Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact on Quantity and Quality of the Resin Secreted

*Kewal Krishan Sharma and Thamilarasi Kandasamy*

## **Abstract**

Lac insects are a specialized group of phytosuccivorous insects (Coccoidea: Tachardiidae) that secret resin of industrial importance having diverse applications. Due to unique biology, host preference and dispersal mechanisms, lac insects are expected to differentiate locally, forming geographic and host races without adequate morphological differentiation. 101 species of lac-insects and over 400 species of lac host plants have been reported but insects belonging to sub-family Tachardiinae are considered important for *laksha-culture* (lac insect farming). With a wide host-plant range and diverse habitat, the insects have developed a specialized ecosystem with multi-trophic complex of flora and fauna. Not only the lac insect but also the host plants and various biotic associations play a significant role in determining the quantity and quality of the produce. This insect being an obligate phloem sap sucker completes its life cycle on host plant species. Phloem sap is nutritionally unbalanced, as it is rich in carbohydrates but deficient in essential amino acids. Due to the scarcity of essential elements in phloem sap, endosymbionts are likely to co-evolve within the insect cell, while fulfilling their nutritional requirement. Implication of these intricate biotic associations on quantity and quality of the lac resin produced merits thorough understanding for sustained lac production.

**Keywords:** lac insect, *Kerria lacca*, Tachardiidae, insect-plant interaction, resin quantity and quality, lac-endosymbionts

## **1. Introduction**

Lac, reputed as the only resin of animal origin is secreted mainly by the Indian lac insect, *Kerria lacca* (Kerr) (Hemiptera: Tachardiidae), which thrives on the tender twigs of specific trees called lac hosts (*kusum*, *palas*, *ber*, *Flemingia etc*.). Since time immemorial, lac farming has been practised for the products of commerce *viz*., lac, a resinous or non-resinous covering substance over their body, dye – a natural crimson/ yellow color in the body fluid and the lac wax present within and above the lac resin. These products find application in diverse areas such as food, pharmaceuticals, cosmetics, paints and varnish industries [1]. Lac insects (Hemiptera: Coccomorpha) specialized coccids (scale insects) belonging to the family Tachardiidae (=Kerriidae) [2, 3] are sap-sucking insects thriving on certain plant species called lac host plants. Almost all the stages of lac insect are sedentary and attached to the host plants except for neonate nymphs (crawlers) and adult male insects. Lac insects can be found in tropical and subtropical regions (between the latitudes 40<sup>0</sup> N and 40<sup>0</sup> S) due to their preference for a warm climate. Lac production is mainly done in some South, East and Southeast Asian countries like India, Thailand, China, Indonesia, Bangladesh, Myanmar, Laos and Vietnam but the product is in demand all over the world. India is the global leader in lac production followed by Thailand.

The family comprises of nine genera and 99 species worldwide [4]; *Kerria* Targioni Tozzetti is the largest genus in the family. Recently two more species *K. destructor* [5] and *K. canalis* [6] have been described taking the total number of species to 101. The family is characterized by sclerotized features of the adult female. The outer lac encrustation, called lac cell, does not always help in recognition, although it provides indicators in some cases, *e.g*., lac is resinous and alcohol soluble in sub-family Tachardiinae, but hard, horny and insoluble in Tachardiininae.

India is endowed with a rich wealth of lac insect resources. The genus *Kerria* includes 28 species worldwide, 21 of which are recorded from India [7]. 27.7% of lac insect biodiversity reported from the world is found in our country under two genera *i.e*., *Kerria* (23 species) and *Paratachardina* (5 species). Species of *Paratachardina* do not produce true lac and are considered pests of economically important plants but have been utilized as bio-control agents for controlling weeds. Other minor species include *K. chinensis* and *K. sharda*. *K. chinensis* is the principal insect in East and Southeast Asian countries. The *kusmi* form of *Kerria lacca*, known for its superior quality of lac and higher productivity is unique to India.

*Strains of lac insects:* Indian lac insect is known to comprise two distinct infra subspecific forms (commonly termed strains), '*Kusmi'* and '*Rangeeni'* [8]. *Kusmi* strain is grown on *Kusum* or on other alternate host plants and the *kusmi* crops are (i) *Jethwi* – summer season (maturing in June/July) and (ii) *Aghani* – winter season (maturing in Jan./Feb.). *Rangeeni* strain thrives on host plants like *palas* but not on *kusum* and it has also two crops; they are (i) *Katki* – rainy season (maturing in Oct. /Nov.) and (ii) *Baisakhi* – summer season (maturing in June/July).

### **2. Factors affecting lac production**

Production of lac per unit area and time depends on various biotic and abiotic factors impinging upon lac insect ecosystem. Important biotic factors, which affect lac productivity are:

#### **2.1 Lac insects**

Species mainly belonging to *Kerria* are exploited for commercial production of lac. Yield of lac insect varies significantly depending upon the lac insect species and its strain. Differences in lac yield also exist between the two strains. Average resin secreted by *kusmi* cells and *rangeeni* cells is 16.96 mg and 8.07 mg, respectively.

*Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact… DOI: http://dx.doi.org/10.5772/intechopen.106902*

Similarly, *kusmi* strain produced twice the quantity of resin (4.99 mg/mm) per unit size of the cell compared to *rangeeni* strain (2.55 mg/mm) [9]*.*

#### *2.1.1 Initial density of settlement and mortality*

Lac insects are gregarious in nature and settle in close proximity. Hence, density of settlement has a bearing on lac yield. Different lac insects showed a varied density of settlement (average 80192 per sq. cm in *rangeeni,* 186242 per sq. cm in *kusmi* and 160–264 per sq. cm in Meghalaya stock) based on the broodlac quantity used [10]. Under the optimal broodlac conditions, *kusmi* strain tends to settle closer compared to *rangeeni* strain, which is evident from their mean density of settlement. However, under excess brood condition 'crowding effect' is exhibited, which is more marked in *rangeeni.* On the other hand, crawlers of *K. chinensis* (Meghalaya collection) tend to settle closer during both the seasons. Settlement of larvae in variance with the desired number affects the production adversely. Higher density of settlement results either in increased mortality due to insufficient availability of space and nutrition or a higher male population, which ultimately affects lac yield. Mortality up to 21 days of inoculation is attributed to non-feeding of the larvae at the time of initial settlement. Higher initial mortality is indicative of non-suitability of the host plant and/or of unfavorable environmental conditions.

#### *2.1.2 Sex-ratio*

Although the contribution of male lac insects to commercial lac production is very little, they are vital for good lac crop and vis-à-vis broodlac production since the rate of lac secretion increases vigorously in the females after fertilization. However, female lac insects are the sole commercial lac producers. Progeny size and sex ratio vary widely in different lac insect stocks as well as when same lac insect is reared on different host plants. Variation in sex ratio in different crops of lac insect vis-à-vis host plant is more pronounced in *rangeeni* strain than in *kusmi,* especially during the summer season. It varied between 18.07–64.39 and 25.59–31.48%, respectively, for summer and rainy season crops of *rangeeni* and between 22.02–29.33 and 25.49– 35.44%, respectively, for summer and winter season crops of *kusmi* strain [10].

Average male population is lower in smaller progenies and higher in larger progenies; suggesting that larger progeny size increases the male population. In general, sex ratio ranges between 20 and 50% depending on various biotic and abiotic factors. However, in smaller colonies, it may vary between 0 and 100 per cent. Sex ratio is found to vary with (i) season [11]; (ii) sequence of emergence [12]; (iii) site of colonization [12]; iv) density of settlement [13, 14]; v) plant-host [15] and plant-host variety [16]. However, the exact reasons for the wide fluctuation of sex ratio in lac insect population need to be investigated further.

#### **2.2 Host-plant**

Although more than 400 species of plants have been reported to support lac insects on them, various biological attributes such as survival, resin production and fecundity differ greatly from the host species used [17]. Coccoids have been found to be very specific not only to different host species but also to specific varieties and even individual phenotypes of host plants due to inter-specific and intra-specific variation in the host plant defense. Srinivasan [18] has indicated preference of lac insects to

specific phenotypes *kariya* over *charka* the later has a lighter colored bark than the former in *palas* (*Butea monosperma*) and *Kusum* (*Schleichera oleosa*); however, the two are botanically inseparable.

*Food quality:* The quality and quantity of food available to the insect are important in determining its survival and reproduction rate. This is particularly true for phloem sap-feeding insects. Passive exudation of sap of phloem bundles has a substantial role in supply of phloem sap to the insects. Quantitative and qualitative differences in the nutrition available from different host plants, cause variation in biological attributes of the lac insect. Variability studies in case of four species of *Flemingia viz*. *macrophylla*, *semialata*, *stricta* and *bractiata* with regard to various attributes of lac insects have shown significant differences [19].

*Sap condition:* Lac insect feeds on the phloem sap of the host plant. The insect inserts its proboscis and feeds on exudation of sap by i) turgor pressure and (ii) capillary action. Turgor pressure of the host changes with the season and phenotypic activity of the plant. The sap pressure is higher during rainy season and considered favorable for the growth of the insect and inverse is true for summer season. 'Sap reaction' and 'sap density' are possibly among the factors, which influence the suitability of the host plant for lac infection. Good host plants have phloem sap of pH ranging between 5.8 and 6.2 [20]. Similarly, sap density of good hosts ranges between 0.140.1728.

#### *2.2.1 Initial density of settlement and mortality*

Initial settlement of crawlers is affected mainly by host plants and the physical characteristics of the twigs where they settle down. Density of settlement on *S. oleosa* (*kusum*) is higher than on *Albizia lucida* (*Galwang*). Some lac host plants support a certain species or strain in a better way and *vice-versa*. Although lac insect crawlers can be made to settle on any plant twigs, they would be able to survive and complete their life cycle only on good hosts. *Rangeeni* lac insect cannot survive on *kusum* and it exhibits very high mortality on *F. semialata*; similarly, *kusmi* strain cannot survive on *B. monosperma* (*palas*), while *Ziziphus mauritiana* (*ber*) supports both strain up to maturity. There is a significant decrease in survival when lac insect was reared on pumpkin fruits (*Cucurbita moschata*) in comparison to *Flemingia macrophylla*. The increase in mortality observed was 67.2% and 104.9% for *kusmi* and *rangeeni* strains, respectively [21].

#### *2.2.2 Sex-ratio*

Chauhan [15] has reported that the Meghalaya lac insect stock showed significant difference in sex ratio on different host plants. It was observed that 72%, 82% and 98% were in favor of males on *F. macrophyll*a, *Cajanus cajan* and *Z. mauritiana*, respectively. Similarly, Sharma and Ramani [21] have also observed that male percentage of *rangeeni* and *kusmi* strains of *K. lacca* on *F. macrophylla* was 39.76 and 37.28%, which increased to 70.05 and 62.65% when they were reared on *C. moschata* fruits.

#### *2.2.3 Effect of host-plant on resin production*

Average mean cell diameter of a *kusmi* female cell was 3.02, 3.16, 3.50 and 3.54 mm on *C*. *moschata*, *F. macrophylla, Acacia auriculiformis* and *S. oleosa,* respectively [9]*.* Resin produced by individual female lac cells varied significantly. It ranged from

*Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact… DOI: http://dx.doi.org/10.5772/intechopen.106902*

6.11 mg on *C*. *moschata* fruits to 22.84 mg on *S. oleosa.* Resin production by individual female lac insects was the highest on *S. oleosa* followed by *A. auriculiformis, F. macrophylla* and *C*. *moschata* fruits (**Figure 1**). Very high intra-strain variations were observed in resin-producing efficiency of lac insect even when cultured on the same host plant.

Similarly, the average mean diameter of a *rangeeni* female cell grown on *A. auriculiformis, C*. *moschata*, *F. macrophylla* and *B. monosperma* was 3.10, 3.17, 3.19 and 3.22 mm, respectively. Average resin secreted by an individual *rangeeni* female cell ranged between 6.00 mg (on *C*. *moschata* fruits) and 9.09 mg (on *A. auriculiformis*)*.* Resin production by single *rangeeni* female lac insect was found to be the highest on *A. auriculiformis* followed by *B. monosperma, F. macrophylla* and the lowest on *C. moschata* fruits (**Figure 2**). Higher values of coefficient of regression in good hosts *S. oleosa* and *A. auriculiformis* for *kusmi* strain and *B. monosperma* in *rangeeni* strain corroborate the fact that a good lac host allows full manifestation of the resinproducing potential of the lac insect. Resin productivity is higher on tree hosts in comparison to *F. macrophylla* (a shrub) and the pumpkin fruit. Though variations in cell size were less prominent, weight of the cell and resin output per female recorded greater variations showing the effect of host on resin productivity of the insect. Lac insect – host plant interaction in terms of lac production and host suitability is reflected in the data provided in **Table 1**.

#### *2.2.4 Host suitability index*

Host preference, length of settlement at crop maturity, lac insect survival at crop maturity and resin production by the insect are the most important attributes affecting the production of lac. Host Suitability Index is calculated for identifying an ideal host plant by using the following formula. By taking the lowest value of Host Suitability Index for a particular host plant as 1.00, relative suitability indices are calculated for the other hosts.

#### **Figure 1.**

*Resin productivity and cell size relationship in* Kerria lacca *(*kusmi *strain) on different host plants during winter season crop.*

#### **Figure 2.**

*Resin productivity and cell size relationship in* Kerria lacca *(*rangeeni *strain) on different host plants during rainy season crop.*


**Table 1.**

*Host suitability index of different lac host plants for rearing Indian lac insect,* Kerria lacca *(Kerr).*

## Host Suitability ¼ Index

%Host Preference � %Length of settlement � %lac insect survival �Mean resin production mg ð Þ� 100

where % host preference � (% of host-plants on which lac insect survived till crop maturity), % length of settlement at crop maturity � (% length of encrustation of available shoot length), % survival of lac insect (per square cm) at crop maturity �

(% surviving lac insect number of the initial density of settlement) and resin produced (mg) by lac insect (average weight of resin produced by fifty randomly collected individual female lac insects).

#### **2.3 Local environment**

Abiotic and biotic components also affect the suitability of the host plant to lac insect. *Ziziphus xylopyra* (*Ghont*) is a good host in Madhya Pradesh but not in Ranchi (India). Similarly, *C. cajan* and *Grewia* spp. are used as lac hosts in North Eastern states but attempts to cultivate lac *Grewia* at Ranchi proved futile. Analysis of weather data of 19842012 of Ranchi (India) revealed that the winter months (December and January) have become colder and pre and post-winter months (November and February) the warmer [22]. These changes in climatic parameters have implications in lac cultivation as it is a critical period of lac insect development (pre-sexual maturity) during the summer season crop. Monsoon and winter rainfall spells and magnitude were also found to affect lac crop performance. Effect of abiotic factors (temperature, rainfall and relative humidity) was correlated with lac production of *rangeeni* crop during 2006–2007 to 2012–2013. It was observed that maximum temperature had a significantly negative (0.911\* and 0.837\*) and RH a positive significant (0.850\* and 0.800\*) correlation with lac production during the critical crop period (March and April) of development in the summer season (*baisakhi*) crop whereas, during rainy season (*katki)* crop, minimum temperature had a significant negative (0.765\*) correlation with lac production. The vulnerability level of lac insect is high during and prior to sexual maturity stage in the summer crop, thus post-winter season is the critical period for lac insect survival and any undesired variability in weather parameters in this stage can impact adversely on lac productivity.

## **3. Factors affecting lac quality**

Lac insects have been designated as crimson, yellow, cream or white depending upon the quality and intensity of the water-soluble pigment (laccaic acids) present in their body. Resin pigment (erythrolaccin) is not soluble in water and is not desirable in some of its application areas. Quality of the secreted resin due to the presence of dye differed significantly among various stocks of lac insects. *Kusmi* strain of *K. lacca* produced the lightest colored resin whereas that produced by *K. chinensis* and *K. sharda* insects was the darkest. Saha [23] has also reported significant differences in physicochemical properties of lac resin secreted by three different lac insect species. Lac insect species and their host plants also affect the quantity of lac dye (erythrolaccin) present in the particular lac insect stock.

Biochemical parameters of the lac host plants get altered after lac insect inoculation. Fengshu *et al.* [24] have analyzed phenolics, sugars, amino acids and some inorganic elements of the host plants of different ages and strains with and without lac inoculation. The biochemical parameters showed variations after lac inoculation and correlated with the seedlac quality and quantity in different seasons. Liu *et al.* [25] have analyzed tannins and phenolics in relation to the quality and size of the winter generation *K. lacca* on different lac hosts such as *Dalbergia szemaoensis, D. hupeana, D. obtusifolia, Pueraria thunbergiana, Zenia insignis, Cassia siamea* and *C. cajan*. Yang *et al.,* [26] have reported that the host tree root secretion and the quality of the lac produced are the indexes to the inter-adaptability between the lac insect and its host

tree such as *Acacia suma, Hibiscus syriacus, Moghania macrophylla and S. oleosa*. Various studies indicated a decrease in the amount yet increase in the variety of amino acids in the host trees on lac inoculation. Lac composition also differed significantly from the lac insects grown on different host trees.

### **4. Associated fauna**

A number of major and minor pests/diseases at times almost destroy the lac crop, thereby, not only reducing the yield drastically but also affecting the quality of the lac. Sessile nature of lac insects makes them more prone to predators and parasitoids.

#### **4.1 Predators**

The losses in lac cultivation due to various insect predators are known to be far greater than what is usually met in other agricultural crops. About 22 predators have been reported to be closely associated with lac insects, of which three are major predators *viz*. *Eublemma amabilis*, Moore (Noctuidae); *Pseudohypatopa pulverea* Meyr. (Blastobasidae) and *Chrysopa* spp. (Chrysopidae)*. E. amabilis* and *P. pulverea* alone are responsible for 3040% of damage to the standing crop [27, 28] of which *E. amabilis* alone causes 2025% damage [29]. These lepidopterous predators cut a hole in the lac and feed on the insect from inside by making a tunnel. *Chrysopa*, though a sporadic pest, sometimes causes havoc, particularly in *kusmi* strain.

Chemical communication between the lac insect-associated products and the lac predators have been evaluated under the laboratory condition and semio-chemicals identified for different stages of lac insect and its associated products [30]. Six major compounds *viz*. decane, dodecane, tetradecane, heptadecane, eicosane and octacosyl acetate constituted 78%, 79% and 85% in lac insect whole body, lac resin and lac wax extracts, respectively. Electroantennogram (EAG) revealed higher responses of adult male and female of lac insect predator, *Eublemma amabilis* to lac insect whole body extract than resin, wax, crawler and lac insect female extracts. Whereas, in *Pseudohypatopa pulverea,* EAG response was significantly higher in females towards lac insect whole body than resin, wax, adult female and crawler extracts than males. Both *E. amabilis* and *P. pulverea* exhibited high level of sensitivity to lac insect whole body extracts with different concentrations ranging from 1000 to 10,000 ppm than the identified semiochemicals *viz*., Decane, Hexadecane, Nonadecane, Eicosane and hexane.

#### **4.2 Parasites**

Thirty different parasites of lac insect have been reported by Varshney [31]. They lay eggs into the lac cell through the anal tubercle in/on the body of lac insect. The grub that hatches feeds only on lac insect.

#### *4.2.1 Inimical parasites*

Of all the parasites associated with lac insect, eight parasites namely, *Coccophagus tschirchii*, *Erencyrtus dewitzi*, *Eupelmus tachardiae*, *Parechthrodryinus clavicornis*, *Tachardiaephagus tachardiae*, *Marietta javensis*,*T. somervillei* and *A. purpureus* are of regular occurrence in the lac ecosystem. Among these,*Tachardiaephagus tachardiae*

*Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact… DOI: http://dx.doi.org/10.5772/intechopen.106902*

and *Aprostocetus purpureus* are the most abundant lac associated parasites. Extent of parasitisation varied between 15.5% in summer season (*baisakhi*) crop and 18.6% in rainy season (*katki*) crop of *rangeeni* strain. While for the *kusmi* strain it was 19.04% in winter season (*aghani*) crop and 22.8% in summer season (*jethwi*) crop [10]. Fecundity of lac insect is adversely affected by parasitisation. Parasitized cells adversely affect the resin production and brood value (fecundity) of the crop. Either, there is no emergence or very low emergence of young ones from the parasitized cells, which ultimately affects the inoculation of the next crop.

*Parasitic losses:* Percentage of parasitism recorded is much higher than the earlier reports of average 4.8 9.9 per cent parasitism based on seven years' data [29]. Reinterpretation of the same data by Srivastava and Chauhan [32], however, revealed that average per cent parasitism for the crop on the basis of females alone worked out to be 20 to 37. In certain years and in some localities, it was as high as 50%. Jaiswal and Saha [33] have found a positive and significant correlation between density of lac insects and number of parasitoids. So it is highly likely that actual per cent parasitism would be higher than recorded since lac insects generally form a continuous encrustation and the present study was confined only to isolated female lac insects.

*Rangeeni* strain is more vulnerable to pest attack than *kusmi* and damage is more in rainy season crop. Parasites lay eggs into the lac cell through the anal tubercle in or on the body of the lac insect. The grub that hatches feeds only on lac insect and not on lac. As a result of parasitisation, fecundity and resin-producing capability *Kerria lacca* is adversely affected. Quality of the resin produced declined by 17.92% and 17.44% while fecundity decreased by 32.55% and 34.71% for *kusmi* and *rangeeni* strains, respectively [9, 34]. Damage caused by parasites varies depending upon the virulence of outbreak and stage of development of the lac insect at which damage is inflicted. Instances of super-parasitism (as many as 19 larvae have been reported from a single mature female lac insect) and multi-parasitism are not uncommon, which further aggravate the problem.

It was found that the decrease in fecundity of lac insect due to parasitisation ranged between 10100%. Thus, proportionately more broodlac would be required as compared to healthy broodlac for inoculating the same number of trees. If parasitized at an early stage, the lac insect is practically eaten up by the developing parasitoid rendering the lac useless for broodlac purposes. Moreover, the broodlac harboring parasites if used for raising next crop would serve as a source of infection to new lac culture.

Lac insect is gregarious in nature and resin secreted by these coalesces to form a continuous encrustation. The thickness of lac encrustation is one of the criteria for assessing quality of broodlac. Size of parasitized as well as healthy cells did not differ much but comparison of weight of the resin secreted by the two revealed that the amount of the resin produced by the parasitized cells was significantly lower. Hence, visual assessment of broodlac quality on the basis of encrustation thickness alone may prove to be deceptive unless weight is also taken into account.

#### *4.2.2 Beneficial parasites*

Several types of insects are hyper-parasitic to the lac insect. Though their natural population constitutes only about 410% of the total fauna associated with lac insect, they act as a bio-control agent in controlling the damage done by inimical insects. Many ants and other insect species feed on the honey dew excreted by lac insect, and these prevent losses by fungus infection.

#### *4.2.3 Effect of fungi on lac production*

In addition to the damage caused by insect pests, lac crop yield suffers significant losses due to other biotic agents, particularly fungi. Few earlier reports suggested that the lac insects had mutualistic relationship with fungi. However, an association of fungi with lac insects is not beneficial always. Lac insects being phloem feeders excrete excess sugar in the form of honey dew, which invites sooty mold to grow over the lac encrustation. Besides, this rainy season crop is also prone to fungal infection when grown on *ber* and *kusum* due to their shady nature. Avoidable losses due to fungi alone were observed to be 40.9% to 59.85 in the *kusmi* strain of lac insect [35]. Similarly, Mishra *et al.* [36] have reported significant reduction (75.05%88.41%) in mortality of second instar lac nymphs with application of different fungicides on the *kusmi* strain of lac insect.

#### *4.2.3.1 Fungi associated with lac insects*

The earliest record of honeydew that drips from colonies of lac insects on the twigs of host trees inviting black mold species of *Capnodium* and *Fumago* is that of Lindsay and Harlow [37]; the presence of pathogenic fungi, *pythium* sp. in female tests causes a heavy mortality in the larvae which fail to enclose satisafactorily and lie dead in clusters within the female resinous cell [38]; sooty mold fungi *Conidiocarpus* (Syn. *Podoxyphium conidioxyphium*) and *Polychaeton* spp. are obligate anaerobes capable of producing endospores and causing 3040% damage to lac insect in Vietnam [39]; 11 species of saprophytic pathogenic fungi causing dark mildew on lac insect have been reported from China [40] and Three species of fungi belonging to family *Eurotiaceae*/ *Aspergillaceae* causing severe damage to lac culture have been reported from India [41]; *Aspergillus awamori* Nakazawa forms black sheet-like covering on lac encrustation, *Aspergillus terricola* Marchal traverse the whole length of the anal tubercle and blocking it leading to disruption of mating and larval emergence and *Penicillium citrinum* Thom (Syn. *Penicillium aurifluum* biourage) blocks the breathing pores of lac insects. Fungal infection in lac cultures causes losses in lac yield by (i) inhibiting respiration, (ii) hindering mating process, (iii) blocking larval emergence and (iv) affecting lac host efficiency.

### **5. Interaction of lac insects with microbes**

Insects and microbes' interaction ranges from obligate mutualism to facultative parasitism. Insects harbor symbiotic bacteria on the integument, in the digestive tract and in some unique structures within their body [42, 43]. Interaction between insects and microbes is one of the important factors, which makes insects the most successful group of organisms on the earth. Insect microbiota plays significant role in growth, development, reproduction and adaptation of the insects to the environment. Endosymbionts assist insects in their survival by nutritional supplementation [44], aiding in digestion of recalcitrant food materials [45], protecting them from predators and parasitoids [46], detoxifying phytotoxins and pesticides [47], imparting resistance against insecticides [48], immune system stimulation [49], inter and intraspecific communication [50] etc.

Symbiosis with bacterial community is obligatory in insects whose diet is imbalanced such as vertebrate blood (by mosquitoes), phloem sap (by sap-sucking insects) and wood (by termites). Lac insect life stages are morphologically and physiologically *Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact… DOI: http://dx.doi.org/10.5772/intechopen.106902*

highly diverse. Besides, their resin production potential also varies with different stages; crawlers and adult males do not secrete resin whereas, settlers and adult females after fertilization secrete resin albeit in different quantities. Lac insects are almost sedentary throughout their life and depend on nutritionally imbalanced sugarrich phloem sap for their survival. It is postulated that lac insects must harbor myriad of endosymbionts for nutrition supplementation, host plant adaptability, defense etc. Earlier works have found that the presence of microbial flora in lac insects was beneficial during rainy season crops for higher lac yield. Some bacteria such as *Micrococcus* spp., *Clostridium* sp. and *Bacillus subtilis* have been reported from lac insects [51]. *Micrococcus varians* and *Micrococcus conglomerates* are associated with various stages of lac insect and are considered beneficial for good yield of lac production.

Till now, bacterial flora associated with lac insects has been identified either based on culturing method [52, 53] or PCR method for specific endosymbionts like *Wolbachia* [54]. Gender-specific bacterial flora has been identified from lac insects [53], and host plant-induced variation was observed in the bacterial composition of lac insects based on culturing method [52].

#### **5.1 Association with** *Wolbachia*

*Wolbachia* are members of the order Rickettsiales, a diverse group of intracellular bacteria that include species having parasitic, mutualistic and commensal relationships with their hosts. *Wolbachia* species are well known for their vast abundance, effect on hosts in terms of reproductive manipulation and mutualism and have potential applications in pest and disease vector control [55]. *Wolbachia pipientis* is the type species of *Wolbachia* genus. Based on the 16S ribosomal sequence and other sequence information, *Wolbachia* spp. have been divided into seventeen different supergroups (A-Q). Two supergroups (C and D) are commonly found in filarial nematodes, whereas other groups are found in arthropods, in which A and B are the most common. *Wolbachia* that participate in symbiotic relationships with arthropods have a range of phenotypic effects on their hosts and generally behave as reproductive parasites. *Wolbachia* manipulate host reproduction through cytoplasmic incompatibility, parthenogenesis, feminization and male-killing [56–58]. Mostly *Wolbachia* undergoes vertical transmission from mother to offspring. However, a horizontal transfer is also reported in nature [58].

In lac insect populations, there is a wide variation of male–female sex ratio, which ultimately affects the lac production as only females can produce lac. Vashishtha and co-workers [54] have found that lac insects are associated with *Wolbachia* based on 16S rDNA and wsp (*Wolbachia* cell surface protein) PCRs. Lac insect-associated *Wolbachia* was termed as wKerlac. Phylogenetic tree revealed it to be a subgroup "ori" of supergroup B, which is predominantly present in arthropods. *Wolbachia* of *K. lacca* was grouped with *Wolbachia* of *Tagosodes orizicolus* and *Ephestia cautella*. *Wolbachia* on both these hosts are responsible for cytoplasmic incompatibility. Further investigations are required on whether the identified *Wolbachia* would have any role in feminization. It is one of the most important factors in attributing lac yield because commercial lac is obtained solely from female lac insects.

#### **5.2 Detection of** *Wolbachia* **phage (WO) in lac insects**

*Wolbachia* species also harbor a bacteriophage called bacteriophage WO or phage WO [59]. Comparative sequence analyses of bacteriophage WO revealed the

possibility of large-scale horizontal gene transfer between *Wolbachia* coinfections in the same host [60]. Molecular mechanism used by *Wolbachia* to manipulate its host in terms of cytoplasmic incompatibility, feminization, parthenogenesis, male killing *etc*. remain elusive and has been speculated due to genes on extrachromosomal factors such as plasmids or bacteriophages [61, 62]. Out of seventeen identified super-groups of *Wolbachia*, named A–Q, WO phage has been reported to infect *Wolbachia* belonging to super-group A, B, F and G [63, 64].

Screening and distribution of *Wolbachia* and WO phage sequences were studied by amplifying and sequencing the partial *ftsZ*, a cell cycle gene involved in cell division and a putative minor capsid protein, *orf* 7, respectively [65]. Two different lines *kusmi* and *rangeeni* each were found to be singly infected by *Wolbachia* belonging to Supergroup B. It was the first report on molecular detection of WO-phage infecting *kusmi* and *rangeeni* infrasubspecific forms of *K. lacca*. Further phylogenetic analysis revealed distinct differentiation of WO between *kusmi* and *rangeeni* infrasubspecific forms. In the phylogenetic tree made based on *orf7* sequences, *rangeeni* and *kusmi* forms clustered with group III and group I, respectively. Although there is a differentiation of *kusmi* and *rangeeni* forms based on *orf7* of WO sequences, the tripartite association of lac insect-*Wolbachia*-WO needs further investigation to implicate their role in such differentiation.

#### **5.3 Association with yeast-like symbionts**

Insects not only possess bacterial symbionts but also yeast-like fungal symbionts (YLS). Although microbiologists observed such yeast-like endosymbionts in 1960s, the identity was not known due to their fastidious nature and the lack of molecular tools at that time. Later, it was found that the phloem sap feeders harbor obligate intracellular yeast-like symbionts, YLS (subphylum Ascomycota, class Pyrenomycetes, family Clavicipitaceae) [66, 67] especially in the mycetocytes formed by fat body cells of abdomen. YLS have also been reported in Hemiptera (aphids, planthoppers, and scale insects) and Coleoptera (beetles) [68, 69]. They grow by budding and are vertically transmitted to the next generation by transovarial infection [70, 71]. YLS appears to play roles in nitrogen metabolism of the host through recycling of uric acid [72, 73], in insect metabolism by synthesizing sterols, the precursor molecule for many hormones (*e.g*., 20-OH ecdysone-a molting hormone), as insects, in general, are unable to synthesize them [74] and in detoxifying the toxic substances to the host [75]. Occurrence of YLS is highly essential for the survival and reproduction of the host insects because they play vital role in development, reproduction and embryonic development [76]. Transmission electron microscopy and PCR-based studies revealed the presence of YLS in lac insects. However, acquisition of YLS in lac insects seems to be different from that of aphids and plant hoppers and a horizontal transfer was also suggested for them [54].

#### **5.4 Sex-specific endosymbionts**

Male and female lac insects'specific bacterial species were identified by Shamim and co-workers [53] based on 16S rDNA PCR and biochemical characterization. Eight different bacterial species were isolated and categorized as endosymbiont, gut bacterium or subsurface bacterium. Three of them were exclusive to males, three to females and two were common to both the sexes (**Table 2**). *Bacillus megaterium*, *A. subterraneus* and *Pantoea ananatis* were found to be the most abundant bacterial

*Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact… DOI: http://dx.doi.org/10.5772/intechopen.106902*

species. Among 13 bacterial isolates found in males, two were present as an internal gut bacterium which may get excreted out with honey dew. Single isolate of *Paenibacillus barengoltzii* was found at subsurface. *P. ananatis* was exclusively and majorly found in males as endosymbiont bacteria. *P. fulva* was also found in males as endosymbiont bacteria. Twelve bacterial species were isolated from the females and 50% populated with *Bacillus* sp. *B. megaterium, Curtobacterium citreum* and *A. subterraneus* were majorly reported in non-crushed samples, therefore, it was thought to be thriving at subsurface but *B. cereus* and *Solibacillus silvestris* were considered as endosymbiont. *A. subterraneus,* found in both the sexes, as endosymbiont in males and at subsurface in females.

Out of these bacterial species, *B. cereus, B. megaterium* and *P. ananatis*, are widespread in occurrence and have also been reported in other insect's body. Some strains of *P. ananatis,* also referred as "ice nucleation-active" bacteria, are used in pest control because when present in the insect gut, they lower the cold resistance [77]. Owing to this property and bacterial abundance in the insect, absence of lac insects in colder regions could be attributed to the association of *P. ananatis*. Description about the bacterial species identified from both the sexes of lac insect [53] is given in **Table 2**.

#### **5.5 Host plant-induced variation of endosymbionts**

Since phloem sap constituents vary for host plants, variation in the endosymbionts of lac insects growing on different host plants is anticipated. Culture-based method was followed to isolate bacteria from lac insects grown on different host plants and 16S rDNA PCR based molecular method was followed [52] to identify them.

From 29 different bacterial isolates, 10 different bacteria were identified. *Bacillus kochii, Bacillus oceanisediminis*, *Bacillus amyloliquefaciens*, *Bacillus nakamurai* and *Enterobacter cloacae* were observed on *kusmi* lac insects collected from *Kusum* trees. *Klebsiella quasipneumoniae subsp. similipneumoniae, Citrobacter amalonaticus*, P*rovidencia vermicola* and *B. nakamurai* were found in bacteria isolated from lac insects collected from *ber* trees*. Enterobacter ludwigii, Enterobacter cancerogenus* and *B. nakamurai* were found in lac insects collected from *semialata*. In most of the cases, different species of *Bacillus* and *Enterobacter* were found. *Bacillus* is a very common genus found in different types of insects, which include *B. subtilis, Bacillus thuringiensis, Bacillus cereus, Bacillus sphaericus, Bacillus popillae, Bacillus circulans, B. megaterium, Bacillus lentimorbus a*nd *Bacillus polymyxa* [78]. Bacteria such as *Enterobacter* spp. *K. quasipneumoniae*, *C. amalonaticus*, *P. vermicola* belong to Enterobacteriaceae family. All these Enterobacters might have come from lac insect gut and belong to proteobacteria, primarily within the γ–subdivision.

*B. nakamurai* the most frequent bacteria identified from lac insects grown on three different hosts might be involved in some vital functions in lac insects. *B. nakamurai* was originally isolated from soil and known to produce black pigment [79].


#### **Table 2.**

*Bacterial species identified from lac insects.*

*B. amyloliquefaciens* is known to control plant pathogens due to its antifungal activity [80]. Hence, antifungal activity may be anticipated for the *B. amyloliquefaciens* strain present in lac insects.

## **6. Conclusion**

#### **6.1 Impact on quantity and quality**

Lac insects feed on host plant phloem sap by passive sucking mechanism through capillary action and turgor pressure of the sap [81]. Passive exudation of phloem sap through the intruded lac insect stylet as a function of phloem turgor pressure has a significant role in the supply of phloem sap to lac insects. Lac insects are sedentary soon after settlement on host plants. As a result, there is no escape mechanism for lac insects from the host plant's defensive chemicals. Hence, lac insects should have ability to detoxify the host plant's defensive chemicals and avoid them which is important for good lac insect host plant interaction. Due to differences in the ability of lac insects to detoxify the defensive chemicals and the nature of defensive chemicals produced by the plants, there is difference in yield of lac on different lac host plant species and different genotypes of the same host plant. Lesser quantities of toxicant ingested when feeding on phloem sap of good host plants rich in nutrients would be detoxified quickly, whereas the larger quantities of toxicant ingested when feeding on poor quality host plant may not be detoxified by lac insects. Haque (1984) observed variations in the quality and quantity of amino acids present in honey dew (anal fluid) of *K. lacca* grown on different host plants such as *Moghania* (=*Flemingia) macrophylla, Ficus glomerata*, *Ficus indica* and *Ficus religiosa* [82], indicating the variations in nutritional composition of these host plants.

Due to sub-optimal nitrogen/carbon ratio in the phloem sap, phloem feeders need to ingest excess phloem sap along with excess organic carbon to obtain sufficient nitrogen for their growth. Non-optimal ratios of essentially required nutrients in the phloem sap warrant the insects to ingest supra optimal quantities of less required nutrients and also the defensive chemicals, which are toxic to them. These factors affect the resin-producing efficiency of lac insects. Resin production is found to be high on tree hosts in comparison with bushy hosts such as *F. macrophylla* and pumpkin fruit. Auclair [83] has reported that the phloem sap exudation rate through excised stylets in aphids is about an order of magnitude higher in woody plants compared to that in herbaceous plants. Since lac insect feeding habit is also similar to aphids and passive in nature, the higher turgor pressure of woody plants leads to greater food ingestion and higher resin production on tree hosts and vice versa in herbaceous hosts. Intraspecific variation in the host plant defense caused the deviation in the susceptibility of a host plant species in *Nuculaspis californica* [84]. Lac insects have the ability to manifest their biological parameters very specifically not only to lac host species but also to varieties, phenotypes of host plants [18, 85] locality and season of cultivation. Periodic host-plant resistance is considered to be a physiological response to meteorological and edaphic conditions, a response usually rendering the plant temporarily unsuitable to coccoid development.

It can be concluded that lac insects, as well as lac-host plants and associated flora and fauna, play significant roles in the quantity and quality of the lac production. The naturally existing high degree variability in lac insects and host plants can be successfully exploited for selection and evolution of high-yielding varieties and lac

insect-host combinations. Quantity and quality of the lac resin can be significantly enhanced with better management of lac insects and their host plants.

#### **6.2 Interaction with microbes**

Crawlers *i.e*., early developmental stage of lac insect consists more of unknown and other bacterial types followed by *Wolbachia* and *Mucilaginibacter*. *Wolbachia* and *Pantoea* are the two important genera found in the adult female lac insects besides unknown bacteria. *P. ananatis* is already reported to be present exclusively in male lac insects based on the culture method [53]; *Pantoea cypripedii* and *Pantoea dispersa* are present in honeydew secreted by lac insects [86]. In insects, *Pantoea* is primary endosymbiont and its association with insects is mostly mutualistic and sometimes commensalistic. In the mutualistic association, insects provide habitat and nutrition to *Pantoea*, whereas *Pantoea* may help insects by hydrolysis of proteins, antagonism of pathogens, breakdown of toxic substances, nitrogen fixation, nutrition and digestion [87]. Since *Pantoea carbekii* genome encodes complete or near-complete canonical pathways for the production of several vitamins and cofactors such as folate, riboflavin, pyridoxal-5<sup>0</sup> -phosphate, glutathione, iron–sulfur clusters and lipoate, it is assumed that *Pantoea* supplements nutrition by providing essential vitamins and minerals to its host stink bug [88]. Taking these things into account, it is plausible to assume that the *Pantoea* spp. present in lac insects may be involved in nutrition supplementation because plant phloem sap on which lac insects feed is not a nutritionally balanced diet.

*Wolbachia* is an obligate endosymbiont of arthropods and nematodes and present in most of the insects wherein they play a major role in the reproductive manipulation of the host. They alter the reproduction of the host insects by the way of male killing, parthenogenesis, cytoplasmic incompatibility and feminization. Generally, the mode of transfer of *Wolbachia* in arthropods and nematodes from one generation to another is transovarial [89]. Due to such reproductive manipulation of the host by *Wolbachia*, the frequency of *Wolbachia* infected females increases in population sometimes at the expense of host fitness [90]. Vashishtha and co-workers [54] have reported the presence of *Wolbachia* in lac insects by 16S rDNA and *wsp* PCRs. *Wolbachia* infection is known to be biased based on sex or may increase as the development progress. Frequency of occurrence of *Wolbachia* was found to be more in female insects compared to crawlers in the current study. Similar results of higher *Wolbachia* incidence in the adult stage compared to an immature stage were obtained in several other insects. Besides manipulating host reproduction, *Wolbachia* may affect host fitness positively by nutrient supplementation. It has been demonstrated in bed bugs, *Cimex lectularius* that riboflavin provision ability of *Wolbachia* can positively impact the host's growth, survival and reproduction [91]. In the scale insect, *Dactylopius coccus*, two species of *Wolbachia* were found to have metabolic capabilities for riboflavin and heme biosynthesis [92]. Besides, reproductive manipulation and nutrient supplementation, an additional function of protecting *Drosophila* from virus induced mortality was attributed by *Wolbachia* infection [93].

As far as lac insects are concerned, females are the productive gender as the commercial lac is obtained only from females but not from male insects. Since, *Wolbachia* can eliminate males, turn them into females, sterilize uninfected females or behave as a mutualistic symbiont [94], their role in lac production needs to be explored thoroughly. Whether the role of *Wolbachia* is restricted up to reproductive manipulation or it is extended to nutrient supplementation and virus protection in lac insects needs thorough investigation in future.

Lac insect endosymbionts are very diverse and supposed to carry out various vital functions in the insects. The available literature describe mainly cultivable bacteria and to certain extent uncultivable microbes. Much more uncultivable bacteria and also stage-dependent and strain-dependent endosymbionts may be anticipated to be present in lac insects. Culture-independent methods such as metagenomics would reveal more number of endosymbionts in lac insects. Different functions such as nutrition supplementation, sex differentiation, strain differentiation and protection from pests and predators are anticipated for the lac insect endosymbionts.

## **Author details**

Kewal Krishan Sharma\* and Thamilarasi Kandasamy ICAR-Indian Institute of Natural Resins and Gums, Namkum, Ranchi, Jharkhand, India

\*Address all correspondence to: kewalkks@gmail.com

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

*Interaction Among the Multi-Trophic Lac Insect Complex of Flora and Fauna: Impact… DOI: http://dx.doi.org/10.5772/intechopen.106902*

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## **Chapter 5**

## The Ability of Insects to Degrade Complex Synthetic Polymers

*Biswarup Mitra and Amlan Das*

## **Abstract**

Insects while feeding, encounter a wide array of hydrocarbon polymers in their diet and the digestive tracts of various insects contain microbial symbionts that aid in the degradation of these polymers. Thus the idea of insects as synthetic polymer bio-degraders was established. Soon various insect, like mealworms, flour beetles, weevils, wax moths etc. particularly from the Coleopteran and Lepidopteran orders, were identified to have remarkable abilities to consume and degrade a wide range of synthetic polymers like polyethylene, polyurethane, polypropylene, polystyrene and polyvinyl chloride into lower molecular weight, simple, and nontoxic molecules which are eventually excreted as fecula. In this review we aim at congregating the diversity of polymer degrading insect fauna and understanding the underlying mechanism in which the insect's digestive enzymes works in synergy with the gut microbiota to digest complex synthetic polymers.

**Keywords:** synthetic polymers, insects, gut microbiota, enzymes, degradation

## **1. Introduction**

The vast majority of eukaryotic biodiversity in terrestrial ecosystems is represented by insects [1]. While eating, insects come into contact with a wide range of hydrocarbon polymers, and the intestinal tracts of some insects contain microbial symbionts that aid in the decomposition of these polymers. Thus, the concept of the insect as a biodegrading organism for synthetic polymers was developed. Various insects of the Coleopteran and Lepidopteran orders have been observed to have remarkable abilities to consume and degrade a wide range of synthetic polymers such as polyethylene (PE), polyurethane (PU), polypropylene (PP), polystyrene (PS), and polyvinyl chloride (PVC) into lower molecular weight, simpler, and nontoxic molecules that are eventually excreted as fecula.

Although microbial biodegradation appears sustainable, it has limits; and compared to plastic trash generation, its efficiency is modest. Furthermore, since biodegradation of a single polymer is usually a complicated process involving numerous enzymes, microbial consortia rather than a single species or strain biodegrade diverse natural and even synthesised polymers. As a result, a microbial assemblage will likely provide a more efficient biodegradation rate [2]. To overcome these limits, there was a need for a niche that would make plastic trash more accessible and bio-available to a dynamic microbial consortium. Recent research has shown that the digestive

tracts of some invertebrates, notably insects, have microbial symbionts that help in the decomposition of various natural polymers that have similar structural arrangements to synthetic polymers [2, 3]. Therefore, the insect gut microbiome offered an efficient alternative for fast plastic degradation, and plastic degrading bacteria operating in concert with gut enzymes revealed increased breakdown inside the gut microbiome.

A better understanding of the function that the insect gut microbiome plays in the breakdown of plastic may be attained by actively force-feeding insects with different antibiotics and examining the variance in the molecular weight of the provided plastic feed between the insect culture with antibiotic suppressed gut microorganisms and the control insect culture without antibiotic treatment. This will allow for the acquisition of a better knowledge of the role that the insect gut microbiome plays in the degradation of plastic.

## **2. Insect's gut anatomy and the path to plastivory**

Even though insects digest a wide range of foods, their digestive systems are largely the same. The adaptation of their diverse feeding guilds is primarily responsible for changes in their digestive tracts. The digestive tract of an insect can be structurally segmented into foregut, midgut, and hindgut. The foregut and hindgut can be divided into separate sections, each of which corresponds to a specific function. For instance, the foregut of insects is divided into pharynx and oesophagus and has a crop or diverticula for temporary food storage in addition to proventriculus for food grinding. The hindgut is separated into various regions, which include fermentation chambers and a separate rectum for retaining faeces before discharge. However, in many insects, the midgut serves as the main organ for digestion and absorption of ingested food materials [4]. Although it lacks an exoskeletal lining, the insect gut has a unique embryonic origin, having originated from endodermal cells. The peritrophic matrix serves as a protective lining for the epithelial cells lining the midgut of many insects. The peritrophic matrix divides the midgut into endo- and ectoperitrophic spaces, preventing microorganisms and abrasive food from coming into direct contact with the midgut epithelium thus preventing it from injury, pathogen infection etc. The peritrophic matrix also deactivates ingested toxins and pollutants such as pesticides and other inorganic or metal elements [5]. Furthermore, this matrix increases digestion efficiency by compartmentalising the digestion process and selectively transporting solutes and enzymes between the ectoperitrophic and endoperitrophic spaces. The peritrophic matrix further increases digestion efficiency by generating a countercurrent flow between the endo and ecto—peritrophic spaces, favouring nutrient absorption and minimising digestive enzyme loss by frass excretion [4, 6].

In this above described structure of the insect gut, resides a consortia of microorganisms which include protists, fungi, archaea, and bacteria. Fungi are common in the guts of insects that consume wood or detritus and are thought to aid digestion. Methanogenic archaea are most commonly associated with insects that feed on wood or detritus, like coleopteran beetles and isopteran termites [7, 8]. Apart from these, the most common organisms found in the almost all insect gut are a huge diversity of bacterial species. Insects that consume primarily wood as part of their diet (a behaviour known as "xylophagy") have gut microbial communities that are capable of taking part in the breakdown of cellulose [9].

*The Ability of Insects to Degrade Complex Synthetic Polymers DOI: http://dx.doi.org/10.5772/intechopen.106948*

Cellulose is a good source of carbon, but it appears in plant cell walls as crystalline or amorphous microfibrils, making it inaccessible to the host [10]. Here the bacteria participates to break down complex cellulose into simpler sugar residues and monosaccharides [11, 12].

The relative importance of microbial and host-derived enzymes varies as per insect species and feeding habits or diet composition. According to this theory, if insects are actively force-fed, they can degrade plastic and synthetic polymers. In general, mandibulate insects have the ability to masticate and consume plastic materials by breaking them down into smaller pieces. Even though plastic fragments are small, they have a greater surface area of contact with gut microorganisms and are therefore mixed with them. Gut microbes use the enzymes responsible for depolymerizing plastic polymers into oligomers, dimers, or monomers, and the depolymerized products are mineralised into CO2, after which limited carbons are assimilated into biomass. Residual fragments and certain microorganisms in the gut are excreted as fecula, allowing for further degradation.

### **3. Synergy between insect gut microorganisms and synthetic polymers**

Insect larvae owing to their capacity of consuming and absorbing synthetic polymers, especially plastic have recently opened a huge scope for researchers seeking the most efficient procedure of plastic biodegradation. Larvae of Coleopterans beetles are reported to consume and degrade plastics. *Tenebrio molitor* [13–18] and *Tenebrio obscurus* [15], the super-worm, *Zophobas asatratus* [14–16, 18], *Tribolium castaneum* [19, 20] and *Plesiophthalmus davidis* [21] etc. are few examples of members of the coleopteran order with this special ability. Besides coleopteran fauna, lepidopteran caterpillars, such as, Indian meal moth, *Plodia interpunctella* [3], the greater wax moth, *Galleria mellonella* [22–26], and the lesser wax moth, *Achroia grisella* [27] are also reported to digest synthetic polymers like polystyrene (PS), polyethylene (PE), polyvinyl chloride (PVC), and polypropylene (PP) (**Table 1**).

These insect larvae use their mandibles to consume plastics or diets that are high in plastic content. The gut symbiont and commensal microbiota of insect larvae undergo alterations when they are forcibly fed or co-fed plastic feed. In general, regardless of insect species or polymer type, consuming plastic alters the relative abundance or diversity of certain Operational Taxonomic Units (OTUs) likely Enterobacteriaceae, Enterococcaceae, and Streptococcaceae in comparison to larvae fed natural, plasticfree diets [15]. These OTUs subsequently follow a three-step process to degrade the ingested plastics: (a) microbial colonisation and biodeterioration, (b) enzymatic depolymerization (breakdown of polymer into simpler monomers) and (c) mineralisation.

The microorganisms initially colonise on the polymer either individually or in consortium (colonisation), which is assisted by various polysaccharides and/or proteins [45]. Following that, the interplaying polysaccharides and cysteine-rich proteins permeate the surface, changing the size of the polymeric pore [46]. These alterations cause biodeterioration. The durability and resilience of the polymer will decrease over time, but its surface area will expand, giving microbes a bigger surface area to adhere to. Various bacterial cells often produce an extracellular slime material that promotes adhesion and resulting in a slow positive feedback by increasing pollutant build-up, allowing for increased microbial proliferation [47]. Various bacterial cells often produce an extracellular slime material that promotes adhesion



#### *The Ability of Insects to Degrade Complex Synthetic Polymers DOI: http://dx.doi.org/10.5772/intechopen.106948*


**Table 1.** *List of various insects and the synthetic polymers they degrade with the interplaying microbes and host enzyme.* and resulting in a slow positive feedback by increasing pollutant build up, allowing for increased microbial proliferation [47]. A number of different microbial enzymes have now initiated the enzymatic degradation process by depolymerizing and bio deteriorating the plastic polymers. Microbial enzymes (exo-enzymes) do biofragment synthetic polymeric structures into shorter chain oligomers, dimers, and monomers. The smaller molecules permeate and pass through the semi-permeable outer bacterial membrane (bio-assimilation) before taking up the depolymerization products (monomers) to obtain energy for cell metabolism and biomolecule production. The larvae can use the depolymerization products in the synthesis of different biomolecules.

Polymeric structures of plastics can be divided into C∙C backbone and C∙O backbone based on microbial breakdowns. PE, PP, PVC, and PS are examples of synthetic polymers with C∙C polymeric backbones that can also be biodegraded. Microbial oxidation begins with the hydroxylation of C∙C bonds and the formation of primary and secondary alcohols after the first breakdown of long-chain polymers to shorter and lower molecular weight carrying oligomers or monomers. This process is aided by the enzyme alkane hydroxylase, which does terminal and subterminal oxidation. Alcohol dehydrogenase further oxidises these alcohols, producing aldehydes and ketones. Aldehyde dehydrogenase then produces carboxylic acids, which increases the number of carbonyl-groups. The final carboxylate molecules, which are chemically identical to fatty acids, are incorporated into the oxidation pathway by microbes that provide bio-assistance for this process. In the case of PS, this generic degradation process shows only slight variation. The phenyl moieties are connected to the alternative backbone atoms of PS, which has a linear carbon backbone. Because of its unusual structure, PS biodegradation is more complicated; the organic product styrene formed after initial polymeric fragmentation is processed under the influence of numerous dioxygenase, isomerase, dehydrogenase, hydrolase, and aldolase enzymes. Easter bonds in the chemical structure of synthetic polymers with C∙O backbones, such as PU and PET, increase their hydrolyzability. Polyurethane (PU) is made up of di- or poly-isocyanate and poly-ols that are linked together by carbamate (urethane) bonds [48]. Carbamate bonds connecting the crystalline stiff segments are vulnerable to attack by microorganisms. Microbial ureases, esterases, and proteases are among the enzymes that interact during PU depolymerization. During the process of PU depolymerization, ureases are responsible for breaking the urea linkage, proteases are responsible for hydrolyzing the amide and urethane linkages, and esterases are responsible for hydrolyzing the easter bonds [49] After depolymerization, the poly-ols are dehydrogenated and oxidised to produce acetyl-CoA, which is then integrated into the TCA cycle or further valorized. Terephthalic acid (TPA) and ethylene glycol (EG) are ester-bonded together to form the polymer polyethylene terephthalate (PET) [50]. The ester linkages are hydrolyzed to produce polar hydroxyl and carboxylic groups by various PET surfacemodifying enzymes such as PET hydrolases after hydrolysis and depolymerization of monomeric constituents such as ethylene glycol (EG), terephthalic acid (TPA), monoethylene terephthalate (MHET), and bis-2-hydroxyethyl TPA (BHET) [51]. The enzyme MHETase is activated to further degrade the intermediate MHET and BHET into TPA and EG, which are then transported into the bacterial cell for further metabolism by dioxygenases and dehydrogenases. Finally, the final metabolites are converted into acetyl-CoA and succinyl-CoA, which enter biochemical cycles for mineralisation processes [52]. Fecula are expelled as residual and undigested particles.

## **4. A brief account of insects degrading synthetic polymer**

## **4.1 Lepidoptera**

The Lepidopteran insects capable of degrading synthetic polymers are discussed and detailed below. Following that, an overview of interplaying gut bacteria (**Figure 1**) that function in synergy with the host gastrointestinal enzyme is included.

## *4.1.1 The Indian meal-moth*

*Plodia interpunctella* (Lepidoptera: Pyraloidea), an adult Indian mealmoth with black-tipped feet and a black to brown small-headed caterpillar, feeds on cereals, fruits, and other similar items. The waxworm caterpillars live as parasites in bee colonies and eat on pollen, cocoons, and beeswax in addition to grain mix [53]. In meal wax, at least two intestinal bacteria were found: gram-positive strain YP1, Bacillus sp., and gram-negative strain YT1, *Enterobacter asburiae* [3]. The YP1 and YT1 strains were found to have roughly 11% and 6% net loss of PE polymers, respectively [3]. During inoculation, bacteria grow on PE sheets and gain weight, resulting in the formation of a liquid suspension within about a month, and finally, the hydrophobicity and tensile strength of PE decline. PE samples become less resistant to microbial destruction as they grow less hydrophobic [54]. Both the YP1 and YT1 bacteria adhere to PE films almost immediately and form biofilms within three hours after being inoculated, indicating that they are ready for biodegradation [55, 56].

#### **Figure 1.**

*Lepidopteran gut morphology and interplaying plastic degrading microbes.*

#### *The Ability of Insects to Degrade Complex Synthetic Polymers DOI: http://dx.doi.org/10.5772/intechopen.106948*

A biofilm, as a non-soluble substrate, permits microorganisms to adhere to it efficiently. The presence of predominantly living bacterial strain cells on biofilm shows that PE metabolism provides these cells with the necessary nutrients [54–56]. The two bacterial strains also cause damage to the physical integrity of PE by changing surface topography, as multiple micro-pits and cavities are identified on the surface of biofilms using scanning electron microscopy (SEM) and atomic force microscopy (AFM) [3]. By increasing the quantity of carbonyl groups, the YP1 and YT1 strains elicit chemical alterations in PE [3]. The presence of the carbonyl group suggests that bacterial strains can oxidise PE materials to produce the carbonyl group, which is an important indicator of PE biodegradation. Furthermore, the weight loss of PE samples inoculated with two bacterial strains increases consistently, but the sample's molecular weight decreases. This process implies that the long-chain structure of PE is depolymerized, resulting in smaller molecular weight fragments. The chemical and physical alterations of injected PE samples show that wax worm gut bacterial strains YT1 and YP1 are capable of decomposing PE. Plastic-chewing insect larvae of the Indian meal moth, P. interpunctella, may thus represent a promising source of plastic-degrading insects.

#### *4.1.2 The greater wax moth*

The greater wax moth, *Galleria mellonella* (Lepidoptera: Pyraloidea), also known as the honeycomb moth, is a lepidopteran insect with brown-grey pigmented forewings and scaly hind wings are also reported for their plastic consuming ability. They are sexually dimorphous, 10–18 mm in length, and are distributed worldwide [57–59]. The larvae of the honeycomb moth are creamy-white, 3–30 mm in length [60], feed more intensely during earlier instars compared to later instars, and undergo eight to ten moulting stages [59]. Its larval stages are extremely damaging due to its voracious feeding habits, especially for bees and bee hives [59, 61]. The larva feeds on pollen, honey, wax, and broods and can tunnel through the comb [59, 62].

Honeycomb larva may devour PE films by generating pores and holes at a rate of more than two holes per hour per worm and can consume approximately 200 mg of PE mass in 24 h at a rate of 0.23 mg/cm2 /h [22, 53]. Ethylene glycol was identified as a metabolic by-product due to PE degradation through FTIR analysis [22] or by treating caterpillars with broad-spectrum antibiotics [23]. The intestinal microbiomes of these caterpillars were found to play a distinct role in the PE degradation process [23, 40, 42]. Additionally, the larvae fed on PE showed the highest microbial abundance in their intestines, demonstrating the intestinal microbiome's favourable response to the PE diet. As a result, the presence of microbe abundance in *G. mellonella's* gut implies that the insect is benefiting metabolically from PE substrate [23]. When microbes from the intestine of *G. mellonella* were cultured in a liquid C-free medium containing PE for 60 weeks, Acinetobacter species ACT126, as well as Enterobacter sp. [42], and the fungus Aspergillus flavus [40], were discovered to be excellent candidates for contributing to the biodegradation process. Moreover, by performing Atomic Force Microscopy (AFM), an apparent change in the topography of the PE surface was observed after treating the PE with the greater wax moth. Following further microbe contact to PE films, the PE surface roughens, facilitating microbe adherence to plastic films [22].

According to the data, *G. mellonella* works naturally with flawless metabolic machinery to biodegrade lengthy hydrocarbon chains [23]. The greater wax moth feeds on beeswax in the natural, which is constituted of a highly diverse variety of lipid compounds, including alkanes, alkenes, fatty acids, and esters, with ethylene being the most common hydrocarbon bond in PE. Although more research into the molecular intricacies of wax biodegradation is needed, it appears that one of the targets of digestion is the C∙C single bond of aliphatic molecules. The presence of holes in PE films exposed to waxworms and the FTIR analysis of damaged PE revealed chemical disintegration of PE, including the breakage of C∙C bonds [22].

Along with PE degradation, the greater wax moth, *G. mellonella* larvae, was also reported to chew and ingest PS after analysis of their frass through GPC, FTIR, and GC-MS analysis [26]. When PS was allowed to feed on *G. mellonella* as a sole diet, the larvae could reduce PS's weight by nearly one gm in three weeks. However, co-dieting with their conventional nutritional food along with PS has resulted in increased PS degradation. The gram-positive lactic acid-producing bacteria, *Enterococcus* sp., facultatively anaerobic gram-positive bacteria, *Bacillus cereus*, and the gram-negative rod-shaped bacteria, *Serratia marcescens*, were isolated from the *G. mellonella* larval gut and were suspected of participating in PS degradation [26].

#### *4.1.3 The lesser wax worm*

An adult lesser wax worm, *Achroia grisella* (Lepidoptera: Pyraloidea), is light brown with golden highlights and black scales with long filiform antennae. Generally, it is 8–13 mm long; females are larger than males [63]. Lesser wax moths are widely distributed in tropical, subtropical, and temperate regions. The caterpillars of *A. grisella* are considered serious pests of beehives as their larvae consume bee wax [64, 65].

Like other pyraloid moths, *A. grisella* has also been reported as a PE-degrading bio-agent. They can degrade PE but less rapidly than the greater wax worm and can complete their lifecycle by consuming PE films [27]. When PE films are left in direct contact with *A. grisella* worms, the lesser wax worms, after chewing the films, make holes in them within a few days, approximately 2 ± 1 holes per worm per hour, and one individual larva can degrade nearly 2 mg of PE film daily [27].

Though the PE diet is not a good source of nutrients to grow and survive, the larvae of *A. grisella*, by consuming PE as a sole diet, live for almost one month and may develop into a second generation [27]. However, when additional nutrients were provided for them, PE degradation increased rapidly, and as a result, the survival and reproduction rate of *A. grisella* increased. In the wild, *A. grisella* larvae consume and digest beeswax, which has strong chemical bonds similar to PE. The ability of *A. grisella* larvae to digest PE plastic might be due to the presence of PE-degrading bacteria within their gut or any other unique extracellular enzymes that have not been discovered yet. The *A. grisella* caterpillar treated PE films showed an increased deviation of PE mass and decreased residual PE, suggesting most larvae consume PE either by disintegrating or assimilating the PE.

FTIR and NMR analyses of frass confirmed that the biodegradation process successfully occurs in *A. grisella* larvae [27, 66]. The presence of new carbonyl and alcoholic groups with the increase in unsaturated hydrocarbons provides evidence for the biodegradation process of PE in the lesser wax worm. However, further research must understand whether this PE biodegradation is gut-dependent or independent.

## **4.2 Coleoptera**

Representatives of Coleoptera, capable of degrading synthetic polymers are discussed and detailed below. An overview of interplaying microbes residing in the coleopteran gut (**Figure 2**) that function in synergy with the host gastrointestinal enzyme to degrade PE, PS, and PP is included.

## *4.2.1 The yellow mealworm*

Adult yellow-meal-worm beetles, *Tenebrio molitor* (Coleoptera: Tenebrionidae), also known as darkling beetles, are black to brown, have moniliform antennae, and complete their life cycle holo-metabolically. The larvae of yellow mealworms typically measure about 2.5 cm or more in length and have lighter body colours than adults, with long and slender structures. Generally, the mealworm feeds on stored grains, vegetation, and dead insects [67].

The larvae of mealworm beetles are capable of chewing and eating PS (Styrofoam) plastic as their sole diet [17]. Other investigations further supported this fact [13, 15, 68]. The larvae were found to degrade almost half of the consumed PS within 12–15 hours in their guts [13]. PS samples inoculated with the *Exiguobacterium* sp. bacterial strain (YT2) were found to lose more than 7% of their weight after two months of incubation [69]. The bacterial strain (YT2) was noticed to cause surface topography changes on PS materials, and as a result, the hydrophobicity of PS decreases, and carbonyl groups form. As a result, PS weight loss is due to molecular weight loss [69, 70]. It has also been opined that besides *Exiguobacterium* sp., a variety of microorganisms play an essential role in the digestion process of mealworms [71, 72].

#### **Figure 2.**

*Coleopteran gut morphology and interplaying plastic degrading microbes.*

The information indicates that PS biodegradation and mineralisation occur within the gut of yellow mealworms [73]. During consumption, the larva generally produce hollows in Styrofoam samples, resulting in a decrease in Styrofoam mass [17] and the resultant small fragments of Styrofoam samples have an increased surface area. As a result, they were subjected to enhanced enzymatic depolymerization [17]. Another strain of mealworm (strain CA) was reported to be capable of biodegrading seven PS wastes [68]. Further investigation using mealworms from 12 different sources showed that mealworms from different regions could eat and digest PS, and those findings support the hypothesis that the capability of biodegradation of Styrofoam by mealworms is independent of their geographic origin and seems to be ubiquitous to the members of this species [68]. From this result, it could be assumed that chewing and consuming PS by yellow meal worms is their adaptive intrinsic behaviour, as they feed upon decaying forest vegetation in the wild [74]. Styrofoam-feeding mealworms had a significant survival rate, implying that Styrofoam feeding did not cause a negative effect on their survival ability [75, 76], but it was obvious that the PS degradation rate could notably be enhanced if the diet was supplemented with conventional sources of nutrition. Mealworms fed on such a diet could reproduce and enter into the second generation, which seemed to have a higher affinity for PS materials [68, 75, 76]. The temperature was also found to have corresponded with the PS degradation rate. It was found that at 250 C, the mealworm degrades PS at a significantly higher rate [68]. Moreover, PS consumption is influenced by the density of the foam materials, which is related to product hardness rather than molecular weight and thus likely to be chewed and consumed by mealworms. FTIR and NMR analysis revealed that due to cleavages at long-chains of PS molecules, they turn into low molecular weight phenyl derivative metabolites in the gut of mealworms [17, 36].

Yellow mealworms fed with PE and PS plastic each as the sole diet were found to cause mass loss of both the plastics. The yellow mealworms can degrade both PE and PS, but the degradation efficiency of PE was noticed to be much higher (48%) than PS (32%) on solo plastic diets. However, in both cases, degradation efficiency can be increased by up to 61% (for PE) and 54% (for PS) if the larvae are fed conventional food in addition to plastics [13]. The difference in mass loss of PE and PS might be due to the differences in density of the plastics, and it is presumed that less dense plastic molecules are ingested at a higher rate [68]. Among the present plastics, PE possessed a higher density than PS, which indicated that there might be other factors responsible for affecting the relative consumption rates of PE-PS plastic combinations. However, no clear evidence has yet been established to get an answer. Analysis involving HT-GPC, FTIR, and NMR studies certified that plastics could be degraded entirely and mineralised in the gut of the mealworms within a month approximately.

It was hypothesised that microbial communities significantly differed from the diets of the caterpillars or larvae. However, most microbial community members do not vary significantly in PE-fed diets among insects, but the composition is distinct in the PS-fed community. For example, *Citrobacter* sp. and *Kosakonia* sp., belonging to the family Enterobacteriaceae, were intensely associated with both plastic diets, viz., PE and PS [42] Both can use oxygen, which proves their participation in plastic degradation, as the biodegradation of both PE and PS is accelerated upon incorporating O2 [32]. On the other hand, another two microbes, anaerobic gram-negative *Sebaldellatermitidis*, and gram-positive *Brevibacterium* sp., were uniquely associated with PE degradation [77]. Seven other microbes, viz., Listeria sp., Nitrospiradefluvii, Pedomicrobium sp., Aquihabitanssp., unclassified Xanthomonadaceae, Saprospiraceae, and Burkholeriales, were found to be

#### *The Ability of Insects to Degrade Complex Synthetic Polymers DOI: http://dx.doi.org/10.5772/intechopen.106948*

associated with PS degradation in mealworm gut, further suggesting the significance of the microbial community in the plastic degradation process [78]. The information regarding the presence of various microbes in the mealworm gut suggests that mixed plastics of PS and PE could be depolymerized within the gut of the same mealworm. Therefore, the mealworm gut is not plastic-specific rather than independent in the degradation of any PE or PS plastics.

#### *4.2.2 The dark mealworm*

Adult dark meal-worm, *Tenebrio obscures* (Coleoptera: Tenebrionidae), also known as mini mealworms similar to yellow mealworms in appearance, are also known for their plastic-consuming ability. The larvae of dark meal worms are 1.5–2.5 cm in length, possess dark black rings on their abdomen, and become dark with maturity. The larvae have higher light sensitivity than yellow mealworms [15]. They usually consume seeds, vegetables, flour, and oats [68].

The larvae of dark mealworms were found to have the ability to degrade PS [15, 32], the depolymerization rate being higher than equally sized yellow mealworm larvae [15]. When dark meal worms were supplied with PS as their sole diet, mass loss of PS was found to be 55% in a month, but the amount of PS degradation was increased by 67% when the larvae were co-fed with supplementary food [15]. The investigation suggests that PS degradation ability can be achieved at a higher tempo when the insects are allowed to feed on a nutrition-rich co-diet. GPC and FTIR analysis supported that PS degradation was found to be operated by the active participation of gut bacteria residing in the dark mealworms. Before feeding PS, the gut microbiome was found to have higher diversity in *T. obscurus* than in *T. molitor* [15]. According to microbial community analysis, bacteria from the Enterococcaceae, Spiroplasmataceae, and Enterobacteriaceae families were particularly associated in their guts for PS depolymerization and degradation [15].

#### *4.2.3 The super worm*

Super worms, *Zophobas atratus* (Coleoptera: Tenebrionidae), also known as blind click-beetles, have very dark elytra on their cover, and after attaining maturity, the beetles become darker and are then called "black beetles." Superworms have mandibulate mouthparts like mealworms, which provide these species the ability to chew and eat plastic.

Super worms are also found to chew and eat Styrofoam as their sole diet [79], and when they were left on Styrofoam samples, they instantly started to ingest and penetrate through the blocks and made hollows in the blocks within an hour [68]. *Z. atratus* can consume up to 0.58 mg of Styrofoam per day, which is four times more than mealworms (0.12 mg/day/worm) [17, 32]. Interestingly, the survival rate of super worms eating Styrofoam was almost equal to that of a regular diet, which indicates that super worms can complete their lifecycle by consuming Styrofoam diets [32]. After passing through their guts, the consumed long-chain PS molecules were degraded into low molecular weight products, styrenes, which were again mineralised into CO2 [32]. Moreover, an antibiotic suppression assay using a combination of gentamycin, rifampicin, and streptomycin indicated that repression of gut microbiota by antibiotics diminished the ability of superworms to degrade PS and, therefore, confirms that the gut microbiota plays an important role in PS degradation in superworms [32, 35, 36]. Three bacterial strains, *Aeromonas* sp. and *Klebsiella pneumonia*

from Enterobacteriaceae [36] and *Pseudomonas* sp. [35] from Pseudomonadaceae have been isolated from *Z. atratus* gut and confirmed their ability to degrade PS.

## **4.3 Orthoptera**

## *4.3.1 Crickets*

Orthopteroid fauna like crickets, such as *Gryllus bimaculatus* are also found to consume polyurethane (PU) plastics [44]. It has been noticed that *G. bimaculatus* is capable of consuming a diet that is 63% more rich in polyurethane (PU) than its usual food. Nine distinct microbial organisms, including bacteria and fungus, were identified in their digestive tracts which might take part in PU digestion. The fungus strain *Aspergillus flavus* G10 was isolated and identified from their gut after PU-degrading activity assays. The fungus was also noted to be responsible for PU degradation [44]. However, more research needs to be done on effective insect species as well as the potent gut microbial organism that are capable to degrade PU.

## **4.4 Other insects**

There are some other insects from coleopteran and lepidopteran order that is seen to degrade synthetic or natural polymers (**Table 1**). Insects like cigarette beetles (*Lasioderma serricorne*), lesser grain borer (*Rhyzopertha dominica*), rice weevil (*Sitophilus oryzae*), saw toothed grain beetle (*Oryzaephilus surinamensis*)—from order coleoptera are among those, that have potential in digesting polyethylene or structurally similar polymers. Other Coleopterans namely red flour beetle (*Tribolium castaneum*), darkling beetles (*Plesiopthalmus davidis*) and lesser mealworm (*Alphitobius diaperinus*) are capable of degrading polystyrene or alike polymers. Other Lepidopterans like Rice moth (*Corcyra cephalonica*) and Isopteran Termites owing to their feeding habits of complex natural polymer have immense possibilities in degrading synthetic polymers.

## **5. Conclusion**

In recent years, there has been a significant increase in the production of plastic due to the proliferation of its usage in areas ranging from the domestic sphere to multiple business spheres. However, improper treatment and management of plastic waste disposal have led to the accumulation of this material in the environment, which poses threats to the health of living species as well as to the health of humans. The most common petroleum-based polymers, PE, PP, PS, and PVC, have been thought to be non-biodegradable for many years. However, recent studies have shown that these polymers can be degraded by the microbial communities either on their own or with the active participation of the microbial activities that are present in the larval guts of certain insects. The knowledge that is currently available about the role that insects play in the breakdown of plastic is quite restricted, and as a result, several questions on the process of plastic degradation via insects are still unclear. It has not yet been determined what the precise processes underlying the degradation process are or what the function of the enzymes should be in this process. However, the good news is that the capability of some insects to degrade compounds that are rarely biodegradable or even non-biodegradable may be employed for the practical applications for the waste management programme, which can be shown to be extremely helpful for the health of the environment.

## **Acknowledgements**

The authors are thankful to Head, Department of Zoology, University of Calcutta for providing necessary facilities to conducting this work. The authors are also thankful to Moumita Mondal, Debjit Safui and Sumit Mondal for helping of this manuscript preparation.

## **Author details**

Biswarup Mitra\* and Amlan Das Entomology Laboratory, Department of Zoology, University of Calcutta, Kolkata, WB, India

\*Address all correspondence to: biswarupmitra1999@gmail.com

© 2022 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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## **Chapter 6**

## Deterrents and Their Effects on the Feeding Behavior and Sensory Physiology of Insects

*Vonnie D.C. Shields*

## **Abstract**

The gustatory system of insects is a prominent model in neuroscience. This important sensory system allows insects to detect, encode, and process gustatory information. This important sensory modality allows insects to perceive their environment. All animals detect and react to chemicals in their environment. Using insects as model systems allows us to obtain fundamental information regarding the processing of sensory information in the brain of the animal. Stimuli, associated with taste and smell, are responsible in insects being able to locate and select food sources, mates, and egg-laying sites. One line of research can be directed to better understanding gustatory cues in the selection of food sources by insects. Experimentally, this will involve feeding behavioral and electrophysiological testing in insects. Examining the structural organization of the gustatory organs using transmission electron and scanning electron microscopy will shed more light on the detailed structure of these taste sensory organs, the sensilla. During feeding, these taste organs sample the plant sap that contains a multitude of phytochemicals. Gustatory sensory input is encoded as patterns of nerve impulses by gustatory receptor cells which are housed in these taste sensory organs. Taste information gathered by these receptor cells will allow the insect to determine if the food is palatable or should be rejected.

**Keywords:** gustation, taste, glucosinolates, deterrent, feeding behavior, insect

## **1. Introduction**

Our ability to taste is crucial for our survival and is central in our nutrition. The sense of taste determines the palatability of food and beverages. It provides early warning alerts for the detection of spoilage. Taste disorders affect the quality of life, daily living, psychological well-being and can change body weight or appetite. Having an appreciation of basic gustatory mechanisms in animals, including humans, allows us to have a better understanding and promises to contribute toward an explanation of taste disorders.

Using caterpillars as insect models allows us to increase our understanding of taste recognition and coding and to unravel some of the principles that govern food selection behavior. One feature that makes these larvae ideal candidates for such studies is their recognizable gustatory behaviors. In addition, they have relatively

simple gustatory systems with relatively low numbers of sensory cells located in sensory organs (sensilla) on their mouthparts that mediate taste mechanisms. These cells are individually identifiable, show strong electrophysiological responses, and are relatively easy to access for experimental manipulation. All these features make the larval gustatory system amenable to structural, behavioral, and electrophysiological approaches, respectively.

## **2. Ultrastructure of main taste sensory organs: styloconic sensilla**

Caterpillars bear a pair of uniporous styloconic sensory organs or sensilla (i.e., lateral and medial styloconic sensilla) on the maxillae, and more specifically, maxillary galeae. These are the main organs involved in feeding and detect plant phytochemicals by being in constant contact with plant sap during feeding. Each sensillum appears as a small cone inserted into a fibrous cuticular socket of a cylindrical projection or style of insensitive cuticle. The cone bears a terminal pore (**Figure 1**) [1]. In each styloconic sensillum, four bipolar taste neurons extend toward the tip of the cone. Receptors bound to these neurons interact with plant sap as the caterpillar is feeding. These receptors respond to salt, one or more sugars, and bitter compounds [2, 3]. One bipolar neuron, a putative mechanosensory neuron, terminates near the base of the cone and lies near the dendritic sheath [4, 5]. Here, many microtubules lie parallel to one another within a dense matrix. This location is thought to be the site of sensory transduction of mechanical stimuli [6]. Each styloconic sensillum bears a single apical or terminal pore. Gustatory sensory input gathered from the receptor cells within the sensory organs is encoded as patterns of nerve impulses which ultimately determine if relevant information is accepted or rejected in the brain of the animal (**Figure 2**).

#### **Figure 1.**

*A–C, Scanning electron micrographs of Lymantria dispar (L.) fifth instar larvae. The specimens were critical point dried. (A) Frontal view, whole head. The arrows denote the galeae, components of the maxillae. Bar = 1 mm. (B) Side view of a medial styloconic sensillum. The cone is inserted into the style or cylinder. The arrow shows the location of the pore at the tip of the cone. Bar = 1 μm. (C) Higher magnification view of the cone (c) from a lateral styloconic sensillum showing the pore (p with arrow) at the tip of the cone. Bar = 5 μm. This figure was adapted from [1].*

*Deterrents and Their Effects on the Feeding Behavior and Sensory Physiology of Insects DOI: http://dx.doi.org/10.5772/intechopen.112735*

#### **Figure 2.**

*Diagrammatic reconstruction of a uniporous styloconic sensillum shown in longitudinal section with five bipolar neurons innervating this sensillum: four gustatory and one mechanosensory. This illustration also shows the electrophysiological tip recording method. This method is useful for recording the neurophysiological responses from taste cells in a styloconic sensillum [7]. A taste stimulus is dissolved in an electrolyte solution (e.g., 0.1 M KCl dissolved in deionized water) contained within the stimulating or recording electrode. This electrode is placed over the tip of the pore of the sensillum. The solution diffuses through the pore. Taste compounds bind to dendritic taste receptors which transduce the quality and quantity of the stimulus into a neural code of action potentials. The indifferent or ground electrode contains a similar electrolyte solution except for the taste stimulus. Each electrode contains a silver wire. The solution and wire allow contact to be made with the internal environment of the insect (e.g., body). The excitatory responses recorded are amplified, digitized, and analyzed using a computer software program. ax, axon; cb, cell body; cs, ciliary sinus; dbb, distal basal body of proximal dendritic segment; dc, dendritic channel; dd, distal dendritic segment; ds, dendritic sheath; f, fibrils; i, inner sheath cell; n, intermediate sheath cell; o, outer sheath cell; pcu, peg cuticle; pd, proximal dendritic segment; po, terminal pore; pbb, proximal basal body of proximal dendritic segment; r, rootlets; scu, style cuticle; ss, sensillar sinus; tb, tubular body. This figure was adapted from [1].*

### **3. Sensory responses to deterrents**

At least one sensory cell is particularly sensitive to substances that cause a deterrent response in some larval Lepidoptera known as the deterrent neuron [8]. Neurophysiological responses from one or more taste cells within the sensillum can be recorded using can be acquired using an electrophysiological tip recording method [7].

Deterrent receptors in caterpillars were thought to have evolved as a proliferation of receptor types. They have an extensive action spectrum sensitive to a large variety of secondary plant compounds, ultimately resulting in appropriate behavioral outcomes that are associated with specific sensory inputs [8]. In 1992, Schoonhoven et al., hypothesized that the deterrent receptor evolved from ancestral nerve cells that retained their sensitivity to noxious plant compounds [9]. It was thought that other chemoreceptor cell types, such as sugar-sensitive cells, developed a relative insensitivity to noxious chemicals. Deterrent cells were thought to respond to compounds not previously experienced in their recent evolution [10]. If the insect transitioned to a new host-plant, a loss of sensitivity by a deterrent receptor could occur [10]. If the ingested deterrent compound, or its metabolic products, are taken up in the blood, they could potentially travel to the chemosensory cells causing a desensitization of response [11]. Over the course of evolution, the deterrent cell in the crucifer specialist, *Pieris brassicae*, became insensitive to sinigrin, probably because of this insect's very close host–plant association with the Cruciferae [12–14]. Another cell in *P. brassicae* is sensitive to glucosinolates, presumably mediates host-recognition, and likely signals acceptance rather than rejection [13].

### **4. Insect-plant interactions: sensory basis of feeding**

The sense of taste plays a key role in the behavior of insects. Insects often rely on gustatory cues from plants to detect and find their food sources. These cues are typically nonvolatile chemicals that are either liquids or solids and can be simple or complex. They can be detected via contact chemoreceptors located on various body parts [15, 16]. Other compounds may be partially volatile [16, 17]. Examples of such compounds include DEET (N,N-diethyl-m-toluamide) [18, 19], ammonia, water, polyamines, and certain acids, pheromones, and fatty acids. Insects, in general, are selective to some extent with respect to the selection of their food choices. Monophagous insects feed on one or a few closely related plant species, whereas oligophagous insects feed on a larger number of hosts, usually confined within a certain plant family. Polyphagous insects consume many plants representing a wide taxonomic range. Insects never feed on all plant groups [20], however.

Tastants have often been grouped into taste qualities: sweet, sour, bitter, and salty [21]. Umami (savory) was added later [22]. While insects can respond to these five canonical taste qualities, taste quality perception in insects may be different than in humans. Sweet, often associated with sugars [23, 24] and sugar alcohols [23, 25–28] are attractive to Lepidoptera, as well as other insects, as are some artificial sweeteners (e.g., acesulfame K) [29]. Bitter, i.e., deterrent tastants, are represented by compounds such as caffeine, denatonium, and quinine. They may be toxic [30–33] and have diverse chemical structures [34]. Sour tastants are associated with certain acids, including acetic acid, citric acid, hydrochloric acid, and lactic acid [35–38]. Salty tastants are associated with sodium and other mineral ions, such as NaCl and KCl [39–41]. Lastly, umami tastants are associated with some amino acids [24, 42–45].

Two theories exist to explain how chemical constituents of plants provide stimuli that determine food-plant preferences. Brues first suggested that insects' "botanical

#### *Deterrents and Their Effects on the Feeding Behavior and Sensory Physiology of Insects DOI: http://dx.doi.org/10.5772/intechopen.112735*

instinct" was based on responses to chemical and physical stimuli originating from plants [46]. Later, Dethier [47] and Fraenkel [48] stated that nutritionally unimportant "token stimuli" or attractants and repellents were primarily responsible for regulating the feeding preferences of phytophagous insects.

Many plant feeders are very specific in their food habits. They are usually restricted to a single order, family, genus, species, or even subspecies of plants. This specificity can be supported by one of two different factors, or possibly their combination: (i) insects will tend to specialize on plants that meet their dietary needs (i.e., protein, carbohydrates, fat, minerals, sterols, and vitamins) and (ii) if dietary composition is very similar for all insects, the guiding factor will be the presence or absence of additional compounds (i.e., "secondary" plant substances); the presence of nutrients would be less important. The "dual discrimination" theory proposes that insects respond to secondary compounds (token stimuli for recognition of host-plant species), as well as nutrients (for recognition of plants of the exact physiological condition nutrient content [49]). The primary "sapid" nutrients in plants act as important taste indicators of a suitable food, in addition to the recognition of secondary plant substances [50]. Host-plant selection by specialist feeders is thought to be largely influenced by the presence of token stimuli, whereas for generalist species, the presence of deterrents plays a major role [51].

Host-plant recognition and utilization, as well as avoidance or rejection of nonhost plants, are generally inherited and cannot be changed by experience [52]. The primary role of secondary plant substances in insect-host-plant relationships is that they form the "fingerprint" (specific signal pattern) or biochemical profile, by which the insect identifies the plant [53]. If the plant biochemistry, as perceived by the insect, fits the expected innate image of "host-plant" to the insect, the plant will be consumed or selected as a location for egg-laying [54].

For an insect to feed, it must (a) recognize and orient to the plant; (b) begin feeding (biting or piercing); (c) maintain feeding, and (d) stop feeding, prior to dispersal. Terms applied to define classes of stimuli should encompass both physical and chemical stimuli. An "attractant" is a stimulus to which the insect responds by moving toward the food source. "Arrestants" cause the insect to stop moving toward the food source. Initiation and maintenance of feeding are separable phenomena. "Feeding incitant" is a stimulus initiating biting or piercing of the plant tissue. Once biting has started, maintenance of feeding is dependent on the presence or absence of feeding stimulants or deterrents [55]. Food selection behavior can be compared to a "key-lock" system, where the key represents a complex sensory pattern [56]. A precise behavioral response will be triggered if the pattern sufficiently corresponds with an innate standard. When the incoming sensory information differs too much from the desired pattern, the food is rejected. Host selection comprises of a series of steps (i.e., keys). Each step unlocks only one behavioral step. The lack of detail in one key will be compensated for by details in another sensory pattern (**Figure 3**) [57, 58].

The gustatory and olfactory systems of lepidopterous larvae distinguish the presence of various chemicals. Sensilla associated with these senses are located and distributed on their antennae, mouthparts, and legs. The styloconic sensilla, located on the mouthparts, are in continuous contact with plant sap during feeding. Four types of gustatory receptors have been classified into four cell types: those sensitive to nutrients, salts, phagostimulating alleochemics, and deterrents (**Figures 4** and **5**) [59, 60].

The sensitivity of chemoreceptors also vary with age, time of day, feeding history, effect of food deprivation, adaptation rate, individual insect, and temperature

#### **Figure 3.**

*A. Mean consumption by fifth instar Mamestra configurata exposed to a diet containing increasing concentrations of sinigrin. B. Mean consumption by fifth instar Trichoplusia ni exposed to a diet containing increasing concentrations of sinigrin. This figure was adapted from [58].*

#### **Figure 4.**

*Representative electrophysiological responses from the lateral and medial styloconic sensilla of fifth instar Mamestra configurata in response to: (a) 50 mM potassium chloride; (b) 2 mM sinigrin (glucosinolate); and from (c) 60 mM sucrose from lateral sensilla; (d) 50 mM potassium choride, and (e) inositol from medial sensilla. Potassium chloride (50 mM) served as the electrolyte. Note the strong firing response to inositol in (e), whereas potassium chloride evoked a very minimal response (a). This figure was adapted from [60].*

[57, 59]. Städler and Hanson demonstrated in *Manduca sexta* larvae, that three of the four chemoreceptive cells, only in lateral styloconic sensilla, possessed a shortrange (0–0.5 mm) olfactory capability to perceive vapors [61]. This would allow the receptors to monitor food without being in actual physical contact with it. The three mechanosensory galeal trichoid sensilla may provide information about the proximity of the food source, permitting the lateral sensillum to gauge the concentration of plant vapors accurately [61].

*Deterrents and Their Effects on the Feeding Behavior and Sensory Physiology of Insects DOI: http://dx.doi.org/10.5772/intechopen.112735*

#### **Figure 5.**

*(a) Dose-response curve showing the sinigrin-sensitive cell in the lateral styloconic sensillum of* Mamestra configurata *when stimulated with various concentrations (mM) of sinigrin. Each point represents 10-23 larvae (cells). Error bars represent standard error of the means. (b) Adaptation curves for the sinigrin-sensitive cell in the lateral styloconic sensillum of* M. configurata *during stimulation with 8 mM (filled squares) and 20 mM (open circles) sinigrin. Each point represents means for 4-6 larvae (cells). Error bars represent standard error of the means. (c) The inset shows the first 10.1 secs. of the adaptation response for both cells. This figure was adapted from [60].*

## **5. Secondary plant compounds and the role of sinigrin as a feeding stimulant or deterrent**

There are four major classes of secondary plant compounds: nitrogen-containing (alkaloids, amines, amino acids, cyanogenic glycosides, and glucosinolates), terpenoids (monoterpenes, sesquiterpenes, diterpenes, saponins, limonoids, cucurbitacins, cardenolides, carotenoids), phenolics (simple phenols, flavonoids including tannins, quinones), and polyacetates (polyacetylenes). These are distributed widely in vascular plants, including Solanaceae, Scrophulariaceae, Cucurbitaceae, Gymnospermae, etc. Secondary plant compounds are found in concentrations varying from, e.g., 0.0002–>40% concentration dry weight [62].

Secondary plant compounds serve as positive compound signals when an insect species becomes adapted to particular plants and uses these cues to recognize their

hosts. Glucosinolates, found in the family Brassicaceae, the mustard family of flowering plants (order Brassicales), composed of 338 genera and 3700 species, appear to be limited to families of dicotyledonous angiosperms occurring in the order Capparales including the families Cruciferae, Capparaceae, Tovariaceae, Resedaceae, and Moringaceae [62]. They are present in every part of the oilseed rape plant [63] and are unlikely to serve a role in the basic metabolism of plants. Food specificity of insects is thought to be based solely on the presence or absence of these compounds [51]. Glucosinolates, or mustard oil glucosides, are derived from amino acids and contain sulfur, as well as nitrogen atoms. They can either be acyclic (e.g., sinigrin) or aromatic (e.g., sinalbin) [62].

Sinigrin (allyl- or 2-propenyl glucosinolate) is a principal crucifer token stimulus and is a widespread glucosinolate in many species of Cruciferae, as well as in other plant families. Glucosinolates are broken down by a glucosinolate-degrading enzyme (myrosinase) when plant tissues are eaten or damaged, thereby releasing toxic hydrolysis products. These products may include isothiocyanates, nitriles, thiocyanates, and oxazolidinethiones [63, 64]. Myrosinase is present in idioblasts (specialized cells in parenchymatous tissue of the green parts of crucifer plants, whereas glucosinolates are stored in vacuoles of leaf cells [64].

The role of mustard oil glucosides acting as feeding attractants, incitants, and stimulants have been studied extensively. Insects, not adapted to a particular plant species, may be repelled or deterred by the plant. For noncruciferous feeding insects, glucosinolates have been implicated as feeding deterrents. Verschaffelt demonstrated the role of a mustard oil glucoside in food-plant selection in two lepidopterous species, namely *Pieris brassicae* and *Pieris rapae* [12]. Experiments showed that these insects could be stimulated to feed on normally rejected plants by treating the plant tissue with juices extracted from crucifers. When solutions of pure sinigrin were applied to unacceptable plants, they rendered them palatable. This was further exhibited with the diamond-back moth (*Plutella maculipennis*) when a solution of either sinigrin or sinalbin (p-hydroxybenzyl glucosinolate) was applied to the leaves of nonhost plants stimulating feeding [65]. Feeding did not occur, however, when leaves were treated with the mustard oil, allyl isothiocyanate, a product of enzymatic degradation from sinigrin. The stimulatory effect of sinigrin and sinalbin was demonstrated in larval *Pieris maculipennis* feeding on a synthetic diet [66], as was the case when the addition of sinigrin strongly promoted feeding in the mustard beetle, *Phaedon cochleariae* on a synthetic diet. [67]. A similar result was observed by other researchers with synthetic diet for larval *P. brassicae* [68–71]. Isothiocyanates, such as allyl isothiocyanate, are effective in attracting larvae of the European cabbage butterly, *P. rapae* [72, 73].

Glucosinolates were found to be both deterrent and toxic to the noncruciferfeeding lepidopterous caterpillar, *Papilio polyxenes*, which normally feeds on Umbelliferae [74]. Breakdown products may be responsible for the potency of sinigrin, as well as other glucosinolates as feeding deterrents [75]. Allyl isothiocyanate is known to be a powerful tissue irritant [64] and may be responsible for sinigrin toxicity to some species due to its release in the gut. Sinigrin may represent an innocuous form of storage in the plant, possibly as a means of avoiding autotoxicity [76]. Recently, some crucifer feeders have been shown to be deterred by some glusocinolates. Work on *Mamestra configurata* and the flea beetle, *Phyllotreta cruciferae* [77] (using the glucosinolate, sinalbin) and *Mamestra brassicae* [78], *M. configurata*, and *Trichoplusia ni* (using sinigrin [58]), clearly demonstrated deterrence in these crucifer-feeding insects.

*Deterrents and Their Effects on the Feeding Behavior and Sensory Physiology of Insects DOI: http://dx.doi.org/10.5772/intechopen.112735*

## **6. Conclusions**

Insects constantly monitor and respond to changes in their internal and external environments to maintain themselves under the most favorable conditions for survival. In the case of lepidopterous larvae, gustatory sensilla (i.e., lateral and medial styloconic sensilla) located on each maxilla can detect phytochemicals present in plants. They act as the first level of environmental perception and play important roles in host–plant selection, as they are in constant contact with plant sap liberated during feeding. The plant material enters each of these sensilla through an apical pore and interacts with four gustatory neurons and their receptors. Receptors bound to the dendrites transduce the chemical stimulus into a code of action potentials reflecting the quality and quantity of the complex plant chemistry. Subsequently, these nerve impulses are sent to the brain of the insect. The responses of these receptors to phytochemicals are key in determining which plants are deemed palatable and which should be rejected. Deterrent substances, such as e.g., some glucosinolates and alkaloids, are important in influencing the food selection of many insects, as they may be potentially toxic. Having a better understanding of the sensory mechanisms by which insects detect plant phytochemicals will help in finding novel biocontrol techniques against insect pests, especially highly polyphagous ones capable of defoliating forests or destroying crops.

## **Acknowledgements**

This work was supported by NIH grants 1R15DC007609-01 and 3R15DC007609- 01S1 to V.D.C.S., a grant from each of the Fisher College of Science and Mathematics, the Towson University Office of Undergraduate Research, and NIH grant DC-02751. The author gratefully acknowledges B.K. Mitchell and T. Maugel.

## **Author details**

Vonnie D.C. Shields Biological Sciences Department, Fisher College of Science and Mathematics, Towson University, Towson, MD, USA

\*Address all correspondence to: vshields@towson.edu

© 2023 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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Section 3
