规格型号：
序号 货 号 型 号
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
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OXFAM - DELAGUA
Portable Water Testing Kit
VERSION 4.3
REVISED 2012
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
Portable Water Testing Kit
Users Manual
Revised 2012
This equipment was designed to test for the critical beters 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 inbation 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
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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 bs
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 bs. 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 - baldehyde 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 b (see Section 5.4.4 [9])
which allows the baldehyde gas to
penetrate all areas of the filter head.
Dry the sample cup
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3. Pour about 1ml (approximaby
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 compleby
burned up (ie. as the flames are dying
down), place the filtration head over
the sample cup and push firmly into
place to b a good seal.
Add 20 drops (approximaby 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 immediaby 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, approximaby 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 approximaby 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 (approximaby 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
perbed 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 compleby. Apply gentle heat if the powder is
slow to dissolve. The culture medium will be b with a bright red colour
when dissolved.
6. Pour a suitable volume of culture medium (approximaby 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 perbed 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 b 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 bs. If this is not
possible, then the following b 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 immediaby 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 bs).
Some kit operators prefer this b 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 b 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 bs 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 b 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
b 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 approximaby 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 b
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 b 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
b 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
colib 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) colib bacteria
and therefore it may not be necessary to carry out thermotolerant colib analysis.
If the results do not meet these criteria, it will be necessary to carry out
thermotolerant colib 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 compleby. 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. Immediaby 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
bies 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 bies with your fingers or with everyday bs 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 colibs 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 colib bacteria, but inhibits the
growth of any other bacteria caught in the filter. To ensure that only thermotolerant
colib bacteria grow, the filter is kept at 44°C in the kit’s incubator. During this
time the colib bacteria multiply many times to b bies that can be seen
with the naked eye. Thermotolerant colibs 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 by-bing
units per 100ml of water (CFU/100ml).
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Thermotolerant colibs 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 b
should be taken to improve contaminated water supplies.
Some users may need to analyse for total colib bacteria, which, although of less
sanitary significance than thermotolerant colibs, can be used to indicate hygiene
problems in large distribution networks. Total colib analysis is carried out using
the same procedure as for thermotolerant colibs, 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 colib and total colib 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 bs 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 b.
5.4.3 Selection of Appropriate Sample Volumes for Colib Analysis
The most appropriate volume to process is that which allows the most accurate
count of the bacterial bies. It is generally agreed that 100 bies is the
maximum that can be counted reliably on a 47mm membrane. Counts above 100
are considered an estimate. If a large number of bies develops on the
membrane you can either divide the plate into sections, count the bies 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 inbation 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 colib bacteria.
For these waters we would recommend using a 100ml sample.
Suggested sample volumes for thermotolerant colib 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 colib 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) Colib 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 (approximaby
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
bs. Replace the bottle cap
immediaby. 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
approximaby 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 b 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 immediaby, without
allowing them to come into contact
with any external bs. 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 bs:
1. Compleby free - the apparatus can be dismantled
when in this b.
2. Loose but not free - all interior surfaces are
exposed to the atmosphere. This is the b
used when sterilising the apparatus.
3. Fully tightened - the funnel bs a tight seal
between the membrane support and the membrane
filter. This is the b 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 b 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 inbation
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 colib 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 bies 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
bies.
2. Count all the yellow bies which
have a diameter of between one and 3
millimetres (an example is shown on
the front cover).
Frequently, 2 or more bies will
merge together. Examine the shape
of the by and it is usually b
how many bies have merged
Count all yellow bies with a diameter
between one and three millimetres
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together. Count each of the sub-bies. Do not count bies 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 colibs.
Colonies may vary considerably in size. Generally, when the membrane
contains a large number of bies, the bies are smaller in diameter.
When bies are fewer, they tend to be larger. This is because the bies
compete for nutrients and will grow larger where there is no competition.
If there are large numbers of yellow bies, count bically using the
horizontal grid lines. In this way it is possible to count up to100 bies on a
membrane. If there are more than 100 bies on the membrane, the number
can be estimated by dividing the membrane into sections and counting the
number of bies in one section. Multiply the result by the number of
sections to obtain an estimate of the total number of bies on the membrane.
3. Convert the count into number of thermotolerant colibs per 100ml and
record the result on the daily report sheet (see Appendix D). The calculation is
made as follows:
Volume filtered Thermotolerant colibs per 100ml
100ml Number of bies x 1
50ml Number of bies x 2
10ml Number of bies x 10
1ml Number of bies 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 compleby. 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 compleby 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 immediaby 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 bl 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.
Approximaby 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
approximaby 50ml of clean water
into the incubator pot (giving a
depth of approximaby 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 compleby immersed in the water.
4. Ensure that the internal battery is compleby 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
approximaby 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
bephone, 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 beters 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 colib count (TC).
44°C - Thermotolerant colib count (TTC).
The analytical procedures for TC and TTC using MLSB are identical except for the
incubation temperature. Similarly, the by characteristics of TC are the same as
TTC, although you may find a more varied by size and differences in the
intensity of the yellow colour.
Many water samples, particularly untreated water samples, will contain bacteria
other than colib bacteria that can grow on MLSB at 37°C. The by
characteristics of these background organisms will vary: only the colib bacteria
will produce yellow bies. However, be prepared for high background counts
when perbing 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 colib bacteria
