Part 3: Session Activities

Transcription

1 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.

2 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 ( ICT bench aid ( LF test strip bench aid ( WHO LF M&E Handbook ( WHO LF Entomology Handbook ( WHO Bench aid for the diagnosis of filarial infections ( WHO Intestinal Helminth Bench Aids for the diagnosis of Intestinal Parasites ( Annexes and additional resources: Annexes 1. ICT SOP

3 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

4 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 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

5 Reference: World Health Organization, 2011; Global Programme to Eliminate Lymphatic Filariasis: Monitoring and Epidemiological Assessment of Mass Drug Administration

6 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

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8 Annex 3: Brugia rapid SOP The Brugia Rapid TM 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 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.

9 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

10 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

11 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.

12 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 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).

13 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.

14 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.

15 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:

16 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.

17 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

18 Hand tally counter Laboratory forms Disinfectants and waste disposal: Disinfectant wipes Medicated soap Methylated spirit Waste container (containing disinfectant) Preparation of Kato Katz Reagents 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 ). Images 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.

19 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 minutes. After that time, the hookworm eggs disappear. Microscopic Examination of S.mansoni and STH 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. Images 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).

20 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 soiltransmitted 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 4 o C 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

21 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): Rinaldi L 1, 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): 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

22 Annex 9: Hemastix test Diagnosis of: Schistosoma haematobium. Equipment for Hemastix test Hemastix test strip and Hemastix pot with scale Scissors Gloves Steps for Reagent Strips Disinfectants and waste disposal Data collection form 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 = +++

23 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.

24 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)

25 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.

26 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.

27 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.