規格型號：

序號 貨 號 型  號

1 10098 DW-10098-TF44

2 11372 DW-11372-TF44

3 10099 DW-10099-TF44&37

4 11433 DW-11433-TF44&37

5 10100 DW-10100-TF44

6 11434 DW-11434-TF44

7 11240 DW-11240-TF-1

8 11241 DW-11241-TF-2

9 11242 DW-11242-TF-3
 
10 15735 DW-15735-TF-4

11 10101 DW-10101-TFZ

12 16477 DW-16477-TFZ

13 14867 DW-14867-TFZ

14 14404 DW-14404-TFZ

OXFAM-DELAGUA Water Testing Kit — Users Manual

1

OXFAM - DELAGUA

Portable Water Testing Kit

VERSION 4.3

REVISED 2012

OXFAM-DELAGUA Water Testing Kit — Users Manual

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OXFAM-DELAGUA Water Testing Kit — Users Manual

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CONTENTS

1 The OXFAM - DELAGUA Kit Components 10

1.1 General 10

1.2 Filtration Apparatus and Components 11

1.3 Contents of the Spares Case 11

1.4 Materials Needed for Testing 12

2 Sampling Programmes 13

2.1 Selection of sites and frequency of sampling on a network supply 13

3 Preparation of the Kit 15

3.1 Sterilisation of the Filtration Apparatus 15

3.2 Preparation of Culture Medium in the Laboratory 17

3.3 Preparation of Culture Medium in the Field 19

3.4 Storage of Culture Medium 20

3.5 Sterilisation of the petri-dishes 20

3.6 Disposal of Contaminated Material 21

3.7 Absorbent Pads and Dispenser 21

3.8 Methanol Dispenser 22

4 Sampling Methods 23

4.1 Sampling from a Tap 23

4.2 Sampling from a Lake, Reservoir or other Surface Water Source 24

4.3 Sampling from an Open Well or Storage Tank 26

5 Processing of Samples using the Kit 27

5.1 Introduction 27

5.2 Analysis of Free Chlorine Residual and pH 27

5.3 Turbidity Analysis 29

5.4 Bacteriological Analysis of Water 31

6 Care and Maintenance of the Kit 45

6.1 The Battery 45

6.2 Electronic Components and the Incubator 46

6.3 Filtration Apparatus 46

6.4 Chlorine and pH Comparator and Turbidity Tubes 46

6.5 Kit Case 46

6.6 Maintenance 47

7 Evaluation and Repair of the Kit 48

7.1 Fault Finding in the Incubator, Battery and Charger 48

7.2 Fault Finding Chart 51

7.3 Checking and Recalibrating the Incubator 52

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Appendix A 56

Incubator electronic circuit diagram.

Appendix B 57

Field Checklist

Appendix C 58

Spares list

Appendix D 60

Daily report sheet

Appendix E 61

Alternative sources of water for media preparation

Appendix F 62

Additional instructions for operating the dual incubator kit

Appendix G 64

Alternative media

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OXFAM-DELAGUA Water Testing Kit — Users Manual

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OXFAM - DELAGUA

Portable Water Testing Kit

Users Manual

Revised 2012

This equipment was designed to test for the critical parameters specified in World

Health Organization (WHO) Guidelines for Drinking Water Quality, Second

Edition, Volume III.

The equipment should only be used by trained personnel familiar with those

guidelines.

For more information about the kit and for technical help and guidance, please

contact DelAgua Water Testing Ltd.

DelAgua Water Testing Ltd, The Old Dairy, Church Lane, Lower Fyfield

Marlborough, Wiltshire, SN8 1PX

Tel: +44 1672 861 198/1672 861 049

Fax: +44 1672 861 724

Email: info@delagua.org

Website: www.delagua.org

Facebook: www.facebook.com/delaguawater

Twitter: www.twitter.com/del_agua

OXFAM-DELAGUA Water Testing Kit — Users Manual

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Copies of this manual are available in several languages. Please consult our website

for current availability. Abridged versions of the manual are available to download.

If you regularly use the OXFAM - DELAGUA water testing kit and have translated

the manual into another language, please send us the translation. Under these

circumstances, we normally organise printing and give free copies to the

programme which provided the translation.

We are continually trying to improve the OXFAM - DELAGUA water testing kit

and because of this, some components may be different from those which appear in

the manual. We also welcome suggestions from users about ways for improving the

kit to meet their own particular needs.

Training

Purchasers of the kit are entitled to participate in a one-day course at the DelAgua

office in the use of the kit. The course is free of charge.

DelAgua also offers one and 2-week training courses overseas that include water

quality testing, sanitary inspection, water supply disinfection, and use and

maintenance of the OXFAM - DELAGUA kit.

Please consult our website for further details or contact us at info@delagua.org

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Instructional CD-ROM

This version of the manual includes a CD-ROM that provides guidance on the use

and maintenance of the kit and includes video sequences that demonstrate the

procedures described in the manual. The CD-ROM will start automatically after it

is inserted into your CD-ROM drive. This is the first time we have produced a

training CD-ROM for the kit and so we would welcome your general comments

and any suggestions that would help us to improve the product in the future.

Note: The main sections of this manual describe the use

and maintenance of the single incubator water test kit.

The additional procedures for the operation of the dual

incubator kit are described in Appendix F.

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1 The OXFAM - DELAGUA Kit Components

1.1 General

The kit also includes an electronic timer with its own instructions for use.

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1.2 Filtration Apparatus and Components

1.3 Contents of the Spares Case

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1.4 Materials Needed for Testing

Before you start to use your OXFAM - DELAGUA water testing kit, you will need

the following materials:

For preparation of culture medium:

1. Pressure cooker, portable steriliser or autoclave.

2. Electric heating element, gas burner, stove or similar to heat the steriliser.

3. Distilled water (for alternatives see Appendix E).

4. Means of measuring distilled water eg. Measuring cylinder, or graduated

beaker.

For using the kit in the field:

1. Methanol (for alternatives see Section 5.8).

2. Paper towels or clean cloths.

3. Wax pencil or marker pen.

4. Report sheets (see Appendix D).

5. Cigarette Lighter (any fluid filled lighter is suitable).

On receipt of a new Oxfam - Delagua kit we would

recommend that you recharge the battery fully

(Section 6.1) and check the operating temperature of

the incubator (Section 7.3).

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2 Sampling Programmes

2.1 Selection of sites and frequency of sampling on a network supply

Samples should be taken from locations

that are representative of the water

distribution network and household

connections.

Where there are several sources and a

mixed distribution system, it is

necessary to take account of the

variation that may exist in the system

and incorporate this into the sampling

programme.

Where there is a branched distribution

system, samples should be taken at

random points evenly spread throughout

the system.

Where there are main branches and a

remote periphery (as shown), greater

attention should be devoted to the main

branches and remote points in the

network.

The recommended minimum frequencies for sampling of both piped supplies and

point sources are shown in the tables below:

Minimum Frequency of Sampling and Analysis of Piped Water Supplies

Population served Minimum frequency of sampling

Less than 5,000 One sample per month

5,000 to 100,000 One sample per 5,000 population per month

More than 100,000 20 samples monthly plus one extra sample per 10,000

population

Location of key sample points on a network distribution

system.

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Minimum Frequency of Sampling and Analysis of Unpiped Water Supplies

Source and

mode of supply

Bacteriological Physical/

Chemical

Remarks

Open well Sanitary protection

measures and

testing only if

situation demands

Once initially

for community

wells

Pollution usually

expected

to occur

Covered well.

Shallow tubewell

with handpump

Sanitary

protection

measures and

testing only if

situation

demands

Once initially.

Thereafter as

situation

demands

Testing needed when

Environmental

conditions

change or when an

outbreak or increase in

waterborne disease

occurs

Deep tubewell

with handpump

Once initially.

Thereafter as

situation

demands

Once initially.

Thereafter as

situation

demands

Testing needed when

environmental

conditions

change or when an

outbreak or increase in

waterborne disease

occurs

Springs and piped

supplies

Once initially.

Thereafter as

Situation demands

Test periodically for

residual

chlorine if water is

chlorinated

Testing needed when

environmental

conditions

change or when an

outbreak or increase in

waterborne disease

occurs

Community

rainwater

collection

systems

Sanitary

protection

measures and

testing only if

situation

demands

Not needed

Source: Adapted from WHO Guide lines for Drinking-Water Quality Volume

III. Second Edition, Geneva, 1985.

We would recommend that you refer to the WHO website (www.who.int) for

the latest advice regarding sampling and analysis of water supplies.

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3 Preparation of the Kit

3.1 Sterilising the Filtration Apparatus

The sample cup and the filtration apparatus must be sterilised before use and resterilised

between samples when analysing water from 2 different sources.

Sterilising the equipment in the field presents some practical difficulties and must

be carried out using simple methods. The most appropriate is the use of methanol,

which is described below. When methanol is burnt in a low oxygen atmosphere -

for example, in the closed sample cup - formaldehyde gas is produced as a byproduct

of combustion.

Formaldehyde gas is a very effective disinfectant. Methanol is expensive to freight

and requires special transport conditions. We would recommend that you first try to

obtain methanol in-country from a pharmaceutical supplier, a local hospital or

university laboratory. If necessary, however, methanol can be supplied by the

DelAguaon request.

If methanol is not available, the filtration apparatus and sample cup can be

sterilised by immersion in boiling water for 10 minutes.

Procedure for sterilising the filtration apparatus using methanol

Note: Methanol is the only alcohol suitable for sterilising the filtration

apparatus; there is no substitute.

1. Carefully dry the sample cup and

filtration assembly with a clean dry

towel or tissue.

2. Using the plastic collar, secure the

filtration funnel in the loose but not

free position (see Section 5.4.4 [9])

which allows the formaldehyde gas to

penetrate all areas of the filter head.

Dry the sample cup

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3. Pour about 1ml (approximately

20 drops) of methanol into the sample

cup.

4. Carefully ignite the methanol in the

sample cup using the cigarette lighter.

Place the cup on a flat surface which will

not be damaged by heat.

Caution: Keep the mouth of the sample

cup away from your face and the hole

uppermost to prevent methanol running

onto your hand.

5. Allow the methanol to burn for several

seconds and, when almost completely

burned up (ie. as the flames are dying

down), place the filtration head over

the sample cup and push firmly into

place to form a good seal.

Add 20 drops (approximately 1ml)

of methanol

Carefully ignite the methanol

Replace the filtration head

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6. Keep the filtration apparatus sealed for at least 15 minutes before use.

Note: It is best to sterilise the filtration apparatus immediately after each analysis

and to keep the filtration apparatus in a sterile condition during transport and

storage. In this way, the filtration apparatus is always ready for use.

3.2 Preparation of Culture Medium in the Laboratory

You will need the following items:

1. 38.1g of Membrane Lauryl Sulphate Broth (MLSB)(a)

2. Distilled water (b). Check that the pH of the water is between 7.0 and

7.8 using the comparator and phenol red tablets (Section 5.2)

3. Ten polypropylene bottles (60ml)

4. Measuring cylinder or graduated flask

5. Clean flask or beaker, approximately 1 litre capacity

6. Pressure cooker, steriliser or autoclave(c)

7. Heating element, stove or burner

(a) The medium is available in 38.1g, pre-weighed amounts from DelAgua

(b) See Appendix E for suggested alternative sources of water

(c) A portable steriliser kit is available from DelAgua

Method

1. Carefully wash the plastic polypropylene bottles in clean, warm water before

use. If necessary, use a little detergent and then rinse well with clean water to

remove all traces of the detergent.

2. Measure out 500ml of distilled water using the measuring cylinder or

graduated flask. Decant approximately half of the water into the clean flask or

beaker.

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3. Add the 38.1g of MLSB powder to the distilled water in the clean flask or

beaker and stir until the powder has dissolved. Gentle heat can be applied if the

powder is slow to dissolve. Add the remaining volume of distilled water and

continue stirring to thoroughly mix the broth.

The culture medium will be a bright red colour when dissolved.

Note: MLSB is a fine, but non-hazardous powder. However, the dust may irritate

the nose or upper respiratory tract if inhaled. Take care to avoid creating excess

dust when handling the powder and cover the nose and mouth with a cloth or dust

mask to reduce exposure. Spillages can be cleaned up using water and an absorbent

cloth.

4. Pour a suitable volume of culture medium (approximately 50ml, but no less

than 40ml) into each of the 10 polypropylene bottles. This provides sufficient

medium in each bottle to carry out 16 tests; the maximum that can be

performed in one day using the Delagua kit.

5. Replace the screw caps on the polypropylene bottles. Make sure the caps are

secure but do not tighten. Leaving the caps slightly loose prevents the bottles

from collapsing during sterilisation.

6. If an autoclave is available, sterilise the bottles at 121oC (equivalent to 1 bar,

or 15 psi steam pressure) for 15 minutes. Tighten the caps carefully once the

medium has cooled.

7. If you do not have access to an autoclave, then a household pressure cooker or

portable steriliser may be used. Place the bottles in a rack inside the cooker

(they may melt if placed directly on the base of the cooker), replace the lid and

heat to full pressure (about 1 bar or 15psi).

Once the cooker has reached full pressure allow steam to issue from the release

valve for 5 minutes, then time the 15 minutes sterilisation cycle using a

stopwatch or clock. At the end of the 15 minutes, switch off the heat and allow

the cooker to cool until it is comfortable to touch. Remove the media bottles

and tighten the caps.

8. Label the bottles to indicate sterilised contents and the date and batch of

medium.

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3.3 Preparation of Culture Medium in the Field

You will need the following items:

1. 38.1g of Membrane Lauryl Sulphate Broth (MLSB)(a)

2. Distilled, or clean water(b)

3. 10, polypropylene bottles (60ml)

4. Measuring cylinder or graduated beaker

5. Portable steriliser(c) or pressure cooker or cooking pot or pan

(a) The medium is available in 38.1g, pre-weighed amounts from DelAgua

(b) See Appendix E for suggested alternative sources of water

(c) A portable steriliser kit is available from DelAgua

Method

1. Carefully wash the plastic polypropylene bottles in clean, warm water before

use. If necessary, use a little detergent and then rinse well with clean water to

remove all traces of the detergent.

2. Use distilled water if possible. If this is not available obtain the cleanest water

possible. DO NOT use water that has been treated with chlorine or any other

chemical disinfectant.

3. Use the comparator and phenol red tablets in the kit to check that the pH of the

water is between 7.0 and 7.8. If it is not, it will be necessary to find an

alternative source of water.

4. Measure out 500ml of clean water in a beaker.

5. Add 38.1g of the MLSB powder to the 500ml of water in the beaker.

Mix to dissolve the powder completely. Apply gentle heat if the powder is

slow to dissolve. The culture medium will be clear with a bright red colour

when dissolved.

6. Pour a suitable volume of culture medium (approximately 50ml, but no less

than 40ml) into each of the 10 polypropylene bottles. This is sufficient medium

in each bottle to carry out 16 tests; the maximum that can be performed in one

day using the Delagua kit.

7. Replace the screw caps on the polypropylene bottles. Make sure the caps are

secure but do not tighten. Leaving the caps slightly loose prevents the bottles

from collapsing during sterilisation.

8. If a pressure cooker is available, sterilise the culture medium as described in

Section 3.2, paragraph 7.

9. If a pressure cooker or portable steriliser is not available, the medium can be

sterilised using a process called Tyndellisation. Note, this

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procedure takes 3 days. Place the bottles of culture medium into a cooking pot

or pan of boiling water, taking care to ensure that the bottles do not come into

contact with the base of the pan (use a rack or stand) or become submerged.

Boil for 20 minutes. Leave the medium to stand for 24 hours at room

temperature (20-30oC) in the dark. On the following day heat the medium in

boiling water for a further 20 minutes and, once again, leave to stand for 24

hours. On the third day repeat the heat treatment. The medium should now be

sterile.

3.4 Storage of Culture Medium

Sterile MLSB will be stable for up to 6 months if stored in a refrigerator (between 4

and 6oC). Alternatively, the medium can be stored for up to 3 months in a cool,

dark place. If the medium has been stored for several days below 6oC a deposit

may form which dissolves when the medium is warmed and gently shaken. The

deposit is caused by the lauryl sulphate coming out of solution.

If signs of deterioration are observed, eg. cloudiness or yellow colouration, the

contents of the bottle must be discarded.

3.5 Sterilising the petri-dishes

1. Wash the dishes in a solution of mild detergent, rinse thoroughly with clean

water and dry.

2. Assemble the dishes into batches of 16 in the straps.

EITHER

3. Sterilise the petri dishes in an autoclave, steam steriliser or pressure cooker at

121oC for 15 minutes (see section 3.2 paragraphs 6 and 7).

OR

4. Place the dishes in a conventional oven at 180oC for 30 minutes.

OR

5. Plunge the bases and lids of the dishes into boiling water for 10 minutes. Pour

away the water and assemble the dishes as they dry, but while they are still hot.

OR

6. Add a few drops of methanol (or ethanol) to a clean cloth and wipe the inside

of the lid and the base of each petri-dish. Assemble the petri-dishes and allow

the alcohol to evaporate before use.

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OR

7. Whenever possible, always use one of the above methods. If this is not

possible, then the following method can be applied. Flame the bases and lids of

the dishes with a lighter or gas burner using the tweezers to hold the bases and

lids. Assemble while still hot.

3.6 Disposal of Contaminated Material

Note: To minimise the risk of infection from contaminated materials, take care not

to touch contaminated membranes directly with your hands. Do not eat, drink or

smoke while handling contaminated materials. Wash your hands immediately after

you have touched any contaminated material and after you have finished your

work.

Contaminated material, such as used membranes and pads, MUST be made safe

before disposal. DO NOT discard non-sterile membranes and pads into the

environment since they pose a major risk to public health.

After you have completed the analysis, stack the petri-dishes in the straps and

sterilise the dishes and contents at 121oC for 30 minutes using an autoclave,

steriliser or pressure cooker. Alternatively, plunge the petridishes, pads and

membranes into boiling water and heat for at least 30 minutes (use a dedicated pan

for this procedure. DO NOT use a pan that will subsequently be used for food

preparation or other domestic purposes). After sterilisation, the used membranes

and pads may be destroyed by incineration.

The petri-dishes must be carefully washed with detergent after use, rinsed with

clean water and dried.

3.7 Absorbent Pads and Dispenser

The pads are supplied sterile in packs of 100 units. A pad dispenser is also supplied

with the kit. Never leave the dispenser without a pack of pads attached as it will

increase the possibility of contamination.

You might find it more convenient to dispense the pads into the petridishes at your

base to avoid the need to take the dispenser and pads into the field.

If it is necessary to dispense pads in the field, take care not to contaminate the

dispenser assembly. If the dispenser is lost or damaged, pads may be dispensed in

the field using the sterile tweezers (see Section 5.4.4[3] for sterilisation methods).

Some kit operators prefer this method to using the dispenser.

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3.8 Methanol Dispenser

The methanol dispenser is supplied with a plastic cap and dispensing nozzle. The

dispenser should be half-filled with methanol using a small funnel, pipette or

syringe to avoid spillage. Do not overfill the methanol dispenser as it may leak in

hot weather.

To dispense methanol, lever the dispensing nozzle into the upright position with the

tip of the tweezers. To seal off the flow of methanol, push the nozzle down into the

recess in the cap. Be sure to close the dispensing nozzle after using the kit as the

methanol will evaporate.

Note: Methanol is highly flammable. Keep methanol away from naked flames.

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4 Sampling Methods

4.1 Sampling from a Tap

1. Remove any attachments from the tap; eg. nozzles,

pipes, etc. Check that the tap does not leak and that

all seals are in good condition.

2. Carefully clean the mouth of the tap with a clean

cloth or tissue to remove any dirt or grease. Open

the tap and leave water running for at least one

minute before taking a sample. Do not adjust the

flow of water during this time. This ensures that

any deposits in the pipes are washed out and the

water sample is representative of the water in the

supply pipes.

Remove attachments from tap

Leave the tap running for at least

one minute

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3. Take a water sample with the

non-sterile vacuum cup. Rinse the cup

twice with the sample water before

taking the sample. Analyse the sample

for chlorine residual and turbidity

using the methods in sections 5.2 and

5.3.

4. If the chlorine residual and turbidity

results suggest that there is a risk of

microbiological contamination (see

Section 5.1), then take a second

sample for bacteriological analysis

using the sterile sample cup.

4.2 Sampling from a Lake, Reservoir or other Surface Water Source

1. Where there is safe and adequate

access to the source it may be

possible to take samples by hand.

2. Grasp the sample cup firmly, keeping

your fingers clear of the top of the

cup to avoid contamination, and dip

the open mouth of the cup into the

water.

3. Submerge the cup about 30cm below

the surface of the water and scoop up

the water sample. This scooping

action ensures that no external

contamination enters the sample cup.

4. Lift the sample cup carefully and place on a clean surface where it cannot be

knocked over.

Take a sample for bacteriological analysis

using the sterile sample cup

Take sample from approximately 30cm

below the surface

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5. In areas where there is a current flow,

eg. rivers and streams, the sample

should be taken against the current

flow.

6. In many cases it may be inconvenient

or dangerous to enter the water. The

sample cup can be lowered into the

water from a firm area of riverbank

or river crossing by fastening the 2m

sample cable to the hole in the lip of

the sample cup.

Note: It is important that you obtain a sample which is representative of the main

body of water. For example, when sampling from a river, do not sample the quiet

or stagnant areas near the bank, as these do not represent the main body of water.

Furthermore, it is vital not to introduce external contamination into the sample. For

this reason it is often better to sample with the help of the cable supplied with the

kit.

In rivers and streams take samples against the

current flow (the arrow shows the direction of

flow)

Lower the sample cup into the water

using the 2m cable

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4.3 Sampling from an Open Well or Storage Tank

1. Fasten the sampling cable to the hole

in the lip of the sample cup by means

of the clip on the end of the cable.

Make sure the clip is closed and

secure.

2. If necessary, increase the length of

the cable by attaching a rope or string

to the sample cable. Take care not to

lose the sample cup.

3. Lower the sterile sample cup into the

well or tank, taking care not to allow

the cup to touch the walls of the

structure where it may pick up dirt.

Submerge the cup to a depth of 30cm.

If possible keep fingers clear of the

cable or rope so as not to contaminate

the water body.

Fasten the sampling cable to the sample cup

Increase the length of the cable if necessary using

string or rope

Lower the sample cup into the well or

tank

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4. Lift the sample cup carefully and place on a clean surface where it cannot be

knocked over.

5 Processing of Samples using the Kit

5.1 Introduction

The first tests that should be carried out on a drinking water sample are the

determination of chlorine residual, pH and turbidity. The results from these tests

will indicate whether or not the water sample is likely to contain living

microorganisms and whether it is necessary to carry out analysis for thermotolerant

coliform bacteria. The sample must be taken in a clean, but not necessarily sterile

cup, eg. the vacuum cup. Rinse the vacuum cup several times with the water that is

to be analysed before taking a sample for analysis (see Section 4).

If the results of the analysis are as follows:

? Free chlorine residual greater than 0.2mg/litre (0.2ppm) and

? Turbidity less than 5TU

it is unlikely that the sample will contain thermotolerant (faecal) coliform bacteria

and therefore it may not be necessary to carry out thermotolerant coliform analysis.

If the results do not meet these criteria, it will be necessary to carry out

thermotolerant coliform analysis. Under these circumstances, samples for analysis

must be taken with the sterile sample cup.

5.2 Analysis of Free Chlorine Residual and pH

1. Wash the comparator cells three times

with the water that is to be analysed

and finally fill both cells with the

sample.

Fill the comparator cells with sample water

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2. Drop one DPD No 1 tablet into the

right hand cell (C12) and one

Phenol Red tablet into the left hand

cell (pH).

3. Replace the lid of the comparator and

push down firmly to seal. Invert the

comparator several times until the 2

tablets have dissolved completely. If

the tablets are slow to dissolve use the

plastic paddle from inside the box of

tablets to crush and mix the tablets in

the sample water. Do not shake the

comparator as this will introduce air.

4. Immediately read the free chlorine residual (mg/litre) and pH by holding the

comparator up to daylight and matching the colour in the cells with the

standard colour scales. If the colour falls between 2 standard colours, then it

will be necessary to estimate the concentration. Record the result on the daily

report sheet (see Appendix D for an example).

5. To test for total chlorine residual, DO NOT discard the liquid in the

comparator. Remove the lid and add one DPD No 3 tablet to the right hand cell

(C12).

Add the DPD No 1 and phenol red

tablets

Invert the comparator to dissolve the tablets

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6. Again, invert the comparator several times to dissolve the tablet (use the

plastic paddle if the tablet is slow to dissolve). Leave the colour to develop for

10 minutes. Read the total chlorine residual (mg/litre) by matching the colour

in the cells with the standard colour scale.

7. Subtract the free chlorine result from the total chlorine result to obtain the

combined chlorine concentration:

Summary

Phenol Red = pH

DPD No 1 = Free chlorine residual

DPD No 1 plus DPD No 3 = Total chlorine residual

Total - Free chlorine = Combined chlorine

5.3 Turbidity Analysis

Note: The turbidity tube covers the range 5 to 2,000 TU

1. Carefully remove the 2 halves of the

turbidity tube from their clips in the

lid of the case. Push the upper tube

(open at both ends) squarely into the

lower tube and align the graduation

marks up the side. Look through the

open end of the tube at the black circle

printed on the yellow base of the tube;

this is the marker. Ensure that there is

good illumination available. Normal

daylight is adequate for this purpose.

Assemble the turbidity tube

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2. Hold the tube vertically and slowly

pour the water sample into the tube

until the marker just disappears when

viewed from the top of the tube. Avoid

creating bubbles, as these may cause

false readings. Do not strain to see the

black circle as this can sometimes

cause biased results.

Alternatively, the procedure can be

carried out in reverse by filling the tube

with the sample and then slowly

pouring out the water.

3. Hold the tube vertically and read the turbidity using the graduations on the side

of the tube. The graduations follow a logarithmic scale with the most critical

values marked on the side of the tube. The result is the value of the line nearest

the water level. This permits a reasonable estimation of the turbidity of the

water sample. Alternatively, you can judge the distance of the water level from

the 2 nearest graduation marks and calculate a more accurate turbidity value.

Remember, however, that you are reading a log scale.

Fill the tube until the marker disappears

Estimate the turbidity using the graduation marks. The tube is shown horizontal for convenience

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5.4 Bacteriological Analysis of Water

Note: Microbiological analysis presents some special risks to the health and safety

of yourself and others. It is vitally important that you take great care when handling

potentially contaminated materials, such as the petridishes, membranes and pads

that have been used for the growth of microorganisms. Although most of the

bacteria that will grow on the membrane are relatively harmless, some of the

colonies may contain pathogenic bacteria. High standards of hygiene should be

applied at all times:

? Never eat drink or smoke when carrying out microbiological tests.

? Do not touch colonies with your fingers or with everyday objects such as pens

and pencils that you may use again for other purposes.

? Always wash your hands thoroughly after handling contaminated, or

potentially contaminated materials.

? Cover wounds with a waterproof dressing.

? Do not carry out microbiological tests in food preparation areas.

? Do not dispose of contaminated materials into the environment.

Always sterilise the items before disposal or cleaning.

? Keep all non-essential personnel, particularly children, away from the work

area when handling contaminated materials.

? Always keep your work area clean and tidy.

? Clean and disinfect (methanol, ethanol or a weak solution of domestic bleach

can be used or this purpose) the work surfaces after you have finished the

analysis.

5.4.1 Introduction

The analysis of water samples for thermotolerant coliforms is carried out by

passing a measured quantity of water through a sterile filter. Any bacteria present

in the water are caught in the filter. The filter is then placed onto a paper pad

soaked in a liquid growth medium which feeds coliform bacteria, but inhibits the

growth of any other bacteria caught in the filter. To ensure that only thermotolerant

coliform bacteria grow, the filter is kept at 44°C in the kit’s incubator. During this

time the coliform bacteria multiply many times to form colonies that can be seen

with the naked eye. Thermotolerant coliforms are recognised by their ability to

produce a colour change (from red to yellow) in the culture medium at 44°C. It is

normal practice in water microbiology to express the results as colony-forming

units per 100ml of water (CFU/100ml).

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Thermotolerant coliforms are of sanitary significance when present in drinking

water supplies. Users should refer to country-specific water quality standards or

guidelines, or to the latest edition of the World Health Organization Guidelines for

Drinking Water Quality (available on-line at www.who.int) to decide when action

should be taken to improve contaminated water supplies.

Some users may need to analyse for total coliform bacteria, which, although of less

sanitary significance than thermotolerant coliforms, can be used to indicate hygiene

problems in large distribution networks. Total coliform analysis is carried out using

the same procedure as for thermotolerant coliforms, the only difference being that

the filters are incubated at 37 °C.

The OXFAM - DELAGUA incubator can be recalibrated to 37°C by following the

recalibration procedure in Section 7.3. However, this is not convenient when

carrying out both thermotolerant coliform and total coliform analysis on a regular

basis. A dual incubator kit (see Appendix F) is available from DelAgua which

allows both tests to be carried out simultaneously.

5.4.2 General Hygiene in the Field

Although all components of the kit should be kept free from dirt and other

contamination, there are some parts of the kit which must always be kept clean and

sterile. These are as follows:

a) All those areas in direct contact with the water sample, eg. the internal surface

of the sample cup, the internal surface of the filter funnel, the upper part of the

filtration base and the surface of the bronze disc.

b) Surfaces in contact with the culture medium, eg. the internal surface of the

petri-dishes and the absorbent pads.

c) Parts in contact with the membrane filters, eg. the filtration apparatus, the

absorbent pads and the tweezers.

Under no circumstances should any of these components be allowed to come into

contact with dirt, dust or external objects which may contaminate them and

interfere with the bacterial count.

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Before handling a membrane filter and after processing a sample, the tips of the

tweezers should be flamed with a cigarette lighter. Hold the tips in the flame for

five seconds and allow them to cool before handling a membrane filter. After

sterilising the tweezers in this way, they should be placed so that the tips do not

touch any other object.

5.4.3 Selection of Appropriate Sample Volumes for Coliform Analysis

The most appropriate volume to process is that which allows the most accurate

count of the bacterial colonies. It is generally agreed that 100 colonies is the

maximum that can be counted reliably on a 47mm membrane. Counts above 100

are considered an estimate. If a large number of colonies develops on the

membrane you can either divide the plate into sections, count the colonies in one

section and multiply the count by the number of sections, or repeat the analysis

with a smaller volume of the sample and then adjust the result to give a count per

100ml of the original sample.

The selection of the most appropriate sample volume for a given source, treatment

plant or distribution system is normally best made in the light of previous

experience. For sites where this information does not exist, the following

paragraphs provide some guidance on sample volumes.

Treated water and water in piped distribution systems

Historically, the microbiological quality of drinking water has been assessed using

the number of bacteria present in a standard volume of

100ml. Treated water and water in a piped distribution network are unlikely to

contain large numbers of thermotolerant coliform bacteria.

For these waters we would recommend using a 100ml sample.

Suggested sample volumes for thermotolerant coliform analysis by the

membrane filtration technique (alternative volumes are shown in brackets).

Source Sample volume

Waters in treatment plants after partial

treatment

50ml (100ml or 10ml)

Waters in treatment plants after full

treatment

100ml

Reservoirs, distribution networks and

household taps

100ml

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Other Water Sources

Recommended sample volumes for each source are shown below.

Suggested sample volumes for thermotolerant coliform analysis by the

membrane filtration technique (alternative volumes are shown in brackets).

Source Sample volume

Lakes, ponds and other surface waters 10ml (1ml*)

Protected groundwater, eg. wells and

springs

100ml (50ml or 10ml)

Unprotected groundwater, eg. open dug

wells and springs

50ml (10ml or 1ml*)

*Note: This volume will require the use of sterile pipettes and dilution water.

Please note, these volumes are only a guide. They do not represent absolute

recommendations to be applied to sampling programmes. It may be useful to

analyse different volumes of the same sample in order to decide the best range in

which to count the bacteria. It is not necessary to sterilise the filtration apparatus

and sample cup between two analyses of the same sample provided that the smaller

volume is processed first.

5.4.4 Sample Processing for Thermotolerant (Faecal) Coliform Analysis.

1. Using the absorbent pad dispenser,

place one pad into each petri-dish (this

operation may be done at base before

leaving for the field). If the dispenser

is damaged, the pads can be dispensed

using the sterilised tweezers.

Dispense one pad into each petri dish

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Pour the medium onto the pad

2. Allow the medium to warm to ambient

temperature before use. Pour enough

culture medium onto the absorbent pad

in the petri dish to soak the pad and

leave a slight excess (approximately

2.5ml). Try not to let the medium run

down the side of the bottle onto the

pad, and do not allow the bottle neck

to come into contact with any external

objects. Replace the bottle cap

immediately. If too much medium

has been added, drain off the excess

and wipe up any spillage with an absorbent cloth or tissue. Always ensure that

a slight excess remains in the petri-dish to prevent the pad drying out during

incubation.

Note: Once the bottle of culture medium has been opened, it is recommended that

the contents are used within one day. It is not advisable to use the medium in one

bottle over several days since this can lead to contamination.

3. Flame the tips of the tweezers

with the lighter for

approximately 5 seconds and

leave to cool.

Flame the tips of the tweezers

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4. Place the heel of the tweezers into the

test kit case as indicated. This ensures

that the tips are kept away from all

sources of contamination whilst

analyses are in progress.

5. Remove the sterile sample cup from

the filtration apparatus. Push the

filtration apparatus firmly onto the

vacuum cup (if this is difficult,

lubricate the black rubber O-ring with

silicone grease; see Section 6 for

maintenance of the kit). Place the

assembly in an upright position in a

convenient place in the kit. Do not

place the apparatus on the ground

where it may become soiled.

6. Unscrew the plastic collar and filtration funnel in order that these may be

easily removed. Do not place these on any surface other than the filtration

base.

Allow the tips of the tweezers to cool

Assemble the filtration apparatus

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7. Using the sterile tweezers, carefully

remove a sterile membrane filter from

the packet.

Hold the membrane only by the edge

and do not let the membrane filter

touch anything while it is being

transferred to the filtration apparatus.

8. With one hand, lift the filtration funnel

and plastic collar above the filtration

base. With the tweezers in your other

hand, place the membrane filter (grid

side facing upwards) onto the bronze

disc filter support. Replace the filter

funnel and collar immediately, without

allowing them to come into contact

with any external objects. Hold the

funnel between the thumb and

forefinger to ensure that the collar will

not slip off and that the fingers do not

come into contact with the interior

surface of the funnel.

9. Screw the plastic collar down tightly

to provide a water tight seal between

the filter membrane and the filter

funnel.

Remove membrane from packet

Place the membrane onto the filter support

Fit the filter funnel and plastic collar

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Note: The plastic collar has 3 adjustment positions:

1. Completely free - the apparatus can be dismantled

when in this position.

2. Loose but not free - all interior surfaces are

exposed to the atmosphere. This is the position

used when sterilising the apparatus.

3. Fully tightened - the funnel forms a tight seal

between the membrane support and the membrane

filter. This is the position for filtration.

10. Rinse the sterile sample cup once with the water before taking the sample.

Take care not to allow external contamination (eg. dirt and debris) to enter the

sample cup.

11. Pour the sample into the filtration

funnel up to the appropriate mark

(10, 50 or 100ml) engraved on the

internal surface of the funnel. To

avoid damaging the membrane, tilt

the filtration apparatus and carefully

pour the first few millilitres of water

down the inside of the filter funnel.

Return the filtration apparatus to the

Upright position and continue adding

the sample. Take care not to allow

external debris to enter the funnel.

Pour the sample into the filtration funnel

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12. Insert the plastic connector of the

vacuum pump into the vacuum

connection on the filtration base.

Squeeze the pump bulb several times

to draw a vacuum, then squeeze as

required to draw all the water through

the membrane filter.

When all the water has passed

through the filter, disconnect the

pump from the filtration apparatus.

Do not allow excess air to be drawn

down through the filter once all the

water has gone through.

13. Unscrew the collar and remove the

funnel and collar with one hand.

Using the sterilised tweezers in the

other hand, lift the membrane

carefully from the filtration base.

Hold the membrane by the edge only.

Attach the vacuum pump and create a

vacuum to draw the sample through the

membrane

Remove the membrane from the filter

head

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14. Remove the lid of a prepared petridish

and place the membrane, grid side

uppermost, onto the absorbent pad

soaked in culture medium.

Start at one edge (it is easier to use the

one at a greater distance from the wall

of the petri-dish) and lower the

membrane on to the pad by ‘rolling’

so as to avoid trapping air bubbles

under the membrane.

15. Replace the lid of the petri-dish and

mark the lid with sample information

eg. volume filtered, source, time and

date; or a code which relates to details

on the daily report sheet. A wax pencil

or marker pen is suitable for this

purpose (the writing needs to be easily

removed after the tests are complete).

16. Place the petri-dish with the lid uppermost

into the carrier (insert the dish at the

bottom so that new dishes can be taken

from the top of the carrier) and return the

carrier to the incubator pot. All 16 petri

dishes should be in the rack during

incubation. This allows for an even

distribution of heat in the incubator.

Replace the incubator lid.

17. Resterilise the filtration apparatus

(Section 3.1).

Lower the membrane on to an adsorbent pad

Replace the lid and label the petri dish

Stack the petri-dishes and place in the

incubator

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5.4.5 Resuscitation of Bacteria

Once the last sample of the day has been taken, wait for a minimum of 60 minutes

before switching on the incubator (resuscitation time). Try to plan the day so that

the time between processing the first and last sample is not more than 3 hours. This

restricts the resuscitation time to a maximum of 4 hours. In cold weather

resuscitation can be achieved by keeping the samples close to the body (eg. In

hands or upright in a pocket but take care to avoid media leaking from the petridishes).

Resuscitation time is particularly important for chlorinated waters or marine water

where the thermotolerant coliform bacteria are ‘stressed’ due to environmental

exposure. For these types of waters it is beneficial to leave processed membranes

for 4 hours after the last sample has been processed before switching on the

incubator.

5.4.6 Sample Incubation

Incubate the samples for 16 to 18 hours. The incubator is designed to maintain a

temperature of 44oC +/- 0.5oC. Always incubate the petri-dishes with the incubator

and case lids closed to reduce heat loss and save battery power (note that it is not

possible to close the kit lid if you are using the double incubator kit, or if you are

powering the kit using the charger or another external power source). Place the kit

on a chair or table to prevent heat loss through the floor and avoid incubating

samples outdoors during cold weather. In order to maximise battery life, do not

leave the incubator on for more than the incubation period, ie. 16 to 18 hours.

There are 3 alternative power sources for the incubator:

1. Mains electricity supply via the charger unit

2. Internal battery

3. External 12v battery (or your vehicle battery)

It is recommended that the mains supply option be used wherever possible.

When used in this way, the charger unit will operate the incubator and at the same

time charge the battery. If the mains electricity fails the internal battery will operate

the incubator.

Using mains electricity or generator via the charger unit

When using mains electricity, the incubator can be operated and the internal battery

charged simultaneously. If the power fails for any reason,

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the internal battery continues the incubation

cycle. When operating from mains

electricity, connect the 3-pin plug to the

socket in the left hand side of the incubator

console.

Plug the battery charger into the mains

electricity socket using an appropriate plug

and switch on the mains. Switch on the

incubator and leave until the incubation

cycle is complete.

Internal Battery

It is possible to obtain up to 5 incubation cycles from the internal battery when it is

new. The number of cycles will reduce as the battery ages. When using the internal

battery in this way, do not use the incubator for more than 5 cycles without

recharging the battery or run the incubator for more than 18 hours during any cycle.

Always recharge the battery fully at every opportunity using mains electricity.

External 12v Battery

If you are planning to work in the field for

more than 5 days, or to work in remote

areas, it is possible to operate the incubator

using an external 12v battery, eg. vehicle

battery, using the connection lead provided

in the spares case.

To operate the incubator from an external

battery, connect the crocodile clips on the

external battery lead to the correct

terminals on the external battery (Red to

Positive or ‘+’, and Black to Negative

or ‘-’). Connect the

Connect the kit to the mains power

supply via the battery charger

Connecting the kit to an external battery

using the crocodile clips connection

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3-pin plug to the left hand side of the incubator console. Switch on the incubator

and check that the ‘Power On’ indicator is lit.

An external battery cannot be used to recharge the internal battery, only to operate

the incubator. Very little current is drawn during incubation and it is usually safe to

operate from a vehicle battery for one incubation cycle without risk of discharging

the vehicle battery excessively. Never run the incubator from a vehicle battery for

more than one cycle if the vehicle is not being driven regularly. Repeated use of the

incubator will drain the vehicle battery.

Note: A poorly maintained external battery may cause the internal battery to

discharge.

5.4.7 Counting Colonies and Recording Results

Note: It is important that counting is completed as soon as possible after the petridishes

have been removed from the incubator (certainly within 15 minutes) as the

positive colonies will change colour on cooling and standing.

1. Once the incubation period is

complete, remove the petri dishes and

their holder from the incubator pot.

Remove the lid of a petri-dish and

observe the surface of the membrane

in good incident light. If necessary,

use a hand lens to examine the

colonies.

2. Count all the yellow colonies which

have a diameter of between one and 3

millimetres (an example is shown on

the front cover).

Frequently, 2 or more colonies will

merge together. Examine the shape

of the colony and it is usually clear

how many colonies have merged

Count all yellow colonies with a diameter

between one and three millimetres

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together. Count each of the sub-colonies. Do not count colonies that are

transparent, red/pink or blue/grey. These are bacteria which do not ferment

lactose and cannot be identified without further study. They are not

thermotolerant coliforms.

Colonies may vary considerably in size. Generally, when the membrane

contains a large number of colonies, the colonies are smaller in diameter.

When colonies are fewer, they tend to be larger. This is because the colonies

compete for nutrients and will grow larger where there is no competition.

If there are large numbers of yellow colonies, count methodically using the

horizontal grid lines. In this way it is possible to count up to100 colonies on a

membrane. If there are more than 100 colonies on the membrane, the number

can be estimated by dividing the membrane into sections and counting the

number of colonies in one section. Multiply the result by the number of

sections to obtain an estimate of the total number of colonies on the membrane.

3. Convert the count into number of thermotolerant coliforms per 100ml and

record the result on the daily report sheet (see Appendix D). The calculation is

made as follows:

Volume filtered Thermotolerant coliforms per 100ml

100ml Number of colonies x 1

50ml Number of colonies x 2

10ml Number of colonies x 10

1ml Number of colonies x 100

5.4.8 Disposal

All contaminated materials should be sterilised before disposal to avoid creating a

risk to the public. DO NOT discard contaminated membranes and filter pads into

the environment. Refer to Section 3.6 for recommended procedures to sterilise

contaminated materials.

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6 Care and Maintenance of the Kit

6.1 The Battery

Never

Allow the internal battery to discharge completely. The useful life of the battery

will be maximised if the battery is always kept in a well-charged state. In order to

ensure this, it is advisable to recharge the battery fully at weekends. If the kit is in

storage, recharge the battery once a month.

Never

Leave the incubator switched on for more than 18 consecutive hours.

Always

Incubate samples with the incubator lid firmly in place and the kit closed.

Always

Operate the incubator in a vehicle or indoors, on a chair or table to prevent heat loss

through a cold floor if possible. Do not operate outside in cold weather.

Always

Recharge the internal battery at the end of a period of work in the field.

Always

Leave the battery in a charged state when the kit is out of use or in storage.

During storage, recharge monthly.

To recharge the battery, connect the small 3-pin plug from the charger to the left

hand side of the incubator console. Plug the charger into the mains electricity

supply and switch on. Check that the incubator is switched off unless it is in use.

The LED on the battery charger will be an orange/amber colour. Continue charging

until the LED light turns green. This process will normally take 3-5 hours with a

new battery, but may take longer with an older or fully discharged battery. When

the battery is completely charged, switch off the charger, disconnect the charger

from the mains electricity supply and the incubator and store in a safe place.

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When using the kit in low temperature environments, eg. less than 10°C, the

maximum number of 18 hour incubation cycles on one battery charge should not

exceed 3.

If any faults or malfunctions are apparent in the kit, refer to the section on ‘Fault

Finding in the Incubator, Battery and Charger’ in Section 7.1.

6.2 Electronic Components and the Incubator

Do not allow water to enter the base of the kit

The electronic components are sealed during construction. This allows a certain

tolerance of moisture. However, always immediately dry any spillage of water or

other liquids inside the kit.

The temperature of the incubator should be checked periodically, eg. Every month,

as indicated in Section 7.3 ‘Checking and Recalibrating the Incubator’.

6.3 Filtration Apparatus

At the end of each day, it is good practice to carefully dry all components of the

filtration apparatus, including the vacuum and sample cups, and to sterilise the

apparatus. This practice prevents corrosion of the metal components of the

filtration apparatus.

6.4 Chlorine and pH Comparator and Turbidity Tubes

Avoid scratching the comparator and turbidity tubes. They rely on an adequate

transmission of light for accurate results.

Keep the surfaces clean and dry and free of residues that may prove difficult to

remove once dry. After use, always wash in clean water.

Approximately once a month wash the comparator and turbidity tubes in a dilute

solution of mild detergent and rinse thoroughly with clean water.

Never use acids or organic solvents.

6.5 Kit Case

The outer case is robust and can withstand a certain amount of harsh treatment.

However, try to avoid abrasion and hard impacts. Periodically clean the case with

warm water and mild detergent.

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6.6 Maintenance

Weekly

1. Wash, rinse and dry the filtration apparatus

2. Apply a smear of silicone grease to the black rubber O-ring

3. Charge the internal battery fully at the end of each week

Monthly

Weekly maintenance, plus:

1. Check the incubator temperature and recalibrate if necessary

2. Clean all components of the kit, including the case

3. Check all components for damage that may affect the operating of the

kit

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7 Evaluation and Repair of the Kit

7.1 Fault Finding in the Incubator, Battery and Charger

1. Connect the battery charger to the incubator. Do not switch on the incubator.

Connect the charger to the mains electricity supply.

Does the LED on the charger unit light up when the mains supply is switched

on?

Yes Go to Step 2

No Go to Step 6

2. Charge the internal battery according to the instructions given in

Section 6.1.

How many hours does it take from switching on the charger to the point

when the LED light turns green, ie the battery is fully charged?

If after 48 hours the LED is not green, the battery is damaged or worn out and

will require replacement. Battery replacement should be carried out only by a

qualified electronics technician. A battery replacement kit is available

DelAgua. Generally, total discharge of the battery is a sign of misuse.

Go to Step 3

3. Prepare the kit for temperature checking and calibration as described in the

section ‘Checking and Recalibrating the Incubator’ in Section 7.3.

? Disconnect the charger from the mains electricity supply.

? Disconnect the charger from the incubator unit.

? Switch on the incubator unit.

Do the two red lights on the incubator console light up brightly?

Yes Go to Step 4

No Go to Step 7

4. Leave the incubator switched on until the temperature reading is stable over a

period of at least 30 minutes. The time taken for the incubator to reach this

point will depend on the ambient temperature, but is usually no more than 3

hours.

Does the incubator hold a temperature of between 43.5 and 44.5°C?

Yes Go to Step 5

No Go to Step 10

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5. Does the incubator hold a temperature of between 43.5 and 44.5 °C for4

incubation cycles of 18 hours each, without needing to recharge the battery?

Note: Between each cycle, leave the incubator to cool for at least 8 hours.

Yes Your incubator and battery charger are in good

condition.

No Go to Step 9

6. If the charger is fitted with a 3-pin UK style plug, the main fuse may have

blown. Ensure that the charger unit is disconnected from the mains electricity

supply. Replace the fuse in the plug. Reconnect the charger to the mains

electricity supply.

Does the LED on the charger light up?

Yes Go to Step 2

No The charger is damaged. Replace with a new unit or

organise repair through DelAgua. Then go to Step 1

Note: The battery charger has been selected to deliver the correct output for the

internal battery. The use of a car battery charger on the equipment will cause

permanent damage to the battery.

7. Reconnect the battery charger to the incubator unit. Connect the charger to the

mains electricity supply. Switch on the incubator.

Do the 2 lights on the incubator console light up brightly?

Yes Go to Step 8

No Go to Step 11

8. Check the reading on the thermometer.

Does the incubator hold a temperature of between 43.5 and 44.5 °C?

Yes Go to Step 9

No Go to Step 10

9. The battery is damaged or worn out. Battery replacement should be carried out

by a qualified electronics technician. A Battery Replacement Kit is available

from DelAgua.

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10. Follow the incubator recalibration procedure in the section ‘Checking and

Recalibrating the Incubator’ in Section 7.3

Does the incubator hold a temperature of between 43.5 and 44.5 °C after

adjustment?

Yes Go to Step 5

No The incubator is damaged. A Repair Kit is available

from Delagua. Contact an electronics technician to carry

out repairs, or return the kit to DelAgua for repair.

11. Disconnect the battery charger from the incubator unit. Connect the incubator

unit to a well charged 12v battery using the lead with crocodile clips supplied

with the kit. Switch on the incubator unit.

Do the 2 lights on the incubator console light up brightly?

Yes Go to Step 12

No The incubator is damaged. A Repair Kit is available

from DelAgua. Contact an electronics technician to

carry out repairs, or return the kit to DelAgua for repair.

12. Check the reading on the thermometer.

Does the incubator hold a temperature of between 43.5 and 44.5°C?

Yes The charger unit is damaged. Replace with a new unit,

then go to Step 1.

No The incubator is damaged. A Repair Kit is available

from DelAgua. Contact an electronics technician to

carry out repairs, or return the kit to DelAgua for repair.

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7.2 Fault Finding Chart

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A damaged or worn out kit can only be repaired by DelAgua if a funding agency

will accept the costs involved, ie. repair, freight and insurance.

Send kits for repair to the address on the back cover of this manual. Before sending

your kit, please remove all loose items from the box such as the turbidity tubes,

chlorine and pH comparator, filtration apparatus, etc. These items may be lost or

damaged in transit, and their additional weight may increase freight costs.

7.3 Checking and Recalibrating the Incubator

The equipment supplied for checking and recalibrating the incubator includes the

following items:

1. Testing incubator lid with centre hole,

2. Thermometer,

3. Trimmer tool (similar to a small screwdriver).

Note: It is recommended that the temperature of the incubator is checked once

every month.

1.3.1 Procedure for checking the incubator temperature

Note: Carry out the following procedure at an ambient temperature of between 15

and 25°C.

1. Remove all contents from the kit

and wipe clean the internal

surfaces with a clean, damp cloth

or paper towel. Pour

approximately 50ml of clean water

into the incubator pot (giving a

depth of approximately 20mm).

2. Push the thermometer through the

hole in the testing lid.

Incubator fitted with testing lid and

thermometer assembly

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3. Replace the incubator lid with the testing lid and thermometer assembly. The

bulb of the thermometer should be completely immersed in the water.

4. Ensure that the internal battery is completely charged, or that the kit is

operating from a mains electricity supply or well-charged external 12v battery.

Switch on the incubator.

5. Check the temperature of the incubator and observe over a period of 30

minutes to make sure that it has stabilised. The incubator normally takes no

more than 3 hours to reach a stable temperature, depending on the ambient

temperature.

6. Once the incubator has stabilised, if the temperature is between 43.5 and

44.5°C, then recalibration is not necessary and the water may be removed with

tissue or a cloth. If the temperature is not within these limits, follow the

recalibration procedure below.

7.3.2 Procedure for Recalibrating the Incubator

1. Leave the testing lid and thermometer assembly in place and keep the

incubator switched on.

2. Insert the trimmer tool into the hole

on the side of the unit and locate

the tool in the calibration screw

(blue trim).

Note: Small adjustments of the

screw result in large changes to the

temperature. A quarter turn (90°)

results in a temperature change of

approximately 1°C.

3. To increase the temperature, turn the adjustment screw anti-clockwise. To

decrease the temperature, turn the adjustment screw clockwise. Make the

adjustments in stages, a little at a time. After each adjustment, leave the

incubator to stabilise for at least 30 minutes. The complete recalibration

procedure may take several hours. Be patient.

?

+

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4. Once the incubator has been recalibrated to read between 43.5 and

44.5°C, leave it switched on for at least 3 hours. Take note of the temperature

at 30 minute intervals to ensure that the temperature is stable.

5. Switch off the incubator and leave to cool. Do not disconnect the incubator

from the mains electricity supply.

6. The following day, switch on the incubator and allow to reach a stable

temperature. If the temperature is not within the correct limit, repeat the

recalibration process detailed in Steps (1) to (4).

7. Dismantle the temperature checking equipment and store in a safe place.

Empty the water out of the incubator and dry the inner surfaces.

Note: The above procedure guarantees an average incubator temperature of 44°C

+/- 0.5°C. After reaching the set temperature, the temperature in the incubator may

vary within +/- 0.5 °C during incubation.

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APPENDICES

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Appendix A

Incubator electronic circuit diagram.

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Appendix B

Field Checklist

Before leaving for the field, check that you have the following items:

Kit

? Filtration apparatus

? Sample cup

? Vacuum cup

? Sample cable

? Vacuum pump

? Turbidity tubes (pair)

? Chlorine/pH comparator

? Tweezers

? Petri dishes

? Spares box (complete)

? Lighter

Consumables

? Culture medium (one bottle per day and a spare)

? Membrane filters

? Absorbent pads and dispenser

? DPD No 1 tablets

? DPD No 3 tablets

? Phenol red tablets

? Methanol

? Daily report sheets (Appendix D)

? Paper towels or clean cloth

? Some clean water to rinse the equipment after use

? A wax pencil or marker pen

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Appendix C

Spares list

The following spares and consumables are available from DelAgua. Please

telephone, fax or email for a current price list.

Alternatively, visit our web site at www.delagua.org

Components

Battery replacement kit containing: Battery 12v 9.5 Ah

Silicone sealant

Temperature check kit containing: Incubator testing lid with hole

Thermometer

Adjuster/trimmer tool

Electrical repair kit containing: Electrical circuit

Temperature chip

Sealant

Foam compound

Adhesives

Filtration apparatus (complete) Spares box (specify empty or

complete)

Filter funnel with plastic collar Tweezers

Filtration base Chlorine/pH comparator

Vacuum cup External battery cable

Sample cup Battery charger

Vacuum pump Silicone grease (2g or 100g)

Sample cable Polypropylene bottles 60ml (x10)

Bronze disc with sealing gasket set Methanol dispenser plastic

Black rubber O-ring Petri-dishes

Turbidity tubes (pair)

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Consumables

Membrane filters and absorbent pads (x200)

Pad dispenser

Culture medium: 38.1g tub for 500ml of growth medium (sufficient for 200 tests)

Culture medium: 500g tub for 6.5 litres of growth medium (sufficient for 2,600

tests)

DPD No 1 Tablets (x250)

DPD No 3 Tablets (x250)

Phenol red Tablets (x250)

Consumables for 200 tests

Optional Extras

Portable conductivity meter

Portable steriliser kit

Other items of equipment and the consumables for the analysis of a range of

chemical parameters can be supplied upon request.

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Appendix D

Daily report sheet

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Appendix E

Alternative sources of water for media preparation

We would strongly recommend that you try to find a source of distilled water to

prepare the medium. However, we appreciate that this may not always be possible

and would suggest the following as suitable alternatives:

? A high quality bottled water that has not been treated with chlorine or any

other disinfectant that has a residual activity.

? Rainwater. Collect a sufficient volume of rainwater in a clean container and

leave it to stand overnight to allow any suspended matter to settle out.

Carefully pour off the water into a separate clean container. Alternatively, the

rainwater can be clarified by filtration through a membrane filter or, more

quickly, through one of the membrane pads (see the relevant paragraphs of

Section 5.4.4).

? Water from a well protected groundwater source. Groundwater from a well

protected source is usually suitable for preparing the medium. However, check

the pH of the water before use to be sure that it is not too acidic or alkaline (see

Section 3.2).

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Appendix F

Additional instructions for operating the dual incubator kit

The dual incubator kit allows you to double the number of tests carried out at one

temperature or test the same sample at 2 temperatures, typically:

? 37°C - Total coliform count (TC).

? 44°C - Thermotolerant coliform count (TTC).

?

The analytical procedures for TC and TTC using MLSB are identical except for the

incubation temperature. Similarly, the colony characteristics of TC are the same as

TTC, although you may find a more varied colony size and differences in the

intensity of the yellow colour.

Many water samples, particularly untreated water samples, will contain bacteria

other than coliform bacteria that can grow on MLSB at 37°C. The colony

characteristics of these background organisms will vary: only the coliform bacteria

will produce yellow colonies. However, be prepared for high background counts

when performing TC analysis.

There are 2 additional procedures that you should be aware of when operating the

dual incubator kit.

Charging and connecting the battery

The dual incubator kit is powered by an external 12V battery that is contained

inside the separate battery pack. On the front of the battery pack, the lead connects

to the dual incubator and the 3-pin socket is used to plug the charger into.

? To charge the battery, connect the battery charger to the 3-pin socket and

charge for 12 hours

? You must use the integral lead on the external battery pack to connect to the

dual incubator. Connect. Note: you must leave the lid of the kit open when the

battery pack is connected to avoid damaging the battery lead.

? The battery pack can be connected to the battery charger and to the kit at the

same time.

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Temperature calibration

The procedure for checking and calibrating the temperature of the 2 incubators is

the same as described in Section 7.3 for the single incubator kit. However, the

corresponding temperature adjuster screws are located underneath the labels

showing the incubator temperatures.

The temperature adjuster screws are located underneath the temperature labels

? ?

+ +

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Appendix G

Alternative types of media that can be used with the Delagua kit for the

isolation of coliform bacteria