REPORT OF THE SECOND ANCAP SUMMER SCHOOL, 2003

 

VENUE: DEPARTMENT OF CHEMISTRY

UNIVERSITY OF NAIROBI,

NAIROBI-KENYA

DATE:  8TH TO 19TH SEPTEMBER 2003

 

 
EXECUTIVE SUMMARY

 

The summer school was conducted from 8th to 16th September of 2003, under the co-ordination of Professor Shem O. Wandiga and four facilitators. Participants of the summer school were drawn from Kenya, Uganda, Tanzania and Ethiopia; mainly MSc, PhD students and technical staff involved in pesticide research and analysis. Two participants were registered from both Uganda and Ethiopia, seven from Tanzania and nine from Kenya, adding up to 19 participants (see list of participants, attachment five).

          

Several topics were covered during the summer school including general survey of pesticide research work in Kenya comprising of field and laboratory studies, field sampling logistics, theory and application of radioisotopes in pesticide research. Further topics covered were application of TLC in qualitative and quantitative analysis of pesticides, gas chromatograph and its application in pesticide research using ECD, TSD and FID detectors, and overview of proposal writing for research grants (See attachment one, two and three).

          

The course conception and design was based on alleviating the current gross state of pesticide pollution in the environment. Systems simulating the natural ecosystem consisting of fish, weeds, sediments and water were set up using aquarium tanks. Distribution, partitioning and dissipation of p,p-DDE and lindane were studied using unlabelled standards, tagged with known activities of respective labeled pesticide compounds. 

               

The practicals aimed at creating awareness of the possible ways of remediating the environment by means of phytoremediation, chemodynamics, and bioaccumulation; extending knowledge of adsorption onto soil surface; and investigating the extent of dissipation of the pesticides from water bodies.       

                

The practical work involved sampling procedures, sample preservation, sample preparation and extraction using soxhlet and orbital shaker, sample treatment and analysis using liquid scintillation counter and gas chromatograph fitted with electron capture detector.

             

Participants were also trained on data handling for liquid scintillation techniques including normalization and calibration of the LSC machine, generation of calibration curve using unquenched standards, data manipulation and interpretation. Application of soxhlet and orbital shaker extraction techniques were set up mainly to determine extraction efficiencies and hence show how this knowledge can be used in selecting the extraction method. This was facilitated by spiking three replicates of soil, weed, sediments and water samples with known concentrations of lindane and DDE; extracting using orbital shaker and soxhlet methods, and analyzing by liquid scintillation counter and GC-ECD.

              

Based on the radioisotope techniques, the participants were informed of other field modelling studies that include mineralization, photolysis, sorption (adsorption-desorption), leaching, bioconcentration, depuration, volatilization, and metabolism among others. Eleven printed papers involving these studies were given to representative institutions to enable the participants for further references. Introduction to the operation of the gas chromatograph, including column fitting, equilibration, injection, and data acquisition and handling were also covered.

                    

Further training on application of TLC in qualitative and quantitative analysis of photosynthesis inhibiting herbicides was conducted using Amaranthus as an example. Training on mobile phase preparation, equilibration, spotting, developing, spraying and scanning were conducted.

 

ACKNOWLEDGEMENT

 

The ANCAP organising committee highly acknowledgeS the grant support from IPICS used for organising the workshop and symposium, for buying chemicals, consumables, and in facilitating the travelling and upkeep of participants from Ethiopia, Uganda, Tanzania and one upcountry Kenyan, who attended the summer school.


DETAILED REPORT

 

PROGRAMME

 

 

Day 1.    

8.30             Registration

9.00             Official opening by chairman, Department of Chemistry

9.30             Introduction to pesticides research in Nairobi- SO WANDIGA

10.30           Tea/Coffee Break

11.00           Field Sampling- V. Madadi

12.00           Group Discussion

12.30           Lunch Break

14.00           Radio Isotope methods- LSC Lecture, V. Madadi/ F. Kengara

15.00           Preparation of model freshwater ecosystem aquarium

16.00           Tea/Coffee Break

16.30           Lab work contd.

17.30           End of Day

Day 2.

8.30              Sample Preparation

10.30           Tea/Coffee Break

11.00           GC Analysis Lecture- J. Makopa

    FID

    ECD

    PND (TSD)

12.30            Lunch Break

14.00            Sample Preparation

16.00            Tea/Coffee Break

16.30            Sample Preparation

17.30                  End of Day

Day3-8.

8.30             Sample Analysis

10.30            Tea/Coffee Break

 

11.00            Group Discussion

12.30            Lunch Break

14.00            Sample Analysis

16.00            Tea/Coffee Break

16.30            Sample Analysis

17.30            End of Day

Day 9-12      ANCAP SYMPOSIUM.  Workshop participants to give short presentations on topics of their choice

.

 TRANSPORT, FATE AND PERSISTENCE OF PESTICIDES

 

DISSIPATION, DISTRIBUTION AND PARTITIONING OF g-HCH AND

p,p-DDE IN ECOSYSTEMS SIMULATING SAGANA DAM FISHERIES

 ABSTRACT

 

Most of the OC pesticides have been banned under the Persistent Organic Pollutants Stockholmn Convention, except where extremely inevitable for public health like malaria control, restricted use is allowed. Some banned or restricted OC compounds are still found in the environment in unexpected concentrations and their eventual elimination is required. The current study was set up to investigate the dissipation, distribution, and partitioning of p,p-DDE and lindane in systems simulation natural ecosystem containing water, sediments, weeds and fish. Dissipation of p,p-DDE and lindane from the aqueous media were described using the first order equation. p,p-DDE exhibited two phases with rate constants k1= 2.02180.0569, k2 = 0.47370.0344 per day. Lindane also exhibited biphasic dissipation rates, but with lower rate constants, k1 = 0.50960.0040, and k2 = 0.17090.0109 per day. Sorption contributed to most of the pesticide losses from aqueous systems, followed by ingestion by fish, and lastly uptake by weeds.

 

OBJECTIVES

   To train in the application of radioisotopes and Gas Chromatographic techniques as appropriate tools for environmental monitoring of pesticide pollution, using models simulating Sagana Dam ecosystem.

   To establish the extraction efficiencies of soxhlet and orbital shaker methods of chemical pesticide extraction.

 

1. INTRODUCTION

 

The application of radioisotopes in research traces its way back to 1934. However, the technique has not been extensively applied compared to the chromatographic techniques like GC, HPLC, and TLC. This course was designed to introduce/ extend the knowledge about the application of Liquid scintillation counting and Gas Chromatographic technique as applied in pesticide research, using g-HCH and p,p-DDE.

 

The commercial formulations of g-HCH [1,2,3,4,5,6-hexachlorocychlohexane] generally contains alpha, beta, gamma and delta isomers of which the gamma isomer is the most insecticidal while the beta isomer is the most persistent. Although the beta isomer is a minor constituent in the commercial formulation of HCH, its entry (through persistence) into the food chain following intensive use has been reported.

 

Hexachloracyclohexane (HCH) has been intensively applied in the tropical rice soils to control the common rice insect pests and in seed dressing. The degradation of the four isomers of HCH takes place faster in non-sterile soil than in sterile soils. Also degradation of gamma-HCH has been noted to occur under flooded soils by means of bacteria (Clostridium sphenoides). The bacterial degradation converts the gamma-HCH to gamma-3,4,5,6-terachloro-1-cyclohexane [TCCH], and the alpha-HCH to delta-TCCH. The gamma and delta-TCCH are further metabolised leading to nearly complete dechlorination of the gamma-HCH, with the formation of chlorine ions in the stoichiometric amounts.

             

Studies on fish samples from Lake Naivasha showed presence of Lindane, DDT and dieldrin (GITAHI, 1999).  Further research work conducted on Lake Victoria showed that lindane, DDT and its metabolites were the most common organochlorine pesticides in the samples (Mitema and Gitau, 1990).  Therefore, more studies were recommended on these pesticides to determine their distribution, partitioning, and transport in natural ecosystem.

 

2. MATERIALS AND PROCEDURES

 

2.1 CHEMICALS AND OTHER ITEMS USED

 

  Unlabeled g-Hexachlorocyclohexane (g-HCH), p,p-dichlorodiphenylethane (p,p-DDE), C-14 p,p-DDE from SIGMA Chemicals USA, and C-14 g-HCH from the institute of radioisotopes, Budapest-Hungary, were used in the study. Sodium chloride (AR), activated carbon were purchased from ZETA chemicals, 2,2p-phenylenebis(5-phenyl oxazole or POPOP, and 2,5-Diphenyloxazole (PPO) were obtained from Kodak, USA, and Fisher Chemicals respectively. GPR methanol, hexane, acetone, HPLC grade toluene, Diethyl ether, dichloromethane, Triton X-100 and florisil (60-100 mesh) were purchased from Kobian Kenya LTD. Anhydrous sodium sulphate (AR) and disodium hydrogen phosphate (AR) were purchased from ZETA chemicals LTD.         

 

 2.2 EXPERIMENTAL PROCEDURES

 

2.2.1 AQUARIUM TANKS PREPARATION

 

The aquarium tanks were set up to study partitioning of the 14C g-HCH and p,p-DDE between water, fish, sediments and weed samples. Fresh water, sediments, fish and weeds collected from Sagana Dam fisheries were used to set up three model ecosystems, comprising of experimental and control tanks. Water was kept in the dark for two weeks before being introduced into the aquarium tanks. This was to kill the plankton organisms that could change the pH of the systems.

 

Two experimental tanks (30x15x15) were filled with 13.5 kg and 10.5 kg of sediments (wet basis) for lindane and p,p-DDE, respectively, and  60 litres of water, 20 (5-10cm long, average weight 20g) Tilapia azilli, and six weed samples added to each tank. Algal growth on the tank walls was scrapped off to maintain the pH of the water at 7.3. The water level, hence its density, salinity and chemical balance of each tank was maintained by adding 200-250ml distilled water daily.

            

The systems were left to stabilise for two weeks, and the water temperature equilibrated to room temperature before spiking solutions of radiolabelled pesticides to give 10 ug/ml of lindane, and 0.1 ug/ml of p,p-DDE. Water, fish, sediments and weed samples were taken at intervals of 0, 1, 2, and 4 days for the determination of g-HCH and p,p-DDE residues in the compartments of the ecosystems.

            

 Control system was set up by filling a third aquarium tank with 60 litres of water, 20 (5-10 cm long) Tilapia azilli and six weed samples. The control tank was not treated with the pesticide. The temperature and pH were maintained at 220C and 7.3 respectively, throughout the study period. The water in the control tank was aerated at a rate of 2900cm3/min using a centrifugal air pump (model Elite 803), whereas the experimental aquarium tanks was aerated at a rate of 1800cm3/min using force1 and AZOO-3500 type air pumps. All systems were illuminated with florescence lamps for 8 hours a day. The fish in the systems were fed on flakes once every three days.

 

2.2.2 SAMPLING AND SAMPLE PREPARATION
 

2.22.1 Sediments

Sampling of sediments was done by vertically scooping out samples using a glass tube (50 cm long and 1.5 cm id) from different spots in the tank to obtain representative samples. The samples were mixed on aluminium foil, and two sets taken in triplicates. One set was used for extraction, and the second used for moisture content determination. Three 10 g replicates of sediment samples were mixed with 4-6 g of anhydrous sodium sulphate, transferred into 150 ml Teflon vials.50 ml portions of triple distilled methanol were added to each vial and placed on orbital shaker for four hours. Each extract was concentrated to 10 ml using a LABCONCO rotor-evaporator, and stored at 40 C for clean up prior to GC analysis and liquid scintillation counting.

             

The extractable residues were determined by counting the radioactivity in 1ml of the extract in a Packard Tricarb 1000 LSC. The counting was done by adding 6 ml of the scintillation cocktail (a solution of 4-g PPO and 0.25 g dimethyl POPOP in 1.0 L of toluene) to 1 ml aliquots of samples in 20 ml glass vials.

 

22.2.2   Fish

Three fish samples were harvested at each sampling time. The samples were rinsed with distilled water, and dried with paper towels. The samples were homogenised by grinding in mortar and pestle before three replicates of 20 g samples were taken. The weighed samples were ground with 4-6 g anhydrous sodium sulphate in a mortar and pestle until they were lumpy, and transferred into 250ml Erlenmeyer flasks. 50 ml portions of triple distilled hexane were added to each sample and placed on orbital shaker for four hours.

             

The extracts were concentrated to 10 ml using LABCONCO rotor-evaporator, and kept at 40C for clean up prior to GC and LSC analysis. 1ml aliquots of extracts were mixed with 6ml scintillation cocktail and counted in Parckard Tricarb 1000 LSC. The fish samples not extracted immediately were wrapped in pre-cleaned aluminium foil, labelled and stored in a deep freezer at temperatures between 200 C and 00 C.

 

2.2.2.3  Water

The determination of radioactivity in the water samples was achieved by taking 1ml aliquots in triplicate, adding 6ml scintillation cocktail and directly counting in the Packard Tri-carb 1000 LSC. The cocktail for aqueous samples was prepared by dissolving 4 g of PPO and 0.1 g of dimethyl POPOP in 1 L mixture of toluene and Triton X-100 in 2:1 ratio.

 

2.2.2.4 Weeds

 

The weed samples from the tanks were harvested, rinsed with distilled water and air-dried overnight in the laboratory. The air-dried samples were chopped, ground and three replicates of 10 g samples taken. The replicates were ground with 4-6 g of anhydrous sodium sulphate in a motar and pestle, and mixed with 50 ml triple distilled methanol. The samples were extracted by placing on the orbital shaker four fours.

 

The extracts were concentrated to 10ml, decolourised with activated charcoal and the extractable residues determined by counting the radioactivity in 1ml aliquots. The remaining extracts were stored in a refrigerator at 40 C for clean up prior to GC analysis.

 

2.2.2.5  Blank samples

The blank samples for each compartment of the aquarium tank were prepared, extracted and analysed following the same procedures as above. The results obtained were used to make background corrections. Sampling done between 0 and 30 minutes was considered as 0 day sampling.

 

2.3    Determination of Extractable 14C-gHCH and p,p-DDE residues by LSC

The radioactivity in 1ml aliquot and final volume of extract were used to calculate the total radioactivity in the sample. Calculating the natural radioactivity in the blank samples (in disintegration per minute, dpm) and deducting from the value obtained for corresponding spiked samples did background corrections.

 

2.4 Moisture content

The moisture content of each sediment sample was determined by heating 2g of the air-dried raw (un-extracted) sample in a GallenKamp oven at 1050 C for overnight. The cleaning of the crucible was done by soaking in general purpose detergent for 2 hours, washing with water and rinsing with acetone. The weights of the crucible with content before and after heating in the oven were recorded and the weight difference used to determine the moisture content of the sample at that time of extraction (UNEP, 1982).

 

2.5 Determination of the pH of sediments and water samples.

Fisher scientific pH meter was used to determine the pH of the sediments and water samples. The pH of sediments was measured by adding 25 ml of distilled water to 10g of blank sample, to form a 2:5 sediment-water suspension. The mixture was shaken for 30 minutes on orbital shaker before an electrode was dipped into the suspension to get pH reading.

 

 pH of water was determined by fetching 50 ml water in a beaker and taking direct reading. All the readings were recorded at 220 C. The buffer solutions of pH 4, 7 and 10 were used to standardize the pH meter before taking the readings.

 

2.6  Recovery of the extractable pesticides from sediments, fish, water and weeds.

The recovery experiments were conducted to estimate the amount of the pesticide that could be recovered as extractable residues from fish, water, sediments and weed samples. This was done by spiking 2000mg of 14C-p,p-DDE (in acetonitrile) to untreated 10 g of sediments, fish, weeds, and 3900 mg to 500 ml of water. Extraction of sediments, weeds, and fish were done in 50ml hexane (for fish) and methanol on orbital shaker for four hours. Extraction for water was done by solvent-solvent extraction method. 500 ml of distilled water was transferred into a 1litre separatory funnel, spiked with the pesticide solution, and mixed. 25 ml of 0.2 M disodium hydrogen phosphate buffer was added to each sample, and pH adjusted by adding drops of 0.1 N sodium hydroxide or 0.1 N HCL solution appropriately to get pH 7. The neutral solutions were treated with 50 g sodium chloride to salt out the pesticides from the aqueous phase, before adding 30 ml of HPLC dichloromethane. The aqueous-organic mixture was shaken for two minutes and allowed to settle for 30 minutes to enhance separation of the phases. The organic layer was collected in 250ml Erlenmeyer flask and kept at 40 C in a refrigerator. Extraction was repeated twice using 30 ml portions of dichloromethane and extracts combined.

 

 Lindane extraction efficiencies were determined by spiking 800 mg of the compound to 10 g of sediments, fish and weeds, and 80 mg of 14C-lindane to 500 ml of water. The same extraction, clean up and analytical procedures were followed as above. Three replicates of 10g sediment, fish and weed samples, and 500ml water samples were taken for extraction.

 

2.7  Clean up of water, fish, aquatic weeds and sediments

The concentrated fish, sediments, weed and water sample extracts were purified from co-extractants by passing through a 50 cm long column (2cm id) packed with 10 g florisil (magnesium silicate, 60-100 mesh), topped with 2 g of anhydrous sodium sulphate. The pigments from weed and sediment samples were removed by adding 0.5 g activated charcoal on top of the anhydrous sodium sulphate.

 

The florisil column was first pre-wet with 40-50 ml hexane, before the pesticide residues were eluted with 200 ml of 6% diethyl ether in hexane. The elutes were concentrated to near-dryness using LABCONCO rotor-evaporator, reconstituted in 5 ml HPLC hexane, and stored at 40 C for GC analysis.

             

The anhydrous sodium sulphate and florisil used in the clean up were activated by baking overnight at 3500 C and 2000 C respectively, and cooled to room temperature in an airtight desiccators before they were used in the clean up process (UNEP, 1982; UNESCO, 1993).

 

2.8  Clean up of glassware and other items

Cleaning of the glassware was accomplished by soaking in a general-purpose detergent for at least two hours, and rinsing with distilled water before drying in the oven at 1050 C. The dry containers were cooled and rinsed with methanol. Rinsing with methanol and baking at 4500 C for two hours did the cleaning of aluminium foil (Gold-Bouchot, 1993).

 

2.9  Gas Chromatographic analysis of pesticide residues

Identification and quantification of the g-HCH and p,p-DDE residues in the sediments, water, fish and aquatic plant samples by chromatographic technique were accomplished using a Varian Star #1 CP-3800 Gas Chromatograph equipped with ECD  at 3000 C, column oven temperature programmed from 1500 C to 2000 C, and Nitrogen gas constant column flow rate of 7.5 ml/min. A CP-SIL 8 CB capillary column of dimensions 10 m x 0.25 mm x 0.25 mm, was used in this study, and injector temperature maintained at 2500 C. 1uL of the cleaned samples extracts in hexane (HPLC grade) were taken for injection after appropriate dilutions .The respective peaks were identified by comparing their retention times with those of the standards run separately, and quantified by drawing a calibration curves.

 

2.10 Mass balance of total 14C-g-HCH and p,p-DDE

Mass balance in the compartments of the aquarium tanks, was done by accounting for the total amount of 14C g-HCH and p,p-DDE introduced to the tanks. The total g-HCH and p,p-DDE residues in each component were evaluated by adding the residues extracted from each compartment at the end of the experiment.

 

 

3. RESULTS

 

Table 3.1: Extraction efficiencies of pesticides in water, fish and sediments

 

Pesticide

Water

Sediments

Fish

 

%

%

%

p,p-DDE

99.83583.7423

89.28322.3309

85.21191.6968

 

 

84.92734.6659*

 

 

 

 

 

LINDANE

96.04139.1638

90.97622.9361

86.43233.3966

 

 

86.42124.2732*

 

 

 

 

 

* Extracted using Soxhlet method

 

Higher recovery rates of the two pesticides were observed in aqueous system compared to sediments and fish. p,p-DDE showed recoveries of 99.83583.7423, 89.28322.3309, and 85.21191.6968 percent for water, sediments and fish samples respectively. Lindane had mean recovery rates of 96.04139.1638, 90.97622.9361 and 86.43233.3966 percent for water, sediments and fish respectively, (Table 3.1). The observed mean recoveries were in the range reported in earlier studies (Lalah, 1993; Mughenyi, 1988). The orbital shaker extraction method was observed to have higher recovery rates compared to the soxhlet extraction (Table 3.1). Higher levels of p,p-DDE were recovered from aqueous samples compared to lindane. This could be accounted for by higher solubility of the former and hence more likely to be retained in the aqueous media.  In general, recovery rates increased from fish to sediments to water for both pesticides.

 

Table3.2: Extractable residues of p,p-DDE from water, sediments, weeds and fish in    ecosystem simulating Sagana Dam fisheries

 

Day

Water

Sediments

Weeds

Fish

 

mg/ml

%

mg/g

%

mg/g

%

mg/g

%

 

 

 

 

 

 

 

 

 

0

0.1017

102.7959

0.0094

1.7746

0.0093

0.0040

0.0005

0.0000

 

0.0012

0.0012

0.0007

0.0081

0.0081

0.0081

0.0054

0.0054

 

 

 

 

 

 

 

 

 

1

0.0133

14.0274

0.3848

69.3547

0.0470

0.0574

0.5652

0.1293

 

0.0008

0,0008

0.0115

0.0115

0.0075

0.0075

0.0188

0.0188

 

 

 

 

 

 

 

 

 

2

0.0060

6.0726

0.4024

70.5464

0.1532

0.1223

7.3476

1.4573

 

0.0001

0.0001

0.0060

0.0060

0.0060

0.0060

0.8009

0.8009

 

 

 

 

 

 

 

 

 

4

0.0030

3.4753

0.4035

69.1437

0.2563

0.1650

9.9379

2.0033

 

0.0004

0.0004

0.0096

0.0095

0.0095

0.0095

0.1446

0.1446

 

 

 

 

 

 

 

 

 

 

Rapid dissipation of p,p-DDE from water was observed during the first 24 hours contributing to over 85 percent loss of the dosed pesticide in water. This was followed by a slow dissipation rate during the following period of the study. On the other hand, there was rapid increase in the concentration of the p,p-DDE residues in the sediments within the first 24 hours and tapered off during the next period of study. The concentration of pesticide residues after the fourth day of study decreased from sediments, water, fish to weeds; giving 69.14, 3.47, 2.00 to 0.17 percent of the introduced pesticide in the aquarium (Table 3.2).

Fig 3.1: Distribution of p,p-DDE in the natural ecosystem simulating Sagana Dam fisheries                                                                         

 

Total residues of p,p-DDE in the compartments of the aquarium tank expressed as a percentage of the total p,p-DDE introduced into the aquarium dropped rapidly within the first day followed by a slower rate. Dissipation of p,p-DDE from water was described using first order equation. Two phases were observed, the first phase with rate constant k = 2.02180.0569, R2 value of 0.99980.0002, and calculated half-life of 0.33990.0112 days. The second phase had rate constant k = 0.47370.0344 per day, R2 value of 0.94520.0289 and half-life of 1.43770.1236 days (Table 3.3). The biphasic first order kinetics was accounted for by the first phase dominated by adsorption of the pesticide on to the soil matrix, and the second phase where both adsorption and de-sorption played significant role.

Table 3.3: Chemodynamic properties of p,p-DDE established in the Experiment

 

P,P-DDE

K (per day)

R2

T1/2  (Days)

1ST PHASE

 

2.02180.0569

0.99980.0002

 

0.33990.0112

 

 

 

 

2ND PHASE

0.47370.0344

0.94520.0289

1.43770.1236

       

 

Table 3.4: Extractable residues of lindane from the natural ecosystem simulating                                                                                             

   Sagana Dam fisheries

 

Day

Water

Sediments

Weeds

Fish

 

mg/ml

%

mg/g

%

mg/g

%

mg/g

%

 

 

 

 

 

 

 

 

 

0

8.9032

89.0323

0.5881

1.3232

0.0474

0.0139

2.2824

0.1522

 

0.0430

0.4301

0.0996

0.2240

0.0040

0.0100

0.1707

0.0114

 

 

 

 

 

 

 

 

 

1

6.0138

60.1376

17.4800

39.3300

0.2005

0.0274

-

-

 

0.0279

0.2788

0.1518

0.3415

0.0186

0.0093

-

-

 

 

 

 

 

 

 

 

 

2

4.4258

44.2583

15.7099

35.3472

0.3734

0.0666

-

-

 

0.0037

0.0373

0.0661

0.1486

0.0128

0.0087

-

-

 

 

 

 

 

 

 

 

 

5

3.0000

30.0001

-

-

0.7161

0.1175

-

-

 

0.0657

0.6571

-

-

0.0540

0.0079

-

-

 

 

 

 

 

 

 

 

 

 

Partitioning, distribution and bioaccumulation of lindane was followed using the aquarium tank experiment. The concentration of lindane decreased from the water with time, whereas the concentration in the sediments, weeds and fish increased. Based on the concentration of the residues in the water and sediments, it was observed that adsorption on the soil surface was the main factor contributing to the fast dissipation rate of lindane from the aqueous media. The concentration of lindane increased up to 39% within the first 24 hours and tapered off.  Slow rate of bioaccumulation of the pesticide was observed in the weeds, amounting to about 0.1% of the introduced pesticide after the 5th day of the experiment. Bioaccumulation of lindane by the fish proceeded up to the first four hours and stopped after all the fish samples died. The amount of the pesticide extracted from the dead fish was 2.28240.1707 mg/mg which was equivalent to 0.15220.0114% of the total residues of lindane introduced in the aquarium tank, and this killed all the fish samples.

 

Fig: 3.2 Distribution and Partitioning of lindane in water, sediments, weeds and fish

              in ecosystem simulating Sagana Dam fisheries

 

Dissipation of lindane from the aqueous media was described using the first order equation. A biphasic dissipation rate was observed, the first phase had rate constant k = 0.50960.0040 per day, R2 value of 0.82380.0274 and half-life of 1.35990.0107 days. The second phase had rate constant k = 0.17090.0109 per day, R2 value of 0.96160.0093 and half-life of 4.06750.2573 days.

 

Table 3.5: Chemodynamic properties of lindane established during the experiment

 

LINDANE

K (per day)

R2

T1/2 (Days)

 

 

 

 

1st Phase

0.5096. 0040

0.82380.0274

1.35990.0107

 

 

 

 

2nd Phase

0.17090.0109

0.96160.0093

4.06750.2573

 

 

 

 

 

 

4. DISCUSSION

 

Rapid dissipation rate of p,p-DDE from the aqueous phase and subsequent accumulation of the pesticide by the sediments was attributed to low solubility of the compound in water that enhanced sorption on to sediments. After attaining the adsorption equilibrium, both de-sorption and adsorption became significant and retarded the process, giving raise to a second phase with low dissipation rate constant. Other factors that could contribute to the fast disappearance of the pesticide from the water include adsorption on the aquarium tank walls, bound residue formation, and volatilization.         

             

Higher and faster rate of bioaccumulation was observed in the fish compared to the weeds. Nevertheless, accumulation of pesticide by biota revealed interesting advances, which are currently being recommended for remediation of the environment. Weeds in particular are likely to give one of the best ways of cleaning the environment through phytoremediation strategies, which are comparatively cheaper and environmentally friendly.

             

The dissipation of lindane from the water showed slower rates compared to p,p-DDE. The main reason explaining the variation could be the differences in the solubility of the two compounds. Lindane is more soluble in water compared to p,p-DDE, and as a result larger amount of lindane residues are expected to remain dissolved in the aqueous phase. 

             

Bioaccumulation by the biota was more pronounced in the fish compared to the weeds (lake cabbage). Fast accumulation of the pesticide residues by the fish was mainly due to direct intake of the compounds through water. At low concentration of 0.1 mg/ml in water, p,p-DDE showed no toxic effects to the fish. Lindane at concentration of 10 mg/ml showed toxic effects manifested by shuddering, darting, side swimming, and ultimately death of the fish samples. The sample extracts from the dead fish revealed pesticide concentrations of 2.28240.1707 mg/g, amounting to 0.15220.0114% of the total pesticide residues introduced in the tank. Higher solubility of the compound reduces the chances of its loss through volatilization and sorption, which were the main factors in the given period of the study. As a result, lindane showed lower rate constants and subsequently longer half-lives.

             

The implication of the results of the study was that lindane is more persistent in the aqueous phase than p,p-DDE. Processes such as adsorption and bound residue formation have been reported to inhibit microbial degradation of the pesticides. Strongly sorbed or bound compounds are likely to persist more than less sorbed or bound compounds. Consequently fast adsorption rate and slow de-sorption would lead to adsorbed pesticides staying on the sorbate for an extended period of time. This will results to sediments contaminated with persistent organic pesticides like lindane and p,p-DDE retaining them for longer periods, and continue to release them to aqueous systems and biota through de-sorption and direct extraction through feeding. The final fate of such compound will also depend on physico-chemical properties and environmental conditions such as the presence of pesticide degrading microorganisms.

 

5. CONCLUSION

 

Dissipation, distribution and partitioning of p,p-DDE and lindane from the water to sediments, weeds and fish showed two phases. Higher dissipation rates were observed in the first phase, and were attributed to adsorption on to the sediments. The second and slower phase was attributed to the contribution of both adsorption and de-sorption processes. Other processes such as volatilization and bound residue formation, adsorption on aquarium walls could also partly account for the loss of the pesticides from the aqueous phase. Accumulation of lindane and p,p-DDE by the weeds is a slow process but likely to contribute to significant remediation of the environment with time.

 

 

 

RECCOMMENDATION

 

The current study was designed to investigate the dissipation, distribution and partitioning of DDE and lindane from water to sediments, fish and weeds. Similar experiments simulating other bodies such as rivers and lakes, either using the same compounds or different pesticide compounds should be carried out. Extension of the study to monitor processes such as metabolism, sorption, volatilisation, degradation, mineralization, leaching, toxicity, bioconcentration, photolysis, and depuration is recommended. Most local farmers use a number of pesticides in combination and hence modelling studies are recommended which involve pesticide mixtures.

 

 

REFERENCES

 

Gitahi, S. M., (1999). Organochlorinated and Organophosphorus pesticide                  concentration in water Sediments and selected organisms of Lake Naivasha. Thesis, Moi University, Kenya.

 

Gold-Bouchot, G., T. Silvia-Harrera and O. Zapata-Perez, (1993). Chlorinated pesticides in the Rio Palizada, Cameche, Mexico. Marine  pollution Bulletin 26 (11), 648-650.

 

Lalah, J. O., (1993). Studies on the dissipation and metabolism of a variety of insecticides

under Kenyan environmental conditions. Ph.D Thesis, Department of Chemistry, University of Nairobi. Nairobi, Kenya.

 

Mitema, E. S., and F. K. Gitau, (1990). Organochlorine residues in fish from Lake Victoria, Kenya. Afri J. Ecol. 28(3):234-239.

 

Mughenyi, J. M., (1988). Persistence of DDT and Lindane in Tropical soils. MSc. Thesis, Department of Chemistry, Universisty of Nairobi, Nairobi, Kenya.

 

UNEP, (1982). Determination of DDTs and PCBs in selected marine   orgainsms by GC. Reference methods for marine pollution studies. No. 14. P. 4-7.

 

UNESCO, (1993). Chlorinated biphenyls in open ocean waters: Sampling, Extraction, clean-up and instrumental determination. IOC manuals and  guides No. 27. 20.

 

 


A LECTURE ON PROPOSAL WRITING

 

Shem O. Wandiga

Department of Chemistry

Colledge of Bilogical and Physical Sciences

University of Nairobi, P. O. Box 30197, Nairobi, Kenya.

Email: sowandiga@iconnect.co.ke

 

OUTLINE OF A PROPOSAL: Each grant application is different and one needs to pay close attention to the requirements outlined in the call for proposals. This outline gives some important features that grant reviewers look for in a proposal. A well-written proposal with these elements outlining high quality scientific content is most likely to succeed.

 

TITLE: The title is the most important first contact with the proposal. It should convey the message of what the proposal is about. It should be short and comprehensive in meaning. It should not only be appealing but should entice one to read more about the proposal. If you could make a short acronym out of the title, do so.

 

TABLE OF CONTENTS: Below the title, list the contents and pages of the proposal.

 

PROJECT SUMMARY: Give a short summary of the proposal. The length of the summary should not be more than half a page. The summary should contain all the major messages in the proposal.

 

INTRODUCTION: Give details of the literature review in this section. It should start with the background of the study site(s) and should also include major scientific elements that have been studied. Give clear indication of what is known and what still needs to be studied. The introduction sets the stage for the rest of the proposal. Knowledge of the field is revealed in this section especially by the literature quoted. Ensure you are up to date with the current literature in the field. Do not forget to quote the works of authorities who are most likely to review your proposal.

 

PROJECT DESCRIPTION: Give clear description of what you want to study. Start with a well thought out conceptual framework. The conceptual framework should show how you are going to use what is known and how they are related to what is to be studied. Use flow charts to outline your concepts. Show how your hypothetical linkages will be tested. The conceptual framework should be followed by a problem formulation.

 

State very clearly the problem. A clearly stated problem will lead to research questions that need to be answered. Give all relevant questions related to the project that you think you will be able to answer through your research. State your working hypothesis. This should not be more than a sentence or two. This section should also contain the objective of the proposal. Give a general objective, which is broad, and specific objective that clearly outline what you will do. If you are going to do field work describe the site, and if your proposal is deskwork describe your department and what you have done in the past that is related to the proposed study. Finally, give a justification of your proposal. Justification should show why the study is important.

 

METHODOLOGY: Outline in this section the methodology according to the specific objectives. This outlined methodology should also be reflected in the work-plan. The methodology should show how your conceptual framework is going to be tested. A detailed description of why, what and how you are going to undertake the study should clearly come out. This should be followed with a detailed work-plan giving number of activity, year, month, activity itself, method, indicator of activity being done, persons responsible for the activity as shown in the table below.

 

DETAILED WORK PLAN

 

No & Yr.

Mo

Activity

Methods

Indicators

Persons

Responsible

 

 

 

 

 

 

 

COLLABORATION AND PARTNERSHIP: Give names, institution, expertise and role of each collaborator. If you are going to contract or subcontract any section of the work give full details of the persons to undertake such contract. Give details of the institutions and the person that will be responsible for administration of the grant contract.

 

EXPECTED OUTPUTS: Provide all the expected outputs that will arise from the research. State the reasons why the project will be relevant to decision making, development or science itself.

 

CAPACITY BUILDING: Capacity building is very important section. Provide all capacity building activities you will undertake during the project execution.

 

REFERENCES: List all references sited giving names of authors, year, and title of papers, journal, volume and pages.

 

PROJECT BUDGET: The budget section should contain the narrative section which summarizes the budget request, the detailed budget showing the activity, cost estimate, collateral funding in each year of the activity.

 

APPENDICES: Present full details of CV of each person involved in project. In a number of cases only relevant publication may be requested instead of all publications of each participant. Follow the instructions given in the call for proposal. Attach maps and other details referred to in the text here.

LIST OF PARTICIPANTS IN ANCAP (2003) SUMMER SCHOOL

 

NAME

ADDRESS

EMAIL $ TEL CONTACTS

1.    S. O. Wandiga

University of Nairobi,

Department of Chemistry,

Box 30197,

Nairobi, Kenya.

sowandiga@iconnect.co.ke

 

254-44446140

2.     Haji  Mwevura

University of Dar-es Salaam,

Chemistry Department,

Box 35061,

DSM-Tanzania.

mwevura@chem.udsm.ac.tz

 

 

 

3.     Juma M. Makopa

University of Nairobi,

Department of Chemistry,

Box 30197,

Nairobi, Kenya.

jmakopa@uonbi.ac.ke

 

4.     John Wasswa

Makerere University,

Box 7062,

Kampala-Uganda.

jnwasswa@chemistry.mak.ac.ug

 

5.   Madadi O. Vincent

University of Nairobi,

Department of Chemistry,

Box 30197,

Nairobi, Kenya

madadivin2002@yahoo.co.

 

6. Fredrich O. Kengara

Maseno University,

Department of Chemistry,

Box 333,

Maseno, Kenya.

fkengara@yahoo.com

 

Tel: 254-0722262159

 

7. Kyarimpa Christine

Makerere University,

Box 7062,

Kampala-Uganda.

christinkyarimpa@avu.org

 

 

8. Geofrey Malisa

University of Dar-es Salaam,

Chemistry Department,

Box 35061,

DSM-Tanzania.

malisa@chem.udsm.ac.tz

 

9. Aviti J. Mmochi

University of Dar-es Salaam,

Institute of Marine Sciences,

Box 668,

Zanzibar-Tanzania.

mmochi@ims.udsm.ac.tz

Tel: 255-242230741

 

10. Orata Francis

University of Nairobi,

Department of Chemistry,

Box 30197,

Nairobi, Kenya

fraora@yahoo.com

 

11. Philip K. Maritim

Moi University,

School of Environmental Studies,

Box 3900,

Eldoret, Kenya.

pkmaritim@yahoo.com

 

 

12. Tarekegn Berhanu

University of Addis Ababa,

Chemistry Department,

Box 1176,

Addis Ababa, Ethiopia.

tarekegnbr@yahoo.com

 

 

13. Lutufyo Mwamtobe

University of Dar-es Salaam,

Chemistry Department,

Box 35061,

DSM-Tanzania.

mwamtobe@chem.udsm.ac.tz

 

 

14. Ahmed Hussen

University of Addis Ababa,

Chemistry Department,

Box 1176,

Addis Ababa, Ethiopia.

ahdekebo@yahoo.com

 

 

15. Andrew A. Andayi

Maseno University,

Department of Chemistry,

Box 333,

Maseno, Kenya.

andrewandayi@yahoo.com

 

 

16. Joseph Ng'ang'a

University of Nairobi,

Department of Chemistry,

Box 30197,

Nairobi, Kenya

jnganga@uonbi.ac.ke

17. V. Muinde

University of Nairobi,

Department of Chemistry,

Box 30197,

Nairobi, Kenya

vmuinde@uonbi.ac.ke

 

 

18. Matobola J. Mihale

University of Dar-es Salaam,

Chemistry Department,

Box 35061,

DSM-Tanzania.

mihale@chemistry.udsm.ac.tz

 

 

19. F. Seme

Government Chem Lab

Box 164,

Dar-es Salaam, Tanzania

gcla@gclago.tz

Tel: 255-22-2113383/4