identification of mosquito biting deterrent constituents from the indian folk remedy plant jatropha...

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Identification of Mosquito Biting Deterrent Constituents from theIndian Folk Remedy Plant Jatropha curcasAuthor(s): Charles L. Cantrell, Abbas Ali, Stephen O. Duke, and Ikhlas KhanSource: Journal of Medical Entomology, 48(4):836-845. 2011.Published By: Entomological Society of AmericaDOI: http://dx.doi.org/10.1603/ME10244URL: http://www.bioone.org/doi/full/10.1603/ME10244

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VECTOR CONTROL, PEST MANAGEMENT, RESISTANCE, REPELLENTS

Identification of Mosquito Biting Deterrent Constituents From theIndian Folk Remedy Plant Jatropha curcas

CHARLES L. CANTRELL,1,2 ABBAS ALI,3 STEPHEN O. DUKE,1 AND IKHLAS KHAN3

J. Med. Entomol. 48(4): 836Ð845 (2011); DOI: 10.1603/ME10244

ABSTRACT An investigation of the Indian folk remedy plant Jatropha curcas L., was performed toidentify the constituents responsible for the mosquito biting deterrent activity of the oil. J. curcas seedoil is burned in oil lamps in India and parts of Africa to repel biting insects, primarily mosquitoes. Theseed oil was thoroughly analyzed by 1H NMR, 13C NMR, high-performance liquid chromatography-refractive index, and gas chromatography-ßame ionization detection to identify the constituents in theoil. IdentiÞed constituents, both free fatty acids and triglycerides, were evaluated for activity inAedesaegypti (L.) (Diptera: Culicidae) biting deterrent assays. Furthermore, an oil condensation trap wasused to demonstrate that free fatty acids or triglycerides are partially volatilized during the combustionprocess. These compounds were found to be responsible for the biting deterrency of the burned oil.SpeciÞcally, oleic, palmitic, linoleic, and stearic acids were all active at 25 nmol/cm2 above that ofsolvent control in Ae. aegypti biting deterrent assays. Oleic, palmitic, and linoleic acids were all moreactive than stearic acid in the same bioassay. Evaluation of the triglycerides containing each of thesefatty acids revealed that tripalmitin, tristearin, trilinolein, and triolein all demonstrated signiÞcantactivity above a solvent control at 10 �g/cm2, whereas tripalmitin was the most active. Due to literaturereports suggesting larvicidal activity of the oil, J. curcas seed oil and its free fatty acid constituents alsowere evaluated against 1-d-old Ae. aegypti larvae up to 500 ppm. Oleic acid was the only fatty acidhaving larvicidal activity against 1-d-old Ae. aegypti larvae, with an LD50 of 47.9 ppm.

KEY WORDS Jatropha curcas, insect repellent, ethnobotany, mosquitoes, Aedes aegypti

There is an urgent need for the development of al-ternative insecticides and insect repellents to manageimportant disease vectors such as Aedes aegypti (L.)(Diptera: Culicidae).Ae. aegypti transmits viral patho-gens to humans, including yellow fever and dengue,both of which can cause severe human morbidity andmortality. One potential source of new insecticidesand insect repellents is natural compounds fromplants. Not only might certain natural plant com-pounds be a source of new insecticides and insectrepellents but also botanical chemical derivatives maybe more environmentally friendly than syntheticchemicals (Cantrell et al. 2010).Jatropha curcas L. extracts and oil are known to be

insecticidal to mosquito larvae (Fagbenro-Beyioku etal. 1998, Georges et al. 2008, Rahuman et al. 2008,Murthy and Rani 2009), house ßies (Sievers et al.1949), and other insects (Adebowale and Aderdire2006, Boateng and Kusi 2008, Sujeetha 2008, Ratnadasset al. 2009). In particular, the phorbol esters known tobe toxic to mammals are also toxic to insects (Gubitzet al. 1998). In India, J. curcas preparations for insectpest management have been traditionally used

(Narayanasamy 2006). Extracts of J. curcas are anti-feedants for some insects (Meshram et al. 1996). Otherspecies of Jatropha have insecticidal constituents.Japodic acid, a aliphatic acid from the roots of J.podagrica, inhibits growth of the insect Helicoverpazea (Boddie) (Aiyelaagbe and Gloer 2008) and leafextract of J. gossypifoliawas toxic to second instars ofSpodoptera litura (F.) (Phowichit et al. 2008). How-ever, not all insects are affected by J. curcas. In fact,some insects sequester the toxic phorbol esters of J.curcas, apparently as a self-defense mechanism (Winket al. 2000).

Very little is known about the mosquito spacialrepellent activity of J. curcas compounds. Seed oil ofthis plant is somewhat toxic to the termiteCoptotermesvastator Light, but it is also an antifeedant and a re-pellent to this insect (Acda 2009). It is also a repellentto the stored grain insect pest Callosobruchus macu-lates (F.) (Boateng and Kusi 2008). Jatropha oil isextensively used in Africa and India as a folk remedyarea mosquito repellent by burning the oil in an oillamp (e.g., http://www.times.co.zm/news/viewnews.cgi?category�8&id�1153510056 http://www.snvworld.org/en/Documents/Case_study_Zambia_oil_seed_and_Jatropha_value_chains.pdf), but we Þnd no scientiÞcstudies which investigated the speciÞc constituents re-sponsible for such activity. Direct communications bylocal residents to one of us (I.K.) during travels to India in

1 USDAÐARS, Natural Products Utilization Research Unit, Univer-sity, MS 38677.

2 Corresponding author, e-mail: [email protected] National Center for Natural Products Research, The University of

Mississippi, University, MS 38677.

2008 also indicated this traditional use. Seed oil from J.curcaswasprovided to I.K.during this visit and lateronanauthenticated sample from the Central Council for Re-search in Unani Medicine, India, was provided that wasused for detailed chemical and biological investigations,whereas the former sample was used in preliminary workonly. The objective of this study was to investigate Aeaegypti repelling and biting deterrent properties of J. cur-cas oil and speciÞcally to identify the constituent(s) re-sponsible for biting deterrent activity of this oil and thevolatiles generated by burning the oil.

Materials and Methods

Jatropha Oil Combustion Trapping Experiment.This experiment was performed to trap the volatilesproduced during the combustion of seed oil. Threemilliliters of J. curcas oil was pipetted into a 4-ml glassvial with a 2.5-cm-long cotton wick from a commercialoil lamp. The vial was placed into a 250-ml two-neckround-bottomed ßask and ignited. Ignition was main-tained by a low ßow of breathing grade air (20.9%oxygen; Nexair, Memphis, TN) through the system(Fig. 1). The ßask was connected to a cold condenserallowing the vapors to condense into a 100-ml round-bottomed ßask. Air ßowing out of the system througha Teßon tube was bubbled through a 150-ml methyl-ene chloride (CH2Cl2) trap. The CH2Cl2 trap wasdried by rotary evaporation almost to dryness (3 ml).The oil condensate collected in the 100-ml ßask wastransferred to a 60-ml separatory funnel and extractedwith 60 ml of CH2Cl2. The CH2Cl2 phase of the oilcondensate was dried by rotary evaporation almost todryness (1 ml). The H2O phase of the oil condensatewas frozen and dried by lyophilization. The entirecombustion setup was rinsed with 200 ml of MeOHand dried by rotary evaporation almost to dryness (3ml). The CH2Cl2 phase of oil condensate was directly

methylated using the procedure described for meth-ylation.Methylation of Jatropha Oil and Preparations. The

method was used for the conversion of fatty acids andtriglycerides (TAGs) into their corresponding fattyacid methyl esters allowing for subsequent analysis bygas chromatograph (GC). One hundred microliters(�80 mg) of Jatropha oil was weighed into a 50-mlTeßon-lined, screw-capped tube. One milliliter of asolution of tricosanoic acid (C23:0; purity 99%) (2.0mg/ml) dissolved with heat in n-hexane was added asan internal standard, followed by 1 ml of methanol and3 ml of 3 N methanolic-HCl (Sigma-Aldrich, St. Louis,MO). The tubes (replicated six times) were cappedtightly and reßuxed in a water bath at 80�C for 1 h.After cooling to room temperature, 8 ml of 0.88%(wt:vol) NaCl solution and 3 ml of hexane were addedto the tube, and the contents were vortexed for 1 min.After centrifugation for 5 min at 3,000 rpm, the toplayer was removed, transferred into a 2-ml vial, andanalyzed on a GC by using ßame ionization detection(FID) detection. All fatty acid and TAG standardswere purchased from Supelco (Bellefonte, PA), andpurities were as follows: oleic acid (99.0%), palmiticacid (99.0%), linoleic acid (99.0%), stearic acid(99.5%), tripalmitin (99.4%), tristearin (99.9%), tri-linolein (98%), and triolein (99.9%).GC-FID Analysis. GC-FID analysis was performed

on a CP-3800 GC (Varian, Palo Alto, CA). The GC wasequipped with a DB-23 (Agilent Technologies, SantaClara, CA) column (60-m by 0.25-mm capillary col-umn; Þlm thickness, 0.25 �m) operated using the fol-lowing conditions: injector temperature, 270�C; col-umn temperature initially set at 130�C and held for 1min, followed by ramping from 130 to 170�C at 6.5�C/min, again followed by ramping from 170 to 215�C at2.8�C/min, held for 12 min and followed by a Þnalramp from 215 to 230�C at 40�C/min and held for 3min; injection volume, 1 �l (split 20:1); 3.0 ml/minconstant ßow of He; and FID temperature, 300�C.Fatty acid methyl esters present in the oil sampleswere speciÞcally identiÞed by injection of commer-cially available fatty acid methyl ester standards (Su-pelco) and subsequent comparison of retention timesof standards with that of unknowns, allowing for un-equivocal identiÞcation.Separation of Triacylglycerols and Free Fatty Acids(FFAs) in Jatropha Oil by Solid-Phase Extraction.This technique was used in the determination of therelative amounts of FFAs and TAGs present in seedoils. A binary solvent was freshly prepared before eachextraction by mixing dichloromethane and n-hexane(1:4 by volume). Solid sodium carbonate (600 mg)prewetted with 1 ml of 0.1 M KOH was added with123.0 mg of J. curcas oil. The triacylglycerols fractionwas then extracted with 10-ml volumes of binary sol-vent (three times) by vortex-mixing, and each extractwas decanted after sodium carbonate was pelleted bybrief centrifugation. The FFAs fraction that remainedadsorbed to the solid sodium carbonate was recoveredby acidiÞcation (to pH �2) of the particles with 6 MHCl and then extracting with three successive 10-ml

Fig. 1. Combustion trap for J. curcas oil showing a vial ofoil containing a wick that is placed inside a two-neck ßask fedby a tank of air. Much of the oil that burns is condensed andcollected as the oil condensate and the air outlet is bubbledthrough DCM.

July 2011 CANTRELL ET AL.: MOSQUITO BITING DETERRENT FROM J. curcas 837

volumes of binary solvent, with vigorous vortex-mix-ing before each extraction. The combined extractswere dried over anhydrous Na2SO4, providing 118.7mg of a TAG fraction and 1.3 mg of an FFA fraction.Separations and Identification of TAGs by High-Performance Liquid Chromatography (HPLC)WithRefractive Index (RI) Detection. HPLC analysis wasperformed using a 1200 series HPLC (Agilent Tech-nologies) with a Zorbax SB-C18 column (5 �m, 4.6 by250 mm). The system also included a degasser, qua-ternary pump, diode array detector, RI detector, andcolumn compartment. The column compartment washeated to 35�C throughout the analysis. The RI opticalunit temperature was set at 35�C with polarity set onpositive and an analog output range of 1.0 V. An iso-cratic method using acetone and acetonitrile (80:20)was used with a ßow rate of 1 ml/min over 30 min.Each standard was analyzed at 10 mg/ml and jatrophaoil at 30 mg/ml in methanol. Standards analyzed in-cluded triolein, tripalmitin, tristearin, and trilinolein.Insects. Ae. aegypti used in these studies were from

a laboratory colony maintained since 1952 at the Mos-quito and Fly Research Unit at Center for Medical,Agricultural, and Veterinary Entomology, USDAÐ

ARS, Gainesville, FL. This colony is maintained since1952 by using standard procedures (Pridgeon et al.2007). We received the eggs and stored these in ourlaboratory to use as needed. Mosquitoes were rearedto the adult stage by feeding the larvae on a diet of 2:1alfalfa pellets (U.S. Nutrition Inc., Bohemia, NY) andhog chow (Ware Milling, Houston, MS). The dietcontents were ground in a grinder and passed throughsieve no. 40, 425 �m (USA Standard Sieve, HumboldtMFGCo.,Norridge, IL).Theeggswerehatchedundervacuum (�1 h) by placing a piece of a paper towelwith eggs in a cup Þlled with 50 ml of deionized watercontaining small quantity of larval diet. Larvae wereremoved from vacuum and held overnight in the cup.These larvae were then transferred into 500-ml cups(�50 larvae per cup) Þlled with deionized water.Larval diet was added every day until pupation and theinsects were kept in an environment controlled room.Both the larvae and adults were maintained at a tem-perature of 27 � 2�C and 70 � 5% RH in a photoperiodregimen of 12:12 (L:D) h. The adults were fed on 10%sucrose solution. Cotton pads moistened with the su-crose solution were placed on the top of screens of4-liter cages. Five- to 9-d-old mated females were usedin these bioassays. Females were deprived off of su-crose solution for 24 h before the test and were fed onwater-soaked cotton. Timings of these tests were cen-tered around 1200 hours.Mosquito Biting Bioassays. Experiments were con-

ducted by using a six-celled in vitro Klun & Debboun (K& D) module bioassay system developed by Klun et al.(2005)forquantitativeevaluationofbitedeterrentprop-erties of candidate compounds for human use. This bio-assay method determines speciÞcally measured biting(feeding) deterrent properties of the chemicals. In brief,theassaysystemconsistsofasixwellbloodreservoirwitheach of the 3- by 4-cm wells containing 6 ml of blood. Asreported previously (Klun et al. 2008), female mosqui-

Fig. 2. GC-FID chromatogram of J. curcas oil after conversion of triglycerides to fatty acid MEs.

Table 1. Mosquito biting deterrent effects of J. curcas oilagainst 5–9-d-old mated female Ae. aegypti

Treatment (n � 80)a ConcnMean proportion not

biting (SE)b

DEET 25 nmol/cm2 0.775 (0.047)aJ. curcas oil 100 �g/cm2 0.650 (0.053)bJ. curcas oil condensate 100 �g/cm2 0.613 (0.054)bcJ. curcas oil 10 �g/cm2 0.550 (0.055)cJ. curcas oil condensate 10 �g/cm2 0.538 (0.056)cEthanol 0.213 (0.046)d

a n is number of females tested for each treatment.b Values within a column not followed by the same letter are

signiÞcantly different (P � 0.05; DMRT).

838 JOURNAL OF MEDICAL ENTOMOLOGY Vol. 48, no. 4

toes feed as well on citrate-phosphate-dextrose-adenine(CPDA-1) � ATP as they do on blood. Therefore, weused the CPDA-1 � ATP instead of human blood.CPDA-1 was prepared by dissolving 3.33 g of sodiumcitrate, 0.376 g of citric acid, 4.02 g of dextrose, 0.28 g ofmonobasic sodium phosphate (Fisher Chemical, Fair-lawn, NJ) and 0.346 g of adenine (Sigma-Aldrich, St.Louis, MO) in 1,026 ml of deionized water. ATP wasadded to CPDA-1 to yield 10�3 M ATP (AABB 2005).CPDA-1 and ATP preparations were freshly made onthe day of the test. N,N-Diethyl-3-methylbenzamide

([DEET]; 99.1% purity) was obtained from Sigma Al-drich and used as a positive control. Molecular biologygrade ethanol was obtained from Fisher Chemical. J.carcus oils (Table 1) were produced as described above,and TAGs and fatty acids were purchased from Sigma-Aldrich. J. curcas oils preparations were made by dis-solving the neat oil in the respective solvent, therebycreating a stock solution that was used to apply theprecise amount of material to the cloth.

The temperature of the solution in the reservoir wasmaintained at 37�C by continuously passing the warm

Fig. 3. HPLC chromatogram of J. curcas oil with RI detection overlaid with triolein, tripalmitin, tristearin, and trilinolein,conÞrming the presence of triolein and trilinolein. The initial peak corresponds to the solvent and additional peaks were notidentiÞed.


water through the reservoir by using a circulatorybath. The reservoirs were covered with a layer ofcollagen membrane. This CPDA-1�ATP solutionmembrane unit simulated a human host for mosquitofeeding. The test compounds were randomly appliedto six 4- by 5-cm areas of organdy cloth and positionedover the membrane-covered CPDA-1�ATP solutionwith a separator placed between the treated cloth andthe six-celled module. A six-celled K & D modulecontaining Þve females per cell was positioned overcloth treatments covering the six CPDA-1�ATP so-lution membrane wells, and trap doors were opened toexpose the treatments to these females. The numberof mosquitoes biting through cloth treatments in eachcell was recorded after a 3-min exposure, and mos-quitoes were prodded back into the cells. These mos-quitoes were then squashed to determine the numberthat has actually engorged the solution. A replicateconsisted of six treatments: four test compounds,DEET (a standard bite deterrent compound) and 95%ethanol- or acetone-treated cloth as solvent control.The 25 nmol DEET/cm2 cloth dose was used as astandard, because it suppresses mosquito biting by80% compared with controls (Klun et al. 2005). A setof replications was conducted on different days byusing new lots of the insects.

Larval Bioassays. Bioassays were conducted by us-ing the bioassay system described by Pridgeon et al.(2009) to determine the larvicidal activity of J. curcasoil and its constituents (palmitic acid 16:0, stearic acid18:0, and oleic acid 18:1n9c). Linoleic acid was nottested in this 1-d assay due to the instability of this acidto oxygen. In brief, the eggs were hatched undervacuum (�1 h) by placing a piece of a paper towelwith eggs in a cup Þlled with 100 ml of deionized watercontaining small quantity of larval diet. Larvae wereremoved from vacuum and held overnight in the cupin a temperature-controlled chamber maintained at atemperature of 27 � 2�C and 70 � 5% RH at a pho-toperiod regimen of 12:12 (L:D) h. Five 1-d-old Þrst-instar Ae. aegypti were added to each well of 24-wellplates placed on illuminated light box by using a dis-posable 22.5-cm Pasteur pipette with a droplet of wa-ter measuring �40 �l. One milliliter of deionized wa-ter and 50 �l of larval diet (2% slurry of 2:1 alfalfapellets and hog chow) were added to each well byusing a Finnpipette stepper (Thermo Fisher, Vantaa,Finland). All chemicals to be tested were diluted indimethyl sulfoxide (DMSO). Eleven microliters of thetest chemical was added to the labeled wells, and incontrol treatments 11 �l of DMSO alone was added.After the treatment, the plates were swirled in clock-

Fig. 4. 1H NMR (600 MHz) spectrum of J. curcas oil overlaid with the spectra of triolein, tripalmitin, tristearin, andtrilinolein.


wise and counter clockwise motions and front andback and side to side Þve times to ensure even mixingof the chemicals. Larval mortality was recorded 24 hafter treatment. Larvae that showed no movement inthe well after manual disturbance of water by a pipettetip were recorded as dead. The larval assays wererepeated with Þve to seven concentrations providinga range of 0 to100% mortality for compounds thatshowed activity in initial screening.Statistical Analysis. Data were analyzed by Proc

analysis of variance (ANOVA) (SAS Institute 2007),and means were separated by DuncanÕs multiple rangetest (DMRT). The LD50 values were calculated byusing PoloPlus Probit and logit analysis software(LeOra Software, Petaluma, CA). Control mortalitywas corrected by using AbbottÕs formula.

Results and Discussion

Given that reports of the traditional folk remedy byusing J. curcas indicate that the oil is burned in an oillamp to repel mosquitoes, a crude setup was con-structed to trap the volatile constituents producedduring this burning process (Fig. 1). It was hypothe-

sized that incomplete combustion could potentiallyrelease volatile constituents responsible for the re-ported mosquito deterrency. Such volatile constitu-ents could potentially condense along with the waterproduced during combustion. By carefully placing a4-ml vial containing 3 ml of oil and an oil lamp wickdirectly in the bottom of a two-necked ßask, the wickcould be ignited while carefully maintaining the air-ßow into the setup. Burning of all 3 ml of oil typicallytakes 30 min, depending on the airßow into the ap-paratus. The resulting oil condensate was collected ina receiving ßask and consisted primarily of water.

The J. curcas oil and the oil condensate were bothevaluated for biting deterrency against Ae. aegypti byusing the K & D module bioassay system (Klun et al.2005). The experimental setup (Table 1) consisted ofDEET at 25 nmol/cm2 as the positive control, ethanolas the solvent control, and concentrations of 100 and10 �g/cm2 for both J. curcasoil and the oil condensate.In a previous report (Cantrell et al. 2005), such con-centrations worked well for the evaluation of plantextracts. All treatments were more active than theethanol control, and no treatments were as effective asDEET (P � 0.05). There was no statistical difference

Fig. 5. 13C NMR (125 MHz) spectrum of J. curcas oil overlaid with the spectra of triolein, tripalmitin, tristearin, andtrilinolein.


between J. curcas oil and oil condensate at either 100or 10 �g/cm2. Due to the low amounts of oil conden-sate obtained and the abundance of J. curcas oil avail-able, it was decided to perform a detailed spectro-scopic and chemical analysis of the J. curcas oil to aidin the identiÞcation of the constituent(s) responsiblefor its activity.

Previous reports on the contents of J. curcas oilindicated that it was composed predominately ofTAGs (Patil et al. 2009) and FFAs with oleic andlinoleic acids being the dominant acids (Akbar et al.2009). Additional reports on minor constituents indi-cate the presence of phorbol esters in the seed oil(Ahmed and Salimon 2009). It is well recognized thatconstituents present in seed oil can vary tremendouslydepending on several environmental factors, such assoil nitrogen and rainfall; therefore, we decided tothoroughly investigate theoil components fromthe lotof oil that we were using in this study rather thanrelying on previous reports on oil composition fromthe same species. Analysis of the bioactive J. curcas oilwas performed by reacting the oils with methanolicHCl followed by analysis of the resulting fatty acidmethyl esters by using GC-FID detection. IdentiÞca-tion of individual fatty acid methyl esters (MEs) was

accomplished by injection of commercially availablefatty acid methyl ester standards and comparison ofretention times with that of unknowns (Fig. 2). Areapercentage calculations indicated that oleic acid ME(41.5%) was the major fatty acid followed by linoleicacid ME (34.0%), palmitic acid ME (16.3%), andstearic acid ME (6.4%), accounting for 98.2% of totalcomponents in the oil by FID.

Unfortunately, this procedure does not adequatelydiscriminate between fatty acid MEs derived fromTAGs and those derived from FFAs. For this purpose,a solid phase extraction technique using sodium car-bonate was used consisting of adsorption of the FFAsonto the sodium carbonate, ßushing the unboundTAGs, followed by recovery of the FFAs. The methodwas based on a previously described procedure (Paiket al. 2009) that resulted in a weight distribution sug-gesting that the oil was composed of 96% TAGs and 1%FFAs.

It is now clear that the composition of the J. curcasoil consists of predominately TAGs and a small amountof FFAs, and the major fatty acids building the TAGsare the two unsaturated fatty acids oleic and linoleicacid and the two saturated fatty acids palmitic andstearic acid. What is still unclear is the exact makeup

Fig. 6. OleÞnic region expansion of the 13C NMR (125 MHz) spectrum of J. curcas oil overlaid with the spectra of triolein,tripalmitin, tristearin, and trilinolein.


of TAGs, each of which is probably composed of acombination of three of these fatty acids attached toa glycerol unit. Because commercially available trio-lein, tristearin, tripalmitin, and triolein were available,we used three methods to conÞrm the presence ofthese four TAGs. The methods used consisted ofHPLC-RI detection and 1H NMR and 13C NMR spec-troscopy. In all of these methods, a comparison wasmade between the chromatograms or spectra of J.curcas oil and that of each of the four commerciallyavailable TAGs. Unfortunately, TAGs containing mix-tures of fatty acids were not available commercially.

TheHPLC-RI separationand identiÞcationofTAGswas performed using a modiÞed method from Supel-coÕs bulletin 787D (T100787D, Sigma-Aldrich). Theisocratic method indicated that both trilinolein andtriolein were probably present in J. curcas oil (Fig. 3)based only on retention time overlaps between com-mercially available standards and the oil constituents.Tristearin and tripalmitin did not seem to be present;however, two additional major peaks remained un-identiÞed but are probably mixed fatty acid TAGs.

1H and 13C NMR analysis of the J. curcas oil and thefourcommercially availableTAGsalsowasperformed.The 1H NMR analysis revealed certain diagnostic sig-nals for TAGs triolein and trilinolein in the oil (Fig. 4).SpeciÞcally, the proton triplet at � 2.75 ppm in trilino-lein also was present in J. curcas oil. Similarly, theproton quartet at � 2.03 ppm in trilinolein was presentin J. curcas oil. Both the quartet at � 1.99 ppm and themultiplet at � 5.33 ppm in triolein also were present inJ. curcas oil. Signals from both tristearin and tripalmi-tin were less diagnostic due to the same signals alsopresent in the others TAGs. From the 13C NMR spec-

tra (Figs. 5 and 6), it is again clear that both trioleinand trilinolein signals are present based on similaritiesin the oleÞnic and aliphatic regions of the spectra.OleÞnic carbons in trilinolein at � 130.2, 130.0, 128.1,and 127.9 ppm are all present in the oil (Fig. 6). TheoleÞnic carbon in triolein at � 129.7 is also present inthe oil. The aliphatic carbons at 25.6 and 31.5 ppmseem to be unique to trilinolein, which is also presentin the spectrum of the oil.

GC-FID analysis for the oil was used to analyze boththe methanol rinse of the distillation condenser andthe dichloromethane (DCM) extract of the aqueouscondensate. Again, analysis was performed by reactingthese fractions with methanolic HCl followed by anal-ysis of the resulting fatty acid methyl esters by usingGC-FID detection (Fig. 7). Clearly, all four of theresulting fatty acids also were present in these frac-tions, implying that incomplete combustion results inthe volatilization of these fatty acids, their TAG coun-terparts, or both.

Finally, what remained was the evaluation of themosquito biting deterrency of both the FFAs andthe TAGs by using the K & D module bioassaysystem. The four primary fatty acidsÑoleic,palmitic, linoleic, and stearic acidÑwere all testedat 25 nmol/cm2 against Ae. aegypti versus solventcontrol and DEET. All four of the fatty acids wereactive above that of solvent control (ethanol) inbiting deterrent assays; however, none of the com-pounds were more effective than DEET at the sameconcentration (Table 2). Oleic, palmitic, and lino-leic acids also were more active than stearic acid inthe same bioassay.

Fig. 7. GC-FID chromatogram of J. curcas oil combustion rinses after conversion of triglycerides to fatty acid MEs. (A)Methanol rinse of the distillation condenser. (B) DCM extract of the aqueous condensate.


Evaluation of the TAGs containing each of thesefatty acids also was performed using the K & D modulebioassay system against Ae. aegypti. The experimentalsetup included the TAGs tripalmitin, tristearin, tri-linolein, and triolein at 10 �g/cm2; the solvent controlacetone; and J. curcas oil at 10 �m/cm2. The assayrevealed that tripalmitin, tristearin, trilinolein, andtriolein demonstrated signiÞcant activity above thesolvent control at 10 �g/cm2, whereas the activity oftripalmitin was signiÞcantly higher than trilinolein andtriolein (Table 3).

Because the J. curcas extract is reported to havelarvicidal activity (Aina et al. 2009, Murthy and Rani2009) against Ae. aegypti and other mosquito species,the constituent(s) also were screened for their larvi-cidal effect. In larval screening bioassay, J. curcas oil,palmitic acid (16:0) and stearic acid (18:0) did notshow any larvicidal activity at a maximum dose of 500ppm, and oleic acid (18:1n9c) was the only compoundactive against 1-d-old Ae. aegypti larvae. The LD50

(95% CL) value for oleic acid against 1-d-old larvae at24 h posttreatment was 47.9 (41.08Ð55.23) ppm. Ainaet al. (2009) reported high LC50s (3,250 and 12,000ppm for ethanol and water extracts of J. curcas, re-spectively) against second-instar Anopheles gambiaeGiles larvae. Murthy and Rani (2009) reported anLC50 (95% CL) value of 850 (680Ð990) ppm of J.curcas leaf extract against third-instar Ae. aegypti.These data showed mortality trends similar toRamesewak et al. (2001), who reported LD50 value of100 ppm against fourth-instarAe. aegypti for oleic acidisolated from Dirca palustris L.

In conclusion, we have demonstrated that bothFFAs and TAGs present in J. curcas oil possess Ae.aegypti biting deterrent activity and are at least par-tially responsible for the reported biting deterrentactivity of the oil. SpeciÞcally, oleic, palmitic, linoleic,and stearic acids were active at 25 nmol/cm2 abovethat of control in Ae. aegypti biting deterrent assays.Oleic, palmitic, and linoleic acids also were more ac-tive than stearic acid in the same bioassay. Hwang etal. (1984), who reported ovipositional repellency ofunsaturated fatty acids, also evaluated both oleic andlinoleic acids against Culex quinquefasciatus Say. Botholeic and linoleic acids were signiÞcantly repellentwith oleic acid being the most active fatty acid eval-uated in the study of unsaturated fatty acids. An ad-

ditional report on repellency of fatty acids, both un-saturated and saturated, against Ae. aegypti furtherindicates evidence of repellency (Skinner et al. 1970).It is important to note that our assay measures bitingdeterrency, and this report seems to be the Þrst onsuch activity of fatty acids, both saturated and unsat-urated.

Evaluation of the TAGs containing each of thesefatty acids revealed that tripalmitin, tristearin, trilino-lein, and triolein demonstrated signiÞcant activityabove solvent control at 10 �g/cm2, whereas tripalmi-tin was the most active. This report is the Þrst on Ae.aegypti biting deterrency of TAGs.

Acknowledgments

We thank Amber Reichley and Solomon Green, III, fortechnical assistance. We thank James J. Becnel (Mosquitoand Fly Research Unit, Center for Medical, Agricultural andVeterinary Entomology, USDAÐARS, Gainesville, FL) forsupplying Ae. aegypti eggs. We thank Prof. Jonathon Gressel(Weizmann Institute of Science, Rehovot, Israel) for sug-gesting this study. This study was supported, in part, byDeployed War-Fighter Protection Research Program Grantfunded by the U.S. Department of Defense through theArmed Forces Pest Management Board.

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Table 2. Mosquito biting deterrent effects of J. curcas oilmajor hydrolysis products against 5–9-d-old mated Ae. aegyptifemales

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Table 3. Mosquito biting deterrent effects of J. curcas oiltriglycerides against 5–9-d-old adult Ae. aegypti females

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Received 9 November 2010; accepted 10 March 2011.
