Materials & Methods

Materials and Methods

3.1 Plant species

Leaves of Eucalyptus camaldulensis Dehnhardt (Myrtaceae), Tylophora indica

(Burm. f.) Merrill (Asclepiadaceae) synonym T. asthmatica, Ocimum sanctum

Linnaeus (Lamiaceae), Lantana camara Linnaeus (Verbenaceae), and rhizomes

of Curcuma longa Linnaeus (Zingiberaceae) were used as test material. Fresh

leaves of E. camaldulensis, T. indica and O. sanctum were collected from

germplasm grown and maintained at TERI’s field station in Gual Pahari

(Haryana). Fresh rhizomes of C. longa were procured from CCSHAU, Hisar.

Lantana camara leaves were collected from plants growing wild in the nature in

or around agricultural land in Delhi. A summary is presented in Table 10.

Dr Virender Kumar retired as Head- Department of Botany, Zakir Hussain

College, Delhi University provided authentic identification. The leaves/

rhizomes were shade dried and ground to fine powder in a mixer grinder. Leaf

and rhizome powder were utilized for further experimental purpose.

Table 3.1 Details of the test plant species

S. No. Plant species Family Plant part

tested

Collection site

1 Eucalyptus

camaldulensis Dehnhardt

Myrtaceae Leaves TERI’s field station at

Gual Pahari (Haryana)

2 Tylophora indica

(Burm. f.) Merr.

Asclepiadaceae Leaves TERI’s field station at

Gual Pahari (Haryana)

3 Ocimum sanctum

Linnaeus

Lamiaceae Leaves TERI’s field station at

Gual Pahari (Haryana)

4 Lantana camara

Linnaeus

Verbenaceae Leaves Wild plants growing in

Delhi

5 Curcuma longa

Linnaeus

Zingiberaceae Rhizomes Research Farm,

CCSHAU, Hisar

3.2 The Insect

Helicoverpa (Heliothis) armigera (Hübner) (Lepidoptera: Noctuidae)

commonly known as cotton bollworm was used as a test insect species.

3

3.2.1 Rearing of Helicoverpa armigera

For initial establishment of the colony, larvae were collected from chickpea,

pigeon pea and cotton growing areas of Delhi and Haryana regions. The larvae

were maintained individually in a sterilized glass tube on the diet suggested by

Singh and Rembold (1992). The culture was kept at 27±1oC temperature, 65-

70% relative humidity and a photophase of 14 hours and 10 hours scotophase.

The colony size was regulated such that 250-300 neonate larvae were available

spread over a period of 15 days in each generation to facilitate bioassays during

different experimental stages. The procedure used for diet preparation and

laboratory culture of H. armigera during different growth stages viz. larva,

pupa, adult and egg were as follows:

3.2.1a Diet preparation

The composition for diet is given in Table 11. Ingredients of Part-A were weighed

accurately and blended thoroughly with a hand stirrer for about 4-5 min to make

a homogenous mixture. Simultaneously agar (4.5 g) in 100 ml distilled water

(Part-C) was boiled in a separate container and the dissolved agar was then

poured over the mixture of Part-A and the contents were mixed vigorously with

the help of a glass rod. The mixture was homogenized for 30 s and allowed to

cool for 1 min. The ingredients of Part-B were added to the above mixture of

Part-A & C and mixed thoroughly until an even consistency was obtained.

Prepared diet was poured immediately into sterilized and shallow perplex trays.

The diet was allowed to cool down till it solidifies after few hours. If not required

immediately, diet trays were kept in a fridge at 4oC for 15 days without quality

deterioration.

3.2.1b Larval culture

Neonate larvae (0-12 h old) were transferred with a very soft, fine hair

paintbrush into plastic boxes (6 cm diameter, 2 cm height). The surfaces of the

plastic boxes were provided with larval diet. 4-5 small holes were pricked in the

lids of these containers to provide adequate aeration. About 30 neonates were

placed in each box. The covers of the boxes were lined with tissue paper to

prevent migration of larvae. After 3-5 days 2nd instar larvae were reared

individually in the glass tubes (25 x 100 mm, Borosil) fitted with autoclaved

cotton plugs till pupation to prevent cannibalism. Larvae were provided with

flakes of fresh diet before they run out of food till pre-pupal stage. Larvae were

checked on alternate days and unhealthy or dead larvae were eliminated from

the culture.

Table 3.2 Constituents of semi- synthetic diet for rearing H. armigera

S. No. Material Quantity Quantity (%)

Part A

1 Chickpea seed powder 60 g 19.74

2 Sucrose 6 g 1.97

3 Yeast 6 g 1.97

4 Formaldehyde (10%) 1.5 ml 0.49

5 Choline chloride (20%) 3 ml 0.98

6 Distilled water 120 ml 39.47

Part B

7 Ascorbic acid 1.2 g 0.39

8 Sorbic acid 0.45 g 0.15

9 Methyl-p- hydroxybenzoate 0.75 g 0.25

10 Streptomycin 0.01 g -

11 Cholesterol 0.06 g -

12 Wheat germ oil + tocopherol

acetate (10:1)

0.06 g -

13 Vitamin capsule 1 no. -

Part C

14 Agar 4.5 g 1.48

15 Distilled water 100 ml 32.89

Total 304 g

Source: Singh and Rembold, 1992

3.2.1c Pupal culture

The pupae were allowed to remain undisturbed till the cuticle was fully

hardened (red-brown) and removed from the pupation cells after 2 days of

formation. These pupae were weighed, segregated into male and female. The

pupae were surface sterilised with sodium hypochlorite solution (0.25%) wash

by dipping the pupae for 5 s followed by 2-3 rinses with distilled water. The

healthy pupae having weight in the range of 300-400 mg were kept in the

culture otherwise discarded. These surface sterilised pupae were then

transferred in to clean jars (500 ml capacity) containing a piece of filter paper to

facilitate moth emergence and were observed daily for adult emergence.

3.2.1d Adult culture

The freshly emerged moths were separated on the basis of the colour of the

forewings (greenish in males and light to dark brown in females) and placed in

separate glass jars (500 ml capacity) covered with muslin cloth. Adults were

provided with cotton swab dipped in 10% honey solution as food. A small

quantity (3-4 drops) of methyl-p- hydroxybenzoate in ethanol (1 mg in 10 ml)

was added to the honey solution (50 ml) to prevent the growth of moulds. The

methyl-p-hydroxybenzoate solution was stored in refrigerator for 1-2 days for

further use. These cotton swabs were recharged daily with fresh honey solution.

Adults in the ratio of 3:3 male/ female were kept for mating after 3rd day of

emergence in a perplex mating cage (20 x 20 x 20 cm) provided with absorbent

cotton as an oviposition substrate as standardized in our laboratory (Kathuria

and Kaushik, 2004). Alternatively, adults were paired in 2 l Borosil beakers (20

x 15 cm), which were covered with a piece of muslin. The adults were allowed to

remain in the closed chambers till death. Care was taken to maintain humidity

above 65% during oviposition by using wet cotton swab.

3.2.1e Egg culture

Following mating, the eggs were obtained on the 4th and 5th day of emergence.

Pieces of muslin or cotton containing eggs were transferred to other glass jars

(500 ml capacity) covered with muslin cloth and monitored for larval

emergence. Moist cotton swab were kept on the muslin cloth for maintaining

humidity. Fertile eggs turned brown to black while infertile eggs remained

yellow and shrivelled after few days. Eggs hatched in 3-5 days after oviposition.

Neonate larvae were reared on semi-synthetic diet as per the method described

above. The neonate larvae obtained from this culture were used for conducting

insect bioassays. Laboratory population was supplemented with fresh culture

from fields after 5-7 generations for maintaining a continuous supply of the test

insect. The culture was maintained throughout the experimental period.

3.3 Chemicals, reagents and reference standards

3.3.1 Chemicals and reference standards

The chemicals used for preparation of insect diet were procured from Sigma,

USA. Other chemicals used in the present study were procured from Indian

companies. Detailed information in this regard is presented in Table 12.

3.3.2 Solvents

The solvents used in extraction and chromatographic procedures were of

commercial grade and of adequate purity (>90%). An account of the solvents

used in the present study accompanied by specifications indicating the source,

grade, etc. has been given in Table 13. Rectified spirit containing 95.6% of

alcohol by weight was used as ethanol solvent after distillation.

3.4 Instruments

Details of the instruments used for the present study are presented in Table 14.

Table 3.3 A list of chemicals and reference standards used in the present study

S. No. Reagent Manufacturer Grade Purity Properties

1 Sucrose Qualigens, India ExcelaR >99 White crystals

2 Yeast extracts powder Himedia, India - - -

3 Agar powder Himedia, India - - -

4 L-Ascorbic acid Sigma, USA - - White powder

5 Sorbic acid Sigma, USA - 99 White powder

6 Choline chloride Sigma, USA - 99 White powder

7 Wheat germ oil Sigma, USA - - -

8 p-Hydroxybenzoic acid

methyl ester

Sigma, USA - - White powder

9 Cholesterol Qualigens, India ExcelaR 99 White powder

10 Streptomycin sulphate Sigma, USA Sigma - White powder

11 Multivitamin capsule GSK, India Becadexamin - Sunset yellow

12 d- -Tocopherol acetate Sigma, India - - Wax like

13 Sodium hydroxide pellets Qualigens, India ExcelaR 98 Off white pellets

14 Sodium hypochlorite

solution

Qualigens, India SQ 4 (w/v) Colourless liquid

15 Silica gel (60-120 mesh) Qualigens, India SQ - White powder

Table 3.4 A list of solvents used in the present study

S. No. Solvent Purity

(%)

Manufacturer Grade Polarity

1 Ethyl acetate (CH3COOC2H5) 98 Qualigens, India SQ 4.30

2 Hexane (Petroleum fraction)

(C6H14)

98 Qualigens, India SQ 0.06

3 Acetone (CH3COCH3) 99.5 Qualigens, India ExcelaR 5.10

4 Butanol (CH3(CH2)3OH ) 99 Qualigens, India SQ 3.90

5 Ethanol (C2H5OH) 95 Procured from

licensed supplier, India

Distilled

spirit

5.20

Table 3.5 Instruments used in the present study

S. No. Name Details

1 Extraction apparatus Soxhlet apparatus, Borosil, India

2 Rotary-vacuum evaporator Buchi type, Khera Instruments Limited, Delhi, India

3 Mixer grinder HL 1606, Philips India Limited

4 Magnetic stirrer with hot plate Khera Instruments Limited, Delhi, India

5 UV chamber 365 nm, Jain Scientific and Glass Wares, Ambala,

India

6 Serological water bath 0-100oC, NSW India

7 Centrifuge HIMAC SCR20 BA, Hitachi, Japan

8 Shaker Kuhner, Lab-Therm Switzerland

9 Weighing balance (electronic) PE-3600, Deltarange, Mettler Instrument AG,

Switzerland

ER-182A, up to 0.01 mg, Afcoset, License A&D

Company Limited, Tokyo, Japan

FX-300, up to 0.001g, Afcoset, License A&D

Company Limited, Tokyo, Japan

3.5 Extraction and fractionation

The protocol followed for extraction and fractionation is given in the form of

flow chart (1 – 3) and the details are provided below.

3.5.1 Preparation of T. indica and E. camaldulensis crude extracts

3.5.1a Hexane extract (I)

The powdered material (50 g) was packed into a thimble made of Whatman

filter paper No. 1 and extracted with 500 ml of hexane solvent using soxhlet

extraction apparatus for 48 h until the solvent extracted no more colour. The

extract was concentrated under reduced pressure using rotary-vacuum

evaporator to yield the crude extract. The viscous solution of extract was

obtained from rotary-vacuum evaporator.

3.5.1b Ethanol extract (II)

The powdered material (50 g) was extracted with ethanol solvent as per the

method described above.

3.5.1c Ethanol sequential (III)

The residue remaining after the hexane extraction as described above was

subjected to ethanol solvent extraction using the same apparatus for 48 h. The

ethanol soluble portion was concentrated using rotary evaporator followed by

water bath drying.

3.5.2 Fractionation of the crude extracts

The concentrated ethanol extracts of T. indica and E. camaldulensis were

fractionated through partitioning with combination of solvents of varying

polarities. In addition to this, ethanol extract of E. camaldulensis was also

subjected to column chromatography.

3.5.2.1 Fractionation of T. indica ethanol extract

The steps followed for the fractionation of T. indica ethanol extract are reported

schematically in Figure 1. The crude ethanol extract weighing 5 g was dissolved

in 100 ml of ethyl acetate solvent and left overnight in the beaker. The ethyl

acetate soluble was collected in a separate flask and the residue was re-extracted

twice with ethyl acetate using 100 ml solvent each time. The ethyl acetate soluble

after 3 successive extractions were combined, filtered in vacuum, and

concentrated under reduced pressure using rotary-vacuum evaporator. The

blackish green viscous extract obtained was termed as ethyl acetate soluble

fraction (II a). The ethyl acetate insoluble portion was separated as brown

viscous extract, which was termed as ethyl acetate insoluble fraction (II b). The

ethyl acetate insoluble was further washed with 100 ml of 70% aqueous acetone

separating soluble and insoluble portions. The 70% aqueous acetone soluble

after 3 washings was concentrated under reduced pressure, thereby producing

greenish brown powder (II c). The 70% aqueous acetone insoluble upon drying

gave greenish black solid (II d).

3.5.2.2 T. indica alkaloid extraction

Crude alkaloids from T. indica were extracted as per the method suggested by

Bhutani et al. (1984). The ethanol extract was prepared as per the method

described in section 3.5.1a. The crude ethanol extract (200 g) was dissolved in

freshly prepared 0.5 M HCl solution. In total, 3600 ml of 0.5 M HCl solution

was used to dissolve the crude extract. After filtration, the HCl insoluble portion

was separated as brownish black viscous extract (IV a).

The HCl soluble portion in different beakers (500 ml capacity) were heated at

50-55 0C in water bath for 10-15 min in order to increase the solubility and left

overnight at room temperature. The 0.5 M HCl soluble portions were combined

and filtered under vacuum. The 0.5 M HCl soluble portion after filtration was

taken in a separatory funnel and subjected to extraction with sufficient amount

of ethyl acetate till it gave colourless washings. Ethyl acetate (1 L x 4) was used

primarily to remove chlorophyll. The leftover aqueous acidic solution was

further acidified to pH 2.0 using 2 M HCl solution. This aqueous acidic solution

having pH 2.0 was further extracted with ethyl acetate (1 L x4) in a separatory

funnel to remove neutral components from the solution mixture. The remaining

aqueous acidic solution was then made alkaline (pH 9.0) with 30% ammonium

hydroxide (NH4OH) solution. The alkaline solution was then repeatedly

extracted with ethyl acetate (1 L x3). The ethyl acetate extracts were combined,

washed with water, dried and evaporated under vacuum to yield crude total

alkaloids as a brown solid (IV b). Schematic procedure for extraction of alkaloids

is presented in Figure 2.

Confirmation of alkaloid content

As suggested by Mukherjee (2002), qualitative chemical analysis was performed

based on the fact that most of these alkaloids in acid solution form precipitates

with heavy metal reagents. These are known as general reagents for alkaloid

analysis. The alkaloid content was confirmed as per the following method:

Dragendorff’s reagent test: The freshly prepared reagent was obtained

from Dr Mohammed Ali, Centre for Pharmacy, Jamia Hamdard, New

Delhi. To the alkaloidal solution when added one drop of the reagent,

produced an orange-red precipitate.

Wagner’s reagent test: This was prepared by taking 1.27 g of iodine and

2 g of potassium iodide in 5 ml of water and the volume was made up to

100 ml with distilled water. To the alkaloidal solution when added one

drop of the reagent, produced reddish brown precipitate.

10% Tannic acid solution: This was prepared by dissolving 10 g tannic

acid in 100 ml of water. The solution produced a buff coloured

precipitate with alkaloids.

3.5.2.3 Extraction of tannins from E. camaldulensis leaves

The steps followed for fractionation of E. camaldulensis ethanol extract are

reported schematically in Figure 3. The crude ethanolic extract (10 g) was

dissolved in ethyl acetate (250 ml x 4). The ethyl acetate extracts were combined

and concentrated using rotary-vacuum evaporator to yield dark brown-green

powder (II a). Ethyl acetate water insoluble (II b) was rejected. This powder was

re-dissolved in 500 ml of 70% acetone (aqueous) and subjected to filtration.

Insoluble green solid powder was obtained on filtration (II c). The left over red-

brown water filtrate (II d) on acetone evaporation was divided into two equal

parts (75 ml each). The first red-brown water filtrate (75 ml) was subjected to

extraction with n-butanol (250 ml x 3) separating n-butanol and water layer.

The n-butanol soluble extracts were combined and concentrated in vacuum

using rotary evaporator producing brown viscous semi solid (II e). Sodium

bisulphite (1.5 g) as suggested for the extraction of high purity tannins

(Anonymous, 1952) was added to the second water fraction (75 ml) and kept

overnight. Sediments were removed by centrifugation at 10, 000 rpm for 5 min

as brown solid (II f). Hydrolysis of the remaining reddish brown water fraction

was done with 2N HCl, placed in a water bath at 80oC and neutralised with 30%

aqueous Na2CO3 solution (w/v). After neutralisation, 3 g of sodium bisulphite

was added again and kept overnight. Sedimentation was collected by

centrifugation as reddish-violet crystals (II g).

Tannins were also extracted directly from leaf powder using traditional method

(Foo and Porter, 1980). Leaf powder (50 g) was subjected to 70% aqueous

acetone (500 ml) in a soxhlet apparatus for 48 h. The 70% aqueous acetone

soluble was filtered and subjected to rotary vacuum evaporator for solvent

evaporation. The left over water fraction was extracted with n-butanol (500 ml

x 3) in a separatory funnel. The n-butanol extracts were combined and

concentrated in vacuum using rotary evaporator. This led to the production of

brown solid powder termed as crude tannins (IV).

Extraction of tannins by WHO recommended procedure

Leaves of E. camaldulensis were shade dried and ground to fine powder in a

mixer grinder. The known amount (25 g) of powdered material was taken into a

conical flask to which 150 ml water was added. The mixture was allowed to heat

over a boiling water bath for 30 min. After heating and subsequent cooling, the

mixture was transferred to a 250 ml volumetric flask and dilute to volume with

water. The mixture was allowed to settle. The liquid was filtered through a filter

paper, discarding the first 50 ml of the filtrate.

Out of this filtrate, 50 ml of the water-soluble extract was concentrated using

rotary evaporator followed by water bath drying. The residue was dried in an

oven at 1050C for 4 h and weighed accurately (T1). Out of the remaining filtrate,

80 ml of the plant material extract was taken in a separate conical flask to which

6 g of hide powder was added. The mixture was allowed to shake for 60 min. The

liquid was then filtered. Following this, 50 ml of the clear filtrate was taken to

dryness. The residue was dried in an oven at 1050C for 4 h and weighed

accurately (T2).

Consequent upon this, 6 g of hide powder was taken in a separate conical flask,

added 80 ml of water and allowed to shake for 60 min. The mixture was filtered

and 50 ml of the filtrate was taken to dryness as per the method described

above. The dried residue was weighed accurately (T0).

The quantity of tannins as a percentage was calculated using the following

formula:

Tannins (%)= [T1 – (T2 - T0)] x 500, where w= weight of the plant material (g)

W

Confirmation of tannins

The tannins thus produced i.e. brown solid powder (II f) and reddish-violet

crystals (II g), the n-butanol layer (IV) and as per WHO recommended

procedure were subjected to standard tests for further confirmation based on

some of their chemical reactions as suggested by Mukherjee (2002).

Accordingly, the following colour reactions were performed taking tannic acid as

a standard for tannin class of compounds.

Ferric chloride test: A small quantity of ferric chloride (5 mg) when

added to an aqueous solution of the tannins (0.1 g in 10 ml water)

produced a bluish green colouration following reaction.

Precipitation by alkaloids: A small quantity of alkaloids (extracted from

T. indica) when added to an aqueous solution (0.1 g in 10 ml) of tannins,

a pale-white precipitate was produced after 3 h, which was not dissolved

on shaking.

Precipitation by heavy metals: A small quantity of lead acetate (5 mg)

when added to an aqueous solution of the tannins (0.1 g in 10 ml water)

produced a pale-yellow precipitate following reaction.

Yield of tannins obtained with different procedures were compared for efficiency

of extraction procedures.

3.5.2.4 Column chromatography of E. camaldulensis

Crude ethanol extract of E. camaldulensis was also subjected to column

chromatography to identify active fraction other than the tannins.

3.5.2.4a Column preparation and loading

The essential part of the apparatus consisted of a long narrow glass tube (100 cm

long and 3.5 cm diameter) with a capacity to hold 200 g column packing

material. Activated silica gel (60-120 mesh) was used as packing material for

this purpose. Activation was done by heating the silica gel in an oven at 120 oC

for 60 min. Slurry of the silica gel was prepared in hexane solvent for

introducing the mixture on to the column. The slurry was poured through the

funnel into a clean dry column clamped vertically and adsorbent was allowed to

settle evenly for 48 h. In order to obtain uniform packing, gentle tapping of the

column was done with a wooden rod. Solvent was allowed to elute and more

slurry was added until required length of the column was obtained. Fresh

solvent was allowed to flow through the column under the hydrostatic pressure

to remove air bubbles, if any, and to avoid the formation of cracks and channels

as this may lead to distortion of adsorption bands. Freshly prepared 20 g crude

ethanol extract evaporated to dryness under reduced pressure was re-dissolved

in 25 ml of ethanol solvent adding column adsorbent equal to 3 times its weight

(60 g silica gel). The extract solution adsorbed evenly on the silica gel and

allowed the solvent to evaporate completely. The adsorbent loaded with crude

extract was then added to the column top and packed into an even layer. After

introduction of the extract on to the column, initial adsorption took place

rapidly and hence considered ready for chromatogram development.

Figure 1: Tylophora indica extraction and fractionation procedure

Shade dried T. indica leaf powder

Soxhlet extraction

with hexane (50 g

in 500 ml)

Soxhlet extraction

with ethanol (50 g

in 500 ml)

Concentratio

n (RE)

Concentratio

n (RE) Residue

Hexane extract (I)

(1.05 g)

Ethanol extract

(II)

(9.50 g) Soxhlet extraction with

ethanol

Concentratio

n (RE)

Ethanol sequential

extract (III) (6.50

g))

Ethyl acetate

extraction

(5g dissolved in 100

ml x 3)

Ethyl acetate

layer

Brown viscous

insoluble

(IIb) (3.4 g) Concentration using rotary

evaporator

Blackish green viscous extract (1.6

g) (IIa)

Washed with 70% aq.

acetone

(100 ml x 3)

Aqueous acetone layer

Greenish

black powder

(0.93 g)

(IId) Water layer

Greenish brown solid (2.4 g)

(IIc)

Filtration

(Evaporation of

acetone using

rotary evaporator)

Filtration

Filtration

Crude ethanol extract of T. indica (200 g)

Dissolved in 0.5 M HCl (3.6 l)

Brown black viscous insoluble

(IVa) (52 g)

Aq. Acidic layer

Extraction with ethyl acetate (1 l x 4)

Aq. layer Ethyl acetate layer

(remove chlorophyll)

Rejected Add 2 M HCl to obtain pH

2.0 (acidification)

Extraction with ethyl acetate (1 l x 4)

Aq. layer Ethyl acetate layer (remove

neutral components)

Rejected Added 30% NH4OH

solution to obtain pH

9.0 (Basification)

Extraction with ethyl acetate (1 l x 3)

Aq. Layer

(Rejected)

Ethyl acetate layer

Washed with H2O dried and

evaporated

Viscous solid (crude alkaloids)

(4.1 g) (IVb)

Figure 2: Procedure for Tylophora indica alkaloid extraction

Hexane extract

(2.35 g) (I)

Soxhlet extraction with

ethanol (50 g in 500ml)

Concentration

Ethanol extract (10 g) (II)

Dissolved with ethyl acetate

(250 ml 4)

Ethyl acetate layer Ethyl acetate water

insoluble (IIb)

Concentration in vacuum using

rotary evaporator

Dark green solid (4.70 g)

(IIa)

Dissolved in 500 ml of

70% acetone

Filtration

Soxhlet extraction with

Hexane (50 g in 500 ml)

Concentration Residue

Soxhlet extraction

with ethanol (500 ml)

Concentration

Ethanol sequential

(7.2g) (III)

Green powder (0.578 g) (II c) Dark brown filtrate (150 ml) (II d)

Divided into 2 equal parts

n-Butanol

Partitioning(250 ml 3) Added sodium bisulphite

(SB) (1.5 g)

Sedimentation (Brown in colour)

Kept Overnight

Removal of sedimentation using

centrifugation (10,000 rpm; 5min)

Brown powder (II f)

(crude tannins) (2.467 g)

(IIf)

Water layer (reddish brown)

Added 2N HCl & heat at 80 C

on a water bath for ½ hr

Neutralized with aq. Na2CO3 Added 3 g of SB

bisulphite

Kept Overnight Sedimentation Centrifugation

(10,000 rpm; 5 min)

Hydrolysed Tannins

(410 mg) (IIg)

n-butanol Water layer

Concentration

Brown viscous semi-

solid (1.88 g) (IIe)

Figure 3. Eucalyptus camaldulensis extraction and tannin preparation

Soxhlet extraction with 70%

acetone (50 g in 500 ml)

Filtration and

Concentration

Partitioning with

n-butanol (500 ml X 3)

Concentration

Brown solid

(crude tannins)

(5.4 g) (IV)

E. camaldulensis leaf powder

Materials & Methods

3.5.2.4b Elution of the column

The ethanol crude extract was chromatographed on silica gel (60-120 mesh).

Column elution was carried out with increasing polarity of hexane and ethanol

solvent mixture in the ratio of 100:0, 90: 10, 80: 20, 70: 30, 60:40, 50: 50, 40:

60, 30: 70, 20: 80, 10: 90, 0: 100 respectively. In total eleven solvent mixtures

were used. One hundred ten fractions (each 45 ml) were collected during the

complete chromatogram development. These fractions were then grouped in t0

28 fractions based on the TLC pattern and then screened individually for their

growth inhibition action against H. armigera larvae by diet incorporation

method. These fractions were concentrated under reduced pressure in rotary-

vacuum evaporator. The weight of each fraction was recorded.

3.5.3 Thin Layer Chromatography (TLC)

The sample fractions were spotted on TLC plates using capillary tubes. The

plates were developed in hexane and ethyl acetate (6: 4) solvent system. After

that, the plates were observed under UV light closed chamber and then with

iodine. The spots were marked on the plates and the retention factor (Rf) values

were determined.

TLC plates of 0.5 mm thickness were used. Commercial grade TLC plates were

procured from Merck (0.2 mm, 20 x 20 cm, Aluminium, Silica gel). The plates

were activated at 120 oC in the oven for 30 min before use.

3.5.4 Yield

To establish the yield of the crude extract and fractions, the quantity was

determined gravimetrically by weighing the resulting crude extract or fraction

following removal of the extracting solvent.

Percent Yield= (weight of extract after solvent evaporation/ initial weight of the

powdered material) x 100

3.6 Insect bioassays

3.6.1 Selection of promising plant species

The powdered leaves/ rhizomes of the test plant species were utilized for further

experimental work as described below.

3.6.1a Chronic feeding bioassays

Experiments on effect of test plant species on growth and development of

H. armigera were conducted by allowing the neonate larvae (0-12 h old) to feed

on semi-synthetic diet containing the powder of the test material at 5% level on

w/w basis. The first instar larvae were released into glass vials (25 x 100 mm,

Borosil) containing treated diet with a soft camel hairbrush and plugged the

tubes with cotton plugs. Observations were taken to record the larval moulting

and mortality counts after every 24 h till adult emergence. Moribund larvae were

counted as dead. Larval weights were recorded on 7th and 10th day of the

treatment. Pupal weight was also recorded. Number of male and female

emerged were also recorded. Diet without any test material was taken as control.

There were 5 replicates for each of the treatment. Each replicate consisted of 10

larvae.

Following growth indices were calculated:

Growth index = Per cent pupation / Average duration of larval period

Survival index = Number of adults emerged on treated diet/ adults

emerged on control diet

Larval weight index = Mean live weight (mg) of the larvae on the treated

diet / weight on control diet

Pupal index = Average pupal weight on treated food / Average weight on

control diet

Larval pupal index = A + B / C + D, where

A= Average larval period on control diet

B= Average pupal period on control diet

C= Average larval period on treated diet

D= Average pupal period on treated diet

Developmental period in days was calculated separately for each stage (first

instar to pupal stage) based on mean value of the replicate using following

formula:

Developmental Period=

[Period in days of individual larva in each instars or pupa] X [number of days in

that stage]

Total number of live units in each instar or pupae

3.6.2 Evaluation of the crude extracts

Based on the comparative results from the studies of Objective 1, Tylophora

indica and Eucalyptus camaldulensis were identified as the promising plant

species and subjected to further evaluation. The crude extracts as prepared in

section 3.5.1 of the promising plant species T. indica and E. camaldulensis were

used for conducting insect bioassays to assess the growth inhibitory,

antifeedant, contact and oviposition deterrent activities.

3.6.2a Growth inhibitory properties

Effect of hexane and ethanol leaf extracts of T. indica and E. camaldulensis on

the growth and development of 0-12 h old neonates of H. armigera in chronic

feeding bioassay was studied by diet incorporation method using concentration

in the range of 0.005 – 3.0% on w/w basis. Extracts were evaporated to dryness,

weighed and taken with semi synthetic diet and mixed uniformly with other

ingredients by using magnetic stirrer. Larvae fed with normal diet served as

control. Larvae were allowed to feed on the test diets till the completion of their

larval period. Observations on the larval weight (7th and 10th day of treatment)

and pupal weight (4th day after pupation) were recorded. Mortality counts, if

any, were also taken into consideration. Neem seed kernel extract (NSKE) in

ethanol was taken as a standard for comparative biological activity. Each

treatment consisted of 3 replications. There were 10 larvae per replicate.

3.6.2b Feeding inhibition action

Soxhlet extracted hexane and ethanol leaf extracts of T. indica and

E. camaldulensis using concentrations in the range of 1-10% w/v were evaluated

for their antifeedancy against H. armigera larvae. 8 replications with 4 larvae

per replication were used for each treatment. These larvae were starved for 4 h

prior to conducting the experiment. The starved larvae were allowed to feed for

24 h on cabbage leaf-discs (12.5 cm2) treated with the test solution by dipping

for 5 s using no-choice bioassay. The solvent was removed by evaporation. Area

consumed by the larvae was recorded. The feeding inhibition (FI) was calculated

for each insect as:

FI = [C – T] X 100

[C + T]

Where ‘FI’ is the feeding inhibition,

‘C’ denotes consumption in control disc and

‘T’ is the consumption in the treated disc.

Percent antifeedant values were subjected to regression analysis and the

effective concentration was calculated from the line of best fit.

3.6.2c Direct contact toxicity

Direct contact toxicity of the test preparations was determined by topical

application method against third instar H. armigera larvae. The test solutions

were prepared with water (w/v) containing 1% emulsifier (w/v) of the total

volume prepared. Test solutions in 5 l dose were applied to the dorsum of

newly moulted third instar larvae using a fine micropipette. Concentrations

ranging from 0.01 to 3.0% were tested. Water alone was taken as control. Ten

larvae per replicate in 3 replications were taken for each treatment. Treated

larvae were reared on artificial diet and observations on mortality counts were

recorded daily at 24 h interval up to 3 days.

3.6.2d Oviposition deterrent (OD) activity

Anti-oviposition activity of the crude extracts was tested against H. armigera

moths. Both the female and male adults after emergence from the pupae were

kept separately for two days. One pair of the moth was released inside a glass

jar, which was covered at the top with a muslin cloth. The test extract solution

was applied on the muslin cloth by dipping the cloth for 5 s and allowed to dry.

Different concentrations of the emulsified test solutions (w/v) were prepared in

water with addition of 1% Triton X-100 (v/v). The concentrations tested were

0.5, 1.0, 1.5 and 2.0%. The jars (five replicates) were kept in the BOD incubator.

The eggs obtained on the 4th and 5th day were collected and taken for the

analysis. The percent reduction in mean number of eggs was calculated in

comparison to control using following formula:

Percent reduction= 100 - [Mean number of eggs obtained in treatment] X 100

[Mean number of eggs obtained in control]

The eggs were monitored for larval emergence. Emulsified water treated muslin

was taken as control.

3.6.3 Evaluation of fractionated extracts

The different extracts obtained after fractionation of the crude ethanol extracts

of T. indica and E. camaldulensis as per the procedure mentioned in section

3.5.2.1, 3.5.2.2 and 3.5.2.3 were screened for their growth inhibition action

following diet feeding bioassays mentioned in section 3.6.2a. Concentrations

ranging from 0.01 to 1.o% were tested under similar conditions. For comparative

analysis, NSKE was taken as a positive control. GI50 values based on 7th and 10th

day larval weight were compared with the GI50 values of crude ethanol extract.

Dimlin was taken as a standard for comparative biological activity.

The relative toxicity values of the isolated fractions were calculated as follows:

Relative efficacy (RE)= GI50 value of fraction/ GI50 value of crude ethanol extract

3.6.4 Evaluation of T. indica alkaloids and E.

camaldulensis tannins

Alkaloids constitute a major class of chemical group present in T. indica (Jain

and Agrawal, 1991), while E. camaldulensis leaves contain tannin class of

compounds (Anonymous, 1952, Mukherjee, 2002). Therefore, alkaloids from T.

indica and tannins from E. camaldulensis, besides growth inhibition, were also

screened for their feeding inhibition and oviposition deterrent activity at

different concentrations following insect bioassays mentioned in section 3.6.2b

and 3.6.2d. In case of feeding inhibition bioassays, crude alkaloids and tannins

were tested with two larvae per replication per treatment in four replications.

3.6.5 Identification of the active fraction(s) of

E. camaldulensis thorough column chromatography

E. camaldulensis crude ethanol extract was also subjected to column

chromatography to identify active fraction other than tannins. Based on larval

weight recorded on 7th and 10th day of treatment following chronic feeding

bioassay on semi-synthetic diet at 0.5% concentration (w/w) using neonate

larvae. Ten larvae were tested for each treatment and percent growth inhibition

values were calculated. Percent survival of the larvae was also recorded

following treatment response. Fractions with more than 80% growth inhibition

of the larvae were considered as active.

3.7 Statistical analysis

Standard deviation values were calculated for larval and pupal developmental

period. For testing the significant differences with respect to control, analysis of

variance (ANOVA), SNKT or t-test (mean comparison) and goodness of fit

(G-test for percent values) were used. Linear regression analysis was performed

for all dose response experimental data. Probit analysis was used for calculation

of LC50 /GI50 / FI50 / LT50 values (Finney, 1971). Data analysis was carried out

using software Costat (CoHort, Berkeley California) and SPSS (version 9).