INTRODUCTION
Insecta (insects) is the most successful class of animals on Earth. Seemingly, associated microorganisms have allowed insects to adapt to almost all ecosystems on the planet (Janson et al., 2008; Feldhaar, 2011). Insects with restricted diets, such as hematophagous insects that feed on blood (Rio et al., 2016) or phytophagous insects that feed on sap, bark, leaves or seeds (Hansen & Moran, 2014) face nutritional deficiencies of nitrogen, vitamins or lipids. Bacteria or fungi can provide insects with nutrients or metabolites that are deficient in their diet (Douglas, 2015; Harris et al., 2010; Hansen & Moran, 2014). Symbionts may inhabit inside insect cells and thus they have been designated “endosymbionts”.
More than 50 years ago, Buchner (1965) published his microscopic observations in a compendium of insect-associated bacteria. In his book “Endosymbiosis of Animals with Plant Microorganisms”, he described the morphology of bacteria that inhabit different tissues of phytophagous insects and he assumed that many insect bacteria have their origins in plants. Earlier microscopic studies of scale insect symbionts (or true cochineals) were performed by Walczuch in 1932 and Buchner included them in his 1965 review. Nowadays there is a large research interest in microorganisms associated with insects and several types of insect symbionts are known:
1) Primary or essential endosymbionts. They reside inside specialized cells that are called bacteriocytes in the case of insect cells containing bacteria (Baumann et al., 2000), or mycetocytes when they harbor fungi. A transcriptional factor essential for bacteriocyte development was discovered in insects (Matsuura et al., 2015) and this sets the basis to further understand the evolution of intracellular symbiosis in insects. Bacteriocytes are sometimes clustered in structures called bacteriomes (Fig. 1) that are found in distinct insect species (Baumann, 2005). Bacteriome location and numbers are variable. There are often two bacteriomes per insect, but there can be one to four. They are located in different regions of the insect. In the so-called “nutritional symbiosis”, primary endosymbionts provide their hosts with nutrients that are deficient in the insect diet, such as essential amino acids and vitamins (Baumann, 2005; Moran et al., 2005).
2) Facultative or secondary endosymbionts. They are not essential for the survival of their host. They may also be found inside bacteriocytes. Some of them protect the insect from the attack of wasps, nematodes, fungi, bacteria and viruses (Xie et al., 2014; Jaenike et al., 2010; Łukasik et al., 2013; Chrostek et al., 2013). They can also help insects in the degradation of plant toxic compounds such as monoterpenes or alkaloids, allowing insects to colonize or parasite plants. Secondary endosymbionts can protect insects from high temperatures (Tsuchida et al., 2004; Burke et al., 2010). Others can manipulate the reproductive capacity of the host (Werren et al., 2008). It is known that secondary symbionts alter cellular functions of hosts, ranging from signal transduction to apoptosis (Bentley et al., 2007; Ikeya et al., 2009). When bacteria do not confer an increased fitness, they may be lost without any negative consequence for the insect (Moran et al., 2005).
3) Gut symbionts in crypts. They have been found only in some insects of the superfamily Pentatomomorpha, which includes bugs. They are often transmitted vertically by colonization of eggs, coprophagia or symbiotic capsules, but some species can be acquired from the environment by food as some of them are free-living. They also provide nutrients, degrade polymeric and toxic plant compounds and recycle nitrogen (Kikuchi et al., 2008).
4) Gut symbionts. They contribute to the breakdown of complex chemical compounds and toxins. Gut symbionts may be vertically transferred from mother to offspring by fecal contamination, coprophagy and trophallaxis that leads to the inoculation of the gut bacteria to the progeny (Salem et al., 2015 ; Abdul Rahman et al., 2015). However, intestinal microbiota may be modified by changes in the diet. Except for termites, the number of species in insect microbiota is smaller than that of mammals. Diversity may vary in females and males and this is the case in monarch butterflies (Servín -Garcidueñas et al., 2014 and unpublished data). Using a functional approach, we have reported a meta-analysis of insect gut proteobacteria (Degli Esposti & Martínez-Romero, 2017).
Figure 1
Diagram of a dorsal section of a scale insect (Monophlebidae and Coelostomidiidae families), in which the location of bacteriomes and endosymbionts are shown. Based on diagrams of Walczuch (1932) and FISH experiments of Matsuura et al., (2009), Dhami et al., (2012), and Rosas-Pérez et al., (2014).
In this review, we will discuss primary and secondary endosymbionts, leaving aside gut and crypt bacteria. In general, a single insect hosts two or more different species of endosymbionts in co-symbiosis. Inheritability and reduced genomes are the most outstanding features of endosymbionts. Most endosymbionts cannot be cultured on standard laboratory media and to acknowledge this, most endosymbionts are designated “Candidatus” before the name of the species. To simplify descriptions here, we will not use the “Candidatus” designation. Some endosymbionts share their evolutionary history with their hosts. Maternal transfer, reduced genomes and co-divergence will be reviewed in this article focusing on scale insect endosymbionts. In addition, we present the diversity of bacterial and fungal symbionts found in two models of scale insects studied in our laboratory.
MATERNAL TRANSFER OF BACTERIAL SYMBIONTS
Vertical transfer refers to the transmission of endosymbionts from mothers to their offspring. It is a mechanism that ensures the permanence of these bacteria in insect populations. Endosymbionts are transmitted from the time the embryos are formed and when insects are born they already carry some bacteria (McFall-Ngai, 2002). By being transferred from one generation to another, bacteria can remain associated with insect species for millions of years. Of all endosymbionts studied so far, it is estimated that the oldest is Sulcia muelleri living for over 260 million years in a group of insects that includes cicadas, leafhoppers and planthoppers (Moran et al., 2005). Sulcia muelleri co-exists with different co-symbionts in different insects (Table I) and a metabolic complementation between the different co-symbionts that inhabit an insect seems to exist (McCutcheon & Moran, 2010). This means that if the genes that encode the biosynthetic pathway of a metabolite do not exist in one of the symbionts, they are probably present in other symbiont bacteria of the same insect.
Table I
Examples of co-symbionts (with phylum indicated) of Sulcia muelleri, the primary endosymbionts of Auchenorrhyncha
| Co-symbiont | Phylum | Genome size (kb) | Host (Family) | Reference |
|---|---|---|---|---|
| Nasuia deltocephalinicola | Betaproteobacteria * | 112 | Macrosteles quadrilineatus(Cicadellidae) | Bennett & Moran, 2013 |
| Zinderia insecticola | Betaproteobacteria | 208 | Clastoptera arizonana(Clastopteridae) | McCutcheon & Moran, 2010 |
| Vidania fulgoroideae** | Betaproteobacteria | ND | Oliarus filicicola(Cixiidae) | Bressan & Mulligan, 2013 |
| Purcelliella pentastirinorum** | Gammaproteobacteria | ND | Oliarus filicicola(Cixiidae) | Bressan & Mulligan, 2013 |
| Baumannia cicadellinicola | Gammaproteobacteria | 686 | Homalodisca coagulata(Cicadellidae) | Wu et al., 2006 |
| Arsenophonus*** | Gammaproteobacteria | ND | Macrosteles laevis(Cicadellidae) | Kobialka et al., 2016 |
| Sodalis-like PSPU | Gammaproteobacteria | 1,380 | Philaenus spumarius(Aphrophoridae) | Koga & Moran, 2014 |
| Hodgkinia cicadicola | Alfaproteobacteria | 144 | Diceroprocta semicincta(Cicadidae) | McCutcheon et al., 2009 |
* All betaproteobacteria co-symbionts might have the same origin in Auchenorrhyncha (as suggested by Bennett & Moran, 2013).
Some bacteria may have recently become endosymbionts acquired from the transfer of endosymbionts from another insect (horizontal transfer) or from the environment. When an insect acquires a new endosymbiont bacterium, the insect may gain new characteristics, such as the ability to feed on another plant (Moran, 2007). It has recently been described that endosymbionts such as Wolbachia can be transferred between insects through the plants on which the insects feed (Li et al., 2017; Sintupachee et al., 2006). Besides diet, endosymbionts can be exchanged by parasitoid wasps that transmit them when they sting different insects (Ahmed et al., 2015). In different species of aphids, males may transfer symbiotic bacteria to females during intercourse (Moran & Dunbar, 2006).
ENDOSYMBIONT REDUCED GENOMES AND GENETRANSFER TO INSECT HOSTS
Since endosymbionts live inside insects and have been transferred vertically for millions of years, they display different characteristics than free-living bacteria. Endosymbionts have lost most of their genome, sometimes retaining even less than 10% of the original one (Bennett & Moran, 2013; Martínez-Cano et al., 2015). They no longer retain genes essential for free living. Genes preserved include those under selective pressure and those related to the synthesis of the nutrients that the insect requires. For this reason, endosymbionts of insects with similar diets have retained similar genes, in an example of convergent evolution.
Experimental evolution studies performed in the laboratory have shown that an initial massive loss of DNA blocks occurs rapidly in bacteria (Nilsson et al., 2005). Comparison between the genomes of endosymbionts and those of free-living bacteria suggests that after deletions have occurred there is a proliferation of repetitive elements that causes chromosomal rearrangements. Later on, there is a slow accumulation of deleterious mutations. Non-functional genes are then lost (Moran, 2003; Manzano-Marín & Latorre, 2016). The presence of pseudogenes in the genomes of some endosymbionts suggests that their genome is still in a reduction process.
Most endosymbionts have A + T rich genomes (Table II). They also have high rates of non-synonymous nucleotide substitutions in protein-encoding genes. Endosymbionts live in restricted environments that prevent recombination with other bacteria. The strict vertical transmission has an effect on the population structure since there is a bottleneck with a limited number of bacteria passing to the next generation. Successive bottlenecks during evolution reduce bacterial diversity and increase their mutation fixation rate (Moran, 2007).
Table II
Bacterial endosymbionts in scale insects.
| Endosymbiont (Class) | Host (FAMILY) | Genome size (pb) | %GC | Number of pseudogenes | Function | Location | Reference |
|---|---|---|---|---|---|---|---|
| Walczuchella monophlebidarum (Flavobacteria)P | Llaveia axin axin (MONOPHLEBIDAE) | 309,299 | 32.6 | 27 | Provides amino acids | Bacteriome & ovaries | Rosas-Pérezet. al. (2014) |
| SodalisTME1 (Gammaproteobacteria)S | 3,400,000 | 55.6 | ND | Provides amino acids & recycles uric acid | Bacteriome & ovaries | Rosas-Pérezet. al. (2017) | |
| Tremblaya phenacola (Betaproteobacteria)P | Phenacoccus avenae(PSEUDOCOCCIDAE) | 170,756 | 42.2 | 178 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| Tremblaya princeps (Betaproteobacteria)P | Maconellicoccus hirsitus (PSEUDOCOCCIDAE) | 138,415 | 61.8 | 136 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| Doolittlea endobia (Gammaproteobacteria)S | 834,723 | 44.2 | 564 | Provides amino acids & vitamins | Tremblaya cytoplasm | Husnik & McCutcheon (2016) | |
| Tremblaya princeps (Betaproteobacteria) P | Ferrisia virgate (PSEUDOCOCCIDAE) | 141,620 | 58.3 | 132 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| Gullanella endobia (Gammaproteobacteria)S | 938,041 | 28.9 | 461 | Provides amino acids & vitamins | Tremblaya cytoplasm | Husnik & McCutcheon (2016) | |
| Tremblaya princeps (Betaproteobacteria)P | Planococcus citri (PSEUDOCOCCIDAE) | 138,927 | 58.8 | 125 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| Moranella endobia (Gammaproteobacteria)S | 538,294 | 43.5 | 419 | Provides amino acids & vitamins | Tremblaya cytoplasm | Husnik & McCutcheon (2016) | |
| Tremblaya princeps (Betaproteobacteria) P | Pseudococcus longispinus (PSEUDOCOCCIDAE) | 144,042 | 58.9 | 134 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| 2Sodalis-allied(Gammaproteobacteria)S | 8,190,816 | 53.9 | ND | Provides amino acids & vitamins | Tremblaya cytoplasm | Husnik & McCutcheon (2016) | |
| Tremblaya princeps (Betaproteobacteria)P | Tryonimus perrisii (PSEUDOCOCCIDAE) | 143,340 | 57.8 | 116 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| Hoaglandella endobia (Gammaproteobacteria)S | 628,221 | 42.8 | 510 | Provides amino acids & vitamins | Tremblaya cytoplasm | Husnik & McCutcheon (2016) | |
| Tremblaya princeps (Betaproteobacteria)P | Paracoccus marginatus (PSEUDOCOCCIDAE) | 140,306 | 58.2 | 124 | Provides amino acids & vitamins | Bacteriome | Husnik & McCutcheon (2016) |
| Hoaglandella endobia (Gammaproteobacteria)S | 352,837 | 30.6 | 273 | Provides amino acids & vitamins | Tremblaya cytoplasm | Husnik & McCutcheon (2016) | |
| Dactylopiibacterium carminicum (Betaproteobacteria)P | Dactylopius coccus (DACTYLOPIIDAE) | 3,589,384 | 62.7 | ND | Fixes N & recycles uric acid | Ovaries | Vera-Ponce de León et al. (2017) |
| Wolbachia bourtzisii wDacA (Alphaproteobacteria)S | 1,170,000 | 35.1 | ND | Provides riboflavin & heme groups | Ovaries | Ramírez-Puebla et al.(2016) | |
| Wolbachia pipientiswDacB (Alphaproteobacteria)S | 1,498,000 | 34.1 | ND | Provides riboflavin & heme groups | Ovaries | Ramírez-Puebla et al.(2016) | |
| Uzinura diaspidicola (Flavobacteria)P | Diaspis echinocacti (DIASPIDIDAE) | 263,431 | 30.2 | 7 | Provides amino acids | Bacteriocytes | Sabree et. al. (2013) |
| Sulcia muelleristr. GWSS (Flavobacteria)P | *Homalodisca vitripennis (CICADELLIDAE) | 245,530 | 22.4 | ND | Provides amino acids | Bacteriome | Husnik & McCutcheon (2016) |
| Baumannia cicadellinicola (Gammaproteobacteria)S | 686,192 | 33.2 | 9 | Provides vitamins | Bacteriome | Wu et al. (2006) | |
| Buchnera aphidicola str. APS (Gammaproteobacteria)P | *Acyrtosiphon pisum (APHIDIDAE) | 640,681 | 22.5 | 13 | Provides riboflavin | Bacteriome | Shigenobu et al. (2000) |
| Carsonella rudii (Gammaproteobacteria)P | *Pachypsylla venusta (PSYLLIDAE) | 159,662 | 16.6 | ND | Provides amino acids | Bacteriome | Tamames et al. (2007) |
| Blochmannia floridanus (Gammaproteobacteria)P | *Camponotus floridanus (FORMICIDAE) | 705,557 | 27.4 | 6 | Provides amino acids, participates in urea detoxification & N recycling | Gut & ovaries | Gil et al. (2003) |
In the evolutionary history of insects, it seems that endosymbiont replacements have occurred several times (Moran et al., 2005; Gruwell et al., 2010; Toju et al., 2013; Sudakaran et al., 2017) and this was shown experimentally with a secondary endosymbiont taking the place of a primary endosymbiont (Koga et al., 2003). The elimination of endosymbionts is also evident when bacterial genes of endosymbionts no longer present are found in the genomes of insects. It is presumed that these genes were transferred laterally by past endosymbionts that are no longer currently living in the insects, but have left a trace of their existence. Some insect genes of bacterial origin are transcribed in the bacteriome and may be required for the endosymbiotic relationship (Husnik et al., 2013; Sloan et al., 2014.). Lateral gene transfer from bacteria to the insect has been studied in four species of the suborder Sternorrhyncha (aphids, psyllids, whiteflies and mealybugs) (Nikoh & Nakabachi, 2009; Husnik et al., 2013; Sloan et al., 2014). Not all the transferred genes are the same in each species but they encode similar functions, such as peptidoglycan and amino acid biosynthesis (Sloan et al., 2014). Large genome fragments of Wolbachia have been found transferred to the chromosomes of mosquitoes (Aedes aegypti), vinegar fruit fly (Drosophila ananassae), beetles (Callosobruchus chinensis) and several species of parasitoid wasps of the genus Nasonia. Some of these genes have been found to be functional (Kondo et al., 2002; Dunning Hotopp et al., 2007; Nikoh et al., 2008; Klasson et al., 2009; Choi et al., 2015).
Extreme examples of reduced genomes are found in Nasuia and Vidania endosymbionts with genomes of 112 and 119 kb, respectively (Bennett et al., 2016). Endosymbionts with extreme genome reduction may need the insect contribution of essential enzymes or proteins as occurs in mitochondria. In some cases, several of these proteins are encoded by genes that were transferred from endosymbionts to the insect (Sloan et al., 2014). Remarkably, it was shown that a protein encoded by a gene of bacterial origin in an aphid, is transported to the endosymbiont (Nakabachi et al., 2014).
SHARED EVOLUTIONARY HISTORIES AMONG SYMBIONTS AND HOSTS
If endosymbiotic bacteria and their hosts remain associated for a long evolutionarily time, they can co-speciate or co-diverge together. In those cases, there is a phylogenetic congruence that can be traced back to a single infection event (Baumann et al., 2000).
Occasionally endosymbionts have evolutionary histories different from the hosts. This is more frequent in secondary co-symbionts due to endosymbiont losses, new acquisitions of environmental bacteria or lateral transfers from one insect species to another. Furthermore, it has been observed that secondary endosymbionts have a higher nucleotide substitution rate than primary symbionts (Bennett et al., 2014). Repeated sequences are found in co-symbionts such as Sodalis (enterobacteria) (Siguier et al., 2014).
It is considered that mitochondria evolved from an ancestral alphaproteobacterium more than a billion years ago and underwent a process of genome reduction (Gray, 1992; Gray, 1999; Martin, 2017) that might have been similar to the evolutionary process followed by insect endosymbionts (Sloan et al., 2014; Dunning Hotopp, 2011; McCutcheon, 2016). Mitochondria have very small genomes, in many cases rich in A + T such as in Insecta and Nematoda (Arunkumar & Nagaraju, 2006). Mitochondria have co-speciated with their hosts and they have transferred lots of genes to their hosts that encode proteins that are necessary for the mitochondria to carry out their metabolic functions (McCutcheon, 2016).
FUNGI SYMBIONTS IN INSECTS
Although bacteria-insect associations are usually the most studied, about eight orders of insects are known to host fungi. Arthropods and fungi have coexisted for around 200 million years and their earliest relationship dates from the Jurassic (Vega & Blackwell, 2005). Their interactions include all types of symbiosis, such as the obligatory mutualism of cicadas (Chen et al., 1981), commensalism of the Attini tribe of ants where fungi are their main food source, and parasitism (Hughes et al., 2016). Fungal symbionts can inhabit cavities within the insect such as the micangio in bark beetles (Jones et al., 1999; Klepzig & Six, 2004; Ganter, 2006), or in mycetocytes that are found in the rice pest Nilaparvata lugens or in Drosophila melanogaster (Cheng & Hou, 2001; Ebbert et al., 2003). Fungi can also inhabit the lumen of the digestive system, the Malpighian tubules, gonads and even venom-producing glands in Comperia merceti wasp (Gibson & Hunter, 2010; Rivera et al., 2009; Ricci et al., 2011; Vera-Ponce de León et al., 2016). Insects acquire fungi from other insects or by mycofagia when the insects feed directly from mycelia or yeasts as several species of Drosophila do (Gibson & Hunter, 2010; Becher et al, 2012). Fungi, like endosymbiont bacteria, can be transferred from the mother to her offspring such as the yeast-like symbionts in the case of N. lugens (Cheng & Hou, 2001) and Wickerhamomyces anomalus yeast in Anopheles stephensi mosquitoes (Ricci et al., 2011). The elimination of fungi causes a fitness decrease in their hosts (Sasaki et al., 1996; Menezes et al., 2015). The effects of fungi on host fitness have been demonstrated in cicadas (Sasaki et al., 1996) and in some beetles (Anobiidae and escolitins) (Ayres et al., 2000; Nasir & Noda, 2003). Fungi can supply insects with nitrogen metabolites and essential lipids lacking in their diet. They also participate in the degradation of high-density biological polymers, uric acid recycling, biotransformation of toxic chemicals in the environment, and pheromone production (D’Ettorre et al., 2002; Nasir & Noda, 2003; Vega & Blackwell, 2005; Vera-Ponce de León et al., 2016).
Bacterial endosymbionts of scaleinsects
Scale insects belong to the superfamily Coccoidea (Hemiptera: Sternorrhyncha) and include many plant pest species that cause enormous economic losses in agriculture (Miller et al., 2005). They all secrete a protective wax cover that gives the insect a cotton or waxy powder appearance (Fig. 2). Most of them feed on the plant’s phloem and are known as “cochineals”. There are about 7,800 species. Most adult females remain sessile once they introduce their stylet into the host plant. The stylet is a specialized mouth organ to suck sap. Some species of scale insects damage plants by transmitting diseases and by excretion of honeydew, a sticky waste substance composed of sugars and minerals that promotes fungal growth. Honeydew is a source of food for birds, mammals and other insects, especially for several species of ants (Kondo et al., 2009).
Figure 2
Photographs of some scale insect species mentioned in the review. Archive images from our group.
Symbiotic bacteria that provide scale insects with nitrogen metabolites, vitamins and co-factors have been described (Table II and Figure 3). Endosymbionts cannot be cultured in media in the laboratory because their highly reduced genomes lack many genes needed for independent living. Thus, their study relies on metagenomic analyses. Symbiont genome sequences deduced from metagenomes allows the inference of their metabolic capabilities and their role in symbiosis (e.g. Shigenobu et al., 2000; Sabree et al., 2013; Rosas-Pérez et al., 2014).
Figure 3
Phylogenetic reconstruction of 18S rRNA genes (570 bp) of scale insects and their relationship to bacterial endosymbionts with outgroup from Aphidoidea superfamily. One representative from each family or clade of scale insects was used. Different colors indicate the taxonomic identity of the endosymbionts: Flavobacteria, green; Alphaproteobacteria, yellow; Betaproteobacteria, blue; Gammaproteobacteria, red; Spiroplasma, purple. Dotted branches show endosymbiont losses. Branches from outside correspond to endosymbionts acquired by horizontal transfer. Asterisk shows the likely loss of Flavobacteria and replacement by other endosymbionts. All Flavobacteria except Brownia rhizoecola are phylogenetically related. The + sign at the nodes indicates a bootstrap value > 50%. Not all endosymbionts are present in all species from the same family. Diagram based on von Dohlen et al., 2001; Zchori-Fein et al., 2005; Gruwell et al., 2007; Matsuura et al., 2009; Gruwell et al., 2010; Rosenblueth et al., 2012; Dhami et al., 2013; Husnik & McCutcheon, 2016; Vera-Ponce de León et al., 2017.
The presence of endosymbionts within bacteriocytes has been detected in ten families of scale insects by culture independent methods (von Dohlen et al., 2001; Zchori-Fein et al., 2005; Gruwell et al., 2007; Matsuura et al., 2009; Gruwell et al., 2010; Rosenblueth et al., 2012; Dhami et al., 2013; Husnik & McCutcheon, 2016) (Figure 3). It is possible to locate these bacteria within bacteriocytes by fluorescent in situ hybridization (FISH) (Fig. 1; Kono et al., 2008; Matsuura et al., 2009; Dhami et al., 2012; Rosas-Pérez et al., 2014). Enterobacteriaceae (gammaproteobacteria), especially those close to the genus Sodalis (Sodalis-like) and Flavobacteria (Bacteroidetes), have been found in six families, namely Monophlebidae, Ortheziidae, Coelostomidiidae, Coccidae, Lecanodiaspididae and Diaspididae of the superfamily Coccoidea of scale insects (Table II; Figure 3). There is evidence that flavobacteria have been associated with scale insects since they appeared on Earth, about 150-250 million years ago. Sodalis has been reported in phylogenetically distant insects like the orders Coleoptera (such as weevils), Diptera (such as tsetse fly), Phthiraptera (like lice), and Hemiptera (like bugs, psyllids and spittlebugs) (Dale & Maudlin, 1999; Lefèvre et al., 2004; Fukatsu et al., 2007; Koga & Moran, 2014; Hosokawa et al., 2015; Morrow et al., 2017).
Apparently, scale insects of the Diaspididae family (Gruwell et al., 2007), and several families of the Arqueococcids (Figure 3; Rosenblueth et al., 2012; Dhami et al., 2013) co-speciated with their flavobacterial endosymbionts. In contrast, a high number of host changes have been reported in the enterobacterial co-symbiont. It has even been shown that several individual insects bear two different species of enterobacteria (Rosenblueth et al., 2012). It seems that several insects that lost their flavobacteria and enterobacteria, acquired back a flavobacterium by horizontal transfer from other scale insects, as is the case of several Neococcids. Flavobacteria have been replaced by other endosymbionts in at least three events, for example in species of the Putoidae family (by Spiroplasma and Wolbachia), Pseudococcidae (by Tremblaya) and Dactylopiidae (by Dactylopiibacterium, Spiroplasma and Wolbachia). The Phenacoccinae subfamily of the Pseudococcidae family, after acquiring Tremblaya, apparently lost it and got back a Flavobacterium. Two clades of Phenacoccinae acquired Flavobacterium, one of them from other scale insects and the other (Brownia rhizoecola) from outside scale insects. Brownia shows a greater identity with flavobacteria from unrelated insects such as those of leafhoppers and of cockroaches (Sulcia muelleri or Blattabacterium spp.) than with flavobacteria of scale insects (Gruwell et al., 2010; Rosenblueth et al, 2012.) (Fig. 3).
The endosymbiont bacteria of the Pseudococcidae family have been extensively studied since 1992 (Munson et al., 1992; Thao et al., 2002). They represent an exceptional case in endosymbiosis. Inside the cytoplasm of Tremblaya princeps (Betaproteobacteria) inhabits an enterobacterium close to the genus Sodalis (Husnik & McCutcheon, 2016).
Bacteria of the genus Wolbachia are secondary endosymbionts of over 40% of all arthropods and also of nematodes of the Onchocercidae family (Darby et al., 2012). In arthropods, some wolbachias manipulate the reproduction of their host, either by parthenogenesis, male feminization, male killing or cytoplasmic incompatibility (Werren et al., 2008). Some wolbachias benefit their hosts by providing them with protection against viral infections (Teixeira et al., 2008; Chrostek et al., 2013), or metabolites as riboflavin (Moriyama et al., 2015) or heme (Brownlie et al., 2009). Wolbachia have also been found in several species of scale insects (Figure 3) but it is not yet known whether they have an effect on the reproduction of their hosts.
Our research group has worked with three species of scale insects, Llaveia axin axin (Monophlebidae), Dactylopius opuntiae, and Dactylopius coccus (Dactylopiidae) (Figure 3). We described the bacterial (Ramírez-Puebla et al., 2010, 2016; Rosas-Pérez et al., 2014, 2017) and fungal (Vera-Ponce de León et al., 2016) communities associated with these two Mexican scale insect species. To illustrate the interactions of scale insects with their bacterial symbionts, we review the biology of these scale insects and the possible effects of their microbiota.
Llaveia axin axin (WAX COCHINEAL) AND BACTERIAL SYMBIONTS
Mesoamerican natives called the cochineal Llaveia axin axin “niij” (Fig. 2). Its annual life cycle presents hemimetabolism (gradual changes of egg, nymph and adult, without passing through the pupa stage). Females go through three stages of growth until they reach the adult state. They spend their entire life feeding from their host plant and at the end of their cycle they lay eggs in soil and then die. Under proper environmental conditions, the eggs hatch, nymphs emerge and disperse by moving or being blown by the wind. When they find an adequate plant, they settle on it. Like all scale insects, L. axin axin is exclusively phytophagous, feeding on the sap of plants such as Acacia cochliacantha, Acaciella angustissima, Jatropha curcas and Spondias sp. (Rincón-Rosales & Gutiérrez-Miceli, 2008). This hemipteran is of economic value in southern Mexico and since pre-Hispanic times a wax extracted from it is used to produce a lacquer to coat handicrafts (Williams & MacVean, 1995). It is worth mentioning that the population of niij has declined in recent years, mainly due to overexploitation, deforestation and the burning of the forests they inhabit (Rincón-Rosales & Gutiérrez-Miceli, 2008; Rosas-Pérez et al., 2014). The biology of this insect is little known. We studied its microbiota with metagenomic approaches.
Two endosymbiotic bacteria, a flavobacterium and an enterobacterium were found in the abdominal bacteriomes in L. axin axin (Fig. 2). DNA and RNA obtained from the bacteriomes were sequenced. Metagenomic and metatranscriptomic analyses of L. axin axin showed that both bacteria provide essential amino acids that the insects do not get from their diet. The flavobacterium, now named Walczuchella monophlebidarum, has a very small genome (309 kb) and its genetic repertoire is very similar to that of other phylogenetically distant flavobacteria, indicating a convergence in the evolution of different flavobacterial endosymbionts (Rosas-Pérez et al., 2014). Due to its reduced genetic repertoire, Walczuchella metabolically depends on Sodalis TME1 and their host.
The enterobacterium co-symbiont of L. axin axin, now named Sodalis TME1 (Rosas-Pérez et al., 2017), has a 3.41 Mb genome with high functional diversity. Its large number of mobile elements is indicative of an ongoing genomic reduction process. Sodalis, as other enterobacteria that are co-symbionts of flavobacteria, is considered a newly acquired endosymbiont (Michalik et al., 2014). Sodalis TME1 has genes that may participate in the suppression of defense mechanisms in the insect. Sodalis encodes a Type 3 secretion system (T3SS) and enzymes for polyamine synthesis, which have been recognized as modulators of the defense system of insects (Dale et al., 2001; Jelsbak et al., 2012).
The expression profiles of bacterial and insect genes in the bacteriome and in the ovary showed that Sodalis TME1 may secrete an effector, possibly injected by T3SS, to achieve the colonization of the bacteriome. While Sodalis actively recycles the insect’s waste nitrogen, the insect activates genes for transporters of different types of nutrients and produces an antimicrobial peptide (drosomycin-like) perhaps to keep the density of endosymbionts under control or to facilitate the metabolic exchange between symbiotic partners through membrane permeabilization (Mergaert et al., 2017).
In the ovary, Walczuchella seems to activate a response to oxidative stress, which the insect may use as a defense mechanism. In the ovary, the insect dramatically increases the production of energy for the development of eggs and activates genes encoding lysozyme and peptidoglycan binding proteins, maybe to control bacterial growth.
FUNGI ASSOCIATED WITH THEWAX COCHINEAL L. axin axin
A 500 bp fragments of fungal 18S rRNA and 26S rRNA ribosomal genes were recovered from three metagenomes from the wax cochineal, two from bacteriomes, and a third from a fully macerated insect (Vera-Ponce de León et al., submitted). These sequences allowed the identification of three phyla, Basidiomycota, Chitridiomycota and Ascomycota (Vera-Ponce de León, submitted). In the meta-assemblies of the metagenomes, genes that encode for triglyceride synthesis, purine metabolism and for uricase enzymes of Aspergillus sp. were found. This suggests that fungi associated with L. axin axin may produce lipids or provide nitrogen by recycling molecules such as uric acid and its derivatives, as has also been observed in beetles, grasshoppers and in the carmine cochineal (Sasaki et al., 1996; Nasir & Noda, 2003; Vera-Ponce de León et al., 2016). Genes encoding enzymes like phenol-monooxygenase, involved in the detoxification of xenobiotic compounds, were found, supporting the hypothesis that fungi may help the insect degrade the chemicals produced by the plants for their protection against insects. For example, Jatropha curcas, one plant on which L. axin axin insects feed, produces phorbol esters in high concentrations and various host plants of the niij produce tannins. It was observed that fungi found in the micangio of Dendroctonus beetles are capable of biotransforming toxic terpenoids and turning them into pine pheromones that attract insects of the same species (Brand et al., 1976).
Insects were collected for several years and using a culture-dependent approach, three fungi were isolated in culture. Phylogenetic analysis of 26S rRNA genes identified the three isolates within the genus Aspergillus spp. (Vera-Ponce de León et al., submitted). Aspergillus sequences were also found in the metagenome analyses and clearly, a larger diversity was shown by a culture-independent approach than by a culture-dependent approach. The former revealed several unknown fungi.
DACTYLOPIUS COCCUS (CARMINECOCHINEAL)
D. coccus is known as the carmine cochineal (Fig. 2), and it is of great economic importance because carminic acid (carmine or red E120) is obtained from these insects. Carminic acid is an anthraquinone glycoside that is industrially used as a red dye for food, textiles, and cosmetics (Deveoglu et al., 2011). The genus Dactylopius comprises 11 species: Dactylopius ceylonicus (Green), D. confusus (Cockerell), D. opuntiae (Cockerell), D. bassi, D. coccus (Costa), D. tomentosus (Lamarck), D. austrinus (De Lotto), D. confertus (De Lotto), D. gracilipilus (Van Dam & May), D. salmianus (De Lotto), D. zimmermanni (De Lotto) (Williams & Ben-Dov, 2015). The first six species are found in Mexico and D. coccus is the only one that has been domesticated and depends on human intervention to complete its reproductive cycle. Due to the high concentration and quality of its pigment, D. coccus has been cultivated since pre-Columbian times (Rodríguez et al., 2005). Individuals have a size of 1-6 mm in length. Like other scale insects, their life cycle consists of a hemimetabolous metamorphosis that lasts approximately 110 days. This species exhibits sexual dimorphism. Males are winged and smaller than females (Pérez-Guerra & Kosztarab, 1992). All individuals, males and females, are covered by a cottony secretion wax that protects them against predators or from the environment (Chávez-Moreno et al., 2009). These insects feed on the sap of Opuntiae and Nopalea cacti (Cactaceae: Opuntioidae) (Chávez-Moreno et al., 2009). The sap is composed of water (85-95%), carbohydrates (3-7%) and fiber (1-2%); however, it is poor in protein (0.5-1%) and lipids (0-0.2%) (Stintzing & Carle, 2005).
BACTERIA ASSOCIATED WITH D. COCCUS AND THEIR ROLEIN INSECT METABOLISM
The diversity of bacteria associated with Dactylopius cochineals has been well studied (Pankewitz et al., 2007; Ramírez-Puebla et al., 2010, 2015, 2016) not only in the domesticated species D. coccus but also in wild species. In D. coccus we found Wolbachia, Spiroplasma, Dactylopiibacterium carminicum (a betaproteobacterium), and diverse fungi (Ramírez-Puebla et al., 2010, 2015, 2016; Vera Ponce de León, 2016).
Two Wolbachia strains were found associated with D. coccus that were classified as Wolbachia bourtzisii wDacA and Wolbachia pipientis wDacB after a general proposal of species names in Wolbachia based on phylogenomic analyses (Ramírez-Puebla et al., 2015, 2016). The sizes of the genomes of Wolbachia (Ramírez-Puebla et al., 2016) are similar to those of other wolbachias and the Spiroplasma genome is also within the size range of its group genomes (1 Mb). All genes required for the synthesis of riboflavin and ubiquinone (vitamin Q) were found in the genomes of Wolbachia, suggesting that it provides these vitamins to its host. One of the Wolbachia species, the most abundant, may cause cytoplasmic incompatibility that affects the sex ratio in the progeny.
A bacterial phylogroup (END-1, now called Dactylopiibacterium carminicum) was the only bacterial species found in D. coccus (Ramírez-Puebla et al., 2010). Dactylopiibacterium is related to Uliginosibacterium gangwonense. From the D. coccus metagenomes we were able to reconstruct the entire genome of Dactylopiibacterium. Its characteristics are different to all betaproteobacterial genomes previously described as endosymbionts associated with Hemiptera and other scale insects (Table I; Table II). The size of the Dactylopiibacterium genome (3.6 Mb) is similar to those of free-living bacteria. This suggests that it is a recent symbiont that has not undergone a long process of genome reduction characteristic of other endosymbionts. Furthermore, the functional annotation of this genome has shown that the whole genetic machinery for biological nitrogen fixation (BNF) is present (Vera-Ponce de León et al., 2017). Acetylene reduction assays and analysis of nif gene expression of Dactylopiibacterium showed that BNF occurs in whole D. coccus insects, ovaries and intestines. There is good evidence that the betaproteobacterium Dactylopiibacterium fixes nitrogen in the cochineal and we suppose it alleviates the nitrogen deficiencies in the sap that the cochineal ingests.
In other insects, it has been observed that bacterial symbionts associated with gut, protists or fungal gardens, are able to fix nitrogen and provide it to the host (Ulyshen, 2015; Pinto-Tomás et al., 2009; Morales-Jiménez et al., 2013). In ant fungus gardens, Klebsiella variicola strains are found. These bacteria are not insect-specialized as they thrive in crop plants and hospital patients (Rosenblueth et al., 2004; Martínez et al., 2004).
Furthermore, sequence analysis of clones of bacterial ribosomal genes (16S rRNA) from amplicons obtained by PCR of total DNA extracted from various species of Dactylopius, revealed alphaproteobacteria close to Sphingomonas, Mesorhizobium, and Hepatincola porcellionum. Twelve ribo-clones of D. confusus were related to Acinetobacter (gammaproteobacteria) and betaproteobacteria such as Herbaspirillum and Massilia were also reported (Ramírez-Puebla et al., 2010). Several of these bacteria had been previously reported as plant endophytes (Rosenblueth & Martínez-Romero, 2006) and may colonize the cochineal gut.
FUNGI IN COCHINEAL CARMINEAND THEIR ROLEIN RECYCLING NITROGEN
The study of the fungal communities of D. coccus, D. confusus, and D. opuntiae by a culture-dependent approach revealed a total of 14 different fungal phylogroups associated with three Dactylopius species (Fig. 4). Although there is not a clear pattern between insect species and their fungal phylum groups, we observed that two yeast species Rhodotorula mucilaginosa and Cryptoccocus saitoi were present in females of all three species and in all locations sampled (Fig. 4). Penicillium fungi were detected in males of three Dactylopius species (Vera-Ponce de León et al., 2016). FISH analysis with probes targeting fungal ribosomal genes revealed the presence of C. saitoi in Malpighian tubules and embryos of D. coccus and D. opuntiae. Fungal ribosomal genes of Ascomycota, Basidiomycota, Chitridiomycota and Glomeromycota classes were found by metagenomic analyzes of hemolymph, ovaries, and guts of D. coccus females. Many sequences did not match to any previously reported sequence showing that there are novel fungi well adapted to live inside the carmine cochineals. Fungal genes involved in the bioconversion of uric acid (UA) to urea were identified in the metagenome. It is known that UA is the major waste metabolite from the nitrogen cycle in terrestrial insects. It has been estimated that this compound constitutes about 80% of the product of nitrogen catabolism in these organisms (Pant, 1988). In D. coccus, concentrations between 4 and 34 ng UA/μg -1 tissue, and uricase activity were detected in intestines and eggs. Importantly, no insect uricase gene was found in the metagenomes (Vera-Ponce de León et al., 2016). It was observed that most fungal phylogroups associated with Dactylopius use UA as their sole nitrogen source (Vera-Ponce de León et al., 2016). In tissues of D. opuntiae treated with antifungals to remove fungi, it was observed that UA concentration increased and uricase activity decreased. Furthermore, a significant decrease in the size and weight of treated insects was observed in comparison to control insects (Vera-Ponce de León et al., 2016). These results suggest that symbiotic fungi play an important role in nitrogen supply in Dactylopius. Such is the case in Nilaparvata leafhoppers where yeasts bio-convert UA to amino acids that can be used by their host.
Figure 4
Heat map indicating the presence (light gray) and absence (dark gray) of different species of fungi found with a culture-dependent approach by sequencing ITS of three Dactylopius species (Vera-Ponce de León et al., 2016).
CONCLUSIONS
It is common for many different insects to harbor various co-symbionts (Table I). This also happens in scale insects. There is a consensus that Flavobacteria are among the oldest symbionts, with largely reduced genomes, that provide essential amino acids to the host; but apparently, they frequently have to be metabolically complemented by other bacteria (Wu et al., 2006). Co-symbionts may confer some other additional benefits to the hosts besides helping to provide the ten essential amino acids we metazoans need. This may explain the co-occurrence of Flavobacteria with enterobacteria in many insects including the wax cochineal. However, these symbionts are not found in the carmine cochineal, in which there is a more diverse community. A comparative summary of some of the characteristic of the carmine and wax cochineal symbiosis is presented in Table III. In the carmine cochineal, we studied in particular Dactylopiibacterium that fixes nitrogen to compensate for the carbon-nitrogen imbalance found in cactus sap on which the insect feeds. Dactylopiibacterium may provide essential amino acids to the carmine cochineal as it has complete pathways for the biosynthesis of amino acids. Among the cochineals that we studied, we also detected differences in the composition of their associated fungi (Vera-Ponce de León et al., 2016). Despite the taxonomic differences, there are common functions in fungi from Nilaparvata, Dactylopius, and the wax cochineal L. axin-axin to recycle uric acid.
Table III
Symbiotic relationships, observed in carmin and wax cochineal.
This review shows the enormous plasticity of insect microbiota in different hosts and conditions. This is outstandingly illustrated in the case of Nilaparvata lugens which is the most destructive rice pest that originally could not attack resistant rice plants that lacked asparagine needed by the insect (Shigematsu et al., 1982), but the insects overcame the resistance when carrying yeast-like symbionts that provided amino acids (Ferrater et al., 2013).
