Safety

Follow universal precautions for the prevention of bloodborne pathogens when working with human serum and other body fluids.  These include:

Wear personal protective equipment such as safety glasses, gloves, laboratory coats. If you have cuts or abrasions on the skin of your hands, cover them with adhesive

dressing.

Use needles and lancets only once, and dispose of them in a “sharps” container for decontamination.

Remove gloves and wash your hands after completing any task involving the handling of biological material.

Specimen Collection

Timing:Whenever possible, specimens should be collected before treatment is initiated.  When malaria and babesiosis are suspected, blood smears should be obtained and examined without delay.  Since the parasitemia may fluctuate, multiple smears might be needed.  These can be taken at 8 to 12 hour intervals for 2 to 3 days.

Microfilariae exhibit a marked periodicity depending on the species involved, therefore the time of specimen collection is critical.  If a filarial infection is suspected, the optimal collection time for demonstrating microfilariae is:

Loa loa—midday (10 AM to 2 PM)Brugia or Wuchereria—at night, after 8 PMMansonella—any timeOnchocerca—any time

Type of Sample:Venous blood samples provide sufficient material for performing a variety of diagnostic tests, including concentration procedures (filariasis, trypanosomiasis).  However, in some parasitic diseases (e.g., for diagnosis of malaria in particular), anticoagulants in the venous blood specimen can interfere with parasite morphology and staining characteristics; this problem can be further compounded by excessive delays prior to making the smears.  In such cases, capillary blood samples are preferable.  If PCR is required, please refer to the molecular diagnosis section for appropriate blood collection procedures.

Capillary blood obtained by fingerstick:

1. Label pre-cleaned slides (preferably frosted-end) with the patient’s name (or other identifier) and date and time of collection.

2. Clean the site well with alcohol; allow to dry.

3. Prick the side of the pulp of the 3rd or 4th finger (alternate sites include ear lobe, or in infants large toe or heel).

4. Wipe away the first drop of blood with clean gauze.

5. Prepare at least 2 thick smears and 2 thin smears.

Venous blood obtained by venipuncture:

1. Label collection tubes and pre-cleaned slides (preferably frosted-end) with the patient’s name (or other identifier) and date and time of collection.

2. Clean the site well with alcohol; allow to dry.

3. Collect the venous blood in a vacuum tube containing anticoagulant (preferably EDTA); alternatively, collect the blood in a syringe and transfer it to a tube with anticoagulant; mix well.

4. Prepare at least 2 thick smears and 2 thin smears as soon as possible after collection.

Video link of smear preparation: http://www.dpd.cdc.gov/dpdx/HTML/Filariasis.htm

Specimen Processing

Preparing Blood SmearsIf you are using venous blood, blood smears should be prepared as soon as possible after collection (delay can result in changes in parasite morphology and staining characteristics).

Thick smearsThick smears consist of a thick layer of dehemoglobinized (lysed) red blood cells (RBCs).  The blood elements (including parasites, if any) are more concentrated (app. 30×) than in an equal area of a thin smear.  Thus, thick smears allow a more efficient detection of parasites (increased sensitivity).  However, they do not permit an optimal review of parasite morphology.  For example, they are often not adequate for species identification of malaria parasites: if the thick smear is positive for malaria parasites, the thin smear should be used for species identification.

Prepare at least 2 smears per patient!

1. Place a small drop of blood in the center of the pre-cleaned, labeled slide. 2. Using the corner of another slide or an applicator stick, spread the drop in a circular pattern until it is the

size of a dime (1.5 cm2).

3. A thick smear of proper density is one which, if placed (wet) over newsprint, allows you to barely read the words.

4. Lay the slides flat and allow the smears to dry thoroughly (protect from dust and insects!).  Insufficiently dried smears (and/or smears that are too thick) can detach from the slides during staining.  The risk is increased in smears made with anticoagulated blood.  At room temperature, drying can take several hours; 30 minutes is the minimum; in the latter case, handle the smear very delicately during staining. 

You can accelerate the drying by using a fan or hair dryer (use cool setting).  Protect thick smears from hot environments to prevent heat-fixing the smear.

5. Do not fix thick smears with methanol or heat.  If there will be a delay in staining smears, dip the thick smear briefly in water to hemolyse the RBCs.

Click here to view a video clip for proper preparation of a thick blood smear (Adobe Flash)

Thin smearsThin smears consist of blood spread in a layer such that the thickness decreases progressively toward the feathered edge.  In the feathered edge, the cells should be in a monolayer, not touching one another.

Prepare at least 2 smears per patient!

1. Place a small drop of blood on the pre-cleaned, labeled slide, near its frosted end. 2. Bring another slide at a 30-45° angle up to the drop, allowing the drop to spread

along the contact line of the 2 slides.

3. Quickly push the upper (spreader) slide toward the unfrosted end of the lower slide.

4. Make sure that the smears have a good feathered edge.  This is achieved by using the correct amount of blood and spreading technique.

5. Allow the thin smears to dry.  (They dry much faster than the thick smears, and are less subject to detachment because they will be fixed.)

6. Fix the smears by dipping them in absolute methanol.

Click here to view a video clip for proper preparation of a thin blood smear (Adobe Flash)

Note: Under field conditions, where slides are scarce, national malaria programs (and CDC staff) prepare both a thick and a thin smear on the same slide.  This works adequately if one makes sure that of the two smears, only the thin smear is fixed.

Special Procedures for Detecting MicrofilariaeBlood microfilariae:

A. Capillary (fingerstick) bloodSince microfilariae concentrate in the peripheral capillaries, thick and thin smears prepared from fingerstick blood are recommended.

B. Anticoagulated (EDTA) venous blood (1 ml) should be concentrated by one of the  following methods:

1. Centrifugation (Knott’s technique)

a. Prepare 2% formaldehyde (2 ml of 37% formaldehyde + 98 ml H2O).

b. Mix 9 ml of this 2% formaldehyde with 1 ml of patient’s venous blood.  Centrifuge at 500 × g for 10 minutes; discard supernatant.  Sediment is composed of WBCs and microfilariae (if present).

c. Examine as temporary wet mounts.

d. Prepare thick and thin smears; allow to dry; dip in absolute methanol before Giemsa staining to enhance staining of microfilariae.

2. Filtration

a. Place Millipore® or Nucleopore® membrane filter (5 µm pore) in filter holder with syringe attachment.

b. Mix 1 ml of venous blood (in EDTA) with 10 ml of 10% Teepol® 610 (Shell Co.); allow to stand for several minutes to allow lysis; transfer to a 10 ml Luer-Loc® syringe; attach the filter apparatus.

c. Force the solution through the 5 µm pore filter, followed by several syringes of water to wash out the remaining blood, then 1 or 2 syringes full of air to clear excess fluid.

d. Prepare a temporary wet mount by removing the filter and placing it on a glass slide, adding a drop of stain or dye and a coverslip.

e. For permanent preparations, pass 2 to 3 ml of methanol through the filter while it is still in the holder; remove filter and dry it on a glass slide; then stain it with Giemsa stain, horizontally (so that the filter does not wash off the slide); coverslip filter before examining.

Shipment

Note: When shipping a specimen to CDC, make sure your package will arrive on a weekday and will not arrive at CDC on the weekend or a federal holiday.

1. In emergencies, call the Epidemiology Branch of the Division of Parasitic Diseases at (770) 488-7760 to make special arrangements.

2. For routine requests, submit the specimen to the appropriate city, county or state health department laboratory (see http://www.aphl.org) for processing and examination.  That facility will refer specimens to CDC if necessary.

3. Figure 1 shows correct labeling and packaging of specimens (http://www.cdc.gov/od/ohs/biosfty/bmbl4/b4acf1.htm).

4. If shipping a specimen, please refer to shipping regulations and guidelines at the following addresses:

1. Guidelines for the Safe Transport of Infectious Substances and Diagnostic Specimens, (World Health Organization)

2. The IATA Dangerous Goods Regulations Manual, (International Air Transport Association) http://www.iata.org/ps/publications/dgr

3. Title 49 Code of Federal Regulations, Parts 100-185.  Hazardous Materials regulations (Department of Transportation) http://www.phmsa.dot.gov/hazmat/regs

4. Title 42 Code of Federal Regulations, Part 72.  Interstate shipment of etiologic agents (Department of Health and Human Services) http://www.cdc.gov/od/ohs/biosfty/shipregs.htm

5. Title 42 Code of Federal Regulations, Part 72.6.  Additional requirements for facilities transferring or receiving select agents (Department of Health and Human Services) http://www.selectagents.gov/

6. Biosafety in Microbiological and Biomedical Laboratories (CDC/NIH) http://www.cdc.gov/od/ohs/biosfty/bmbl/bmbl-1.htm

Shipping blood smears for microscopic examination

1. Place labeled dried blood smears (stained and unstained) in a slide box, with grooves to separate the slides.

2. Pack the slide box inside another box, cushioned so that the slides are protected from breakage.

3. Include information:

a. Submitter’s name, address and phone number

b. Physician’s name, address and phone number

c. Patient’s name, travel history (places and dates) and treatment information

d. Specimen collection date

e. What organism is suspected

4. Ship via mail or package carrier. 

Shipping whole blood for isolation of parasites (culture or animal inoculation), molecular diagnosis or drug level

1. Place labeled tube of anticoagulated (EDTA) blood in enough absorbent material to contain any leakage, and place in a sealed plastic bag or 50 ml screw cap centrifuge tube.

2. Pack this bag or container in a box, cushioned so that the blood tube is protected from breakage.

3. Include information:

a. Submitter’s name, address and phone number

b. Physician’s name, address and phone number

c. Patient’s name, travel history (places and dates) and treatment information

d. Specimen collection date

e. What tests are requested and what organisms are suspected

4. Ship via courier or overnight delivery to permit optimum recovery of parasites; refrigeration during shipment may be necessary and should be discussed beforehand with the receiving laboratory.

Staining

Staining Blood SmearsStain only one set of smears, and leave the duplicates unstained.  The latter will prove useful if a problem occurs during the staining and/or if you wish later to send the smears to a reference laboratory.

Wright (Wright-Giemsa) stainUsed in hematology, this stain is not optimal for blood parasites.  It can be used if rapid results are needed, but should be followed up when possible with a confirmatory Giemsa stain, so that Schüffner’s dots can be demonstrated.

Giemsa stain - Recommended for detection and identification of blood parasites.

1. Stock 100× Giemsa Buffer 0.67 M

   Na2HPO4 59.24 g   NaH2PO4H2O 36.38 g   Deionized water 1000.00 ml   Autoclave or filter-sterilize (0.2 µm pore).  Sterile buffer is stable at room temperature for one year. 2. Working Giemsa Buffer 0.0067M, pH 7.2   Stock Giemsa Buffer 10.0 ml   Deionized water 990.0 ml   Check pH before use.  Should be 7.2.  Stable at room temperature for one month.3. Triton X-100 5%    Deionized water (warmed to 56°C) 95.0 ml

   Triton X-100 5.0 ml Prewarm the deionized water and slowly add the Triton X-100, swirling to mix.

4. Stock Giemsa stain (Giemsa stain is available commercially, but the following formulation gives more constant results and does not expire)

   Glass beads, 3.0 mm 30.0 ml    Absolute methanol, acetone-free 270.0 ml    Giemsa stain powder (certified) 3.0 g    Glycerol 140.0 ml

1. Put into a 500 ml brown bottle the glass beads and the other ingredients, in the order listed.  Screw cap tightly.  Use glassware that is clean and dry.

2. Place the bottles at an angle on a shaker; shake moderately for 30 to 60 minutes daily, for at least 14 days.

3. Kept tightly stoppered and free of moisture, stock Giemsa stain is stable at room temperature indefinitely (stock stain improves with age).

4. Just before use, shake the bottle.  Filter a small amount of this stock stain through Whatman #1 filter paper into a test tube.  Pipet from this tube to prepare the working Giemsa stain.

5. Working Giemsa stain (2.5%): make fresh for each batch of smears

   Working Giemsa buffer 39 ml   Giemsa Stain Stock 1 ml   5% Triton X-100 2 drops

Staining

1. Prepare fresh working Giemsa stain in a staining jar, according to the directions above.  (The 40 ml fills adequately a standing Coplin jar; for other size jars, adapt volume but do not change proportions).

2. Pour 40 ml of working Giemsa buffer into a second staining jar.  Add 2 drops of Triton X-100.  Adapt volume to jar size.

3. Place slides into the working Giemsa stain (2.5%) for 45-60 minutes.

4. Remove thin smear slides and rinse by dipping 3-4 times in the Giemsa buffer.  Thick smears should be left in buffer for 5 minutes.

5. Dry the slides upright in a rack.

Note: As alternates to this 45-60 minutes in 2.5% Giemsa stain, the smears could be stained for shorter times in more concentrated stains.  One alternate is 10 minutes in 10% Giemsa; the shorter stains yield faster results, but use more stain and might be of less predictable quality.

Staining Procedure: Quality ControlTo ensure that proper staining results have been achieved, a positive smear (malaria) should be included with each new batch of working Giemsa stain.  Since good quality control smears are not available commercially, they may be prepared from a patient’s blood and stored for future use in the following manner:

1. Choose a patient blood specimen, anticoagulated with EDTA, that has enough parasites so that at least one is found in every 2 to 3 fields.

2. Make as many thin smears as possible, preferably within one hour after the blood was drawn from the patient.

3. Allow the smears to dry quickly, using a fan or blower at room temperature.

4. Fix the smears in absolute (100%) methanol; allow them to dry.

5. Place them, touching front to back, in a box without separating grooves.

6. Label the outside of the box with the species, date and “Giemsa control slides.”

7. Store at -70°C (or colder) if the purpose is to make quality control slides.

8. Just before use, remove the smear from the box and allow the condensation to evaporate; label the slide “+ malaria” and the present date.  The smear is now ready for staining since it was previously fixed.

Microscopic Examination

Examining thick smearsSince the erythrocytes (RBCs) have been lysed and the parasites are more concentrated, the thick smear is useful for screening for parasites and for detecting mixed infections.

1. First screen the entire smear at a low magnification (10× or 20× objective lens), to detect large parasites such as microfilaria.

2. Then examine the smear using the 100× oil immersion objective lens.  Select an area that is well-stained, free of stain precipitate, and well-populated with white blood cells (WBCs) (10-20 WBCs/field).

3. If you see parasites, make a tentative species determination on the thick smear and then examine the thin smear to determine the species present.  Most often, the thin smear is the appropriate sample for species identification.

4. Determination of "No Parasites Found" (NPF): For malaria diagnosis, WHO recommends that at least 100 fields, each containing approximately 20 WBCs, be screened before calling a thick smear negative.  Assuming an average WBC count of 8,000 per microliter of blood, this gives a threshold of sensitivity of 4 parasites per microliter of blood.  In nonimmune patients, symptomatic malaria can occur at lower parasite densities, and screening more fields (e.g., 200, 300, or even the whole smear) might be warranted, depending on the clinical context and the availability of laboratory personnel and time.  NCCLS standards recommend examination of at least 300 fields using the 100× oil immersion objective.

Examining thin smearsThin smears are useful for species identification of parasites already detected on thick smears, screening for parasites if adequate thick smears are not available, and a rapid screen while the thick smear is still drying.

1. Screen at low magnification (10× or 20× objective lens) if this has not been done on the thick smears.

2. Carefully examine the smear using the 100× oil immersion objective lens.  NCCLS standards recommend examination of at least 300 fields using the 100× oil immersion objective.

Quantifying parasitesIn some cases (especially malaria) quantification of parasites yields clinically useful information.  If this information is needed by the physician, malaria parasites can be quantified against blood elements such as RBCs or WBCs.

To quantify malaria parasites against RBCs, count the parasitized RBCs among 500-2,000 RBCs on the thin smear and express the results as % parasitemia.

% parasitemia = (parasitized RBCs/total RBCs) × 100

If the parasitemia is high (e.g., > 10%) examine 500 RBCs; if it is low (e.g., <1%) examine 2,000 RBCs (or more); count asexual blood stage parasites and gametocytes separately. Only the former are clinically important and gametocytes of P. falciparum can persist after elimination of asexual stages by drug treatment.

To quantify malaria parasites against WBCs: on the thick smear, tally the parasites against WBCs, until you have counted 500 parasites or 1,000 WBCs, whichever comes first; express the results as parasites per microliter of blood, using the WBC count if known, or otherwise assuming 8,000 WBCs per microliter blood.

Parasites/microliter blood=(parasites/WBCs) × WBC count per microliter<or 8,000>

Results in % parasitized RBCs and parasites per microliter blood can be interconverted if the WBC and RBC counts are known, or otherwise (less desirably) by assuming 8,000 WBCs and 4,000,000 RBCs per microliter blood.

Detection of blood parasites using fluorescent dyesFluorescent dyes that stain nucleic acids have been used in the detection of blood parasites.  In the Kawamoto technique, blood smears on a slide are stained with acridine orange and examined with either a fluorescence microscope or a light microscope adapted with an interference filter system.  This results in a differential staining of nuclear DNA in green and of cytoplasmic RNA in red, which allows recognition of the parasites.  The method has been applied to malaria parasites (and to a lesser extent, African trypanosomes).

In the Quantitative Buffy Coat (QBC®; Becton Dickinson) method, blood samples are collected in a special tube containing acridine orange, an anticoagulant, and a float, and then are centrifuged in a microhematocrit centrifuge.  After centrifugation, the tubes are examined using a fluorescence microscope with a stage adapter, or a light microscope with a customized fluorescence attachment.  Malaria parasites concentrate below the granulocyte layer in the tube.  The QBC method is reported to have a good sensitivity for detection of malaria parasites, and has also been applied (albeit to a lesser extent) to other parasites such as trypanosomes, microfilaria and Babesia spp.

Molecular Diagnosis

Specimen CollectionMicroscopic examination of stained blood smears is considered the gold standard for diagnosis of malaria and babesiosis.  When species determination cannot be made by microscopic examination, analysis by polymerase chain reaction (PCR) is helpful.  Collect a 1-5 ml blood sample in Vacutainer® EDTA tubes prior to anti-parasitic therapy and ship at 4°C to a reference laboratory.  Alternatively, blood can be collected on filter papers (e.g., products available through Whatman® http://www.whatman.com).  Punching the spots may increase the risk of cross-contamination among specimens.  Spot the paper directly from whole blood or finger stick.  Follow all shipment guidelines and requirements.  Blood smears should always accompany the EDTA blood sample.  The blood smears will be examined first; PCR will be performed only if species determination cannot be made from the blood smears.

The following procedure describes how a specimen will be accepted for PCR analysis at CDC.  Prior arrangements should be made to determine the appropriateness of PCR as an

adjunct for the diagnosis of malaria and babesiosis.  At this time, PCR analysis takes approximately one week for completion.

DNA has to be extracted from the blood specimens for PCR detection.  Click to view the DNA extraction protocols recommended for molecular diagnosis of malaria and babesiosis.

Species-specific PCR for Diagnosis of MalariaPlasmodium sp. genomic DNA is extracted from 200 µl whole blood using the QIAamp Blood Kit (Cat. No. 29106; Qiagen Inc., Chatsworth, CA) or a similar product that can yield the comparable concentration of genomic DNA from the same volume of blood.  Detection and speciation of Plasmodium is done with a two step nested PCR using the primers of Snounou et al 1993.  In the first step (PCR1), 1 µl of extracted DNA is amplified using genus specific primers; in the second step (PCR2), 1 µl of PCR1 amplification product is further amplified using primers specific for each Plasmodium species.  Ten microliters of each PCR2 amplified DNA product is electrophoretically resolved on a 2% agarose gel, stained for 15 min with ethidium bromide and visualized by UV illumination for analysis of results.

Species-specific PCR for Diagnosis of BabesiaBabesia sp. genomic DNA is extracted in the same way as Plasmodium sp. DNA (see above).  Detection of Babesia microti is done with a two step nested PCR using the primers of Persing et al.  In the first step (PCR1), 1 µl of extracted DNA was amplified using B. microti specific primers, Bab1 and Bab4; in the second step (PCR2), 1 µl of PCR1 amplification product was further amplified using internal primers, Bab2 and Bab3.  Ten microliters of PCR2 amplified DNA product was electrophoretically resolved on a 2% agarose gel, stained for 15 minutes with ethidium bromide and visualized by UV illumination for analysis of results.

References:

1. Persing D, Mathiesen D, Marshall WF, Telford SR, Spielman A, Thomford JW, Conrad PA. Detection of Babesia microti by Polymerase Chain Reaction. J Clin Microbiol 1992;30:2097-2103.

2. Snounou G, Viriyakosol S, Zhu XP, Jarra W, Pinheiro L, do Rosario VE, et al. High sensitivity detection of human malaria parasites by the use of nested polymerase chain reaction. Mol Biochem Parastiol 1993;61:315-320.

Detection of Parasite Antigens

Rapid diagnostic tests for malaria have been developed that employ immunochromatographic methods based on the detection of malarial antigens present in peripheral blood.  Most tests use monoclonal antibodies and detect particular malarial antigens in blood specimens.  Tests have been developed that detect the histidine-rich protein II (HRP-II), parasite lactate dehydrogenase (pLDH), or both HRP-II and LDH.  These tests generate results within 15 minutes and do not require skilled microscopists.

Commercially available kits for HRP-II detect P. falciparum HRP-II only and therefore diagnose only P. falciparum malaria.  The HRP-II antigen is synthesized and released by trophozoite and immature gametocyte stages and persists in peripheral blood.  Therefore, HRP-II tests can remain positive for up to 2 weeks following chemotherapy and parasite clearance, as confirmed by microscopy.  These tests have low sensitivities for detecting infections with low level parasitemias (<100 parasites/µl) and mature gametocytes.  In contrast, trained microscopists can diagnose infections with parasitemias as low as 5-10 parasites/µl.  The reported specificities of these tests are high (>90%).  Early tests reported false positives due to cross-reactions with rheumatoid factor, but these issues have reportedly been addressed and corrected.

Parasite lactate dehydrogenase (pLDH) is produced by asexual and sexual stages (gametocytes) of malaria parasites.  Test kits that are currently available detect pLDH from all four species of Plasmodium.  They can distinguish P. falciparum from the non-falciparum species, but cannot distinguish between P. malariae, P. ovale, and P. vivax.  Tests that detect pLDH do not generate persistent positive results following chemotherapy, like the HRP-II test.

Although these assays have advantages over microscopic examination of Giemsa stained blood smears, because of their limited usefulness in detecting low-level parasitemias (< 100 parasites/µl) their use is not indicated for diagnosing infection in most clinical presentations of malaria in the US.  These tests are not FDA approved for diagnosis of malaria in the United States and are available only for research studies from vendors outside the United States.

For more information on rapid diagnostic tests for malaria, visit the WHO site.

Organism  Kit name Manufacturer - distributora Type of Testb

Plasmodium Malaria-Ag Cellabs EIA

  OptiMal Flow Rapid (LDH)

  MAKROmed malaria test MAKROmed Manufacturing, LTD

Rapid (HRP2)

  Paracheck Pf Orchid Rapid (HRP2)

  Visitect Malaria Pf Omega Diagnostics LTD Rapid (HRP2)

Wuchereria bancrofti

ICT Filariasis Binax Rapid

  Filariasis Ag-CELISA Cellabs EIA

a Cellabs, P O Box 421, Brookvale, NSW 2100, AustraliaFlow, Inc., 6127 SW Corbett, Portland, OR 97201MAKROmed Manufacturing, LTD, P O Box 28928, Kensington 2101, South AfricaOrchid, 4390 US Route One North, Princeton, NJ 08540Omega Diagnostics, LTD, Omega Hense, Carsebridge Court, Whins Road, Alloa, FK10 3LQ, Scotland, United KingdomBinax, Inc., 217 Read Street, Portland, ME 04103

bEIA = enzyme immunoassay; Rapid = rapid immunochromatographic test

Special Tests: MQ Testing

Mefloquine is recommended by CDC as a prophylactic against malaria.  When individuals who are presumably on mefloquine prophylaxis exhibit signs of malaria, blood samples are collected and analyzed for the presence of the drug.  The drug is extracted from the blood and the concentration is determined by high-performance liquid chromatographic methods.  Determining the level of mefloquine in the blood helps assess if the individual was adherent with his/her medication.  This procedure is also useful to determine treatment failure due to a mefloquine resistant form of malaria.