Module 4-Speciemn collection.pptxghjfffghh

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1. Module 4 Specimen Collection and Preparation of Blood films; preparation of stains; and blood film staining 1

2. Specimen Collection and Preparation of Blood films 2

3. Learning Objectives At the end of this module , the learner should be able to: Describe the types of blood films used in routine malaria microscopy Collect capillary blood for the preparation of thick and thin blood films. Make a good thick and thin films of blood taken from people with suspected malaria in a single slide 3

4. Learning … Dry the blood films appropriately Correct labeling of blood films Identify good quality of thick and thin blood films Identify the causes of common mistakes in the preparation of both thick and thin blood films 4

5. Content Outline Materials required Types of blood films Blood sample collection Blood film preparation Qualities of good thick and thin films Common mistakes in making blood films 5

6. Materials Required  6

7. Types of blood films 1. Thick films Consist of a thick layer of lysed RBCs . Used to detect and quantify malaria parasite More sensitive than thin blood film The blood elements are more concentrated (~ 30x) than in an equal area of a thin film.   Best for detection of low levels of parasitemia . Used for rapid detection of the parasite Challenge Does not permit an optimal review of parasite morphology.  7

8. 2. Thin films Consist of blood spread in a single layer. In fixed thin film the parasites are found intact inside the RBC. Assist in the identification of the Plasmodium species Provides greater specificity Challenges low-density infections can be missed Requires more time to read. 8

9. Preparation of Slide Use new slides with frosted end to make patient identification easy. Slides must be free of scratches, grease, dust, and acid or alkali contamination. Store slides in a closed dry container. 9

10. Blood Sample Collection : Capillary Blood The ideal sample as the density of trophozoites or schizonts is greater in blood from capillary-rich area. Obtained by pricking a fingertip or big toe For adults: the lateral side of the 3 rd or 4 th finger is best. F or infants : the big toe is preferred. 10

11. Procedure for collecting finger prick blood Wear gloves during the procedure Keep complete record of patient identification in a logbook . Prepare pre-cleaned slides in a clean surface. Handle slides by the edges to avoid fingerprints . 11

12. Procedure… 4. S elect the 3 rd or 4 th finger and c lean the area with a alcohol swab and Allow to dry. 5. Puncture using a sterile lancet. 6. Apply gentle pressure to allow blood drop to ooze out. Wipe away the first drop of blood 12

13. Transferring of blood to slide: For thin film Applay gentle pressure to the finger and collect a single small drop( 2µl ) of blood in the centre of the slide. Release the pressure immediately to allow recirculation of blood. 13

14. For thick film Apply gentle pressure again to transfer more blood and collect one bigger (6µl )drop on the slide, about 1 cm from the drop intended for the thin film or 1 cm from the end of the slide. 14

15. Thick and thin blood films are made as follows: Thin film Take clean slide with smooth edges and use it as a spreader. Hold the end of a spreader slide against the surface of the first slide at an angle of 30-45 degree. Make thin film by bringing in contact with edge of the spreader with the blood drop on the middle of the original slide. Wait until the blood spreads along the entire width of the spreader Push it forward rapidly and smoothly until the film is formed. 15

16. Thin film… Make sure that the spreader is in even contact with the surface of the slide all the time the blood is being spread. If the blood is anemic, hold the spreader at a greater angle to make the blood film slightly thicker. The spreader slide may now be used for the next patien t. 16

17. Thick film Using the corner of the spreader, quickly join the drops of blood and spread in a circular/rectangular manner with 3 to 6 movements to make a round Do not make the film too thick or it will fall off the slide. 17

18. Preparation of thin and thick film 18

19. Qualities of Good Thin Blood film   Uniformly spread over the slide Thin enough so that It is tongue shaped Consists of a single layer of RBCs with feathered end                           19

20. Qualities of Good Thick Blood film   It should be 10 mm away from the edge of the slide Rectangular or round in shape with a diameter of about 10 mm Its thickness contains 10 layers of RBCs At least 10-12 WBCs should be visible / 100x field.   20

21. Common Mistakes in Making Blood Films 1. Too much blood On thick film After staining the background will be too blue . Result too many WBCs per thick film field, and these could obscure malaria parasites . On thin film RBCs will be on top of one another impossible to examine them properly after fixation . 21

22. Common mistakes 2.Too little blood Not able to examine enough blood in the standard examination . False negative result is likely. 22

23. Common mistakes… 3.Blood films spread on a greasy slide The blood films will spread unevenly . Some of the thick film will probably come off the slide during the staining 23

24. Common mistakes… 4. Edge of spreader slide chipped the thin film spreads unevenly, is streaky and has many “tails”. The spreading of the thick film may also be affected. 24

25. Common Mistakes… 5.Badly positioned blood films B lood films should be correctly sited on the slide. If not, it may be difficult to examine the thick film. 25

26. Common mistakes… 6. Thin film too big, thick film in the wrong place The thick film will be out of place and may be so near the edge of the slide that it can’t be seen through the microscope. During staining or drying, portions of the thick film will scraped off by the edges of the staining trough or drying rack. Very difficult to position the thick film on the microscope stage. 26

27. Common technical mistakes in collection and preparation of blood film 27 Mistake Effect Pricking of not well dried finger The parasites and host cells may be fixed by the alcoholic detergent solution Use of dirty slides The blood will not be spread evenly. Generates artifacts Delay in making blood film once you transfer the drops of blood to the slide The blood film will not be spread evenly due to the beginning of the coagulation process Too much blood for thin films Erythrocytes are laid on multiple layers. Observation is impossible

28. Common mistakes… … 28 Mistakes Effects Too little blood used for thin films Parasites may be virtually absent if parasitaemia is low Not labeling slides immediately Confusion may arise leading to unidentified positive or negative slides Slides wrapped together before thick films properly dried The slides stick to one another

29. Common mistakes… Mistake Effects Inappropriate washing of stain from the slide Stain deposits may render the observation d ifficult Long time elapses before staining of thick films Causes Auto fixation which result in difficulty of hemolysis during staining Exposure of thick films to excessive heat Autofixation occurs and haemolysis is Impossible 29

30. Preparation of Stains and Buffered water 30

31. Learning Objectives At the end of this module the trainees should be able to Prepare Giemsa staining solution required for malaria microscopy. Describe the importance of using quality chemicals for reagent preparation. Describe how to safely store Giemsa stain. 31

32. Content Outline Principles of Romanowsky stains Preparation of Giemsa stain Storage of Giemsa stain. 32

33. Romanowsky stains Polychrome stain Used to stain blood cells and organisms. Comprise two staining components : Eosin Y and Azure B, obtained by oxidation of methylene blue. Eosin stains chromatin dot and stippling shades of red or pink. Methylene blue , stains parasite cytoplasm blue. 33

34. Romanowsky stains: Examples Giemsa stain Field’s stain Leishman’s stain Wright’s stain… 34

35. Giemsa stain Gemsa stain is an alcohol-based Romanowsky stain. Is a mixture of eosin, which stains parasite chromatin and stippling shades of red or pink, and methylene blue, which stains parasite cytoplasm blue. White-cell nuclei stain blue to almost black, depending on the type of white cell. The recommended stain for identification of malaria parasite. 35

36. Giemsa stain Reagent and materials required Giemsa Powder Absolute methanol Glycerol Glass beads 50 pcs Brown bottle  Analytical balance 36

37. Preparation of Giemsa Stain To make about 500 ml: Giemsa Powder 3.8g Absolute methanol 250ml Glycerol 250ml 37

38. Procedure… 1. Weigh the Giemsa and transfer to a dry brown bottle of 500 ml capacity which contains a few glass beads. 2. Using a cylinder measure the methanol and add to the stain. Mix well. 3. Using the same cylinder measure, the glycerol and add to the stain. Mix well. 38

39. Procedure… 4. Tightly stopper the bottle. 5. Shake the bottle for 2-3 minutes. 6. Add measured glycerol and repeat the shaking. 7. Continue shaking for 2-3 minutes at 30 minutes intervals at least 6 times. 8. Keep the bottle for 2-3 days; shaking it 3-4 times each day. 9. Keep small amount of the stock in a small bottle. 10. Label with the name of the reagent and date of preparation and mark “inflammable”. Store at room temperature in the dark. 39

40. What you should do after preparation of Giemsa stock solution Keep the stopper screwed tightly. Must be diluted with distilled water (Buffered water) with pH of 7.2. Should be tested for proper staining reaction. Must be Protected from Moisture and direct sunlight. Stored in a cool dry place in a dark bottle. Measure a small quantity of stain into a smaller bottle for one or two days’ use. 40

41. What you should not do after the reagent preparation Never add water to the stock Giemsa solution. Do not shake the bottle of stain before use: you will re-suspend very small, un dissolved crystals of stain. Never return unused stain to the stock bottle. 41

42. Quality controls of Giemsa stain Done to ensure the staining quality and performance of Giemsa stain. Use known positive and negative films with each new batch of working Giemsa stain. Blood films can be prepared using EDTA anticoagulated blood from a patient’s. Note : An ideal blood sample has at least one parasite in every 2–3 fields. 42

43. Quality control … Allow the blood films to dry quickly. Fix the films using absolute methanol. Place them, touching back to back, in a box with separating grooves. Label the outside of the box with the species, date and “Giemsa control slides”. 43

44. Quality control … The slides can be stored at room temperature for a minimum of 1 week but will last longer if stored at -20 C or below –70 °C. Just before use, remove the slide from the box and allow the condensation to evaporate. Label the slide with the date and “Positive control”. The blood film can then be stained and examined. 44

45. Buffer solutions for malaria staining A phosphate buffer solution, balanced to pH 7.2 is important to prepare Giemsa working solution. Check the pH using narrow range pH papers or pH meter and store at room temperature. The buffer is stable for several months. 45

46. Materials, Reagents and Equipment Required Materials 1. Wooden spatula 2. Beaker 250 ml capacity 3. Conical flask 1000ml capacity 4. Filter papers 5. Brown bottle 6. Measuring cylinder 100ml capacity 7. Filter papers 8. Labels   Reagents 1. KH 2 PO4 0.7g 2. NA 2 HPO4 1.0g Distilled or de-ionized water 1000ml Equipment: Analytical Balance 46

47. Preparation of buffered water : Procedure 1.Measure 0.7g KH 2 PO 4 and 1g of Na 2 HPO 4 . 2. Add 0.7g KH 2 PO 4 to the beaker. 3. Add 150ml of water. 4. Stir with the spatula until the salt is dissolved. 5. Add 1.0g of Na 2 HPO 4 to the beaker. 6. Stir with the spatula. 47

48. Procedure… 48 7. When dissolved add the fluid from the beaker to the conical flask. 8. Fill the fluid in the conical flask with water until it is made up to 1ml. 9. Add small quantities of 2% Na 2 HPO 4 if the pH is below 7.2 (acidic) or 2% KH 2 PO 4 if the pH is above 7.2 (basic). Quality Control Check pH of buffered water, and add appropriate correcting fluid.

49. Buffer tablet Buffer tablets that produce a solution of pH 7.2 when dissolved are ready available from laboratory suppliers but are rather expensive. 49

50. Preparation of 2% correcting fluids Used to maintain the Buffer at pH 7.2 . Materials 1. Wooden spatula 2. Beaker 250 ml capacity 3. Conical flask 1000ml capacity Filter papers 5. Brown bottles 6. Measuring cylinder 100ml capacity 7. Filter papers Labels   Reagents 1 . KH 2 PO4, 2g 2. NA 2 HPO4, 2g 3. Distilled or de-ionized water, 1000ml Equipment Analytical balance 50

51. Preparation of 2% correcting fluid: Procedure Weigh 2g NA 2 HPO4 and add it to 100ml of water in the beaker. Stir with the wooden spatula. Pour the solution into one of the glass bottles. Label the bottle “2% disodium hydrogen phosphate” Repeat steps 1 and 2 above weighing out 2g of KH 2 PO4 Pour the solution into the second glass bottle and label it correctly. 51

52. Quality Control of Buffered water Prepare a buffer reagent carefully. Check the PH regularly. Store buffered reagents at 2-8 C. Avoid leaving the reagents in sun light (encourages the growth of algae). Check for contamination at regular intervals . 52

53. Fixation and Staining of Blood Film 53

54. Learning Objectives At the end of the unit, the learner should be able to: Demonstrate appropriate blood film fixation and staining techniques   54

55. Content Outline Blood film fixation Staining blood film 55

56. Fixation of Blood film: Materials Absolute Methanol Dropper bottle Absorbent cotton wool/Gauze Drying rack 56

57. Fixation Used to preserve cellular and parasitic morphology. Done When the blood films are completely dried. Absolute methanol is recommended fixative solution Note: Auto-fixation may also occur spontaneously with time if thin films not fixed immediately after dried (7 to 15 days, varying with humidity and temperature of the atmosphere). 57

58. Fixation methods Done for 10-20 seconds using one of these three methods: Dropping absolute methanol while holding the slide with the thin portion down Dipping in a jar containing absolute methanol. 3. D abbing it with cotton wool dampened with methanol Note: Avoid methanol, or its fumes encountering the thick film. 58

59. Staining blood films There are two methods of staining with Giemsa stain: The rapid (10%) method The slow (3%) method. The rapid method is used in laboratories where a quick diagnosis is an essential part of patient care. The slow method is used for staining larger numbers of slides, such as those collected during cross-sectional or epidemiological surveys and field research. 59

60. Staining Blood Film 1. The rapid (10%)methods Filter the stock Giemsa solution Prepare fresh a 10 % (1:10 ratio) working Giemsa stain solution Add 10ml of Giemsa stock solution to 90 ml of buffered distilled water or 5ml to 45 ml of buffered distilled water Pour the diluted solution into a staining jar Immerse completely dried slides in a jar. Make sure that the stain cover the entire surface of films on the slide. 60

61. Staining Blood film Leave the slides in the stain for minimum of 10 minutes. Wash the stain in a jar containing water.  Remove the slides and clean the back of each slide with dry gauze. 61

62. Staining … Place the slides with the film side down wards in a drying rack to drain and dry at room temperature. Make sure that the thick film does not touch the edge of the rack. 62

63. Staining of blood films The slow (3%) method This method is less appropriate when a quick result is needed but is excellent for staining large numbers (20 or more) of slides. It is ideal for staining blood films from surveys or research work or batches of slides for teaching. It performs best when slides have dried overnight. The method is economical because much less stain is used (3% rather than 10%). 63

64. Quality Control A blood film, thick and/or thin, is run once each month or with each staining procedure The blood film does not necessarily required to be positive for blood parasites. Good color differentiation of red and white cells is an indication of a good quality stain. 64

65. Field’s stain Preparation and staining Useful for rapid detection of malaria parasites particularly for thick films. Schuffener’s dots are not always stained with field’s stain. Made up of Field’s stain A and Field’s stain B. 65

66. Preparation of Field’s A stock solution Can be prepared in two different ways: I. From prepared powder Add 0.5 g of fields satin A powder to 600ml of hot (~ 60 C ) distilled water. Mix until it dissolves. Filter when cool . 66

67. II. From original stains and chemicals Dissolve 10.0 g of anhydrous Na 2 HPO 4 and 12.5 g of KH 2 PO 4 in 1000 ml of distilled water. Pour half of this solution into 1 liter bottle containing a glass beads. Add 1.6g of methylene blue and 1.0 g of azur 1 and mix well. Add the remainder of the phosphate solution Mix well and filter 67

68. Preparation of Field’s stain B- stock solution Prepared in two different ways, From prepared powder Add 4.8 g Field’s stain B powder to 600ml of hot (~60 C) distilled water. Mix until dissolved Filter when cool.   68

69. From original stains and chemicals Dissolve 10.0 g of anhydrous Na 2 HPO 4 and 12.5 g of KH 2 PO 4 in 1000 ml of distilled water. Add 2 g of eosin (yellow, water soluble). Mix until dissolved. Filter. 69

70. Field’s staining Procedure for staining thin films Fix film in methanol for 1 minute Wash off methanol with water. Using a pipette, cover the film with diluted Field’s stain B (1 part by volume of stock solution plus 4 volumes of distilled water buffered at pH 7.2) 70

71. Procedure… Immediately add an equal volume of Field’s stain A solution and mix well by tilting the slide. Wash off stain with clean water Place slide up right in a draining rack to air dry. 71

72. 72 Thank you ! !