Shifting from clonal to sexual reproduction in aphids: physiological and developmental aspects


  • Gaël Le Trionnaire,

    1. INRA Rennes, Agrocampus Rennes, Université Rennes 1, UMR 1099 BiO3P (Biology of Organisms and Populations Applied to Plant Protection), BP 35327, F-35653 Le Rheu, France
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  • Jim Hardie,

    1. Imperial College London, Faculty of Natural Sciences, Department of Life Sciences, Division of Biology, Silwood Park Campus, Ascot, Berkshire SL5 7PY, U.K.
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  • Stéphanie Jaubert-Possamai,

    1. INRA Rennes, Agrocampus Rennes, Université Rennes 1, UMR 1099 BiO3P (Biology of Organisms and Populations Applied to Plant Protection), BP 35327, F-35653 Le Rheu, France
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  • Jean-Christophe Simon,

    1. INRA Rennes, Agrocampus Rennes, Université Rennes 1, UMR 1099 BiO3P (Biology of Organisms and Populations Applied to Plant Protection), BP 35327, F-35653 Le Rheu, France
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  • Denis Tagu

    Corresponding author
    1. INRA Rennes, Agrocampus Rennes, Université Rennes 1, UMR 1099 BiO3P (Biology of Organisms and Populations Applied to Plant Protection), BP 35327, F-35653 Le Rheu, France
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To whom correspondence should be addressed (email


Developmental biology is one of the fastest growing and fascinating research fields in life sciences. Among the wide range of embryonic development, a fundamental difference exists between organisms with sexual or asexual development. Aphids are unusual organisms which display alternative pathways of sexual and asexual development, the orientation of the pathway being determined by environmental conditions. These insects offer an adapted system in which to study developmental plasticity, because a side-by-side comparison of sexual and asexual development can be made in individuals with the same genotype. In this review, we describe the developmental mechanisms that have evolved in aphids for alternative sexual and asexual reproduction. In particular, we discuss how environmental cues orientate the reproductive mode of aphids from signal perception to endocrine regulation, and propose a comparative analysis of sexual and asexual gametogenesis and embryogenesis, which has been possible due to the development of molecular methods. As a result of the recent development of genomic resources in aphids, we expect these species will permit major advances in the study of the genomic basis underlying the choice of developmental fate and multiple reproduction strategies.

Abbreviations used:

juvenile hormone


RNA interference




One central question of modern biology is how developmental pathways of organisms are determined and how the machinery that regulates body plan and organ development has evolved. Among the wide range of shapes and forms found in the living world, a fundamental difference exists between organisms with sexual or asexual development. Sexual development is initiated following the fusion of male and female gametes produced through meiosis, whereas asexual development of a new individual is achieved without meiosis and fertilization. While eukaryotic life is dominated by sexual organisms, a substantial number of species develop and reproduce asexually (e.g. budding in yeasts). Aphids share the ability of alternative pathways of sexual and asexual development with a few other animal groups (e.g. cynipid wasps, cladocerans, ostracods and trematodes), the orientation of the pathway being determined by environmental conditions. These species offer adapted systems to study developmental plasticity, because a side-by-side comparison of sexual and asexual development can be made in otherwise clonal individuals.

Aphids are small insects and constitute a major threat to agriculture by ingesting plant sap for nutrition and by transmitting virus diseases to many crops. They belong to the Order Hemiptera (sucking bugs) and are placed with scales, psyllids and whiteflies in the Sub-Order Sternorrhyncha. Aphids form a monophyletic group that diverged around 280 million years ago (Grimaldi and Engels, 2005). Approx. 4400 species of aphids have been described and all share apomictic parthenogenesis (clonal or asexual reproduction) as the main or exclusive mode of reproduction. However, sexual reproduction in each generation was the ancestral mode from which they evolved. Their typical annual life cycle is characterized by a succession of parthenogenetic generations and a single sexual one (Moran, 1992; Simon et al., 2002). Therefore, by using the same set of genes, aphids are able to develop sexually or asexually—the two most extreme developmental modes. This phenotypic dichotomy is usually triggered by photoperiodic cues and will be referred to as reproductive polyphenism (Figure 1).

Figure 1.

Different morphs of the pea aphid

(A) A parthenogenetic female giving birth by viviparity to a larva stage 1, its genetic clone. (B) Copulation of sexual female and male. Reproduced with permission from Bernard Chaubet (UMR BiO3P, INRA, Rennes, France).

Reproductive polyphenism is a spectacular demonstration of the high level of phenotypic plasticity displayed by aphids. Understanding how a single genotype can develop such different phenotypes (clonal females that reproduce sexually or asexually and which also differ in morphology, behaviour, physiology and biochemistry) still remains a challenge for researchers. This reflects the exceptional flexibility of aphid genomes at both structural and expressional levels (the fluid genome; Casacuberta, 2004) and raises questions of its evolutionary and ecological significance.

In the present review, we discuss how aphids alternate their reproductive mode from viviparous parthenogenesis in the spring and summer (short nights) to oviparous sexual reproduction in the autumn (long nights), and thus cope with seasonal changes of the environment (Figure 2). Aphids are able to measure night length in the cephalic region and to transduce the signal to the target cells located in the ovaries. There, either haploid gametes are produced that require fertilization (sexual reproduction) or diploid oocytes initiate embryogenesis in the absence of male gametes and meiosis (parthenogenesis). In particular, we will review: (1) gametogenesis and embryogenesis in sexual and asexual morphs, and (2) processes by which environmental cues direct reproductive mode from signal detection to endocrine regulation. In light of the recent development of genomic resources in aphids (see Brisson and Stern, 2006; Tagu et al. 2008), we expect these species will permit major advances in the study of the genomic basis underlying the choice of developmental fate and multiple reproduction strategies.

Figure 2.

Life cycle of an aphid (e.g. the pea aphid)

In spring, eggs hatch and young aphid larvae develop into the adult stage. These adults are parthenogenetic females which reproduce by parthenogenesis during spring and summer when days are long. Each adult female can give birth by viviparity to approx. 80 clonal progeny in approx. 10 days. This exponential and high demographic expansion partly explains the severe damage aphids can cause to crops. In autumn, when the day length shortens, parthenogenetic aphids produce sexual morphs which mate. Fertilized females lay eggs over winter that are resistant to low and freezing temperatures.

Gametogenesis and embryogenesis in sexual and asexual morphs

Reproductive system of sexual and asexual females

The reproductive tracts of parthenogenetic or sexual aphid females are similar in their general organization and embryonic development. The female reproductive tract comprises two ovaries each containing several ovarioles. Each ovariole consists of an apical germarium joined to follicular chambers (Figure 3). In the ovarioles, each germarium contains germ cells and nurse cells (or trophocytes). Trophocytes are connected to the following follicular chambers by trophic cords. Aphid ovarioles are thus of the telotrophicmeroistic type. Follicular chambers contain either developing embryos (asexual reproduction) or eggs (sexual reproduction). The sexual female ovaries additionally possess spermathecae and accessory glands. The embryonic development of the female reproductive system is similar in asexual and sexual females. Germ cells form spherical cysts which divide in 32 oogonial cells contained in the fully-formed germarium (Blackman, 1987). These 32 cells are probably formed by synchronous divisions and are interconnected; approximately half of them develop into nurse cells and the other half into oocytes (Büning, 1985). In the pea aphid (Acyrthosiphon pisum), one germarium contains approx. 20 nurse cells for 11 future oocytes (Miura et al., 2003). Germaria in sexual females are larger than in asexuals (Tsitsipis and Mittler, 1976). Nuclei of nurse cells in the germarium drastically enlarge by endomitosis only in sexual females. Sexual females undergo oogenesis, and the future oocytes at the basal part of their germaria enter meiosis. By contrast, asexual females are viviparous, and the future oocytes of their germarium do not enter meiosis (see below).

Figure 3.

Representation of aphid ovarioles

Two ovaries form the female genital tractus in aphids. Each ovary is formed of several ovarioles. Each ovariole is a chain of different follicles with a germarium at the top, followed by a vitellarium. The germarium contains the future oocytes, as well as nurse cells or trophocytes. (A) Ovariole of a parthenogenetic viviparous female. (B) Ovariole of a sexual oviparous female. de, developing embryo, fc, follicle cells; gc, germarium cell; gl, germarium lumen; me, mature embryo; o, oocyte; pb, polar body; po, pre-oocyte; sb, syncytial blastoderm; tc, trophic cord; te, ye, yolky egg. Adapted from ‘Reproduction, cytogenetics and development’ in ‘Aphids: their Biology, Natural Enemies and Control’ (Minks, A.K. and Harrewijn, A.P., eds), 1987, vol. 2A, pp. 163–195, Elsevier, Figure 3.8 © R.L. Blackman, and ‘The endocrine control of polymorphism in aphids’ by A.D. Lees in ‘Endocrinology of Insects’ (Downer, R.G.H. and Laufer, N., eds), 1983, pp. 369–377, Figure V-5-2, © Alan R. Liss.

Oogenesis and embryogenesis in sexual females

During the formation of the germarium of the future sexual females, the future oocytes remain blocked in metaphase I (Blackman, 1976). They are released in a follicle chamber, where they enter a growth phase with accumulation of yolk in their cytoplasm (Figure 3). The paired chromosomes become condensed until the fertilization of the fully grown oocyte. Aphid chromosomes are holocentric and homologous chromosomes are associated along their total lengths with many chiasmata. Oocytes are fertilized when they pass into the common oviduct beside the spermathecae, before being oviposited. At this stage, they still are in metaphase of the first maturation division, and the two maturation divisions, releasing two polar bodies, are completed within the egg after spermatozoid penetration and before the haploid nuclei fuse (see Blackman, 1987).

Eggs are less than 1 mm in length and filled with yolk. They usually darken after fertilization and oviposition by forming a serosal cuticle. Initial rounds of cell division result in a syncytium, and a group of nuclei migrates to the periphery of the egg, whereas others remain within the yolk. The density of nuclei within the blastoderm (18 to 24 h after egg laying) is high in the mid-periphery, lower at the anterior and even lower at the posterior pole. After fertilized eggs are laid (2 to 3 days), the germ band falls into the yolk, and by the end of day 15 the embryo has completed anatrepsis (first inversion of anterior—posterior and dorsal—ventral axes of the embryo) and is fully segmented (Miura et al., 2003). At this stage, the embryo enters diapause, a period of arrested development, which it undergoes in response to photoperiodic cues. In the pea aphid, the early stages of sexual embryogenesis progress at a temperature-independent rate, but later stages progress at a temperature-dependent rate. The period of temperature-independent development ends with katatrepsis (recovering of the anterior—posterior and dorsal—ventral orientation as before anatrepsis, approx. 50 days after oviposition). Thus, as in other insects, diapause in the pea aphid is a mechanism that controls embryonic development, rather than a blocker of development (Shingleton et al., 2003). Pea aphid eggs maintained at 16°C showed considerable embryonic malformations with accelerated development of some, but not all parts, of the body (Shingleton et al., 2003). This contrasts with parthenogenetic embryogenesis (see below), which is accelerated by high temperatures (within range limits) without causing any developmental defects.

Development of embryos in asexual females

In parthenogenetic females, embryos develop within the ovariole from a diploid oocyte after a single modified meiosis II division in the absence of chromosome recombination. The future oocytes, located in the posterior part of the germarium, rapidly differentiate one by one. The several morphological modifications occurring in these cells have been described previously by Blackman (1987): within the germarium, the cells are arrested at an early prophase stage, but the future oocyte selected to differentiate increases in size and enters prophase of the meiosis-II-like division. This oocyte is then released from the germarium, forms a new follicular chamber and its chromosomes rapidly condense (Figure 4). As this cell grows, its nucleus migrates to the periphery of the cytoplasm and enters metaphase. A single maturation division occurs, giving a polar body and a diploid oocyte, which undergoes a series of mitotic divisions to initiate embryo development (Figure 3). During this single maturation division there is no exchange of chromosomal material (for discussion, see Sloane et al., 2001), even though this absence of recombination was strongly debated until the 1980s (Blackman, 1987). Immediately after the release of one oocyte from the germarium, another oocyte starts to differentiate. Thus, within an ovariole, there is a sequential chain of a germarium, differentiating oocyte-like cells and developing embryos, each included in a different follicular chamber (Figure 3). The two first follicular chambers are connected to the trophocytes within the germarium by trophic cords, whereas differentiating and developing embryos of the subsequent follicle chambers are probably supplied with nutrients via follicular cells surrounding the embryos (Tsitsipis and Mittler, 1976). During the life of an adult, approx. six embryos (in the pea aphid, for example) are produced per ovariole, with an average of 72 embryos per asexual female. This number is much higher than the number of differentiating oocytes in sexual females, with only one (occasionally two) oocyte per ovariole.

Figure 4.

Oocyte development in the parthenogenetic pea aphid

Microtubules (green) were revealed by staining with antibodies against β-tubulin, and DNA (red) was visualized with propidium iodide. (A) A germarium containing future oocytes with condensed DNA content. A selected oocyte has already left the germarium to form a new follicle. A trophic cord extends from the germarium to this follicle. (B) Condensed chromosomes of the parthenogenetic oocyte during the unique meiosis-like division. (C) Synchronous mitotic divisions forming the syncytium with self-organization of cortical microtubules into asters. Confocal microscopy on whole-mount preparations of ovarioles. f, follicle; g, germarium; m, mitosis; tc, trophic cord; o, oocyte. Reproduced with permission from G. Callaini and M.G. Riparbelli (University of Siena, Siena, Italy).

In sexual organisms, oocytes usually lack centrosomes and one copy of the centrosome is given by the male gamete during fertilization (Rodrigues-Martins et al., 2007). By contrast, oocytes of parthenogenetic aphids are able to self-organize microtubule-based asters, which are necessary for the first mitotic spindle (Riparbelli et al., 2005), in the absence of male centrosomes. Thus aphid parthenogenetic oocytes can assemble new centrosomes. However, this assembly de novo generates more than the two necessary centrosomes, with the risk of provoking cell division failures. In aphids, only two of the several asters are used for the formation of one single mitotic spindle, preventing future abnormal development of the embryo.

Using cross-reacting antibodies against the germline markers Vasa and Nanos of Drosophila melanogaster, it has been showed that germ-plasm specification in the pea aphid occurs very early during oocyte differentiation (Chang et al., 2006). In Drosophila, Nanos is a transcription factor involved in the establishment of the posterior organizing centre in the embryo, whereas the Vasa protein is required to direct the Nanos maternal mRNA into the posterior part of the egg. In parthenogenetic pea aphids, Nanos was identified in the cells designated as oocytes within the germarium, whereas Vasa was localized during the second nuclear division of the oocyte within the follicle chambers. This corresponds to Stages 0 and 1 respectively of the different embryonic stages that have been defined by Miura et al. (2003). At Stage 3, 16 nuclei of the syncytium are uniformly distributed, and, at Stage 4, some of the 32 nuclei migrate to the periphery. Aphids house symbiotic bacteria (Buchnera) in their abdomen which are transmitted vertically to the offspring during embyrogenesis by bacterial cell migration. Bacteria from the mother migrate and are incorporated at the 32-nuclei stage, when cell membranes begin to form around the 32 nuclei. From Stages 8 to 15, the asexual embryo undergoes successively anatrepsis and katatrepsis, as for sexual embryogenesis. During the later stages, the main body parts differentiate (e.g. eyes, muscles, appendages etc.). Germ-cell migration has been monitored during embryogenesis using the Vasa marker (Chang et al., 2007). To achieve this, the Vasa gene of the pea aphid was cloned and used for RNA in situ hybridization in ovarioles. During gastrulation (Stages 8–10), germ cells are associated with the germ band, then migrate towards the mid-region of the egg chamber together with the endosymbiotic bacteria.

These anatomical and morphological observations indicate that sexual and parthenogenetic embryogeneses progress through two strongly contrasted pathways (Table 1). Gene expression analyses in silico, performed with different cDNA libraries of the pea aphid, showed that parthenogenetic embryos express many genes of unknown functions compared with other fully sexual insects (Sabater-Muñoz et al., 2006). This might indicate specialized differentiation processes specific to parthenogenetic embryogenesis in aphids. Interestingly, two transcripts encoding proteins homologous with Drosophila proteins involved in regulation of sex determination (Transformer-2) and dosage compensation for X-chromosome-expressed genes [MSL3 (male-specific lethal-3)] were detected as being specifically expressed in parthenogenetic embryos. However, it remains unknown whether sex determination and dosage compensation mechanisms in aphids are similar to those of Drosophila. We expect that the construction of EST (expressed sequence tag) libraries that are specific to parthenogenetic and sexual embryogenesis will allow advances in the understanding of alternative developmental pathways in aphids.

Table 1.  Comparative analysis of asexual and sexual development in aphids (from Braendle et al., 2003; Miura et al., 2003)
OocytesOocytes blocked in metaphase of meiosis I in the germarium and the vitellariumPre-oocytes blocked in prophase, within the germarium
 Meiosis resumes after fertilization (expulsion of two polar bodies)First mitotic division (expulsion of one polar body) occurs within the first follicle chamber
Eggs/embryosOne or two eggs per ovarioleSix to ten embryos per ovariole
 Egg=1 mm longEmbryo=60 μm long
 Large amount of yolkLittle if any yolk. No serosal cuticle, vitellogenesis or chorion
 Duration of the syncytium stage longer 
DevelopmentRate of development: 100 daysRate of development: 10 days
 Discontinuous: a rapid early step and a late longer stepContinuous in duration
  Trophic cords thinner
Primary symbiotic bacteriaIntegration by the posterior pole through multi-opening in the follicular epithelium. Bacteria are then integratedIntegrated after cellularization
Buchnera bacteria are housed in specialized aphid cells called bacteriocytes, located close to the ovariesIntegrated before fertilizationBacterial invasion at the posterior pole of the embryo

The switch between parthenogenetic and sexual development

The fate of the pre-oocytes during the formation of aphid germarium is controlled by external factors and, in particular, night length. Photoreceptors present in the brain detect day and night, and internal factors—probably endocrine signals—are induced to act on the germarium. The control of sexual morph production by night length was first observed by Marcovitch (1924) in aphids, and this was the first case of a photoperiodic response recorded in animals. Since then, it has been clearly demonstrated that photoperiodic changes direct reproductive mode in aphids and that temperature can modulate this response, so that high temperatures can override short days (Lees, 1959). Biological processes regulated by seasonal photoperiodism require, first, photoreceptors, then, secondly, a clock mechanism to measure the variation of day or night length (photophase and scotophase respectively) and, thirdly, a counter (or memory) to record the number of days that lie either side of a critical night length (night length that results in 50% of aphids giving birth to sexual or asexual forms). In host-alternating aphids, a pre-sexual winged female is induced by short days which flies to the winter-host plant where it produces the sexual females, or in some species both sexes (see, for example, Blackman, 1975). In aphids, the mechanism of the photoperiodic clock is still controversial (Saunders, 2005). Aphids clearly measure the length of the scotophase (Lees, 1973; Searle and Mittler 1982; Mittler and Matsuka, 1985); typically, sexual reproduction is induced for scotophases longer than 9–10 h, with a range of variation depending on species, clone and temperature. This threshold corresponds to early autumnal conditions. The counter mechanism has not yet been not identified, but at least approx. 10 consecutive light/dark cycles (again depending on species and temperature) with a ‘long’ scotophase are required to trigger a switch from asexual to sexual reproduction (Hardie and Vaz Nunes, 2001). The question whether an endogenous circadian rhythm is involved in the measurement of the scotophase in aphids has been widely debated. Several hypotheses are still under debate; the circadian model proposes a clear circadian oscillator involved in photoperiod measurement, whereas, in the hour-glass model, the duration of the dark phase is involved in photoperiodism measurement (Hardie and Vaz Nunes, 2001; Shiga and Numata, 2007). The availability of genomic resources for aphids, along with appropriate biological experiments, should allow verification of whether circadian clock genes are involved in the response to seasonal photoperiodism.

Detection of photophase by aphids is performed in the head (Lees, 1964); cauterization of eyes or optic lobes did not affect the photoperiodic response (Lees 1964; Steel and Lees 1977), and localized illumination extending a short day to a long day showed that the centre of the dorsum of the head is photosensitive, suggesting a location of putative photoperiodic receptors in the brain (Lees, 1973). Removal of the antennae of aphids reared in long-night conditions did not affect their capacity to switch from parthenogenesis to sexual reproduction, suggesting that antennae are not involved in this process (G. Le Trionnaire, unpublished data). Lesions caused by microcautery in the anterior region of the protocerebrum (which destroy the medial Group I neurosecretory cells; see below) disabled the production of asexual individuals under long-day conditions in the vetch aphid, Megoura viciae (Steel and Lees, 1977). A summary of cauterized lesion studies indicated that brain regions lateral to the Group 1 cells also affected morph determination, and Steel (1976) proposed that the clock and photoreceptors might be located here. Analysis of action spectra involved in the photoperiodic control of reproductive polyphenism indicated that blue light (450–470 nm) wavelengths are most effective, and the shape of the curve is compatible with carotenoprotein receptors (Hardie et al., 1981). Immunocytochemistry (Gao et al., 1999) was consistent with an opsin-based photoreceptor located in the anterior dorsal region of the protocerebrum, but attempts to sequence the aphid brain opsin photoperiodic photoreceptor were unsuccessful (Gao et al., 2000). In order to reach a brain photoreceptor, light must penetrate the head capsule and this is achieved through a semi-transparent window in the dorsal head capsule and brain tissue (Hardie et al., 1981). Recently, transcriptomic analyses performed on heads of the pea aphid showed that different cuticular protein genes (proteins localized within or associated with the cuticle) were differentially expressed under short or long days (Le Trionnaire et al., 2007). Most arthropod cuticular proteins share conserved domains called RR (Rebbers—Riddiford) consensus sequences, which are classified into three sub-groups (Willis et al., 2005). On the basis of structural models of protein interactions, it has been proposed that both RR1 and RR2 domains are chitin-binding structural motifs. Among the pea aphid cuticular protein genes regulated by night-length increase, the three types of cuticular proteins were found, suggesting a remodelling of the head cuticle during a switch of photoperiodic regimes that might accompany photoperiodic signal transduction. The involvement of cuticular proteins in response to the increase of the scotophase was not expected. However, as the light reaching the receptors is subjected to the filtering action of the cuticle and any overlying tissues, we can expect that changing night length could generate modifications in cuticular protein composition related to a perception of a different quantity of light.

Early transduction of the photoperiodic signal probably occurs through two groups of approximately five neurosecretory cells (Group I), which are located in the anterio—dorsal part of the protocerebrum. Ablation of these cells provoked the production of sexual forms, even under long-day conditions (Steel and Lees, 1977; Hardie, 1987a). The chemical nature of the secreted material and its target cells are unknown. Axonal projections of Group I cells pass ventrally and follow the dorsal neuropile tracts to the thoracic ganglion mass (Hardie, 1987a). Axonal projections may terminate in the vicinity of the ovarioles, but it is not clear whether they are directly connected to the ovaries (as discussed by Steel, 1977) to release a putative neurohormone on to the target cells in the ovarioles. Alternatively, they might influence the corpora allata (glands distal to the brain) (Hardie, 1987b) and regulate the production of JH (juvenile hormone), a potent regulator of oocyte differentiation and parthenogenetic reproduction (Hardie, 1981) (see below).

Endocrine regulation of reproductive mode

There is support for JHs as a main factor in the transduction of the photoperiodic signal from the brain to the ovarioles. JHs (which are sesquiterpene compounds) are involved in many physiological functions of insects, including metamorphosis and reproduction. Several types of JHs exist in insects, and only JHIII has been reported in aphids in small amounts (Hardie et al., 1985; Westerlund and Hoffmann, 2004). Thus it is not yet possible to assess directly whether JHs are involved in the regulation of reproductive polyphenism. Despite these difficulties and large intra-individual variations, it has been shown that, in the vetch aphid, the concentration of one form of JH was higher in short-night- (parthenogenetic development) than long-night (sexual development)-reared insects (Hardie et al., 1985). In order to study the role of JHs in the regulation of reproductive mode, Mittler et al. (1976) ectopically applied kinoprene (an analogue of JHs) to long-night-reared insects and observed that they produced viviparous, instead of the expected oviparous, aphids. Thus JH might interfere with the photoperiod effect by mimicking long days. Hardie and Lees (1985) showed that the ovaries of such kinoprene-treated aphids were mixed, with some ovarioles being of the viviparous type and some of the oviparous type. Within an ovariole, the follicle chamber can be either of the sexual or parthenogenetic type. Kinoprene is a potent insecticide (acting as a JH agonist) and its application has serious side effects, such as the number of larvae produced and the duration of each larval stage between two moults. Thus Hardie (1981) and Corbitt and Hardie (1985) applied naturally occurring JHs on aphids reared under long-night conditions, and demonstrated that JH could mimic short-night conditions by inducing parthenogenetic forms. Although judicious, prolonged topical application of JHs or JH analogues to females that should have produced only sexual females could produce seemingly normal parthenogenetic aphids, it is possible that the switch of reproductive mode is tightly regulated by JHs in combination with other still unknown effectors.

Many points remain unsolved, such as the type of JHs involved in reproductive orientation (see above), the regulation of their concentration in relation with the photoperiodic signal detection, the site of action (possibily the developing germarium or the follicle cells surrounding the follicle chambers), their mode of action (JH receptors have still not been identified in insects) and other molecules that JH might interact with (for example, ecdysteroids, which are insect hormones that control moulting). However, new molecular techniques, such as transcript profilings or gene expression modification by RNAi (RNA interference), should help in deciphering the molecular basis of JH action in aphid polyphenism.


As a response to seasonal changes of their environment, the evolution of a spectacular phenotypic plasticity of their reproductive mode has enabled aphids to survive in adverse conditions in the egg stage and to reproduce rapidly through viviparous parthenogenetic generations. This adaptation results in fundamental differences in the development and differentiation of sexual and asexual aphids, although it is achieved by using the same set of genes. But mechanisms of reproductive polyphenism and development in aphids are far from fully depicted. The photoperiodic receptors are not known, the regulation of photoperiodism by circadian clock is still debated, the demonstration of the JH involvement in transduction still awaits molecular data, and genes and proteins involved in parthenogenetic compared with sexual embryogenesis have not yet been identified. The lack of breakthrough in the understanding of mechanisms of reproductive polyphenism can be explained in two ways. First, aphids have not been considered as model species for developmental studies, and it is only very recently that genomic resources have been developed (Brisson and Stern, 2006; Tagu et al., 2008). Owing to the creation of the International Aphid Genomics Consortium, a large collection of ESTs (expressed sequence tags) has been published (Sabater-Muñoz et al., 2006; Ramsey et al. 2007), several sets of microarrays have been used (Wilson et al., 2006; Brisson et al., 2007; Le Trionnaire et al., 2007), a database has been launched (Gauthier et al., 2007) and the genome has been sequenced and is under going annotation (http:www.hgsc.bcm.tmc.eduprojectsaphid). Thus, in the next few years, molecular tools will be available for large functional genomics analyses on sexual compared with asexual reproduction and development in aphids. Secondly, significant advances in developmental biology are possible only when mutants are available. Genetic transformation is not available in aphids, and will be probably very difficult to achieve, as it is for the majority of non-model insects; the low number of eggs produced and the long diapause of embryogenesis are the two main drawbacks of this technique. RNAi protocols now exist for aphids and allow transient knock-down of specific mRNA for approx. 4 days (Mutti et al., 2006; Jaubert-Possamai et al., 2007) This method has not yet been adapted to promote effective and high-throughput mutagenesis in aphids. Thus, in our opinion, it is now of prime importance to develop a mutagenesis system in aphids in order to decipher the molecular mechanisms involved in the regulation of sexual and asexual reproduction and development.


We thank Michel Philippe (Université de Rennes 1, Rennes, France), whose curiousity about this topic led to him suggesting that we write a review about it. We acknowledge Bernard Chaubet (INRA Rennes, UMR BiO3P, Rennes, France) for the photographs. This work was supported by ‘ANR (Agence Nationale pour la Recherche) Exdisum’.

While every effort has been made to obtain permission to reproduce or adapt copyrighted material in this paper, it is possible that in some instances this has not been done. Copyright holders who have not been suitably acknowledged should contact the publishers so that the situation can be rectified.


  1. Apomictic parthenogenesis: No reduction division occurs, so that the diploid offspring is produced without crossing over and has the same genetic constitution as the mother.

  2. Polyphenism: The occurrence of different phenotypes within a species where the development of the phenotype is governed by environmental conditions.

  3. Morph: An individual phenotype resulting from developmental plasticity (e.g. winged versus wingless, parthenogenetic versus sexual).

  4. Ovariole: The female reproductive system consists of a pair of ovaries. Each consists of a number of egg tubes or ovarioles, which are comparable with the testis follicles in males. Each ovariole consists of a distal germarium and a more proximal vitellarium. The germarium contains oocytes produced from oogonia. The vitellarium consists of a succession of follicles containing different stages of maturation of eggs or development of embryos. In meroistic ovarioles, nurse cells called trophocytes are present. In teletrophic ovarioles, nurse cells remain in the germarium and are connected to the follicles by nutritive cords.

  5. Germarium: see definition for Ovariole.

  6. Trophocyte: see definition for Ovariole.

  7. Teletrophic: see definition for Ovariole.

  8. Meroistic: see definition for Ovariole.

  9. Spermatheca: This is an organ of the female reproductive tract in insects which receives and stores sperm from the male.

  10. Holocentric chromosomes: In holocentric chromosomes, spindle fibres are attached all along one side of each sister chromosome. Centromeres are thus dispersed along the chromosomes.

  11. Serosal cuticle: The serosa is the membrane outside the yolk formed during embryogenesis and containing chitin.

  12. Anatrepsis: The movement of the developing embryo when the posterior end of the germ band is flexed upwards into the yolk. The embryo comes to lie with its head-end towards the posterior pole of the egg.

  13. Diapause: This is a delay of development that has evolved in response to recurring periods of adverse environmental conditions (e.g. cold and frost).

  14. Katatrepsis: The movement of the developing embryo following anatrepsis. The embryo moves round to lie with its head towards the anterior pole.

  15. Photophase: This is the light period of a photoperiodic cycle of light/dark.

  16. Scotophase: This is the night period of a photoperiodic cycle of light/dark.

  17. Critical night length: This results in 50% of aphids giving birth to sexual or asexual forms.

  18. Circadian oscillator: A daily rhythmic activity cycle, based on approx. 24-h intervals, that is exhibited by many organisms.

  19. Protocerebrum: This is the biggest region of the insect brain to which nerves from eyes, antennae and many other organs are connected. It is bi-lobed and is continuous laterally with optic lobes. It occupies the dorsal position in the head and contains the pars cerebralis, which is rich in neurosecretory cells.

  20. Corpora allata: These are paired or fused ganglion-like bodies in the head of insects. The bodies secrete juvenile hormones.

  21. Juvenile hormones (JHs): These hormones are sesquiterpenes produced by the corpora allata. In insects, they mainly regulate growth, metamorphosis and reproduction.

  22. Reproductive polyphenism: The different reproductive morphs (sexual and asexual) produced by one genotype under environmental changes.