sap. Although some early literature has
suggested a role of micro-organisms in osmoregulation and
in the sugar relations of phloem-feeding insects (reviewed
in Douglas, 1989), the balance of current evidence is that
micro-organisms have no direct role. In particular, aphids
treated with antibiotics to eliminate Buchnera maintain
their capacity to osmoregulate their body fluids to a constant value over a wide range of dietary sucrose
(Wilkinson et al., 1997).
Phloem sap as a variable resource
Phloem sap varies in composition over multiple timescales,
from the diurnal cycle to the season, with the developmental
Phloem-sap feeding by animals
B
2.5
%oligosaccharides
2.0
1.5
1.0
100
80
60
40
0.0
0
honeydew
20
haemolymph
0.5
phloem sap
osmotic pressure
(MPa) mean + s.e.
A
751
0.00
0.25
0.50
0.75
1.00
dietary sucrose
concentration (M)
Fig. 2. The sugar barrier to phloem sap utilization: osmotic relations of the pea aphid Acyrthosiphon pisum. (A) The osmotic pressure of aphid
honeydew is similar to that of aphid haemolymph (body fluids) and lower than that of the phloem sap of Vicia faba plants on which they were feeding.
(B) The oligosaccharide content of aphid honeydew is strongly dependent on the sucrose concentration ingested from the chemically-defined diet.
(Data from Wilkinson et al., 1997).
age of the plant, and with abiotic factors such as temperature and water availability (Douglas, 1993; Geiger and
Servaites, 1994; Kehr et al., 1998; Ponder et al., 2000;
Corbesier et al., 2001; Karley et al., 2002). The capacity
of phloem-feeding insects to respond to long-term changes,
for example, associated with plant development or season,
has been amply demonstrated for aphids. This has been
achieved by the use of chemically-defined diets with a
range of different compositions that reflect different phloem
sap compositions (Karley et al., 2002).
By contrast, there is very little information on the
response of insects to diurnal variation in phloem sap
composition, even though this variation can be considerable. Phloem sugar levels may differ between day and night
by 10–20% to 2-fold, varying with irradiance, temperature,
and other factors (Geiger and Servaites, 1994), and diurnal
variation in amino acid composition has also been reported
(Winter et al., 1992; Corbesier et al., 2001). This short-term
variation cannot be mimicked readily using chemically
defined diets, which are of fixed composition. Much of the
insect response may be behavioural, through variation
in the rate of food uptake according to the nutrient content
and osmotic pressure. Where compositional changes are
very rapid or large, the insect has the option to withdraw the
stylets and probe for a different sieve element. [Phloem
nutrient composition is far from uniform across sieve
elements, even in plants reared under tightly controlled
conditions, as illustrated by the 3–5-fold variation in amino
acid concentration among replicate samples of exudates
from severed stylets (Girousse et al., 1996; Telang et al.,
1999).] Rapid post-ingestive responses may also be involved, including changes in gut sucrase activity and the
function of transporters in sugar and amino acid assimilation, but the underlying biochemical and molecular processes remain to be established.
There is a real possibility that the capacity of an insect to
utilize a plant is influenced by the scale of diurnal variation
in phloem sap composition, as well as by the composition
quantified at any one point in the diurnal cycle. In this
context, datasets on phloem sap composition obtained
during the light period, and usually at a fixed time, may
not reflect accurately the total nutritional inputs to phloemfeeding insects.
Phloem feeding by proxy
After digestion and assimilation of ingested phloem sap
in the hemipteran gut, the residue is voided via the anus
as honeydew. Honeydew is often produced in copious
amounts. For example, first instar larvae of the willow
aphid Tuberolachnus salignus produce more honeydew
per hour than their body weight (Mittler, 1958).
Honeydew is used as a food by various animals which
can be considered as secondary or proxy phloem feeders
that exploit the capacity of hemipterans to access plant
sieve elements. Many insects, including flies, wasps, bees,
beetles, butterflies, and moths, as well as nectarivorous
birds and flying foxes, consume honeydew that has fallen
onto plants or other surfaces; and some animals take
honeydew droplets directly from the anus of the hemipterans. This behaviour, called tending, is widely displayed
among ants, especially among the dolichoderines and
formicines, and also by polybiine wasps, silvanid beetles
and, remarkably, some Madagascan geckos that specifically
tend planthoppers (Folling et al., 2001). Most research has
concerned ant-tending relationships. They are mutualistic:
the tending ants gain food and the tended phloem-feeding
hemipteran is protected from natural enemies by the ants.
It has recently been argued that the evolution of tending
for hemipteran honeydew contributed to the ants ‘breaking
752 Douglas
out’ from their lifestyle as ground predators to colonize
arboreal habitats and exploit phloem sap by proxy some
40–50 million years ago, triggering their subsequent
dramatic diversification (Wilson and Holldobler, 2005).
A vivid demonstration of the quantitative significance to
ants of honeydew feeding comes from stable isotope
analysis, specifically of 15N:14N (d15N). The 14N isotope
is lost preferentially in catabolic reactions and therefore
herbivores are predicted to be enriched in 15N relative to
plants, their predators enriched relative to herbivores, and
so on through the trophic levels (Griffiths, 1998). In
a detailed analysis of ants in tropical rainforests of Peru
and Brunei, Davidson et al. (2003) established that d15N
of ant species tending hemipterans is similar to that of
phloem sap-feeding hemipterans and chewing herbivores,
but lower than that of predatory ants (Fig. 3). In ecological
terms, tending ants are herbivores, gaining access to
phloem sap through mutualism with hemipterans.
The conclusion that hemipteran honeydew is a quantitatively important component of the diet of many ants
raises the issue of the nutritional suitability of honeydew as
a food for animals. Honeydew is nutritionally distinct from
phloem sap because of the enzymatic and assimilatory
capabilities of the hemipteran gut. In particular, the sugars
are modified by hydrolysis and transglucosidation, and the
amino acid profile is altered by differential assimilation.
With respect to the sugar barrier to phloem feeding, the
osmotic challenge posed by high phloem sugar is negated
by the osmoregulatory capabilities of phloem feeders,
making honeydew an osmotically-neutral foodstuff of
complex and variable sugar composition. (The osmotic
6
pressure of honeydew is, however, expected to rise with
increasing time after deposition through the evaporative
loss of water, posing osmotic problems for animals feeding on honeydew on plant surfaces etc). Furthermore, the
capacity of ants and other honeydew feeders to digest the
complex honeydew sugars, and its enzymological basis,
remain to be investigated in detail.
The significance of the nitrogen barrier to honeydew
feeding depends critically on the amino acid assimilation
patterns of hemipterans. In the aphid species studied in this
laboratory, the amino acid composition of honeydew is
generally more balanced than in phloem sap because
aphids preferentially assimilate non-essential amino acids
(Adams, 1997). For example, when the black bean aphid
Aphis fabae, which is facultatively tended by ants, feeds
from Vicia faba, the essential amino acids in its honeydew
account for 30% of the total amino acid content, about
2-fold higher than their percentage-contribution to phloem
sap, at 13% (Fig. 4). In this dataset, all of the essential
amino acids and just two of the non-essential amino acids
(serine and aspartic acid) are proportionately enriched in
honeydew, relative to phloem sap. However, it would be
inappropriate to generalize from these data because there
could be considerable variation among plant–hemipteran
relationships. Also, ant-tending could affect amino acid
assimilation patterns, and so honeydew composition.
Is the nitrogen nutrition of honeydew-feeding ants
promoted by microbial symbionts? This possibility is
suggested by the presence in some ants of an intracellular
bacterium allied to Buchnera and known as Blochmannia
sp. The genome of Blochmannia sp. has been sequenced
and includes the full gene complement for synthesis of
all nine essential amino acids (Gil et al., 2003). In principle, therefore, Blochmannia could provide ants with these
4
2
0
predatory
(n=8)
hemipteran
tending
(n=8)
leaf
foraging
(n=22)
-2
A
B
C
D
foraging mode
Fig. 3. Mean d15N values (&) of ant species in a Borneo rainforest,
classified according to predominant feeding mode, and calibrated against
d15N of plants (A), hemipterans (B), chewing herbivorous insects (C),
and arthropod predators (D). [Redrawn with permission from Fig. 2 of
Davidson DW, Cook SC, Snelling RR, Chua TH. 2003. Explaining the
abundance of ants in lowland tropical rainforest canopies. Science 300,
969–972. Copyright 2003 AAAS omitting species with mixed and
uncertain foraging strategies.]
amino acid content
(% of total)
in aphid honeydew
δ15 N
15
10
nonessential
amino acids
essential
amino acids
5
0
0
5
10
15
amino acid content
(% of total)
in plant phloem sap
Fig. 4. Amino acid content of phloem sap and aphid honeydew for the
black bean aphid Aphis fabae feeding on Vicia faba, expressed as
percentage of the total amino acids. Asparagine (63% and 47% of phloem
sap and honeydew amino acids, respectively) is not included in the figure.
[Data of Adams (1997).]
Phloem-sap feeding by animals
nutrients. Consistent with this interpretation, ant species
with Blochmannia have been cited to ‘show a preference
for honeydew and other sweet secretions’ (Zientz et al.,
2004), although a detailed analysis of the incidence of
Blochmannia in ants with different nutritional ecologies
remains to be conducted. If 15N:14N stable isotope analysis
(as used in Fig. 3) were used as an index of nutritional
ecology, interpretation would require great care because
the amino acid biosynthetic function of the microbial
symbionts would tend to deplete the 15N content of the
ants, so underestimating ant dependence on honeydew or
other plant products.
Mutualisms are a recurring theme in animal utilization
of sugar-rich plant products. Phloem sap feeding differs
from nectar and fruit feeding in that it is generally an
antagonistic animal–plant interaction (see Introduction),
but it does involve other mutualisms both between animals
(the phloem feeders and their tenders) and between animals
and micro-organisms. Furthermore, on some plants, ants
both tend phloem-feeding hemipterans and protect the plant
from herbivory (Heil and McKey, 2003), thereby transforming an antagonistic relationship between plant and
phloem feeders into a crucial element to a three-way mutualism. As an example, the Crematogaster ants that form
nests in hollow stems of Macaranga species that lack extrafloral nectaries both tend scale insects and protect the
plant from natural enemies (Heckroth et al., 1999).
Future research directions
Recent research summarized in this article has identified
two key elements underpinning phloem sap utilization by
animals: possession of symbiotic micro-organisms that
provide essential amino acids; and carbohydrases with
transglucosidase function that reduce the osmotic pressure
of ingested phloem sap. However, these explanations are
partial in several ways. In particular, the molecular basis of
these capabilities is largely unknown. The transformations
of dietary sucrose in the insect gut are not understood at the
levels of either enzymological or gene function and, to my
knowledge, the only published study of insect utilization of
phloem sugars other than sucrose was conducted 50 years
ago (Duspiva, 1955). Similarly, the molecular basis of the
nutrient and signal exchange underpinning the production
and release of essential amino acids by Buchnera in aphids
and by the symbiotic micro-organisms in other phloemfeeding hemipterans is still unknown. The other key
limitation to our understanding is the narrow perspective
of current research, with near-exclusive focus on aphid
utilization of the most abundant phloem constituents,
sugars and amino acids. This is unsurprising given the
amenability of aphids to experimental manipulation and
analysis of minor constituents of phloem sap. However,
other potentially important phloem constituents are lipids,
the phloem mobility of which is poorly known, and
753
minerals. Recent developments in molecular and analytical
approaches to study the molecular physiology of hemipterans (Douglas, 2003) and to dissect plant sieve element
function and phloem sap composition (see the other papers
in this Focus section) offer exciting new opportunities to
study the traits of hemipteran insects which promote the
phloem-feeding habit. Only then will it be possible to
resolve the fundamental question in this field: why has
utilization of phloem sap as dominant or sole diet uniquely
evolved in hemipteran insects?
Acknowledgements
I thank Professor Doyle McKey, Professor Diane W Davidson, and
Dr Steven C Cook for helpful comments on a draft of this
manuscript. Previously unpublished data presented in this article
was obtained in research funded by BBSRC grant 87/S16725.
References
Adams D. 1997. Host plant effects on an aphid–bacterial symbiosis.
PhD thesis, University of York, UK.
Ashford DA, Smith WA, Douglas AE. 2000. Living on a high sugar
diet: the fate of sucrose ingested by a phloem-feeidng insect, the
pea aphid Acyrthosiphon pisum. Journal of Insect Physiology 46,
335–342.
Baumann L, Thao ML, Hess JM, Johnson MW, Baumann P.
2002. The genetic properties of the primary endosymbionts of
mealybugs differ from those of other endosymbionts of plant
sap-sucking insects. Applied and Environmental Microbiology 68,
3198–3205.
Birkle LM, Minto LB, Douglas AE. 2002. Relating genotype and
phenotype for tryptophan synthesis in an aphid-bacterial symbiosis. Physiological Entomology 27, 1–5.
Botha CEJ, Malcolm SB, Evert RF. 1977. An investigation of
preferential feeding habit in four Asclepiadaceae by the aphid
Aphis nerii B. de F. Protoplasma 92, 1–19.
Brudenell AJP, Griffiths H, Rossiter JT, Baker DA. 1999. The
phloem mobility of glucosinolates. Journal of Experimental
Botany 50, 745–756.
Buchner P. 1965. Endosymbiosis of animals with plant microorganisms. Chichester, UK: Wiley.
Byrne DN, Hendrix DL, Williams LH. 2003. Presence of
trehalulose and other oligosaccharides in hemipteran honeydew,
particularly Aleyrodidae. Physiological Entomology 28, 144–149.
Cristofoletti PT, Ribeiro AF, Deraison C, Rahbe Y, Terra WR.
2003. Midgut adaptation and digestive enzyme distribution in
a phloem feeding insect, the pea aphid Acyrthosiphon pisum.
Journal of Insect Physiology 49, 11–24.
Corbesier L, Havelange A, Lejeune P, Bernier G, Perilleux C.
2001. N content of phloem and xylem exudates during the
transition to flowering in Sinapis alba and Arabidopsis thaliana.
Plant, Cell and Environment 24, 367–375.
Dadd RH. 1985. Nutrition: organisms. In: Kerkut GA, Gilbert LI,
eds. Comprehensive insect physiology, biochemistry and pharmacology, Vol 4. Oxford, UK: Pergamon Press, 313–391.
Dailey GC, Ahrlich PR, Haddad NM. 1993. Double keystone
bird in a keystone species complex. Proceedings of the National
Academy of Sciences, USA 90, 592–594.
Davidson DW, Cook SC, Snelling RR, Chua TH. 2003. Explaining
the abundance of ants in lowland tropical rainforest canopies.
Science 300, 969–972.
754 Douglas
Dolling WR. 1991. The Hemiptera. Oxford, UK: Oxford University
Press.
Douglas AE. 1988. Sulphate utilization in an aphid symbiosis. Insect
Biochemistry 18, 599–605.
Douglas AE. 1989. Mycetocyte symbiosis in insects. Biological
Reviews 69, 409–434.
Douglas AE. 1993. The nutritional quality of phloem sap utilized by
natural aphid populations. Ecological Entomology 18, 31–38.
Douglas AE. 1998. Nutritional interactions in insect–microbial
symbioses. Annual Reviews of Entomology 43, 17–37.
Douglas AE. 2003. Nutritional physiology of aphids. Advances in
Insect Physiology 31, 73–140.
Duspiva F. 1955. Enzymatische Prozesse bei der Honingtaubuldung
der Aphiden. Vergleichen de Deutsche Zoologie Gesellschaft
18, 440–447.
Fisher DB. 2000. Long-distance transport. In: Buchanan B,
Gruissem W, Jones R, eds. Biochemistry and molecular biology
of plants. Rockville, USA: American Society of Plant Physiologists, 730–784.
Febvay G, Liadouze I, Guillaud J, Bonnot G. 1995. Analysis of
energetic amino acid metabolism in Acyrthosiphon pisum: a
multidimensional approach to amino acid metabolism in aphids.
Archives of Insect Biochemistry and Physiology 29, 45–69.
Febvay G, Rahbe Y, Rynkiewicz M, Guillaud J, Bonnot G. 1999.
Fate of dietary sucrose and neosynthesis of amino acids in the
pea aphid, Acyrthosiphon pisum, reared on different diets. Journal
of Experimental Biology 202, 2639–2652.
Folling M, Knogge C, Bohme W. 2001. Geckos are milking
honeydew-producing planthoppers in Madagascar. Journal of
Natural History 35, 279–284.
Geiger DR, Servaites JC. 1994. Diurnal regulation of photosynthetic carbon metabolism in C-3 plants. Annual Reviews of
Plant Physiology 45, 235–256.
Gil R, Silva FJ, Zientz E, et al. 2003. The genome sequence of
Blochmannia floridanus: comparative analysis of reduced genomes. Proceedings of the National Academy of Sciences, USA
100, 9388–9393.
Girousse C, Bournoville R, Bonnemain J-L. 1996. Water deficitinduced changes in concentrations in proline and some other amino
acids in the phloem sap of alfalfa. Plant Physiology 111, 109–113.
Goodchild AJP. 1966. Evolution of the alimentary canal in the
Hemiptera. Biological Reviews 41, 97–140.
Griffiths H. 1998. Stable isotopes. Oxford, UK: BIOS Scientific
Publishers.
Hartmann T. 1999. Chemical ecology of pyrrolizidine alkaloids.
Planta 207, 483–495.
Heckroth HP, Fiala B, Gullan PJ, Maschwitz U, Azarae HI. 1999.
The soft scale (Coccidae) associates of Malyasian ant-plants.
Journal of Tropical Ecology 14, 427–443.
Herrera CM, Pellmyr O. 2002. Plant–animal interactions. Oxford:
Blackwell Science.
Heil M, McKey D. 2003. Protective ant–plant interactions as model
systems in ecological and evolutionary research. Annual Reviews
of Ecology, Evolution and Systematics 34, 425–453.
Hunt JH. 2003. Cryptic herbivores of the rainforest canopy. Science
300, 916–917.
Karley AJ, Douglas AE, Parker WE. 2002. Amino acid composition and nutritional quality of potato leaf phloem sap for aphids.
Journal of Experimental Biology 205, 3009–3018.
Kehr J, Hustiak F, Walz C, Willmitzer L, Fisahn J. 1998.
Transgenic plants changed in carbon allocation pattern display
a shift in diurnal growth pattern. The Plant Journal 16, 497–503.
Merritt SZ. 1996. Within-plant variation in concentrations of amino
acids, sugar, sinigrin in phloem sap of black mustard, Brassica
nigra (L.) Koch. Journal of Chemical Ecology 22, 1133–1145.
Mittler TE. 1958. The excretion of honeydew by Tuberolachnus
salignus (Gmelin). Proceedings of the Royal Entomological
Society (London) 33, 49–55.
Passamani M, Rylands AB. 2000. Feeding behaviour of Geoffrey’s
marmoset (Callithrix geoffroyi) in an Atlantic forest fragment of
south-eastern Brazil. Primates 41, 27–38.
Ponder KL, Pritchard J, Harrington R, Bale JS. 2000. Difficulties
in location and acceptance of phloem sap combined with reduced
concentration of phloem amino acids explain lowered performance
of the aphid Rhopalosiphum padi on nitrogen deficient barley
(Hordeum vulgare) seedlings. Entomologia Experimentalis et
Applicata 97, 203–210.
Shigenobu S, Watanabe H, Hattori M, Sakaki Y, Ishikawa H.
2000. Genome sequence of the endocellular bacterial symbiont of
aphids Buchnera sp APS. Nature 407, 81–86.
Suh SO, Noda H, Blackwell M. 2001. Insect symbiosis: derivation
of yeast-like endosymbionts within an entomopathogenic filamentous lineage. Molecular Biology and Evolution 18, 995–1000.
Tamas I, Klasson L, Canback B, Naslund AK, Eriksson A-S,
Werenegreen JJ, Sandström JP, Moran NA, Andersson SGE.
2002. 50 million years of genomic stasis in endosymbiotic
bacteria. Science 296, 2376–2379.
Telang A, Sandström J, Dyreson E, Moran NA. 1999. Feeding
damage by Diuraphis noxia results in nutritionally enhanced
phloem diet. Entomologia Experimentalis et Applicata 91,
403–412.
Thao ML, Moran NA, Abbot P, Brennan EB, Burckhardt DH,
Baumann P. 2000. Cospeciation of psyllids and their primary
prokaryotic endosymbionts. Applied and Environmental Microbiology 66, 2898–2905.
Thompson GA, Schulz A. 1999. Macromolecular trafficking in the
phloem. Trends in Plant Sciences 4, 354–360.
Van Ham RCHJ, Kamerbeek J, Palacios C, et al. 2003. Reductive
genome evolution in Buchnera aphidicola. Proceedings of the
National Academy of Sciences, USA 100, 581–586.
Walters FS, Mullin CA. 1988 Sucrose-dependent increase in
oligosaccharide production and associated glycosidase activities
in the potato aphid Macrosiphum euphorbiae (Thomas). Archives
of Insect Biochemistry and Physiology 9, 35–46.
Wilkinson TL. 1998. The elimination of intracellular microorganisms from insects: an analysis of antibiotic-treatment in the
pea aphid (Acyrthosiphon pisum). Comparative Biochemistry and
Physiology A 119, 871–881.
Wilkinson TL, Adams D, Minto LB, Douglas AE. 2001.
The impact of host plant on the abundance and function of
symbiotic bacteria in an aphid. Journal of Experimental Biology
204, 3027–3038.
Wilkinson TL, Ashford DA, Pritchard J, Douglas AE. 1997.
Honeydew sugars and osmoregulation in the pea aphid Acyrthosiphon pisum. Journal of Experimental Biology 200, 2137–2143.
Wilkinson TL, Douglas AE. 2003. Phloem amino acids and the host
plant range of the polyphagous aphid, Aphis fabae. Entomologia
Experimentalis et Applicata 106, 1–11.
Wilson EO, Holldöbler B. 2005. The rise of ants: a phylogenetic
and ecological explanation. Proceedings of the National Academy
of Sciences, USA 102, 7411–7414.
Winter H, Lohaus G, Heldt HW. 1992. Phloem transport of amino
acids in relation to their cytosolic levels in barley leaves. Plant
Physiology 99, 996–1004.
Ziegler H. 1975. Nature of transported substances. In: Zimmerman
MH, Milburn JA, eds. Encyclopedia of plant physiology, New
series, Vol. 1. Berlin: Springer-Verlag, 59–100.
Zientz E, Dandeker T, Gross R. 2004. Metabolic interdependence
of obligate intracellular bacteria and their insect hosts. Microbiology and Molecular Biology Reviews 68, 745–770.