A Review of Resistance Mechanisms of Synthetic Insecticides and Botanicals, Phytochemicals, and Essential Oils as Alternative Larvicidal Agents Against Mosquitoes.

Mosquitoes are a serious threat to the society, acting as vector to several dreadful diseases. Mosquito management programes profoundly depend on the routine of chemical insecticides that subsequently lead to the expansion of resistance midst the vectors, along with other problems such as environmental pollution, bio magnification, and adversely affecting the quality of public and animal health, worldwide. The worldwide risk of insect vector transmitted diseases, with their associated illness and mortality, emphasizes the need for effective mosquitocides. Hence there is an immediate necessity to develop new eco-friendly pesticides. As a result, numerous investigators have worked on the development of eco-friendly effective mosquitocidal compounds of plant origin. These products have a cumulative advantage of being cost-effective, environmentally benign, biodegradable, and safe to non-target organisms. This review aims at describing the current state of research on behavioral, physiological, and biochemical effects of plant derived compounds with larvicidal effects on mosquitoes. The mode of physiological and biochemical action of known compounds derived from various plant families as well as the potential of plant secondary metabolites, plant extracts, and also the essential oils (EO), as mosquitocidal agents are discussed. This review clearly indicates that the application of vegetal-based compounds as mosquito control proxies can serve as alternative biocontrol methods in mosquito management programes.

Introduction

Vector borne diseases account for more than seven million deaths annually (World Health Organization [WHO], 2017), among which mosquito borne diseases are the most threatening due to their wide spread occurrence, consequently featuring a higher frequency of disease transmission (Lounibos, 2002; Tyagi et al., 2015). Among different mosquito families, Culicidae is a large family (3,300 Service species-41 genera) comprising Toxorhynchitinae, Anophelinae (anophelines), and also Culicinae (culicines) sub-families (Service, 1996; Senthil-Nathan et al., 2005b). Among the 31 genera, Anopheles, Culex, and Aedes are the most detrimental. Anopheles species, are carriers of major life-threatening diseases (malaria and filariasis-transmitting agents, such as Wuchereria bancrofti, Brugia malayi, and Brugia timori) and also of a few arboviruses (Kalaivani et al., 2012; Benelli et al., 2018; Thanigaivel et al., 2019; Vasantha-Srinivasan et al., 2019).

The discovery of DDT’s insecticidal properties in late 1930s/beginning of 1940s and the following progress of organochlorine invention and organophosphate insecticides concealed biological pesticide merchandise-research since the responses to mosquito regulation were supposed to have remained established (Shaalan et al., 2005; Senthil-Nathan et al., 2006a, b). The ranges of many of the mosquito species were not limited and keep expanding, thereby up surging the rates of disease incidence. Until recently, the use of several of the earlier synthetic-insecticides, such as permethrin and malathion, along with other organophosphates in vector control programes has been partial. This is due to absence of unique-insecticides, expense of synthetic-insecticides, apprehension for ecological sustainability, damaging influence on human health, besides further non-target populations, their persistent nature, greater amount of “biological magnification” through ecosystem and also the development of insecticide resistance (Ghosh et al., 2012). The emergence of DDT resistance in Aedes species (Ae. tritaeniorhynchus and Ae. sollicitans) lead to numerous drawbacks in mosquito control programs (Brown, 1986). Several categories of Mosquitocides are being implemented in malaria control programs (BHC, organophosphorus, carbamate, and pyrethroid). The ability of mosquitoes to evade the insecticidal action of these synthetic compounds are attributed to the increase in the rate of synthesis of detoxifying enzymes such as monoxygenases (MFOs), glutathione-S-transferases (GST) and carboxyl-cholinesterase (CCE). MFOs are often associated with metabolic resistance to pyrethroids, such as permethrin, while GSTs are usually associated with organochloride resistance such as DDT. Resistance to pyrethroids, organophosphates and carbamates, such as bendiocarb are incurred by the magnification of CCE activity (Hemingway and Ranson, 2000). Added insecticides, benzylphenyl urea and the larvicide, Bacillus thuringiensis israelensis (Bti), have partial use against mosquitoes. Unpredicted natural or anthropogenic associated ecological variations that modify the original habitats severely affect the vector biology thereby positively influencing their existence and disease incidence, thus constraining the frame-work of mosquito control strategies.

Biological Management of Mosquitoes

Several phytochemicals from several plant families are identified with larvicidal activities against different mosquito species (Table 1). Plant extracts with their augmented phytochemical elements have a recognized potential as a substitute to conventional mosquito control agents (Sukumar et al., 1991; Tripathi et al., 2009; Tehri and Singh, 2015). The main strategy for mosquito control deals with the restriction of the vector population. As a promising biocontrol agent, the compounds from the plants of the family Meliaceae such as neem Azadirachta indica A. Juss (Senthil-Nathan et al., 2005b; Senthil-Nathan, 2013), Indian white cedar, Dysoxylum malabaricum Bedd. (Senthil-Nathan et al., 2006a), D. beddomei and chinaberry tree, Melia azedarach L. (Senthil-Nathan et al., 2006b) were effective against An. stephensi (Senthil-Nathan et al., 2008). “Secondary metabolites” from Eucalyptus tereticornis Sm. (forest redgum, Myrtaceae) exhibited effective mosquitocidal activities against An. stephensi as reported by Senthil-Nathan (2007). Also, the crude metabolic extracts of Acanthospermum hispidum leaves were active against An. stephensi, Ae. Aegypti, as well as Cx. quinquefasciatus as reported by Vivekanandhan et al. (2018a, b). A study conducted on testing the mosquitocidal activity of Justicia adhatoda L. (Acanthaceae) leaf extracts revealed the potential of natural larvicidal agent against Ae. Aegypti (Thanigaivel et al., 2012, 2017a,b).

TABLE 1

Phytochemicals identified from the specific plant families and their larvicidal activity on the mosquito species.

Family and plant species	Major constituents	Mosquito species	References
Acanthaceae
Andrographis paniculata	Andrographolide	Aedes aegypti	Edwin et al., 2016
Alangiaceae
Alangium salvifolium	Asarinin, sesamin and (+)-xanthoxylol-γ,γ-dimethylallylether, Hexadecanoicacid,1 hydroxymethyl-1,2-ethanediyl ester	Aedes aegypti	Thanigaivel et al., 2017a
Amaranthaceae
Chenopodium ambrosioides	α-Terpineol	Aedes aegypti	Leyva et al., 2009b
Amaryllidaceae
Alium macrostemon	Methyl propyl disulfide; mimethyl trisulfide	Aedes albopictus	Liu et al., 2014a
Alium monanthum	Dimethyl trisulfide; dimethyl tetrasulfide	Aedes aegypti	Moon, 2011
Anacardiaceae
Pistacia terebinthus	α-Pinene; cyclopentane	Culex quinquefasciatus	Cetin et al., 2011
Spondias purpurea	Caryophyllene oxide and α-cadinol	Aedes aegypti	Lima et al., 2011
Annonaceae
Cananga odorata	Benzyl acetate, linalool, methyl benzoate	Aedes aegypti	Vera et al., 2014
Guatteria blepharophylla	Caryophyllene oxide	Aedes aegypti	Aciole et al., 2011
Guatteria friesiana	β-Eudesmol	Aedes aegypti	Aciole et al., 2011
Guatteria hispida	β-Pinene and α-pinene	Aedes aegypti	Aciole et al., 2011
Rollinia leptopetala	Spathulenol	Aedes aegypti	Feitosa et al., 2009
Apiaceae
Angelica purpuraefolia	4’-Chloro-4,4-dimethyl-3-(1-imidazolyl)-valerophenone, 1-Dodecanol,	Aedes aegypti	Nagella et al., 2012
Anethum graveolens	Limonene, carvone	Aedes albopictus	Seo et al., 2015
Apium graveolens	R-+-Limonene	Aedes aegypti	Pitasawat et al., 2007
	Limonene, carvone	Aedes albopictus	Seo et al., 2015
Bupleurum fruticosum	α-Pinene; β-pinene	Culex pipiens	Evergetis et al., 2009
Carum carvi	Carvone	Aedes aegypti	Pitasawat et al., 2007
Conopodium capillifolium	α-Pinene; sabinene	Aedes aegypti	Evergetis et al., 2009
Coriandrum sativum	Linalool, 2,6-octadien-1-ol, 3,7- dimethyl-, acetate, E-	Aedes aegypti	Nagella et al., 2012
Cuminum cyminum	ρ-cymene, β-pinene, cuminaldehyde	Aedes albopictus	Seo et al., 2015
Daucus carota	Carotol	Aedes albopictus	Seo et al., 2015
Elaeoselinum asclepium	α-Pinene; sabinene	Aedes aegypti	Evergetis et al., 2009
Foeniculum vulgare	trans-Anethole, Limonene	Aedes aegypti	Rocha et al., 2015
Heracleum pastinacifolium	Octyl acetate, Hexyl	Aedes aegypti	Tabanca et al., 2012a
Ligusticum chuanxiong	octadecenoic acids	Aedes aegypti, Culex quinquefasciatus	Evergetis et al., 2009
Oenanthe pimpinelloides	γ-Terpinene; o-cymene	Aedes aegypti	Pavela, 2015
Pimpinella anisum	Trans-anethole, α-Pinene; sabinene, β-phellandrene	Aedes aegypti	Pavela, 2015
Petroselinum crispum	β-phellandrene,myristicin, α & β-pinene, myrcene	Anopheles culicifacies	Evergetis et al., 2012
Pe. Sativum	Myristicin,1,8-cineole, 1,3,8-p-menthatriene	Aedes albopictus	Seo et al., 2015
Trachyspermum ammi	Thymol	Anopheles stephensi	Pandey et al., 2009
	ρ-Cymene, γ-Terpinene	Aedes albopictus	Seo et al., 2015
Apocynaceae
Cionura erecta L.	Edren-9-one, alpha cadinol, eugenol and alpha muurolene	Anopheles stephensi	Mozaffari et al., 2014
Araliaceae
Dendropanax morbifera	γ-Elemene	Aedes aegypti	Chung et al., 2009
Aristolochiaceae
Aristolochia indica	Aristolochic acid I and II	Aedes aegypti	Pradeepa et al., 2015
Asarum heterotropoides	Methyleugenol and safrole	Aedes aegypti	Perumalsamy et al., 2009
Asteraceae
Achillea millefolium	Eucalyptol, β-pinene, borneol, sabinene, camphene	Aedes albopictus	Conti et al., 2010
Artemisia absinthium	(Z)-β-ocimene, (E)-β-farnesene (Z)-en-yn-dicycloether	Aedes aegypti, Culex quinquefasciatus, Anopheles stephensi	Govindarajan and Benelli, 2016
Ar. dracunculus	Hexanal, isovaleric acid, (Z)-3-hexenol,	Anopheles stephensi	Pour et al., 2016
	Hexadecanol
Artemisia vulgaris	Camphor, Linalool, terpenen-4-ol, a-and bthujone, b-pinene	Aedes aegypti	Bora and Sharma, 2011
	camphor, alpha-thujone, betacaryophyllene, gammamuurolene, camphene
Artemisia vulgaris	Myrcene, limonene, cineol	Aedes aegypti	Sujatha et al., 2013
Ar. Nilagirica	Capillin	Aedes aegypti, Aedes albopictus	Bora and Sharma, 2011
Blumea densiflora	Borneol, germacrene D, β-caryophyllene, γ-terpinene, sabinene, β-bisabolene	Anopheles anthropophagus	Zhu and Tian, 2011
Blumea mollis	Linalool, γ-elemene, copaene, estragole, Allo-ocimene, γ-terpinene Alloaromadendrene	Culex quinquefasciatus	Senthilkumar et al., 2008
Chamaemelum nobile	α-pinene	Aedes aegypti, Culex quinquefasciatus	Amer and Mehlhorn, 2006
Chrysanthemum indicum	verbenol, 1,8-cineole, α-pinene, camphor, borneol, bornyl acetate	Aedes aegypti	Shunying et al., 2005
		Aedes aegypti	Wu et al., 2010
Eupatorium betonicaeforme	β-Caryophyllene	Aedes aegypti	Albuquerque et al., 2004
Matricaria recutita	α-bisabolol	Aedes aegypti	Heuskin et al., 2009
Pectis oligocephala	p-Cymene and thymol	Aedes aegypti	Albuquerque et al., 2007
Tagetes erecta	Piperitone	Aedes aegypti	Marques et al., 2011
Tagetes filifolia	trans-Anethole	Aedes aegypti	Ruiz et al., 2011
Tagetes lucida	Methyl chavicol	Aedes aegypti	Vera et al., 2014
Tagetes minuta	Trans-ocimenone	Aedes aegypti	Ruiz et al., 2011
Tagetes minuta	5E-ocimenone	Aedes aegypti	Maradufu et al., 1978
Tagetes patula	Limonene and terp	Aedes aegypti	Dharmagadda et al., 2005
Bignoniaceae
Cybistax antisyphilitica	quinone	Aedes aegypti	Rodrigues et al., 2005
Boraginaceae
Auxemma glazioviana	α-Bisabolol, α-cadinol, and T-muurolol	Aedes aegypti	Costa et al., 2004
Cordia curassavica	Cordiaquinones J and K	Aedes aegypti	Ioset et al., 2000
	α-Pinene	Aedes aegypti	Santos et al., 2006
Cordia leucomalloides	δ-Cadinene and E- caryophyllene	Aedes aegypti	Santos et al., 2006
Cucurbiataceae
Bryonopsis laciniosa	Goniothalamin	Culex pipiens	Kabir et al., 2003
Cupressaceae
Callitris glaucophylla	Guaiol & citronellic acid	Aedes aegypti	Shaalan et al., 2006
Chamaecyparis formosensis	Myrtenol	Aedes aegypti	Kuo et al., 2007
Cryptomeria japonica	16-Kaurene and elemol	Aedes aegypti, Aedes albopictus	Cheng et al., 2009c
Cunninghamia konishii	Cedrol, α-Pinene	Aedes aegypti	Cheng et al., 2013
Cupressus arizonica var. glabra	α-Pinene & epi-zonarene	Aedes aegypti	Ali et al., 2013
Cupressus arizonica	Limonene, umbellulone α-pinene	Anopheles stephensi	Sedaghat et al., 2011
Cupressus benthamii	Limonene; umbellulone	Aedes albopictus	Giatropoulos et al., 2013
Cupressus macrocarpa	Sabinene; α-Pinene; terpinen-4-ol	Aedes albopictus	Giatropoulos et al., 2013
Cupressus sempervirens	α-Pinene; δ-3-carene	Aedes albopictus	Giatropoulos et al., 2013
Cupressus torulosa	α-Pinene; δ-3-carene	Aedes albopictus	Giatropoulos et al., 2013
Chamaecyparis	Myrtenol; myrtenal	Aedes aegypti, Aedes aegypti	Kuo et al., 2007
formosensis	Limonene; oplopanonyl acetate; beyerene	Aedes albopictus	Giatropoulos et al., 2013
Chamaecyparis lawsoniana	α-Pinene; sabinene; δ-3-carene	Culex pipiens	Vourlioti-Arapi et al., 2012
Juniperus communis ssp.
Hemisphaerica	α-Pinene; limonene	Culex pipiens	Vourlioti-Arapi et al., 2012
Juniperus drupacea	Sabinene; 4-methyl-1-1-methylethyl-3-cyclohexen-1-ol	Culex pipiens	Vourlioti-Arapi et al., 2012
Juniperus foetidissima	Myrcene; germacrene-D; α-Pinene	Culex pipiens	Vourlioti-Arapi et al., 2012
Juniperus oxycedrus L. ssp.
oxycedrus	α –pinene	Culex pipiens	Vourlioti-Arapi et al., 2012
Juniperus oxycedrus L.
subsp. Macrocarpa	α-Pinene; δ-3-carene; β-phellandrene; α-terpinyl acetate	Aedes albopictus	Giatropoulos et al., 2013
Juniperus phoenicea
Tetraclinis articulate	α-Pinene; bornyl acetate	Aedes albopictus	Giatropoulos et al., 2013
Dioncophyllaceae
Triphyophyllum peltatum	dioncophylline A	Anopheles stephensi	François et al., 1996
Euphorbiaceae
Croton nepetaefolius	Methyleugenol	Aedes aegypti	Morais et al., 2006
Croton regelianus	Ascaridole & p-Cymene	Aedes aegypti	Torres et al., 2008
Croton zehntneri	E-anethole, p-anisaldehyde	Aedes aegypti	Morais et al., 2006
Fabaceae
Copaifera multijuga	β-caryophyllene	Anopheles darling, Aedes aegypti	Trindade et al., 2013
Hymenaea courbaril	α-Copaene, spathulenol	Aedes aegypti	Aguiar et al., 2010
	Germacrene D and β-caryophyllene
Myroxylon pereirae	Benzyl benzoate	Aedes aegypti	Yenesew et al., 2003
Millettia dura	Rotenoids, deguelin and tephrosin caryophyllene oxide; phenol,4-3,7-dimethyl-3-ethenylocta-1,6-dienyl; caryophyllene	Culex quinquefasciatus	Dua et al., 2013
Psoralea corylifolia	Citronellol	Aedes aegypti	Benelli et al., 2017
Geraniaceae		Culex quinquefasciatus	Cavalcanti et al., 2004
Pelargonium graveolens	Neral; geranial	Culex quinquefasciatus
Gramineae	α-Pinene	Aedes aegypti	Cetin et al., 2011
Cymbopogon citratus	Thymol	Culex pipiens	Govindarajan et al., 2013
Hypericaceae
Hypericum scabrum	Δ-3-carene, 1,8-cineole, β-caryophyllene, bicyclogermacrene	Culex tritaeniorhynchus, Aedes albopictus, and Anopheles subpictus	Araújo et al., 2003
Lamiaceae
Coleus aromaticus	β-caryophyllene, bergamotene, and terpinolene	Aedes aegypti	Jaenson et al., 2006
Hyptis martiusii
Hyptis suaveolens
Lavandula gibsoni	α-Terpinolen and thymol	Aedes aegypti, Anopheles stephensi	Kulkarni et al., 2013
		Culex quinquefasciatus.
Lavandula stoechas	Fenchone, 1,8-Cineole	Culex pipiens	Traboulsi et al., 2002
Lippia origanoides	Carvacrol	Aedes aegypti	Mar et al., 2018
Mentha longifolia	Piperitenone oxid	Aedes aegypti	Pavela et al., 2014
M. microcorphylla	Piperitenone, Pulegone, Piperitenone oxide	Culex pipiens	Traboulsi et al., 2002
M. spicata	Carvone	Aedes aegypti	Govindarajan et al., 2012
Nepeta cataria	E,Z-Nepetalactone and Z, E-nepetalactone	Aedes aegypti	Zhu et al., 2006
Ocimum americanum	E-Methyl-cinnamate	Aedes aegypti	Cavalcanti et al., 2004
Ocimum basilicum	Linalool; methyl eugenol	Aedes aegypti	Govindarajan et al., 2013
Ocimum gratissimum	Eugenol	Aedes aegypti	Cavalcanti et al., 2004
Ocimum sanctum	Methyleugenol	Culex pipiens	Gbolade and Lockwood, 2008
O. syriacum	Carvacrol, Thymol	Aedes aegypti	Traboulsi et al., 2002
Perilla frutescens	oleic, S-limonene, perillaldehyde	Aedes aegypti	Pohlit et al., 2011
Plectranthus amboinicus	Carvacrol	Aedes aegypti	Lima et al., 2011
Plectranthus mollis	Piperitone oxide, fenchone	Aedes aegypti	Kulkarni et al., 2013
Pogostemon cablin	Patchouli alcohol, Seyshellene, α-bulnesene, Norpatchoulenol	Aedes aegypti	Lima-Santos et al., 2019
Pulegium vulgare	Pulegone; carvone	Aedes albopictus	Pavela, 2015
Rosmarinus officinalis	1,8-Cineole; camphor	Aedes aegypti	Giatropoulos et al., 2018
Satureja hortensis	γ-Terpinene; carvacrol	Culex pipiens	Pavela, 2009
Thymus capitatus (L.)	Thymol, alpha-Amyrin, Carvacrol + beta-		Mansour et al., 2000
Hoffm. & Link	Caryophyllene	Culex pipiens
Thymus leucospermus	p-Cymene	Culex pipiens	Pitarokili et al., 2011
Thymus satureoides	Thymol; borneol	Culex pipiens	Pavela, 2009
Thymus teucrioides	p-Cymene; γ-terpinene; thymol	Aedes albopictus	Pitarokili et al., 2011
Thymus vulgaris	a-terpinene,carvacrol, thymol		Giatropoulos et al., 2018
	p-cymene, linalool, geraniol	Aedes aegypti
Vitex agnus castus	Trans-caryophyllene; 1,8 cineole	Culex quinquefasciatus	Niroumand et al., 2018
Vitex trifolia	Methyl-p-hydroxybenzoate	Aedes aegypti	Kannathasan et al., 2011
Lauraceae
Cinnamomum camphora	1,8-Cineole	Anopheles sinensis	Zhang et al., 2018
C. cassia	Cinnamaldehyde	Aedes aegypti	Zhu et al., 2006
C. impressicostatum	Benzyl benzoate and α-phellandrene	Aedes aegypti	Jantan et al., 2005
C. japonicum	Borneol	Anopheles sinensis	Zhang et al., 2018
C. microphyllum	Benzyl benzoate	Aedes aegypti	Jantan et al., 2005
C. mollissimum	Benzyl benzoate	Aedes aegypti	Jantan et al., 2005
C. osmophloeum	trans-Cinnamaldehyde and cinnamyl acetate	Aedes aegypti	Cheng et al., 2004
C. pubescens	Benzyl benzoate	Aedes aegypti	Jantan et al., 2005
C. rhyncophyllum	Benzyl benzoate	Aedes aegypti	Jantan et al., 2005
C. scortechinii	β-Phellandrene and linalool	Aedes aegypti	Jantan et al., 2005
C. sintoc	Safrole	Aedes aegypti	Jantan et al., 2005
C. subavenium	Eugenol	Anopheles sinensis	Zhang et al., 2018
C. szechuanense	1,8-Cineole	Anopheles sinensis	Zhang et al., 2018
Laurus nobilis	1,8-cineole, linalool	Culex pipiens	Patrakar et al., 2012
Lindera obtusiloba	α-Copaene; β-caryophyllene	Aedes aegypti	Pavela, 2015
Magnoliaceae
Magnolia salicifolia	Trans-anethole, Methyl eugenol, isomethyl eugenol, Costunolide, lactone and parthenolide	Aedes aegypti	Kelm et al., 1997
Malvaceae
Abutilon indicum	β-sitosterol	Aedes aegypti,	Rahuman et al., 2008a
	Azadirachtin, salannin, deacetylgedunin, gedunin, 17-hydroxyazadiradione and deaceytlnimbin
Meliaceae
Azadirachta indica	Saponins	Anopheles stephensi,	Senthil-Nathan et al., 2005a
	23-O-methylnimocinolide
	6α-O-acetyl-7-deacetylnimocinol	Culex quinquefasciatus	Ansari et al., 2005
	Nimocinolide; 7-O-deacetyl-23-O-methyl-	Aedes aegypti	Siddiqui et al., 1999
	7α-O-senecioylnimocinolide		Banerji and Nigam, 1984
	desfurano-6α-hydroxyazadiradione		Naqvi, 1987
	22,23-dihydronimocinol	Aedes aegypti	Siddiqui et al., 2002
	1α-acetyl-3α-propionylvilasinin	Aedes aegypti	Siddiqui et al., 2003
	Meliatetraolenone	Aedes aegypti	Siddiqui et al., 2003
	azadirachtin, salannin, deacetylgedunin,	Culex quinquefasciatus	Siddiqui et al., 2003
	gedunin, 17- hydroxyazadiradione
	deacetylnimbin	Anopheles stephensi	Senthil-Nathan et al., 2005a
	3β,24,25-trihydroxycycloartane	Anopheles stephensi
Dysoxylum malabaricum	Beddomei lactone	Aedes aegypti	Senthil-Nathan et al., 2009
D. beddomei	Caryophyllene epoxide	Aedes aegypti	Senthil-Nathan et al., 2009
	cis-Caryophyllene
Guarea humaitensis	1α,7α,11β-triacetoxy-4α-carbomethoxy-	Aedes aegypti	Magalhães et al., 2010
G. scabra	12α-(2-methylpropanoyloxy)-14β,15β-epoxyhavanensin	Aedes aegypti	Magalhães et al., 2010
Turraea floribunda	1α,11β-diacetoxy-4α-carbomethoxy-7α-	Aedes aegypti	Ndung’u et al., 2004
	hydroxy-12α-(2-methylpropanoyloxy)-15-	Aedes aegypti	Ndung’u et al., 2004
	oxohavanensin; 1α-acetyl-3α-	Culex pipiens	Ndung’u et al., 2004
	propionylvilasinin	Culex pipiens	Ndung’u et al., 2003
Turraea wakefieldii	11β,12α-diacetoxyneotecleanin	Culex pipiens	Ndung’u et al., 2003
	11β,12α-diacetoxy-14β,15β-	Aedes aegypti	Ndung’u et al., 2003
	epoxyneotecleanin	Aedes aegypti	Ndung’u et al., 2003
Myrtaceae
Eucalyptus benthamii	α-Pinene	Aedes aegypti	Lucia et al., 2012
E. botryoides	p-Cymene, α-eudesmol, and 1,8-cineol	Aedes aegypti	Lucia et al., 2012
E. camaldulensis	1,8-Cineol, p-cymene and β-phellandrene	Aedes aegypti	Lucia et al., 2008
E. citriodora	Citronellal; citronellol;	Aedes aegypti	Vera et al., 2014
	α-humulene isopulegol
E. dunnii	1,8-Cineol and γ-terpinene	Aedes aegypti	Lucia et al., 2008
E. fastigata	p-Cymene	Aedes aegypti	Lucia et al., 2012
E. globulus	1,8-Cineol	Aedes aegypti Anopheles arabiensis	Massebo et al., 2009
E. grandis	α-Pinene	Aedes aegypti	Lucia et al., 2007
E. gunnii	1,8-Cineol and p-cymene	Aedes aegypti	Lucia et al., 2008
E. nobilis	1,8-Cineol	Aedes aegypti	Lucia et al., 2012
E. radiata	1,8-Cineol	Aedes aegypti	Lucia et al., 2012
E. robusta	α-Pinene	Aedes aegypti	Lucia et al., 2012
E. saligna	1,8-Cineol and p-cymene	Aedes aegypti	Lucia et al., 2008
E. tereticornis	β-Phellandrene and 1,8-cineol	Aedes aegypti	Lucia et al., 2008
E. urophylla	1,8-Cineol	Aedes aegypti	Cheng et al., 2009b
E. melanadenia	1,8-Cineol	Aedes aegypti	Aguilera et al., 2003
Myrtus communis	1,8 Cineole, α-Pinene, Linalool	Culex quinquefasciatus	Traboulsi et al., 2002
M. dissitiflora	Terpinen-4-ol	Aedes aegypti	Park et al., 2011
M. leucadendron	1,8-Cineol, α-pinene, and α-terpineol	Aedes aegypti	Leyva et al., 2008
M. linariifolia	Terpinem-4-ol and γ-terpinene	Aedes aegypti	Park et al., 2011
M. quinquenervia	1,8-Cineol and E-nerolidol	Aedes aegypti	Park et al., 2011
Pimenta dioica	Eugenol, linalool	Aedes aegypti	Pereira et al., 2014
P. racemosa	Terpinem-4-ol and 1,8-cineol	Aedes aegypti	Aciole, 2009
P. guajava	1,8-Cineol and β-caryophyllene	Culex pipiens	Leyva et al., 2009a
	1,8-Cineol	Aedes aegypti	Lima et al., 2011
P. rotundatum	Eugenol	Aedes aegypti	Aguilera et al., 2003
Syzygium aromaticum	Eugenol	Aedes aegypti	Costa et al., 2005
Orchidaceae
Vanilla fragrans	4-ethoxymethylphenol, 4-butoxymethylphenol, vanillin, 4-hydroxy-2-methoxycinnamaldehyde and 3,4-dihydroxyphenylacetic acid	Culex pipiens	Sun et al., 2001
Pinaceae
Cupressus L.,	limonene, α & β-pinene,	Aedes aegypti	Burfield, 2000
Juniperus L.	3-carene	Aedes aegypti	Burfield, 2000
Pinus brutia	α-Pinene and β-pinene	Aedes albopictus	Koutsaviti et al., 2015
P. halepensis	β-Caryophyllene	Aedes albopictus	Koutsaviti et al., 2015
P. kesiya	α-Pinene, β-pinene, myrcene and germacrene D.	Aedes aegypti, Culex quinquefasciatus,	Govindarajan et al., 2016
		Anopheles stephensi
P. longifolia	k-terpineol	Culex quinquefasciatus, Anopheles	Ansari et al., 2005
		culicifacies
P. stankewiczii	Germacrene D α-Pinene and β-pinene	Aedes albopictus	Koutsaviti et al., 2015
P. sylvestris	Eugenol 3, Cyclohexene-1-methanol, α-4-	Aedes aegypti, Culex quinquefasciatus	Fayemiwo et al., 2014
	trimethyl
Piperaceae
Piper auritumP. betleP. capenseP. decurrens	SafroleCitronellal2,3-Dihydro-2-(4′-hydroxyphenyl)-3-methyl-5(E)-propenylbenzofuran (conocarpan), 2-(4′-hydroxy-3′-methoxyphenyl)-3-methyl-5(E)-propenylbenzofuran (eupomatenoid-5), 2-(4′-hydroxyphenyl)-3-methyl-5(E)-propenylbenzofuran (eupomatenoid-6), 2,3-dihydro-5-formyl-2-(4′-hydroxyphenyl)-3-methylbenzofuran (decurrenal), and 3,7,11,15-tetramethyl-2(E)-hexadecen-1-ol (trans-phytol)	Aedes aegyptiAedes aegyptiAedes atropalpusAedes aegypti	Leyva et al., 2009bWahyuni, 2012Chauret et al., 1996de Morais et al., 2007
P. gaudichaudianum	Caryophyllene oxide, β-selinene	Aedes aegypti	de Morais et al., 2007
P. hostmanianum	Asaricin and myristicin	Aedes aegypti	de Morais et al., 2007
P. humaytanum	β-selinene, caryophyllene oxide	Aedes aegypti	de Morais et al., 2007
P. klotzschianum	1-Butyl-3,4-methylenedioxybenzene,	Aedes aegypti	do Nascimento et al., 2013
P. longum	limonene, and α-phellandrene	Culex pipiens	Lee, 2000
	Pipernonaline	Aedes aegypti	Yang et al., 2002
		Aedes aegypti	Costa et al., 2004
P. marginatum	Isoelemecin, apiole	Aedes aegypti	Autran et al., 2009
	(Z)-Asarone	Aedes aegypti	Autran et al., 2009
P. permucronatum	(E)-Asarone, patchouli alcohol	Aedes aegypti	de Morais et al., 2007
	Dillapiole and myristicin
Plumbaginaceae
Plumbago zeylanica	Plumbagin	Aedes aegypti	Pradeepa et al., 2016
Poaceae
Cymbopogon citratus	Geranial	Aedes aegypti	Cavalcanti et al., 2004
Cymbopogon flexuosus	citral a-pinene	Aedes aegypti	Syed and Leal, 2008
Cymbopogon nardus	Geranial; neral	Aedes aegypti	Vera et al., 2014
	Girgensohnine	Aedes aegypti	Carreño-Otero et al., 2018
Vetiveria zizanioides	Citronellal	Aedes aegypti	Fradin and Day, 2002
	khusimol, isonootkatool, β-vetivenene, α &	Aedes aegypti	Vera et al., 2014
	β-vetivones
Papilonaceae
Neorautanenia mitis	Neotenone, neorautanone, pterocarpans neoduline, nepseudin,4-methoxyneoduline	Culex quinquefasciatus, Anopheles gambiae	Joseph et al., 2004
		Aedes aegypti, Aedes albopictus
	Elemol, Eudesmols	Culex quinquefasciatus	Zhu et al., 2006
Rutaceae
Chloroxylon swietenia	Heptacosanoic acid	Aedes aegypti, Culex quinquefasciatus	Balasubramani et al., 2015
Citrus aurantifolia	Geijerene, Limonene, Germacrene D	Aedes aegypti, Anopheles stephensi	Kiran et al., 2006
Citrus hystrix	α-terpineol
Citrus limon	β-Pinene; d-limonene; terpinene-4-ol	Culex pipiens	Sutthanont et al., 2010
	Limonene	Aedes aegypti
Citrus reticulata	D-Limonene; γ-terpinene	Culex quinquefasciatus	Michaelakis et al., 2009
Citrus sinensis	Limonene	Aedes aegypti	Sutthanont et al., 2010
	Limonin, Nomilin, Obacunone	Culex quinquefasciatus	Jayaprakasha et al., 1997
	Geijerene; limonene; germacrene D	Aedes aegypti	Vera et al., 2014
Chloroxylon swietenia
			Kiran et al., 2006
Clausena excavate	Safrole and terpinolene	Aedes aegypti, Aedes albopictus	Cheng et al., 2009a
Feronia limonia	Estragole and β-pinene	Aedes aegypti	Senthilkumar et al., 2013
F. limonia	n-hexadecanoic acid	Culex quinquefasciatus	Rahuman et al., 2000
Ruta graveolens	Undecan-2-one	Aedes aegypti	Tabanca et al., 2012b
Swinglea glutinosa	β-Pinene; piperitenone;	Aedes aegypti	Vera et al., 2014
	α-Pinene
Toddalia asiatica	Linalool	Aedes aegypti	Nyahanga et al., 2010
Zanthoxylum armatum	Linalool	Aedes aegypti	Tiwary et al., 2007
Z. articulatum	Viridiflorol	Aedes aegypti	Feitosa et al., 2007
Z. avicennae	1,8-Cineole	Aedes albopictus	Liu et al., 2014b
	Limonene	Aedes aegypti	Pitasawat et al., 2007
	Methyl heptyl ketone	Aedes aegypti	Borah et al., 2012
Z. piperitum	Asarinin, sesamin and (+)-xanthoxylol-γ,γ-dimethylallylether	Aedes aegypti, Culex pipiens	Kim and Ahn, 2017
Z. monophyllum	Germacrene D-4-ol and a-Cadinol	Aedes albopictus, Culex quinquefasciatus, Anopheles stephensi	Pavela and Govindarajan, 2017
Santalaceae
Santalum L. spp.	α-santalol	Aedes aegypti, Culex pipiens	Jones et al., 2007
Santalum album	Guaiol, elemol, and eudesmol	Anopheles stephensi,	Amer and Mehlhorn, 2006
Schisandraceae		Aedes aegypti
Illicium verum	Eugenol, α-Terpinyl acetate, Eucalypt, ol, (E)-anethole	Culex quinquefasciatus	Kimbaris et al., 2012
Scrophulariaceae
Capraria biflora L.	α-Humulene	Aedes aegypti	Souza et al., 2012
Stemodia maritima	β-Caryophyllene and caryophyllene oxide	Aedes aegypti	Arriaga et al., 2007
Tiliaceae
Microcos paniculata	N-Methyl-6b-(deca-l’,3’,5’-trienyl)-3b-methoxy-2bmethylpiperidine	Aedes aegypti	Bandara et al., 2000
Verbenaceae
Duranta repens	β-amyrin and 12-oleanene 3β, 21β-diol,	Culex quinquefasciatus	Nikkon et al., 2010
Lantana camara	Bicyclogermacrene and E-caryophyllene	Aedes aegypti	Costa et al., 2010
	Eucalyptol, caryophyllene,
Lippia alba	Carvone; limonene	Aedes aegypti	Santiago et al., 2006
L. gracilis	Carvacrol	Aedes aegypti	Santiago et al., 2006
L. origanoides	Carvacrol; p-cymene	Aedes aegypti	Vera et al., 2014
L. javanica	Allopurinol,camphor, Limonene, a –terpeneol, verbenone	Aedes aegypti	Mwangi et al., 1992
L. microphylla	1,8-cineole, thymol, α-pinene	Aedes aegypti	Santiago et al., 2006
L. nodiflora	Camphor, p-cymene, γ−terpinene	Aedes aegypti	Santiago et al., 2006
L. sidoides	Thymol	Aedes aegypti	Costa et al., 2005
Zingiberaceae
Alpinia purpurata	β-Caryophyllene and β-pinene	Aedes aegypti	Santos et al., 2012
Curcuma aromatic	1H-3a,7-Methanoazulene and curcumene	Aedes aegypti	Choochote et al., 2005
	Turmerone, curcumene, and zingiberene
Curcuma longa	1,8-Cineol and p-cymene	Aedes aegypti	Leyva et al., 2008
Curcuma zedoaria	Dodecanal	Aedes aegypti	Pitasawat et al., 2007
Hedychium coccineum	1,8-Cineol and β-pinene	Aedes aegypti	Sakhanokho et al., 2013
Hedychium sp.	1,8-Cineol	Aedes aegypti	Sakhanokho et al., 2013
Kaempferia galanga	Ethyl trans-p-methoxycinnamate	Aedes aegypti	Munda et al., 2018
Kaempferia galanga	Ethyl cinnamate	Aedes aegypti	Munda et al., 2018
Zingiber officinale	4-Gingero	Aedes aegypti, Culex quinquefasciatus	Rahuman et al., 2008b
Zingiber officinale	6-Dehydrogingerdione	Aedes aegypti, Culex quinquefasciatus	Rahuman et al., 2008b
Zingiber officinale	6-Dihydrogingerdione	Aedes aegypti, Culex quinquefasciatus	Rahuman et al., 2008b
Zingiber zerumbet	α-Humulene; zerumbone	Aedes aegypti	Sutthanont et al., 2010

Besides secondary metabolites, essential oils (EOs) from plants were also recorded with effective mosquitocidal potentials. The EOs from the plants of Lamiaceae and Zingiberaceae were proved with bioactivity against Ae. aegypti (Kalaivani et al., 2012). The fern Actiniopteris radiata was testified with novel mosquitocidal activity against larvae of Ae. aegypti and An. Stephensi (Kamaraj et al., 2018). The seed oil extract of Acacia nilotica possessed robust larvicidal action against major mosquito vectors (Vivekanandhan et al., 2018a). A remarkable biological activity of EOs against Dengue vectors has been extensively reviewed by Chellappandian et al. (2017, 2018, 2019). Plant volatile oils were also conveyed with mosquitocidal potentials. As studied by Vasantha-Srinivasan et al. (2018), the crude volatile oil (CVO) from Piper beetle leaves possessed significant larvicidal, ovipositional, and repellency effects against Ae. Aegypti.

Derivatives of plants are enriched with active molecules with exceptional mosquitocidal properties and can be advanced as low cost environmentally friendly bio-pesticides. Many botanical extracts along with their chief constituents showed effective insect metabolism inhibition or stimulation of digestive enzymes (Senthil-Nathan et al., 2009; Napoleão et al., 2012; Senthil-Nathan, 2013). Unlike synthetic chemicals, previous literature on plant compounds doesn’t provide any indication for the emergence of resistance so far. This is most likely due to the blend of several bioactive compounds with different mechanisms of action and therefore it is difficult for mosquito vectors to develop resistance (Mulla and Su, 1999; Shaalan et al., 2005).

Impact of Phytochemicals on the Physiology of Mosquito Larvae

As in general, plant secondary metabolites are evolved as protection mechanism against herbivory. When these toxic substances are encountered by the mosquitoes, a relatively unambiguous response is triggered that has a non-specific influence on a wide range of molecular targets such as proteins, nucleic-acids, bio-membranes, besides added cellular components. Consequently, the physiology is disrupted at numerous receptor sites, eventually causing an abnormality in the nervous system. Plant metabolites affect several vital physiological functions that include inhibition of “AChE” as well as “GABA-gated” chloride channel, disruption of Na–K ion exchange besides constricting the cellular respiration. As a subsequent event, the alteration of these enzyme levels gives rise to several anomalies that include the obstruction of nerve cell membranes and octopamine receptors along with calcium channel blockage, resulting in hormonal imbalance, mitotic poisoning, and also modifications of the molecular basis of morphogenesis (Rattan, 2010).

Synthetic insecticides generally increase the level of detoxifying enzymes. Phytochemicals target the mentioned cellular mechanisms and potentially disturb their functions (Figure 1; Zibaee and Bandani, 2010; Zibaee, 2011; Kaur et al., 2014; Senthil-Nathan, 2015). Physiological effects of phytochemicals are discussed below.

FIGURE 1

Mode of action of phytochemicals in insect body (modified after Ghosh et al., 2012).

Impact of Phytochemicals on Detoxifying Enzymes

The antioxidant and detoxification enzymes of mosquito vectors are vital in detoxification of reactive oxygen species (ROS) synthesized by the toxic chemicals (Rattan, 2010). Esterase and phosphatase of the mosquito vectors plays a key role in several physiological events (Koodalingam et al., 2014). Excessive usage of toxic chemicals on mosquito control caused insecticide resistance through sodium channel mutations, activation of detoxification enzymes, and upregulation of key genes and other regulatory components like MicroRNAs (miRNAs). The CYP450s, GSTs, SOD, and esterase gene families are recognized as the foremost four enzymes accountable for the metabolic-resistance of the insects (Hemingway et al., 2004). Generally, detoxifying enzymes are involved in digestion, reproduction, juvenile hormone metabolism, neuronal conduction, moulting, and more importantly detoxification of toxic chemicals (Koodalingam et al., 2014). Phosphatases are involved in tissue development, cellular differentiation, carbohydrate metabolisms, and synthesis of ATP (Koodalingam et al., 2014). Mainly these two major classes of detoxifying enzymes are considered for evaluating the impact of toxic chemicals on physiological or biochemical events of arthropod vectors.

Carboxyl-esterases (EC3.1.1.1) are non-specific omnipresent enzymes that are associated to the major “endogenous” functions in insects, which hydrolyze a different carboxylic-acid ester (Lija-Escaline et al., 2015). Generally, the metabolic pathway of these enzymes was targeted by the chemical pesticides, especially the fourth generation class of Pyrethroids, which acts on the voltage sensitive sodium channels and blocks the mosquito nervous system (Hong et al., 2014). Esterases can also target by sequestering the insecticide through rapid binding and slowly releasing the insecticide metabolites (Karunaratne et al., 1993). This latter type of resistance requires the presence of increased quantities of esterase due to the 1:1 stoichiometry of the reaction and decreases the metabolic breakdown time.

Plant extracts and their derivatives have been widely reported to decrease the levels of carboxylesterase (α- β-carboxylesterase) level in the Ae. aegypti larva (Koodalingam et al., 2014; Lija-Escaline et al., 2015). Besides exhibiting larvicidal activity Alangium salvifolium, also substantially reduced the levels of α, β-carboxylesterase as well as superoxide dismutase (SOD) in Ae. Aegypti (Thanigaivel et al., 2017a). Myrrh commiphora molmol (oil and oleo-resin extract) instigated biochemical changes in Cx. pipiens that affected the cell proteins, as well as loss of enzyme activity (Massoud et al., 2001).

Higher rates of enzyme activities, such as SOD (Agra-Neto et al., 2014; Lija-Escaline et al., 2015) and physiological enzymes like esterase (Wheelock et al., 2005; Lija-Escaline et al., 2015), phosphatases (Walter and Schütt, 1974; Urich, 1994) are recorded with increasing developmental stages and these are considered responsible for increased pyrethroid resistance. The Mosquito vectors that established resistance to Temephos have been found to possess genes that insensitized ACHE on exposure to pesticides. Insects were also characterized by the over expression of varied forms of detoxifying enzymes (GST, SOD, and esterases) (Larson et al., 2010).

Glutathione-S-transferases are a class of detoxification enzymes considered to play a vital role in the existence of insects exposed to toxic metabolites. Increased GST activities are connected with the expression of metabolic resistance toward insecticides (Clark, 1990). GSTs can break down a broad range of substances; amplified GST activity is possibly as a response to an environmental stress. Generally, Cytochrome P450s (CYP450) displayed upregulation when induced by plant secondary metabolites in diverse insect pests especially against the vectors of human diseases (Caballero et al., 2008) and have members which are considered as major elements conferring resistance against insecticides (i.e., CYP2, CYP4, and CYP6) (Sun et al., 2001). The upregulation of GST enzymes usually at the exposure of a prominent dosage of plant compounds suggests the activity of a major detoxification process (Edwin et al., 2016). Consequently, the levels of GST expression may be used as a biomarker to detect the development of resistance (Jukic et al., 2007).

CYP450 group of enzyme family are also designated as key indicators of metabolic resistance besides susceptibility to insecticides (David et al., 2013). Many previous research outcomes proved alteration or inhibition in the expression of major detoxifying enzymes exposed to plant chemicals. Thanigaivel et al. (2017a) showed increase in the rate of GST activity in IV instar larvae of dengue mosquito exposed to methanolic leaf extract of J. adhatoda with their major derivative 3-hydroxy-2,3-dihydropyrrolo[2,1-b]quinazolin-9(1h) one (26.37%). Likewise, carboxylesterase activities differed significantly in Ae. aegypti post treatment with the leaf extracts of P. nigrum with their major derivatives thymol (20.77%) (Lija-Escaline et al., 2015). Correspondingly, the activity of major enzymes (esterases, GST, and CYP450) of dengue mosquito severely affected post treated with dynamic plant compound andrographolide derived from Andrographis paniculata (Acanthaceae) at the maximum dosage of 12 ppm (Edwin et al., 2016). DDT resistance in the mosquito An. gambiae is correlated elevated glutathione transferase (GST) E2 activity (AgGSTE2) (Enayati et al., 2005). The DDT resistant An. gambiae evades the insecticidal activity by the dehydrochlorination of DDT to its non-insecticidal metabolite DDE. Muleya et al. (2008) reported that compounds -epiphyllocoumarin (Tral-1), knipholone anthrone, isofuranonaphthoquinones (Mr 13/2, Mr13/4), and the polyprenylated benzophenone (GG1) were potent inhibitors of AgGSTE2.

Besides the botanical extracts, EO derived from the plants also have strong inhibition of detoxifying enzymes of arthropod vectors (Pavela, 2015). EOs may provide substitute sources of vector control since they are enriched with diverse phyto-molecules with insecticidal properties (Cheng et al., 2013). Insecticide phytochemicals from EOs belong to terpenoids chiefly and Phenylpropanoids to a limited extent. In which, Terpenoids includes monoterpenes and sesquiterpenes as the major compositions of EOs (Chellappandian et al., 2018). Lee et al. (2003) specified that volatile and lipophilic monoterpenoids infiltrate insect body, where they afflict physiological processes, and hence their mode of action is hard to elucidate. Previous research of Vasantha-Srinivasan et al. (2017) showed that the CVO derived from Piper betle (L.) (Pb-CVO) showed upregulation in the level GST and CYP450 and down regulate the expression of Carboxylesterases activity against the field and laboratory strains of Ae. aegypti. Moreover, the above results also showed that the changes in the level of enzymes are steady in both field and laboratory strains compared to the chemical pesticides. Due to enriched chemical diversity and potential mosquitocidal activity, CVO have acquired greater interest from researchers looking for new besides natural replacements to chemical-pesticides in controlling medically challenging pests (Pavela, 2015). Correspondingly, EO constituent’s nootkatone and carvacrol from Alaskan yellow cedar tree inhibits 50% of acetylcholine esterase activity in Ae. aegypti compared to the carbaryl, a known acetylcholinesterase inhibitor (Anderson and Coats, 2012). The impact of major plant molecules against the mosquito larvicides was tabulated (Table 1). Hence, expression of these molecules on detoxifying and metabolic enzymes is considered an important biomarker to evaluate the mosquitocidal potential of bio-rational plant metabolites.

Pradeepa et al. (2014) have reported the antimalarial activities from the compound plumbagin, identified from the rhizome of Plumbago zeylanica against An. stephensi. Also, it was revealed that plumbagin constrains the vector AchE enzyme, An. Stephensi in a dose dependent manner and also can be considered for controlling resistant vectors whose insecticide resistance is associated to an increased SOD activity (Pradeepa et al., 2016). The detection of SOD activity in the anal gills of An. stephensi larvae could be associated with their resistance provided against damaging oxygen products (Nivsarkar et al., 1991). The sensitivity of an insect to an insecticide can hence be increased by identifying certain compounds that can deactivate these enzymes (Larson et al., 2010).

Impact of Phytochemicals on Midgut Tissues

The midgut of the mosquito larvae is the chief interface of exterior environment and chip in major process like digestion, ion transport, absorption, and osmoregulation process (Bernick et al., 2008; Elumalai et al., 2016). Generally, gut region is the target of numerous insecticidal complexes and its integrity is dynamic for digestion and conferring of resistance against toxins (Stenfors Arnesen et al., 2008). With the insect midgut being the important site for synthesis of digestive enzymes, plant derived molecules primarily targets thee gut epithelium layer (EL) (Senthil-Nathan et al., 2008). This might be the significant cause for condensed metabolic rate in addition to a reduced enzyme activity (Selin-Rani et al., 2016). The peritrophic membrane (pM) gaurds the EL from the surrounding the gut lumen (GL) (Lija-Escaline et al., 2015). Phyto-chemicals are proven to exert a serious impact on the digestive epithelial cells and further decrease the growth rate of arthropods (Yu et al., 2015). Neira-Oviedo et al. (2008) stated that plant compounds flow into the gastric caeca and the malpighian tubules thereby affecting the midgut epithelium. For instance, extracts of M. azedarach have been reported to cause extensive harm on the EL and pM of filarial vector Cx. quinquefasciatus (Al-Mehmadi and Al-Khalaf, 2010). The pM may influence the growth and development of parasites vectors by creating a mechanical barrier to invasion by ookinetes (Rudin and Hecker, 1989). Plant extracts and their metabolites are crucial for the impairment of pest mid-gut epithelium (Rey et al., 1999). The compound catechin isolated from Leucas aspera affects the mid-gut of the three mosquito larvae Ae. aegypti, An. stephensi, and Cx. quinquefasciatus (Elumalai et al., 2016). Previous photomicrographic study on the midgut tissues of the dengue mosquito (Field and laboratory strains of Ae. aegypti) treated with the CVO of P. betle displayed severe injuries to the GL and EL (Vasantha-Srinivasan et al., 2018). Correspondingly, leaf extracts of Aristolochia indica L. (Aristolochiaceae) and their derivatives aristolochic acid I and II showed severe damage on the midgut vacuolated gut epithelial columnar cells (epi), GL, and pM (Pradeepa et al., 2015). Likewise, methanolic leaf extracts of P. nigrum severely affected the midgut cellular organelles of Ae. aegypti at the minimal dosage of 10 ppm (Lija-Escaline et al., 2015). Similarly, Vasantha-Srinivasan et al. (2018) reported that P. betle CVO derived from P. betle at the sub-lethal dosage damage the pM, and major alteration in the alignment of EL and GL of dengue mosquito comparable to the control. Previous research on Andrographolide a major derivative of A. paniculata against dengue mosquito gut cells proved that there was an unembellished collapse in the mid-gut pM, in addition to a chief variation in the El and GL alignment (Edwin et al., 2016). Selin-Rani et al. (2016) reported that the active plant molecules may damage the gut epithelium is the vital reason for concentrated metabolic rate and decrease in the enzyme-activity. Midgut cell damage is directly linked to the digestive and detoxifying enzymes dysregulation (Senthil-Nathan et al., 2008). This was also confirmed by histological studies of the mosquitoes that displayed midgut cell damage, post treatment with various botanical compounds (Yu et al., 2015). Further, treatment with plant compounds were also associated with altered protein (Fallatah, 2014) and biochemical profiles in mosquitoes (Senthilkumar et al., 2013).

Biochemical studies on Cx. pipipens exposed to Allium satvium, Citrus limon, and Bti were observed by Saeed et al. (2010). Results revealed that the use of plant oil extracts and Bti have great effect on total protein content of treated mosquito larvae. Fallatah (2010) reported the effect of water extract of fenugreek have high larvicidal effect against Cx. quinquefasciatus, causing noticeable effects on numerous body tissues together with the midgut and nervous system as well as total protein content. Aristolochic acids isolated from A. indica Linn, mainly affected the midgut EL and secondly the larval muscles and cells (Pradeepa et al., 2015). Similar results were also observed in mosquitoes treated with plant extracts (Costa et al., 2012). The orientation of the cytoplasmic protrusions of the apical surfaces of columnar cells toward the lumen suggests the secretion of apocrine and/or apoptosis.

Al-Mekhlafi (2018) reported the effect of Arum copticum (Apiaceae) extract against Culex pipiens larvae. Apart from exhibiting larvicidal activity, the extract was able to display cytopathological alterations of the midgut epithelium. EO and enriched fraction of Peumus boldus displayed larvicidal activity against Cu. Quinquefasciatus. The treated larvae displayed morphological changes in the midgut cells (de Castro et al., 2016). Velu et al. (2015) tested the peel extract of A. hypogaea against Aedes aegypti and Anopheles stephensi. The histopathological studies exposed midgut tissue damage and cuticle injury. Costa et al. (2012) reported similar aberrations in Ae. aegypti larvae (III instar) treated with Annona coriacea extract. Ae. aegypti larvae exposed to squamocin from Annona mucosa Jacq. (Annonaceae) displayed larvicidal and cytotoxic action with changes in the midgut epithelium and digestive cells by increasing the expression of autophagy genes (Costa et al., 2014, 2017). da Silva Costa et al. (2018) also reported that squamocin affected the osmoregulation and ion-regulation of Ae. aegypti larvae which resulted in a lethal effect caused by the development of a great vacuolization in the anal papillae wall.

The histopathological study of Ae. aegypti treated with methanol extract derived from seaweeds Sargassum binderi showed that larvae treated with seaweed extracts had cytopathological alteration of the midgut epithelium. The morphological observation revealed that the anal papillae and terminal spiracles of larvae were the common sites of aberrations (Yu et al., 2015). Phytochemicals (oleic, linoleic, linolenic, palmitic, and stearic acids) and their respective methyl esters were tested against fourth instar Cu. quinquefasciatus larvae. The compounds were found to affect its metabolism and the morphology of midgut along with their fat body (de Melo et al., 2018).

Impact of Phytochemicals on the Insect Behavior

With the development of resistance by this time attained to almost all available chemicals, strategies integrating “plant derived” compounds to influence “semiochemical”-mediated behaviors by means of interruption of mosquito-olfactory sensory system have substantially developed (Muema et al., 2017). As a consequence, the physiological status related to the olfactory sensory system is disrupted. The phytochemicals will bind to these odorant chemoreceptors and subsequent flight orientations of the mosquitoes are hindered (Bohbot et al., 2010). Henceforth the physiological status for instance “circadian-regulated appetitive stimulus” or “gonotrophic status” that triggers olfaction in pursuit of nutritious sources, mates and oviposition sites are disturbed. Plant-based semiochemicals can be exploited to lure the mosquitoes to an insecticide trap, thereby forming an integral part of an integrated vector control programe (Kamala-Jayanthi et al., 2015). Rice volatiles on evaluation with BioGent (BG) sentinel traps elicited antennal responses that stimulated long range oviposition site seeking behavior. Also, p-cresol, from Bermuda grass hay infusion was reported with avoidance response to gravid An. Gambiae (Eneh et al., 2016).

Future Perspectives

Higher rates of anthropogenic activities that are expected to expand with the population increase will increase the incidence of vector borne diseases. Additionally, the development of resistance among the vector population against the synthetic chemical insecticides along with their persistence in the environment and toxicity for non-target organisms are reducing the efficiencies of vector management practices globally. Hence novel plant-based compounds that are safe and effective are being focused for the development of improved management of vectors.

The research has now moved on from the isolation of bioactive compounds with anti-vector potentials to formulate novel application methods. Apart from the direct application of plant metabolites in vector control, nanoparticles (NPs) synthesized from plants using green technology are emerging as a new trend. Nanotechnology is presently “revolutionizing” the manufacture of commercial pesticides. Production of green NPs and nanoencapsulation compounds upsurges the permanence of EOs through “slow-release” phenomenon deliberating sustained fortification against mosquito bites. As reported by Jinu et al. (2018), silver nanoparticles (AgNPs) from Cleistanthus collinus Karra and Strychnos nux-vomica Linn nux-vomica presented highest larvicidal activity against A. stephensi and A. aegypti. Murugan et al. (2018a, b) proved the efficacy of zinc oxide NPs fabricated using the brown macroalga Sargassum wightii Greville ex J. Agardh. against An. stephensi. In another study reported by Murugan et al. (2018b), Poly (Styrene Sulfonate)/Poly (allylamine hydrochloride) encapsulation of TiO2 NPs were found to enhance their toxicity against mosquito vectors of Zika virus.

Conclusion

Mosquito vector borne diseases are a major human health problem in all countries. There has been an alteration toward plant-based insecticides to overcome the problems related with the use of synthetic mixtures in mosquito control programe. Botanicals can be used as mosquitocides for killing both larvae and adult mosquitoes. However, only very few botanicals have moved from laboratory to the field use, which may be due to the light and heat variability of phytochemicals compared to synthetic insecticides. Further these botanicals have been widely explored, but only a comparatively small number of patents have been filed with the persistence of regulating the formulations for use against mosquito species in the field level.

Although the activity of phytochemicals are generally attributed to some specific compounds, but there is increasing evidence that the combination of botanicals and biopesticides will result in an increased bioactivity compared to single phytochemicals (Senthil-Nathan et al., 2005a; Senthil-Nathan and Kalaivani, 2005, 2006).

At present, botanical insecticides make <1% of the world’s pesticide market (Sola et al., 2014). Isolation of active principles and synthesis of secondary metabolites of botanicals against mosquito threat are very important for the management of vector borne diseases. The positive results of initial studies on larvicidal potential of botanicals encourage further interest to investigate the bioactive compounds. Identifying botanical insecticides that are effective as well as appropriate and adaptive to overcome ecological hazards, biodegradable, and have a broad spectrum of larvicidal properties will work as a new defense in the arsenal of insecticides and it may act as an appropriate alternative product to fight against vector-borne diseases.

Thus, the present review collects important information on plant extracts along with their active molecules as agents affecting the physiology and behavior of medically threatening mosquito vectors. Now collective efforts are needed to take advantage of the accumulated knowledge on phytochemical action on mosquitos in order to integrate their application in integrated pest management programs.

Author Contributions

SS-N collected all the information and wrote the review.

Conflict of Interest

The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.