clinical parasitology-hemoparasites.pptx

1. Collection, Preservation,
Artefacts and Examination Of
Blood Parasite Of Human
•ABUZAR KHAN (30)
•HAMMAD AHMAD (29)
•SAMEERA YAQOOB (09)
• SAIF UR REHMAN (27)
•QUDSIA ASIF (05)

2. Collection Of
Hemo Parasites

3. Hemo Parasites:
• Hemoparasite is a group of organisms that live and multiply
within the bloodstream of their hosts.
• They can infect various vertebrate animals, including
humans, and are typically transmitted through the bite of
infected arthropod vectors such as mosquitoes, ticks, and
flies.
• These parasites can cause a range of diseases, leading to
significant health problems if left untreated .

4. • Here is a detailed point note on the collection of blood
samples for parasitic examination:
1. Patient Preparation
2. Gather Necessary Supplies
3. Informed Consent
4. Selecting the Sample Type
5. Skin Disinfection
6. Tourniquet Application
7. Tube Labeling

5. • Collecting blood samples for parasitic examination is a crucial
step in diagnosing various parasitic infections. Proper
collection and handling of blood samples are essential to
ensure accurate and reliable results.
Type of Sample:
• Venous blood obtained by venipuncture
• Capillary blood obtained by fingerstick

6. Venous blood obtained by venipuncture:
Material Required:
• Syringe
• Needle
• Vacuum Tubes
• Alcohol Swab or Antiseptic Wipe
• Sterile Gauze or Cotton Balls
• Tourniquet
• Adhesive Bandage or Tape
• Disposable Gloves
• Sharps Container
• Patient Identification Labels

7. Procedure:
• Gather necessary materials needed for venipuncture.
• Ensure comfort and informed consent. Position the patient and provide easy
access to the chosen vein.
• Use a tourniquet to temporarily stop blood flow, making the veins more visible.
• Select a suitable vein, typically in the arm. Clean the skin over the chosen site
with an alcohol swab.
• Wear disposable gloves for hygiene and safety.
• Hold the syringe with the attached needle and insert it into the vein at a slight
angle.
• Withdraw blood into the syringe or directly into vacuum tubes if using a syringe
and needle combination.
• Remove the needle gently once blood collection is complete.
• Apply sterile gauze or cotton ball over the puncture site. Secure with adhesive
bandage or tape.
• Safely dispose of the needle and syringe in a sharps container

9. Capillary blood obtained by fingerstick:
Material required :
• Alcohol Swab or Cotton Ball with Alcohol
• Sterile Lancet or Lancet Device
• Gauze or Cotton Ball:
• Microcollection Tube or Capillary Tube
• Disposable Gloves
• Labeling Materials

10. Procedure:
• Wash hands and wear disposable gloves.
• Choose the ring or middle finger.
• Clean with an alcohol swab.
• Prepare the lancet Remove the protective cover or follow device
instructions.
• Puncture the side of the fingertip.
• Gently massage to encourage blood droplet formation.
• Position the collection tube or capillary tube.
• Allow blood to be drawn into the tube through capillary action.
• Apply gentle pressure to the puncture site.
• Safely dispose of the lancet in a sharps container.

12. PRESERVATION OF
BLOOD

13. OVERVIEW
DEFINITION:
• Blood preservation is a critical aspect of modern healthcare, involving the
application of advanced techniques to maintain the viability of blood and
its components.
• When blood is collected from a donor, it must be processed to remove red
blood cells and plasma (the liquid part of blood), which are used for
transfusions.
IMPORTANCE:
• Prolonging the shelf life of blood ensures a stable supply for transfusions,
surgeries, and emergency medical interventions.

14. FUNDAMENTAL PRINCIPES OF
BLOOD PRESERVATION
Oxygenation Control:
• Importance of minimizing oxygen exposure to prevent oxidative
damage.
• Techniques: Inert gas flushing and vacuum sealing.
Temperature Control:
• Refrigeration and cryopreservation.
• Impact on cellular metabolism and integrity.

15. TYPES OF BLOOD PRESERVATION
TECHNIQUES
Refrigeration:
• Standard method for short-term storage.
• Temperature range and its impact on red
blood cell preservation.
Cryopreservation:
• Freezing at ultra-low temperatures. (-
80°C)
• Challenges and benefits in preserving
platelets and other cellular components.
Additive Solutions:
• Use of specialized solutions to enhance
preservation and extend shelf life.
• Balanced electrolyte solutions and
plasma extenders.

16. COMMONLY USED PRESERVATIVES
HEPARIN:
• Inhibits the formation of
thrombin from prothrombin.
• Quantity: 2 mg/10 mL of blood
EDTA:
• Has chelating action on calcium
ions and prevents coagulation.
• Quantity: 20 mg/10 mL

17. CALCIUM OXALATE:
• Acts by precipitating calcium ion
• Quantity: 30 mg/100 mL
SODIUM CITRATE:
• Converts calcium into non
ionized form
• Quantity: 30 mg/ 10 mL

18. BLOOD BANKS AND
PRESERVATION PROTOCOLS
BLOOD BANK OPERATIONS:
• Collection, testing, processing, and storage.
• Quality control measures to ensure viability.
TECHNOLOGY INFRASTRUCTURE:
• Refrigerators, freezers, and cryogenic storage units.
• Monitoring systems for temperature and environmental conditions.

19. Preparing Blood Smears
• If you use venous blood, blood smears should be prepared as soon
as possible after collection (delay can result in changes in parasite
morphology and staining characteristics).

20. The Ideal Blood Smear
1. An ideal slide is neither too thin nor too thick.
2. Free from holes caused by grease or dirt on the slide.
3. Good smear is tonged shape with a smooth tail.
4. It should end about two-thirds of the way down the
slide.
5. Thick at one end, thinning out to a smooth rounded
feather edge and there should be no streaks at the

21. The Ideal Blood Smear

22. Thick smears
• In thick smear microscopy, blood smears consist of a thick layer of red
blood cells. The blood elements (including parasites, if any) are more
concentrated and allow for more efficient detection of parasites and
increased sensitivity of the test, compared to thin smear microscopy.
Thick smears are used to determine parasite prevalence in malaria-
related household surveys.
• It is more sensitive and allows for the detection of a higher number of
parasites.

23. Thick Smear
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 (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.Allow the smears to dry thoroughly. Insufficiently dried smears or smears that are too
thick can detach from the slides during staining.
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 hemolyze the RBCs.

24. Thick Smear

25. Thin Smear
• Thin 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.
• The thin smear allows for better observation of the morphology of
the parasites, aiding in species identification and assessment of
parasite life cycle stages.

26. Thin Smear
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.
5.Allow the thin smears to dry.
6.Fix the smears by dipping them in absolute methanol.

28. THICK VS THIN SMEAR

29. THICK VS THIN SMEAR

30. Blood smears – common errors

31. Giemsa Staining
• Giemsa stain is a gold standard staining technique that is used for
both thin and thick smears to examine blood for malaria parasites, a
routine check-up for other blood parasites and to morphologically
differentiate the nuclear and cytoplasm of Erythrocytes, leucocytes
and Platelets and parasites.

32. •Rapid (10% working solution) method
1.Commonest method for staining 1-15 slides at a time.
2.Used in outpatient clinics and busy laboratories
3.Efficient method but costly (as more stain is consumed)
•Slow (3% working solution) method
1.Used for staining a larger number of slides (>20)
2.Ideal for staining blood films collected during cross-sectional or
epidemiological surveys, field research, or for preparing batches of slides for
teaching
3.Time-consuming method, so less appropriate when a quick result is needed
4.Less expensive compared to the rapid method as it requires much less stain.

33. Preparation of Giemsa Working Solution
• Prepare either 10% or 3% Giemsa working solution, depending on your
need. About 3 mL of stain is required for each slide with a blood film.
1.Place 90 mL of prepared buffered water, pH 7.2, into a clean beaker or tube.
2.Filter the Giemsa stock solution through paper Whatman #1 and transfer it
to a 25 to 50 mL container.
3.Add 10 mL of Giemsa stock solution using a clean, dry pipette. Do not take
the aliquot from the large bottle containing the Giemsa stock solution to
avoid contaminating it.
4.Prepare the Giemsa working solution just before staining the blood film(s),
and use it within 15 minutes of preparation. Discard any unused stain.
• To prepare 3% Giemsa working solution, follow the procedure mentioned
above, but mix 97 mL of buffered water with 3 mL of Giemsa stock solution.

34. Staining of the Slides
• For Thin blood smears
1.Fix air-dried film in absolute methanol by dipping the film briefly
(two dips) in a Coplin jar containing absolute methanol.
2.Remove and let air dry.
3.Stain with a working solution of Giemsa stain
4.Wash by briefly dipping the slide in and out of a Coplin jar of
buffered water (one or two dips).
Note: Excessive washing will decolorize the film.
5.Let air dry in a vertical position. Observe under the microscope first
at 40X and then using an oil immersion lens

35. Staining of the Slides
• For Thick blood smears
1.Allow the film to air dry thoroughly for several hours or
overnight.
2.DO NOT FIX.
3.Stain with diluted Giemsa stain
4.Wash by placing the film in buffered water for 3 to 5 min.
5.Let air dry in a vertical position, observe under the microscope
at 40X, and then use an oil immersion lens

36. Staining of the Slides

37. CELL COMPONENTS- COLOR
OBSERVED POST STAINING
• Red Blood Cells: Mauve-pink
• Neutrophils: Reddish purple
• Eosinophils: Purple nuclei & red to orange granules
• Basophils: Purple nuclei & blue coarse granules
• Lymphocytes: Dark blue nucleus
• Platelets: Violet color granules
• Nuclei of host cells: Dark purple
• Nuclei of WBCs: Dark purple
• The cytoplasm of host cells: Pale blue
• The cytoplasm of white cells: Pale blue or grey blue
• Malaria parasite: Red or pink nucleus and blue cytoplasm

38. CELL COMPONENTS- COLOR
OBSERVED POST STAINING

39. EXAMINATION OF
BLOOD SMEAR

40. Different Blood Parasites
RBC Intracellular
parasites
WBC Intracellular
parasites
Extracellular parasites
Plasmodium Leishmania Trypanosoma
Babesia Theileria Filarial worm
Theileria Schistosoma
Anaplasma

41. DIRECT MOUNT METHOD
• We use this method to see the live motility of haemoparasite
(blood parasite).
• In this method ,we put a blood drop on slide.
• After putting cover slip on blood drop on slide, we observe slide
under microscope.

42. Trypanosoma brucei
Filarial worm( W .bancrofti )

43. Microscopic Character of Good Blood
Smear
• Must consist of Head , Body and Tail.
• Tail consist of evenly distributed blood
cell.
• Minimum clumping of RBCs.
• Consist of evenly distributed white
blood cell.
• WBCs must be present in every field .
• Consist of minimum artefacts

44. Examination Of Smear
Since 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.
• First screen the entire smear at a low magnification (10× or 20×
objective lens), to detect large parasites such as microfilaria.
• 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).
• 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.

45. Types Of Blood cell
• Red Blood Cells (Erythrocytes)
• White Blood Cells (Leucocytes)
• Platelet(Thrombocytes)
Size Of RBCs Haemoglobin in RBCs
Normocytic (normal size) Normochromic (Normal Hb)
Microcytic (small size) Hypochromic (Low Hb)
Macrocytic (large size) Hyperchromic (High Hb)

46. Morphology Of RBCs

47. Normal White Blood Cell

48. Artefacts Of Blood Slide
• Bubbles
• Clumping of stain
• Unidentified object (fungus) confused for amastigotes
of Leishmania
• Nucleated red blood cells
• Howell – jolly bodies
• Basophilic stippling
• Pappenheimer bodies

49. Fungus stain with
Giemsa stain
Spore of fungus Howell – jolly bodies
Nucleated
RBCs
Basophilic
stippling

50. Identification of Parasites
• Plasmodium ……………………….. Ring form merozoites.
• Anaplasma ……………….............Dot form in RBCs.
• Babesia and Theileria ……………Pear shaped schizonts.
• Leishmania ………………………….. Amastigotes in monocytes.

52. Blood Film With Falciparum

53. Babesia sp. stained with Giemsa.

54. Amastigotes Of Leishmania
(A) Amastigote form [arrows shows inside a macrophage (Giemsa stain) (B)
Smear shows promastigote form (Giemsa stain).

55. Parasitemia
• 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.
• If the parasitemia is high (e.g., > 10%) examine 500 RBCs.
• If it is low (e.g., <1%) examine 2,000 RBCs (or more).
% parasitemia = (parasitized RBCs/total RBCs) × 100

56. • Hypochromic...+3 (40%). +2
(10%),+1 (5%).
• Neutrophils (3 cell seen in
this field)
• Burr cell 55%
• Howell Jowell bodies (0%)
• Schistocytes (0%)
• Basophilic stippling (0%)
• Tear drop(2%)
• Agglutination(0%)
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