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Artefacts in hemat part 2

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hematologyical procedure done in Lab and various types of errors

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Artefacts in hemat part 2

  1. 1. 1 Presented by Dr. Shrikant Sonune Guided by Dr Ashok Patil, Dr Shilpa Kandalgaonkar, Dr Suyog Tupsakhare, Dr Mahesh Gabhane. Dr. Gaurav Agarwal Investigations & Artifacts in Hematology
  2. 2. Differential count is the percent distribution of various white cells in the peripheral blood. It is determined from a blood smear stained with a polychromatic stain. After examination of the stained smear by using oil immersion objective. The number of each type of white cells is then expressed as a percentage of the total number of cells.
  3. 3. The stained blood smear also helps to study abnormal morphology of leukocytes and red cells. Study of blood smear helps in the diagnosis of various types of anemia, leukemia and detection of blood parasites.
  4. 4. Three major steps involved in differential count are- a) Preparation of blood smear. b) Staining of blood smear c) Microscopic examination of stained smear.
  5. 5. Requires 1. Slide 2. Spreader 1. Slide  Should be clean  Should be free from dust  Should be wipe immediately before use 2. Spreader  Should have smooth edge.  Should be narrower in breadth than the slide  Almost having 2/3 width
  6. 6. A small drop of blood is placed in the central line of a slide about 1-2 cm from one end. The spreader is placed at an of 450 to the slide then move back to make contact with the drop. Drop should be spread out quickly along the line of contact of the spreader with the slide.
  7. 7. The movement this occurs the film should be spread by a rapid, smooth, forward movement of spreader. The drop should be of such a size that the film is 3-4 cm in length.
  8. 8. After smear is prepared then it is fixed with alcohol After fixation time is over add distilled water. Then add stain over it for recommended time. Let it be dry Wash under running tap water. Keep it for drying.
  9. 9. Leishmens Stain 1. Eosin – acidic dye 2. Methylene- Blue. 3. Acetone – free & water free absolute methyl alcohol
  10. 10.  It should be tongue shaped  2/3 width of slide  2/3 length of the slide  Thick at one end, thinning out to a smooth rounded feather edge.  Should not touch any edge of the slide.  Should be margin free, except for point of application.  It should be continuous  A properly stained slide has a pink tint.
  11. 11.  Irregular film (A)  Too long (B)  Too short (C)  Irregular spread (B)  Improper shape (D)
  12. 12. Discontinuous smear  Improper cleaning of slide.  Improper cleaning of finger may incorporate dust on the slide. Thin film  Overzealous force  Less than 30º angulations  Blood drop too small
  13. 13. Thick film-  Very light forces.  More 30º angulations  Blood drop Large. Edge formation  Improper force  Spread of blood drop  Improper direction
  14. 14. Irregular film 1. Spreader slide pushed across the slide in a jerky manner. 2. Failure to keep the entire edge of the spreader slide against the slide while making the smear. 3. Failure to keep the spreader slide at a 30° angle with the slide.
  15. 15. Failure to keep the spreader slide at a 30° angle with the slide. Failure to push the spreader slide completely across the slide. Less than 30° too long film More than 30° too short film.
  16. 16.  Irregular spread with ridges and long tail: Edge of spreader dirty or chipped; dusty slide  Holes in film: Slide contaminated with fat or grease  Cellular degenerative changes: delay in fixing, inadequate fixing time or methanol contaminated with water
  17. 17. Faint staining / thinned smear Dark staining / thickened smear
  18. 18. Too faint  Staining time too short  Excessive washing after staining Stain deposit:  Stain solution left in uncovered jar or tray  Stain solution not filtered  Dirty slides
  19. 19. Excessive blue coloration…………… Causes 1. Excessive staining time 2. Buffer in stain is alkaline 3. Old blood smear 4. Blood smear is too thick
  20. 20. The stain must be free of water, which induces RBC artifacts. Water artifacts may be avoided by fixation of slides or cover slips in anhydrous methanol before staining.
  21. 21. First should see under low power & high power such that area for counting should be selected. Then counting should be done under 100X Using z technique.
  22. 22. • Film quality • Stain quality • Cell distribution • Select the area for counting of WBC
  23. 23. 1. Made one reference point (starting point ) counting is started. 2. Using z technique the counting is done.
  24. 24. Neutrophils (PMN,s) • Nucleus 3-5 lobes. • Diameter 10-14 µm • 50-70% WBC • Numbers rise with all manner of acute infections , especially bacterial. The normal feature seen are
  25. 25. Eosinophil • Bilobed nucleus • 1-5% of WBC • Diameter about 10-14 µm • Contains: eosinophilic pink colour granules The normal feature seen are
  26. 26. • Lymphocyte • Agranular -No specific granules • 20-40% of WBC • Diameter 8-10 µm • 2 types  Small  Large The normal feature seen are
  27. 27. MONOCYTE  Large nucleus that tends to be oval or kidney bean- shaped.  2-8%  Entering peripheral tissue to become tissue macrophage
  28. 28. BASOPHILS  Have numerous granules that stain darkly with basic dyes.  Less than 1% of circulation leukocyte population.  Smaller than neutrophils
  29. 29. NORMAL SMEAR THICK AREA, PLUS DRYING ARTIFACTS
  30. 30. NORMAL SMEAR TRUE ROULEAUX
  31. 31. NORMAL SMEAR DRYING ARTIFACTS
  32. 32. a. Cold agglutinin - RBCs will clump together. Warm the blood at 37° C for 5 minutes, and then remake the smear. c. Rouleaux - RBC’s will form into stacks resembling coins. There is nothing to correct this.
  33. 33. 1) Packed cell volume (PCV). 2) Mean corpuscular volume (MCV). 3) Mean corpuscular hemoglobin concentration (MCHC). 5) Erythrocyte sedimentation rate (ESR). 6) Reticulocyte count. 7) Absolute eosinophil count.
  34. 34. PCV Packed cell volume is the most accurate and simplest of all test in clinical hematology for detecting the presence of degree of anemia or polycythemia.
  35. 35. Instruments required for Venepuncture Wintrobes tube Pasteur pipette
  36. 36. 5ml of blood. Mix with anticoagulant. Keep it for the centrifugation at 3000 rpm for 30 min. Record the height of PCV.
  37. 37. Hemolyzed cells parts, other unviable cells also incorporated into PCV.  Actually measures RBC concentration & not RBC mass Trapping of plasma in RBC column.
  38. 38. Faulty readings due to  Inappropriate concentration of anticoagulants.  Poor mixing of samples.  Insufficient centrifugation.
  39. 39. • Hematocrits calculated by automated instruments depend on correct red cell counts and red cell volumes to arrive at an accurate hematocrit. • Hence, anything affecting the red cell count or volume measurement will affect the hematocrit. • This method is not as sensitive to the ratio of blood to EDTA as the centrifuged hematocrit
  40. 40. The mean corpuscular volume is the mean value of single red cell expressed in cu micrometers To calculate mean corpuscular volume two basic values are required 1. Red cell count in million/ cubic mm 2. Packed cell volume in 100ml blood.
  41. 41. Mean corpuscular volume =PCV(packed cell volume) X 10 RBC(106 / mm3 )
  42. 42. Average hemoglobin content (weight of Hb) in a single red blood cell expressed in picogram To calculate the basic values required are 1. RBC count in million/ cb mm. 2. Hb in g percent.
  43. 43. = Hb in gm% X10 RBC count in million/ mm cb.
  44. 44. Relationship between the red cell volume & its degree or percentage saturation with Hb. It is volumes of red cell occupied by Hb. It represents actual concentration of Hb in red cells only.
  45. 45. Formula MCHC= Hb per 100ml of blood X 100 PCV per 100ml of blood
  46. 46. It is determined by the direct micrometric measurement of the red cells in a stained film. The range is 6.9 to 8 micrometer.  Average of 7.5 micrometer.
  47. 47. Rate at which red blood settle or sediment. Part of complete blood test. The determination is useful to check the progress of the disease.
  48. 48. ESR is increased in all conditions where there is tissue breakdown or where there is entry of foreign proteins in the blood, except for localized mild infections. The changes of ESR are not diagnostic of any specific disease.
  49. 49. Westergren’ method Wintrobe method
  50. 50.  Blood is collected by Venepuncture app 2ml blood is sufficient.  0.5ml 3.8 % sodium citrate is added as an anticoagulant.  Westergren’s pipette is filled with the help of rubber teat.  The pipette is stabilized at stand for one hour at the end of which readings are noted.
  51. 51. Normal Range: - MALE: 0-15 mm after 1st hour - FEMALE : 0-20 mm after 1st hour
  52. 52.  Blood is collected by Venepuncture app 5ml blood is sufficient.  0.5ml 3.8 % sodium citrate is added as an anticoagulant.  Wintrobe’s pipette is filled with the help of rubber teat.  The pipette is stabilized at stand for one hour at the end of which readings are noted.
  53. 53. Normal Range: - MALE : 0-9 mm/after 1st hour - FEMALE: 0-20 mm/after 1st hour
  54. 54. Faulty readings due to improperly clean tubes. Incorporation of air bubbles Disturbances of tube when placed on stand Calculation errors
  55. 55.  Cold agglutinins - low red cell counts and high MCVs can be caused by a increased number of large red cells or red cell agglutinates.  If agglutinated red cells are present, the automated hematocrits and MCHCs are also incorrect.  Cold agglutinins cause agglutination of the red cells as the blood cools.
  56. 56. Cold agglutinins can be present in a number of disease states, including infectious. If red cell agglutinates are seen on the peripheral smear, warm the sample in a 37 degrees C heating block and mix and test the sample while it is warm. Strong cold agglutinins may not disperse and need to be redrawn in a pre-warmed tube and kept at body temperature.
  57. 57.  Fragmented or very microcytic red cells  These may cause red cell counts to be decreased and may flag the platelet count as the red cells become closer in size to the platelets.  Cause an abnormal platelet histogram.  The population is visible at the left side of the red cell histogram and the right end of the platelet histogram.
  58. 58. Platelet clumps and platelet satellitosis: these cause falsely decreased platelet counts. Platelet clumps can be seen on the right side of the platelet histogram. Decreased platelet counts are confirmed by reviewing the peripheral smear. Always scan the edge of the smear when checking low platelet counts.
  59. 59. 4. Giant platelets: these are platelets that approach or exceed the size of the red cells. They cause the right hand tail of the histogram to remain elevated and may be seen at the left of the red cell histogram. 5. Nucleated red blood cells: these interfere with the WBC on some instruments by being counted as white cells/lymphocytes .
  60. 60. 1) Bleeding Time. 2) Clotting Time. 3) Prothrombin time. 4) Partial Thromboplastin Time. 5) Determination of Thrombin Time. 6) Determination Fibrinogen.
  61. 61. The process of stoppage of bleeding after blood vessels are punctured, cut or injured. Involve 4 process 1. Vasocostriction 2. Platelet plug formation 3. Formation of blood clot 4. Fibrinolysis
  62. 62. It is the time interval between the skin puncture & spontaneous, unassisted (i. e. without pressure) stoppage of bleeding.
  63. 63. Materials Equipment for sterile finger prick Clean filter paper Stopwach Normal value 1-5min
  64. 64. Method Finger prick & start the stop watch Absorb/ remove the blood drops every 30sec. Note the time when bleeding stops. This is the end point.
  65. 65. 1. Improper blood collection 2. Squeezing of finger 3. Time to start stopwatch 4. Pressure applied during touching the blotting paper
  66. 66. It is the time interval between the entry of blood into the glass capillary tube, or a syringe and formation of fibrin thread. Normal range 3-6min
  67. 67. Finger prick Absorb first 2 drops After large drop is formed Dip one end of capillary so that blood will rise & timer is started simultaneously Gently break off 1cm bits of glass tube for each 30 sec interval The time at which the fibrin thread is formed is the result.
  68. 68. Improper finger prick. Inadequate filling of capillaries. Air bubble incorporation. Improper pressure during breaking the capillaries. Time to start stopwatch.

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