Urine Filtration

The urine filtration test is used to identify and quantify S. haematobium eggs in urine; this

tool is commonly used in SCH control programmes as it is one of the few methods available

in the field.

The tool is a fast and easy technique which uses 10mL of urine, the sample is passed

through a nucleopore filter which can then be viewed under a microscope at 40x with iodine

to determine the intensity of the S. haematobium infection. It is important to note that the

reading should be recorded as the number of eggs per the volume used; this will usually be

number of eggs/10mL.

Part 3: Session Activities

Activity 1: Sharing experiences

Participants will form into groups of 4-6 individuals and will discuss their experiences with the

current laboratory and diagnostic tools that they have used in the past and are familiar with.

Groups should highlight the following:

The tools they have used

The challenges associated with each tool

The gaps that they feel exist in NTD laboratory and diagnostic tools

After 10 minutes each group will discuss the key points that they have highlighted with the

facilitators and other participants.

The aim of this activity is for participants to become familiar with the challenges that other

individuals in the NTD community have faced in regard to laboratory and diagnostic tools.

Activity 2: Case study

In DRC, in the provinces of Equateur and Bas Congo, most of the districts are co-endemic to

LF, Loa loa and Onchocerciasis. While completing mapping for LF, Loa loa and

Onchocerciasis, could you describe the diagnostic methods that should be used to confirm

endemicity to LF as we know that MDA with Ivermectin could not be implemented in districts

where LF and Loa loa are endemic?

Which methods will you carry out to map LF?

Which diagnostic methods will you use to map Loa loa?

Which additional laboratory and diagnostic methods will you use to confirm endemicity of LF

and Loa loa in most of the districts?

For each diagnostic and lab methods please explain your choice.

Part 4: Summary Job aide related to this module (include where applicable) Key words (maximum 10)

Key action points for district level personnel

Part 5: References and Additional Resources

References:

CDC Onchocerciasis website (http://www.cdc.gov/parasites/onchocerciasis/)

ICT bench aid (http://www.ntdsupport.org/resources/immunochromatographic-card-test-ict-

bench-aid)

LF test strip bench aid (http://www.ntdsupport.org/resources/filariasis-test-strip-bench-aid)

WHO LF M&E Handbook

(http://whqlibdoc.who.int/publications/2011/9789241501484_eng.pdf)

WHO LF Entomology Handbook

(http://apps.who.int/iris/bitstream/10665/87989/1/9789241505642_eng.pdf?ua=1)

WHO Bench aid for the diagnosis of filarial infections

(http://whqlibdoc.who.int/publications/1997/9241544899_eng.pdf)

WHO Intestinal Helminth Bench Aids for the diagnosis of Intestinal Parasites

(http://apps.who.int/iris/bitstream/10665/37323/1/9789241544764_eng.pdf?ua=1)

Annexes and additional resources:

Annexes

1. ICT SOP

2. LF Test Strip SOP

3. Brugia SOP

4. Thick blood film SOP

5. Sedgwick Counting Chamber SOP

6. Skin Snip SOP

7. Kato Katz SOP

8. Flotac SOP

9. Hemastix SOP

10. Urine filtration SOP

Additional Resources:

1. WHO Bench aid for the diagnosis of filarial infections

2. CDC Onchocerciasis Website

3. WHO LF M&E Handbook

4. WHO LF Entomology Handbook

5. WHO Intestinal Helminth Bench aid

Annex 1: Immuno chromatographic test (ICT ) card standard operating

procedures (SOP)

Guidelines

Storage and Transportation

1. Cards have an approximate shelf life of 3 (WHO, 2011) - 6 months when stored at

30° and up to 9 months when stored at 4°.

2. Test 2 cards from each lot using a positive control, if they appear negative do not use

the lot.

3. When transporting cards to the study locations do not expose them to extreme heat

for prolonged periods of time.

Sample Collection

1. Put on gloves.

2. Clean the site of the finger prick using a disinfectant wipe.

3. Draw a small amount of blood using a lancet to prick the participant’s finger.

4. From this collect 100uL of blood using a capillary tube (supplied with ICT card).

5. Add blood sample to the white portion of the sample pad.

Record the time the blood was added to the card.

DO NOT add blood directly to the pink portion.

DO NOT close the card before the sample migrates to the pink portion; takes roughly

30 seconds.

6. Results are ready to read after 10 minutes; record results by marking the card as positive or

negative

DO NOT read the results at any other time as it can increase the chance of false

positives

7. Safely dispose of the ICT card, capillary tube and any remaining blood found in the capillary

tube

Reference:

World Health Organization, 2011; Global Programme to Eliminate Lymphatic Filariasis: Monitoring

and Epidemiological Assessment of Mass Drug Administration

Annex 2:

LF strip test SOP

Storage and Transportation

1. The kits should be storied at temperatures between 2° and 37°; they should NOT be

frozen. The expiry date that is located on the outer packaging must be followed;

once the data has passed the kits should NOT be used as they are no longer stable.

2. Test 2 strips from each shipment using a positive control, if they appear negative do

not use the lot.

3. When transporting cards to the study locations do not expose them to extreme heat

for prolonged periods of time.

Sample Collection and Strip Usage

Annex 3:

Brugia rapid SOP

The Brugia RapidTM test is an immunochromatographic antibody assay in a cassette format.

These tests are used in LF control programmes to detect the presence of recombinant

protein (BmR1) and specific human IgG4 to both Brugia malayi and Brugia timori infections.

Guidelines

Storage and Transportation

1. The test has a shelf life of 18 months when stored between 20°-25°C and can be

extended in stored at 4°C. They should NOT be frozen.

2. When transporting cards to the study locations do not expose them to extreme heat

for prolonged periods of time.

Sample Collection

1. Put on gloves.

2. Clean the site of the finger prick using a disinfectant wipe.

3. Draw a small amount of blood using a lancet to prick the participant’s finger.

4. From this collect 35uL of blood using a capillary tube (or 25uL of serum/plasma).

5. Once collected the sample and one drop of the chase buffer should be added to the #1 well

on the test kit. (see step 1 on below diagram)

6. Carefully add 3 drops of the provided chase buffer the #2 well. (see step 2 on below diagram)

7. Pull on the clear tab until resistance is felt and add 1 drop of the chase buffer to the square

well.

8. Results are ready to read after 25 minutes for blood samples (or 15 minutes for serum or

plasma samples); record results by marking the card as positive or negative

DO NOT read the results at any other time as it can increase the chance of false

positives

9. Cards must be read in a well-lit location, faint lines can be difficult to read if lighting is poor.

10. Safely dispose of the Brugia Rapid card, capillary tube and any remaining blood found in the

capillary tube

Reference:

World Health Organization, 2011; Global Programme to Eliminate Lymphatic Filariasis: Monitoring

and Epidemiological Assessment of Mass Drug Administration

Annex 4:

Thick blood film SOP

TITLE: PREPARATION OF BLOOD FILM FOR EXAMINING MICROFILARIA DURING

NIGHT BLOOD SURVEYS

Introduction

Night blood surveys of lymphatic filariasis sentinel site targets population aged >5 years and is used

to determine the prevalence and density of microfilariae. Prior to night collection, the local community

should be adequately sensitized and encouraged to assemble at a designated area where the

collection will be carried out. Adequate provision should be made by the technical staff to keep the

participants awake by showing movies and other visuals relevant to the disease on a projector.

Purpose

The purpose of this SOP is to use a properly stained blood smear to detect W. bancrofti microfilariae.

Principle

Night blood survey is normally achieved by collecting night blood, preparing blood films, staining and

microscopic examination of the slides. A properly stained blood slide is an inexpensive method used

for detecting whether a person has microfilariae in the peripheral blood. Wucheraria bancrofti

microfilariae appears in the blood with a marked nocturnal periodicity so blood collection has to be

done between 10.00pm and 2.00 am.

Handling precautions

Personal Protection Equipment (laboratory coat and gloves) should be worn during

this procedure.

Discard all sharps in the sharps boxes.

Materials

Slides

Cotton wool (or lint)

Lancet

Collection tube

Calibrated capillary tube

Micropipettor

Micropipette and tips

Giemsa

Microscope

Labeling pens and pencils

Gloves

Lab coat

Slide racks

Sharp containers

Waste container

70% ethanol

Distilled water

Methanol

Recording form for results

Storage conditions

All dry slides should be stored in the slide rack for transportation to the laboratory for further

processing.

Procedure/ methods

Note: Put on PPE (lab coat and gloves)

1. Reassure the client and make him/her comfortable

2. Clean slide with an alcohol swab to remove lint and oil residue by wiping the slides gently.

3. Label the edge of the slide with a pencil stating the clients identification details.

4. With the client’s left hand palm upwards, select the third or fourth finger.

IMPORTANT -The big toe can be used with infants. The thumb should never be used for adults or

children.

Use cotton wool lightly soaked in ethanol to clean the finger – using firm strokes to remove dirt and

grease from the ball of the finger (Figure 1). Dry the finger with a clean piece of cotton wool (or lint).

Figure 1: Cleaning of the finger before collection of the blood sample (WHO, 2011).

5. With a sterile lancet, puncture the internal side of the finger (Figure 2) using a quick rolling action.

By applying gentle pressure to the finger, express the first drop of blood and wipe it away with dry

cotton wool. Make sure that no strands of cotton wool remain on the finger. Discard the lancet into a

sharp’s container.

Figure 2: Puncture the tip of the finger using a lancet (WHO, 2011).

6. Working quickly and handling clean slides only by the edges, collect the blood as follows:

a. Apply gentle pressure to the finger and collect at least 60 μl of blood into a blood

collection tube or calibrated capillary tube.

b. It helps to hold the capillary tube horizontal (flat) as you collect the blood.

c. Try not to get air bubbles into the capillary tube. If you do, fill the blood past the line to

compensate.

d. Wipe the remaining blood away with cotton wool. Ask the client to hold the cotton

wool firmly on the finger until it stops bleeding.

7. Film preparation: Always handle slides by the edges, or by a corner, to make the blood film as

follows:

a. Use a micropipette to measure 20μl of blood from the collection tube, and prepare

three parallel lines of blood (20μl each) along the length of the slide (Figure 3).

b. Air dry the blood film thoroughly for 24–72 hours. Carefully, load the slides into the

staining racks.

c. Dehaemoglobinize the blood film for approximately 5 minutes in tap water, distilled

water or normal saline.

Note: Caution must be exercised at this time because the smear is fragile, and rough washing or

agitation can result in its floating off the slide.

Figure 3: A prepared blood film, ready to be examined (WHO, 2011).

8. Fixation of slides in methanol: Although fixation in methanol is not absolutely necessary, it results in

better staining of the microfilaria.

a. Air dry the slides. This can be done in the staining racks.

b. Fix in methanol 3–5 minutes by placing the dehaemoglobinized blood film in a

staining trough containing the methanol.

c. Stain with 3% Giemsa by placing slides in a coplin jar containing the giemsa solution

for 50 minutes.

d. Air dry the slides.

9. Examine the preparation under the microscope. Use the x 10 objective first to locate the

microfilaria; then identify the filarial species using the x 40 and x 100 objectives. Note positive slides

and count the number of parasites per slide.

Interpretation and recording of the results

On your recording form, record the presence or absence of microfilaria against each client ID.

Figure 4:Wuchereria bancrofti microfilariae

Reporting of results

All positive results should be discussed with the departmental head.

Archiving and storage of results

All results should be recorded in an excel sheet on a computer. The forms should be filed and stored

properly in the designated cabinet in the laboratory.

Reference

World Health Organization, 2011; Global Programme to Eliminate Lymphatic Filariasis: Monitoring

and Epidemiological Assessment of Mass Drug Administration

Annex 5: Sedgewick counting chamber SOP

Protocol for Counting Chamber for quantitative estimation of microfilaria

The detection of microfilaria in blood requires that the samples are taken during the evening between

the hours of 2100 (9pm) and 0200 (2am). Research teams will be required to test individuals who

showed positive results from the ICT card.

Sample Collection

1. Put on gloves.

2. Clean the site of the finger prick using a disinfectant wipe.

3. Draw a small amount of blood using a lancet to prick the participant’s finger.

4. From this collect 100uL of blood using a capillary tube.

5. Add blood sample (100uL) to sample tubes (small plastic tubes/vials, 2-4 ml, with lid

containing 900uL 3% Acetic Acid)

Rotate the tube gently, end over end to dissolve the blood in the liquid. The acetic

acid will fix and preserve the microfilaria allowing for storage and examination months

later.

Analysis of Sample in Laboratory

1. Transfer all the fluid from the sampling tube to the counting chamber with a pipette through

the opening created by the slanted cover. Remove any air bubbles before examination (this

can be done using a needle)

Follow instructions that come with the Sedgewick Rafter Counting Chamber

2. Place the counting chamber with the specimen under the microscope. Leave the specimen

quietly for approximately 3-5 minutes, to let the microfilaria settle at the bottom of the

chamber. Then examine the specimen under 40 x magnifications.

3. Recorded the number of microfilaria that are present in the counting chamber on the M&E

form with the correct participants identifier as well as the demographic questionnaire.

4. After the examination, the specimen can be transferred back into the specimen tube and kept

for later reference.

5. Clean counting chamber by spraying it out with tap water or distilled water. After drying it can

be re-used.

Reference:

Denham, D.A., Dennis, D.T., Ponnudurai, T., Nelson, G.S., Guy, F., 1971, Comparison of

Counting Chamber and Thick Smear Methods of Counting Microfilaria. Transactions of

the Royal Society of Tropical Medicine and Hygiene 65(4)

McMahon, J. E., Marshall, T.F., Vaughan, J. P., Abaru, D.E., 1979, Bancroftian filariasis: A

comparison of microfilaria counting techniques using counting chamber, standard

slide and membrane (nuclepore) filtration. Annals of Tropical Medicine and Parasitology

73: 457-464.

Annex 6: Skin snip for Onchocerca volvulus

Material

The following materials are required for the skin snip biopsy to be conducted:

Gloves

Disinfectant wipes

Sclerocorneal biopsy punch or needle and scalpel

Specimen tube

Saline

Sample Collection

1. Put on gloves.

2. Clean the site of sample collection using a disinfectant wipe.

Recommended taking up to 6 samples from the iliac crest, the scapula or the lower

extremities

3. Using the Sclerocorneal biopsy punch collects the sample. This will result the removal of

approximately 2mg of tissue.

OR

Using a needle raise a small cone of skin (approximately 3mm in diameter) and shave off with

a scalpel. This will result the removal of approximately 2mg of tissue.

4. Place the tissue sample into a pre-marked specimen tube that is filled with saline.

Incubate this at ambient temperature for 24 hours to allow the microfilariae to emerge

from the sample.

5. Review saline solution to identify microfilariae microscopically; this should be

prepared in the traditional wet preparation manner.

Identify and record the number of microfilariae that are detected in each

sample.

Reference:

CDC Website; Onchocerciasis webpage.

http://www.cdc.gov/parasites/onchocerciasis/health_professionals/index.html

Annex 7: Katokatz test

Diagnosis of: Schistosoma mansoni, Trichuris trichiura, Ascaris lumbricoides,

Ancylostoma duodenale and Necator americanus

General Principle: people infected with STH or intestinal Schistosomes pass the eggs of

the worms through their faeces. By examining a stool specimen under a microscope it is

possible to count the number and the type of eggs that are present.

Safety precautions

The stool should be considered potentially infectious.

Wear gloves and lab coats whenever handling stool samples.

Benches, instruments and equipment should be routinely decontaminated with

disinfectants after use.

Materials contaminated with infectious waste should be disinfected before disposal.

Drinking or eating during laboratory procedures is prohibited.

Appropriate disinfectant(s) should be used for disposal of contaminated materials,

wooden spatulas and specimen containers and for cleaning of workbenches.

Used specimen containers must be disinfected before washing.

Equipment for Kato Katz

Kato-Katz:

Stool sample in container

Glass slides

Cellophane sheets

Malachite green

Glycerol

Metal Sieve (Endecott Sieve) with 212um aperture

Slide Boxes

Newspapers

Wooden Applicators

Forceps

Kato-Katz Kit:

o 400 plastic templates with a hole of 6mm on a 1.5mm thick template (delivering

41.7mg of faeces)

o A roll of hydrophilic cellophane (34um thick, 20m)

o 400 plastic applicator stick/spatula

o A roll of nylon screen sieving mesh (20m)

Microscopic examination:

Microscope

Hand tally counter

Laboratory forms

Disinfectants and waste disposal:

Disinfectant wipes

Medicated soap

Methylated spirit

Waste container (containing disinfectant)

Preparation of Kato Katz Reagents Images

Step 1: Weigh out 0.5g of Malachite green powder.

Step 2: Dilute it in 100ml of distilled water (this is the “stock

solution”).

Step 3: Dilute 50ml of glycerine in 50ml of distilled water.

Step 4: Take 1 ml of Malachite stock solution and add it to

100ml of the 50% glycerine solution (this is the “working

solution”).

Step 5: Cut cellophane into 25mm x 30mm pieces and soak

them overnight in the working solution.

Kato-Katz Steps Images

Step 1: Label a glass slide with the sample number and then

place a plastic template on top of it.

Step 2: Place a small amount of the faecal specimen on a

newspaper and press through the underside of the metal

sieve. Using a spatula, scrape the sieved faecal material

through the sieve so that only the debris remains.

Step 3: Scrape up some of the sieved faeces to fill the hole

in the template, avoiding air bubbles and levelling the faeces

off to remove any excess.

Step 4: Carefully lift off the template and place it in a bucket

of water mixed with concentrated detergent so that it can be

reused.

Step 5: Place one piece of the cellophane, which has been

soaked overnight in methylene blue glycerol solution, over

the faecal specimen.

Step 6: Place a clean slide over the top and press it evenly

downwards to spread the faeces in a circle. If done well, it

should be possible to read newspaper print through the stool

smear.

Step 7: If hookworm is present in the area, the slide should

be read within 30–60 minutes. After that time, the hookworm

eggs disappear.

Microscopic Examination of S.mansoni and STH Images

Step 1: To read the slide, place it under the microscope using x400 magnification objective.

Step 2: Read ALL fields of the slide using the vertical ‘zig zag’ scheme and use the tally counter to record how many eggs are seen under the slide as it is read.

Step 3: Record the number and the type of each egg on a recording form alongside the sample number. If no eggs are seen, record “0”.

Step 4: Remove the faeces and cellophane using a tissue into the waste container and place all slides used when conducting Kato-Katz into the disinfectant. These slides should be cleaned and used again for the survey.

Note:

The quality control when reading the Kato-Katz slides is important. For example, confirming

the agreement % for laboratory technicians to ensure quality (see the agreement % on a

specimen collection).

Annex 8: Flotac test

1. Introduction

Faecal egg count (FEC) techniques are widely used for parasitological diagnosis in humans and animals.

They assess the number of parasitic elements (eggs, larvae, oocysts) present in the faecal samples,

expressed per gram of faeces. The FLOTAC apparatus is also very useful in order to recover parasitic

elements after flotation. In this protocol the two flotation solutions should be used concurrently to maximize

yield of parasitic elements.

2. Purpose

The purpose of this SOP is to describe the use of the FLOTAC technique for the diagnosis of soil-

transmitted helminthiasis in monitoring and evaluation (M&E) programmes.

3. Precaution

1. Stool samples are potentially hazardous and should therefore be handled with care.

2. PPE must be worn at all times during the process

3. The solutions used can be hazardous

4. Avoid spillage

4. Materials

1. Fill – FLOTAC

2. Mini – FLOTAC (Fig 1)

3. Flotation solutions:

i. saturated Sodium chloride, S.G = 1.20 for STH

ii. Zinc sulphate; s.g. = 1.35, for S. mansoni and intestinal protozoa

4. 5% formalin solution

5. Tally counter

6. Gloves

7. Hand sanitizers

8. Detergent

9. Bleach

10. Plastic waste bags

5. Storage conditions

Unprocessed stool samples must be stored at 4oC or polyvinyl alcohol (PVA) solution added as

preservative.

6. Procedure

1 Take 1 gram* of stool and put into fill-FLOTAC container

2. Add 1ml of 5% formalin and screw the cap

3 Shake vigorously to homogenize

4. Open screw cap and add the flotation fluid to reach 20ml (1:10 sample dilution)

5. Fit back screw cap and shake vigorously to homogenize

6. Filter and fill the two flotation chambers

7. Wait for 5-10 min, translate and examine under microscope (using a maximum of x400

magnification and count parasite elements individually

Note: * X gram of stool requires X ml of 5% formalin

7. Result and Test interpretation

The intensity of infection was calculated in eggs per gram (EPG).

8.Storage and Archiving

The responsible personnel is to ensure safe storage and proper archiving of results.

9. Measurement of uncertainty

Despite their high sensitivity, a main limitation of the FLOTAC techniques is the complexity of the method

which involves centrifugation of the sample with a specific device, equipment that is often not available in

laboratories in developing countries. To overcome this bottleneck, a new simplified device has been

developed, namely the mini-FLOTAC.

10. Reference

1. Cringoli G. (2006) FLOTAC, a novel apparatus for a multivalent faecal egg count technique.

Parassitologia.;48(3):381-4.

2. Rinaldi L1, Maurelli MP, Musella V, Santaniello A, Coles GC, Cringoli G. (2010) FLOTAC: an

improved method for diagnosis of lungworm infections in sheep. Vet Parasitol.;169(3-4):395-8.

3. Barda BD, Rinaldi L, Ianniello D, Zepherine H, Salvo F, et al. (2013) Mini-FLOTAC, an Innovative

Direct Diagnostic Technique for Intestinal Parasitic Infections: Experience from the Field. PLoS

Negl Trop Dis 7(8): e2344. doi

Annex 9: Hemastix test

Diagnosis of: Schistosoma haematobium.

Equipment for Hemastix test

Hemastix test strip and Hemastix pot with scale

Scissors

Gloves

Disinfectants and waste disposal

Data collection form

Steps for Reagent Strips Images

Step 1: Collect a fresh urine specimen in a clean plastic container. Ensure

that the urine is tested in the field within 2 hours of collection. If there is a

delay, refrigerate the specimen if possible.

Step 3: Remove one strip from its bottle (you can cut the strip in two to

save resources) and label the strips with the patient identification.

Step 4: Completely immerse the reagent areas of the strip into the urine

specimen for a few seconds.

Step 5: When removing the strip, run its edge against the rim of the

container to remove any excess urine.

Step 6: Put the strip horizontally on the table so that the chemicals do not

mix together.

Step 7: Read the strip between 1 and 2 minutes after it has been dipped in

the urine specimen.

Step 8: Match the colour of the strip with the

colour chart on the bottle label and record the

results on the monitoring form. Record “0” if

the result is negative.

1= trace haemolysed

2 = trace non-haemolysed

3 = +

4 = ++

5 = +++

Important Note:

DO NOT LAY THE STRIP ON THE COLOUR CHART AS THIS WILL SOIL THE

CHART

It is extremely important to read the strip 1-2mins after it has been dipped in the urine

sample. Any colour changes that occur after 2 minutes are of no diagnostic value and

should be ignored.

Annex 10: Urine filtration SOP

Diagnosis of: Schistosoma haematobium

Safety precautions

The urine should be considered potentially infectious.

Wear gloves and lab coats whenever handling urine samples.

Benches, instruments and equipment should be routinely decontaminated with

disinfectants after use.

Materials contaminated with infectious waste should be disinfected before disposal.

Drinking or eating during laboratory procedures is prohibited.

Appropriate disinfectant(s) should be used for disposal of contaminated specimen

containers and for cleaning of workbenches.

Used specimen containers must be disinfected before washing

Equipment

General use:

Gloves

Laboratory Forms

Urine Filtration:

Urine pots (250ml)

Swinnex Filter Holder

Tweezers/Forceps

Syringe, plastic, 10ml

Nucleopore Membrane Filter, diameter 13mm and pore size of 12µm

Microscope glass slides

Lugol’s Iodine (5% solution)

Microscopic examination:

Microscope

Hand tally counter

Disinfectants and waste disposal:

Bucket (to discard urine)

1% hypochlorite solution (domestic bleach)

Methylated Spirit

Medicated soap

Rubber washing gloves

Disinfectant wipes

Waste container (containing disinfectant)

Sample collection:

The number of ova in the urine varies throughout the day, with the highest between 10am and

2pm. The specimen should be taken between these times and consist of a single urine sample.

Since eggs are more often found at the end of a urine flow, at least 10ml should be collected at

the end of urination (the terminal urine). The easiest way to ensure a terminal urine sample is to

ask individuals to ‘try to fill’ a large pot, e.g. 250ml. Note that some children, particularly those

who are heavily infected with schistosomiasis, may not be able to provide 10ml of urine. Do not

discard these smaller samples, but note the volume (ml) of urine provided. Specimens should

be examined as soon as possible after collection as the eggs may hatch and then become

invisible, or crystals may form, making a correct diagnosis more difficult.

IMPORTANT NOTE: To increase the volume of urine provided during sample collection, it

would be advisable to promote fluid intake and physical exercise prior to micturition (e.g. provide

the children with 2 glasses of water, one hour before urine collection, and request the children to

participate in 10 minutes of exercise) (Doehring et al. 1983).

Steps for Urine Filtration Images

Step 1: Unscrew the filter holder and insert a nucleopore filter between the two

parts of the filter holder. Make sure it is correctly held in place before screwing

the unit together again.

Step 2: Shake and mix the urine specimen before drawing a 10ml specimen into

the syringe. Then attach the filter unit.

If less than 10ml urine sample is available, withdraw all urine in the sample

pot and note the quantity of urine (ml) on the laboratory form next to the ID

number. Do not discard the urine sample if it is less than 10ml.

Step 3: Keeping the syringe and the unit in a vertical position, press the plunger

down to push all the urine through the filter and out into a bucket.

Step 4: Carefully detach the syringe from the filter unit. Draw air into the syringe,

reattach the syringe to the filter unit holder and expel the air again. This is

important as it removes any excess urine and ensures that the eggs are firmly

attached to the filter.

Step 5: Unscrew the filter holder and use a pair of tweezers to remove the filter

and place it inverted, onto the glass microscope slide. The top side of the filter,

where the eggs were captured, should be face-up on the slide.

DO NOT DISCARD THE FILTER HOLDER OR SYRINGE.

Step 6: Add one drop of Lugol’s iodine and wait 15 seconds for the stain to

penetrate the eggs. This makes the eggs more easily visible.

Step 7: Immediately examine the whole filter under a microscope at a low power

(x40). Schistosome eggs can be seen clearly because they stain orange.

Infection loads are recorded as the number of eggs per 10ml of urine.

Step 8: At the end of the day, wash all reusable equipment (forceps, filter

holders, syringes, urine containers, glass slides) for use next day, discard used

filters and clean the workbench.

Where two urine samples are required: Repeat Steps 1-7 to prepare a second duplicate filter

from the same urine sample, and place it on the glass slide next to the first filter, or on another

slide labeled with the same ID code. The syringe can be re-used for this second filtration on the

same urine sample. However, ensure that a clean syringe is used for each different urine

sample (i.e. from two individuals). Two filters from the urine sample should be read by two

independent laboratory technicians.

IMPORTANT: Read the slide within an hour of the urine sample being taken otherwise the

eggs may be non-viable and become translucent. Do not leave the samples exposed to

the sun.