Published in Microbiology
Monday, 31 July 2017 18:06
Routine care and proper maintenance of microscope will ensure good performance over the years. In addition to this, a properly maintained and clean microscope will always be ready for use at any time. Professional cleaning and maintenance should be considered when routine techniques fail to produce optimal performance of the microscope.
Cleaning and maintenance supplies
Dust cover: When not in use, a microscope should be covered to protect it from dust, hair, and any other possible sources of dirt. It is important to note that a dust cover should never be placed over a microscope while the illuminator is still on.
Lens tissue: Lint-free lens tissues are delicate wipes that would not scratch the surface of the oculars or objective. Always ensure that you are using these types of tissues. Never substitute facial tissue or paper towel, as they are too abrasive.
Lens cleaner: Lens cleaning solution assists in removing fingerprints and smudges on lenses and objectives. Apply the lens cleaner to the lens tissue paper and clean/polish the surface.
Compressed air duster: Using compressed air to rid the microscope of dust particles is far superior to using your own breath and blowing onto the microscope. Compressed air is clean, and avoids possible contamination of saliva particles.
Maintenance tips
  1. Whenever the microscope is not in use, turn off the illuminator. This will greatly extend the life of the bulb, as well as keep the temperature down during extended periods of laboratory work.
  2. When cleaning the microscope, use distilled water or lens cleaner. Avoid using other chemicals or solvents, as they may be corrosive to the rubber or lens mounts.
  3. After using immersion oil, clean off any residue immediately. Avoid rotating the 40× objective through immersion oil. If this should occur, immediately clean the 40× objective with lens cleaner before the oil has a chance to dry.
  4. Do not be afraid to use many sheets of lens tissue when cleaning. Use a fresh piece (or a clean area of the same piece) when moving to a different part of the microscope. This avoids tracking dirt/oil/residue to other areas of the microscope.
  5. Store the microscope safely with the stage lowered and the smallest objective in position (4× or 10×). This placement allows for the greatest distance between the stage and the objective. If the microscope is bumped, the likelihood of an objective becoming damaged by the stage surface will be greatly minimized.


Published in Hemotology
Monday, 31 July 2017 17:06
Automated hematology analyzers work on different principles:
  • Electrical impedance
  • Light scatter
  • Fluorescence
  • Light absorption
  • Electrical conductivity.
Most analyzers are based on a combination of different principles.
(1) Electrical impedance: This is the classic and timetested technology for counting cellular elements of blood. As this method of cell counting was first developed by Coulter Electronics, it is also called as Coulter principle (see Figure 811.1). Two electrodes placed in isotonic solutions are separated by a glass tube having a small aperture. A vacuum is applied and as a cell passes through the aperture, flow of current is impeded and a voltage pulse is generated.
Figure 811.1 Coulter principle of electrical impedance
Figure 811.1 Coulter principle of electrical impedance
The requisite condition for cell counting by this method is high dilution of sample so that minimal numbers of cells pass through the aperture at one point of time. There are two electrodes on either side of the aperture; as the solution in which the cells are suspended is an electrolyte solution, an electric current is generated between the two electrodes. When a cell passes through this narrow aperture across which a current is flowing, change in electrical resistance (i.e. momentary interruption of electrical current between the two electrodes) occurs. A small pulse is generated due to a temporary increase in impedance. This pulse is amplified, measured, and counted. The height of the pulse is proportional to cell volume. The width of the pulse corresponds with the time required for the cell to traverse the aperture. Cells that do not pass through the center of the aperture generate a distorted pulse that is not representative of the cell volume. Some analyzers use hydrodynamic focusing to force the cells through the central path so that all cells take the same path for volume measurement.
An anticoagulated whole blood sample is aspirated into the system, divided into two portions, and mixed with a diluent. One dilution is passed to the red cell aperture bath (for red cell and platelet counting), and the other is delivered to the WBC aperture bath (where a reagent is added for lysis of red cells and release hemoglobin; this portion is used for leukocyte counting followed by estimation of hemoglobin). Particles between 2-20 fl are counted as platelets, while those between 36-360 fl are counted as red cells. Hemoglobin is estimated by light transmission at 535 nm.
(2) Light scatter: Each cell flows in a single line through a flow cell. A laser device is focused on the flow cell; as the laser light beam strikes a cell it is scattered in various directions. One detector captures the forward scatter light (forward angle light scatter or FALS) that is proportional to cell size and a second detector captures side scatter (SS) light (90°) that corresponds to the nuclear complexity and granularity of cytoplasm. This simultaneous measurement of light scattered in two directions is used for distinguishing between granulocytes, lymphocytes, and monocytes.
(3) Fluorescence: Cellular fluorescence is used to measure RNA (reticulocytes), DNA (nucleated red cells), and cell surface antigens.
(4) Light absorption: Concentration of hemoglobin is measured by absorption spectrophotometry, after conversion of hemoglobin to cyanmethemoglobin or some other compound. In some analyzers, peroxidase cytochemistry is used to classify leukocytes; the peroxidase activity is determined by absorbance.
(5) Electrical conductivity: Some analyzers use conductivity of high frequency current to determine physical and chemical composition of leucocytes for their classification.
Further Reading:


Published in Hemotology
Sunday, 30 July 2017 18:20
Automation is a process of replacement of tasks hitherto performed by humans by computerized methods.
Until recently, hematological tests were performed only by manual methods. These methods, though still performed in many peripheral laboratories, are laborintensive, and involve use of hemocytometers (counting chambers), centrifuges, Wintrobe tubes, photometers, and stained blood smears. Hematology cell analyzers can generate the blood test results rapidly and also perform additional tests not possible by manual technology.
Both manual and automated laboratory techniques have advantages and disadvantages, and it is unlikely that one will completely replace the other.
Advantages of Automated Hematology Analyzer
  • Speed with efficient handling of a large number of samples.
  • Accuracy and precision in quantitative blood tests.
  • Ability to perform multiple tests on a single platform.
  • Significant reduction of labor requirements.
  • Invaluable for accurate determination of red cell indices.
Disadvantages of Automated Hematology Analyzer
  • Flags: Flagging of a laboratory test result demands labour-intensive manual examination of a blood smear.
  • Comments on red cell morphology cannot be generated. Abnormal red cell shapes (such as fragmented cells) cannot be recognized.
  • Erroneously increased or decreased results due to interfering factors.
  • Expensive with high running costs.
Automated hematology analyzers are of two main types:
  • Semi-automated: Some steps like dilution of blood sample are performed by the technologist; can measure only a few parameters.
  • Fully automated: Require only anticoagulated blood sample; measure multiple parameters.


Published in Hemotology
Saturday, 29 July 2017 19:18
These are listed in Table 809.1
Table 809.1 Causes of erroneous results with hematology analyzer
Parameter Interfering factors
  Erroneous increase Erroneous decrease
0. All parameters
  • Clotted sample
1. WBC count
  • Nucleated red cells
  • Large platelet clumps
  • Unlysed red cells (some abnormal red cells resist lysing)
  • Cryoglobulins
  • Clotted sample
2. RBC count
  • Very high WBC*
  • Large numbers of giant platelets
  • Clotted sample
  • Microcytic red cells
  • Autoagglutination
3. Hemoglobin
  • Clotted sample
4. MCV
  • Very high WBC
  • Hyperglycemia
  • Autoagglutination (cold agglutinins)
  • Cryoglobulins
  • Hyperlipidemia
  • Autoagglutination (cold agglutinins)
  • Very high WBC
6. Platelets
  • Microcytic red cells
  • WBC fragments
  • Cryoglobulins
  • Platelet satellitism
  • Platelet clumping
*: WBCs are counted along with RBCs, but normally their number is statistically insignificant
‘Flags’ are signals that occur when an abnormal result is detected by the analyzer. Flags are displayed to reduce false-positive and false-negative results by mandating a review of blood smear examination.


Published in Hemotology
Saturday, 29 July 2017 17:54
Parameters measured by hematology analyzers and their derivation are shown in Tables 808.1 and 808.2. Most automated hematology analyzers measure red cell count, red cell indices (mean cell volume, mean cell hemoglobin, mean cell hemoglobin concentration), hemoglobin, hematocrit, total leukocyte count, differential leukocyte count (three-part or five-part), and platelet count.
Table 808.1 Parameters measured by hematology analyzers
Parameters measured by most analyzers Parameters measured by some analyzers
  • RBC count
  • Hemoglobin
  • Mean cell volume
  • Mean cell hemoglobin
  • Mean cell hemoglobin concentration
  • WBC count
  • WBC differential
  • Platelet count
  • Red cell distribution width
  • Reticulocyte count
  • Reticulocyte hemoglobin content
  • Mean platelet volume
  • Platelet distribution width
  • Reticulated platelets
Table 808.2 Parameters reported by hematology analyzers
Parameters measured directly or derived through histogram Parameters measured through calculation
  • RBC count
  • Mean cell volume (Derived from RBC histogram)
  • Red cell distribution width (Derived from RBC histogram)
  • Hemoglobin
  • Reticulocyte count
  • WBC count
  • Differential WBC count (Derived through WBC histogram)
  • Platelet count
  • Mean platelet volume (Derived from platelet histogram)
  • Hematocrit
  • Mean cell hemoglobin
  • Mean cell hemoglobin concentration
Estimation of Hemoglobin
Hemoglobin is measured directly by a modification of cyanmethemoglobin method (all hemoglobins are converted to cyanmethemoglobin by potassium ferricyanide; cyanmethemoglobin has a broad absorbance peak at 540 nm). Some analyzers use a nonhazardous reagent such as sodium lauryl sulphate. A non-ionic detergent is added for rapid red cell lysis and to minimize turbidity caused by cell membranes and plasma lipids.
Estimation of Red Blood Cell Count and Mean Cell Volume (MCV)
Red cell count and cell volume are directly measured by aperture impedance or light scatter analysis. In a red cell histogram, cell numbers are plotted on Y-axis, while cell volume is indicated on Xaxis (see Figure 808.1). The analyzer counts those cells as red cells volume of which ranges between 36 fl and 360 fl. MCV is used for morphological classification of anemia into microcytic, macrocytic, and normocytic types.
Figure 808.1 Diagrammatic representation of red cell histogram obtained by aperture impedance
Figure 808.1 Diagrammatic representation of red cell histogram obtained by aperture impedance. The analyzer counts cells between 36 fl and 360 fl as red cells. Although leukocytes are present and counted along with red cells in the diluting fluid, their number is not statistically significant. Only if leukocyte count is markedly elevated (>50,000/μl), histogram and the red cell count will be affected. Area of the peak between 60 fl and 125 fl is used for calculation of mean cell volume and red cell distribution width. Abnormalities in red cell histogram include: (1) Left shift of the curve in microcytosis, (2) Right shift of the curve in macrocytosis, and (3) Bimodal peak of the curve in double (dimorphic) population of red cells
Estimation of MCH, MCHC, and Hematocrit (HCT/PCV)
These parameters are obtained indirectly through calculations.
MCH (pg) = Hemoglobin (g/l)
                     RBC count (10⁶/μl)
MCHC (g/dl) = Hemoglobin (g/dl)
                         Hematocrit (%)
Hematocrit (%) = Mean Cell Volume (fl)
                              RBC count (10⁶/μl)
Estimation of Red Cell Distribution Width (RDW)
RDW is a quantitative measure of variation in sizes of red cells and is expressed as coefficient of variation of red cell size distribution. It is equivalent to anisocytosis observed on blood smear. It is derived from red cell histogram in some analyzers. RDW is usually elevated in iron deficiency anemia, but not in β-thalassemia minor and anemia of chronic disease (other causes of microcytic anemia). However, this distinction is not absolute and there is a significant overlap between values among patients. Raised RDW requires examination of blood smear.
Among the red cell values generated by the analyzer (red cell count, hemoglobin, hematocrit, MCV, MCH, MCHC, and RDW), most important for decision-making are hemoglobin, hematocrit, and MCV.
WBC Differential
Difference between 3-part and 5-part hemotology analyzer...
Hematology analyzers can either generate a 3-part differential (differential count reported as lymphocytes, monocytes, and granulocytes) or a 5-part differential (lymphocytes, monocytes, neutrophils, eosinophils, and basophils). The 3-part differential counting is based on electrical impedance volume measurement of leukocytes. In volume histogram for WBCs, approximate numbers of cells are plotted on Y-axis and cell size on X-axis. Those cells with volume 35-90 fl are designated as lymphocytes, cells with volume 90-160 fl as mononuclear cells, and cells with volume 160-450 fl as neutrophils (see Figure 808.2). Any deviation from the expected histogram is flagged by the analyzer, mandating review of blood smear. A large proportion of 3-part differential counts are ‘flagged’ to avoid missing abnormal cells.
Instruments measuring a 5-part differential work on a combination of different principles, e.g. light scatter, impedance, and electrical conductivity, a combination of light scatter, peroxidase staining, and resistance of basophils to lysis in acid buffer, etc.
Figure 808.2 Diagrammatic representation of WBC histogram
Figure 808.2 Diagrammatic representation of WBC histogram. WBC histogram analysis shows relative numbers of cells on Y-axis and cell size on X-axis. The lytic agent lyses the cytoplasm that collapses around the nucleus causing differential shrinkage. The analyzer sorts the WBCs according to the nuclear size into 3 main groups (3-part differential): Cells with 35-90 fl volume are designated as lymphocytes, cells with 90-160 fl volume are designated as monocytes, and cells with 160-450 fl volume are designated as neutrophils. Abnormalities in WBC histogram include: (1) Peak to the left of lymphocyte peak: Nucleated red cells, (2) Peak between lymphocytes and monocytes: Blast cells, eosinophilia, basophilia, plasma cells, and atypical lymphocytes, and (3) Peak between monocytes and neutrophils: Left shift
Platelet Count
Platelets are difficult to count because of their small size, marked variation in size, tendency to aggregation, and overlapping of size with microcytic red cells, cellular fragments, and other debris. In hematology analyzers, this difficulty is addressed by mathematical analysis of platelet volume distribution so that it corresponds to lognormal distribution. Platelets are counted by electrical impedance method in the RBC aperture, and a histogram is generated with platelet volume on X-axis and relative cell frequency on Y-axis (see Figure 808.3). Normal platelet histogram consists of a right-skewed single peak. Particles greater than 2 fl and less than 20 fl are classified as platelets by the analyzer.
Figure 808.3 Diagrammatic representation of normal platelet histogram
Figure 808.3 Diagrammatic representation of normal platelet histogram: Counting and sizing of platelets by electrical impedance method occurs in the RBC aperture. The counter designates particles between sizes 2 fl and 20 fl as platelets. Abnormalities in platelet histogram result from interferences such as cytoplasmic fragments (peak at left end of histogram) or severely microcytic red cells and giant platelets (peak at right end of histogram)
Two other platelet parameters can be obtained from platelet histogram using computer technology: mean platelet volume (MPV) and platelet distribution width (PDW). Some analyzers can generate another parameter called as reticulated platelets.
MPV refers to the average size of platelets and is obtained from mathematical calculation. Normal MPV is 7-10 fl. Increased MPV (> 10 fl) results from presence of immature platelets in circulation; peripheral destruction of platelets stimulates megakaryocytes to produce such platelets (e.g. in idiopathic thrombocytopenic purpura). Decreased MPV (< 7 fl) is due to presence of small platelets in circulation (in conditions associated with reduced production of platelets in bone marrow).
PDW is analogous to RDW and is a measure of variation in size of platelets (normal <20%). Increased PDW is observed in megaloblastic anemia, chronic myeloid leukemia, and after chemotherapy.
Some analyzers measure reticulated platelets or young platelets that contain RNA (similar to reticulocytes). Increased numbers of reticulated platelets are seen in thrombocytopenia due to peripheral destruction of platelets.

Reticulocyte Count
Various fluorescent dyes can combine with RNA of reticulocytes; the fluorescence then is counted in a flow cytometer. More immature reticulocytes fluoresce more strongly as they contain more RNA.
Reticulocyte hemoglobin content is a parameter that estimates hemoglobinization of most recently produced red cells. It is a predictor of iron deficiency.
WBC Cytogram (Scattergram)
In the scattergram, each dot represents a cell of a given volume and density, and the positions of dots in the graph are determined by the degree of side scatter, degree of forward scatter, light absorption by the cell, and cytochemical staining (if used). The forward angle light scatter (FALS) is represented on Y-axis, and the side scatter (SS) is represented on X-axis. Low FALS and low SS are indicative of lymphocytes; with subsequent increasing FALS and SS, monocytes, neutrophils, and lastly eosinophils are designated in the graph. Counting of basophils is based on a different technology.
Further Reading:


Published in Hemotology
Saturday, 29 July 2017 15:37
Box 807.1 Properties of a cell measured by a flow cytometerFlow cytometry is a procedure used for measuring multiple cellular and fluorescent properties of cells when they flow as a single cell suspension through a laser beam by a specialized instrument called as a flow cytometer. Flow cytometry can analyze numerous cells in a short time and multiple parameters of a single cell can be analyzed simultaneously. From the measured parameters, specific cell populations are defined. Cells or particles with size 0.2-150 μm are suitable for flow cytometer analysis.
Flow cytometry can provide following information about a cell (Box 807.1):
  • Cell size (forward scatter)
  • Internal complexity or granularity (side scatter)
  • Relative fluorescence intensity.
A flow cytometer consists of three main components or systems: fluidics, optics, and electronics.
(1) Fluidics: The function of the fluidics system is to transport cells in a stream to the laser beam for interrogation. Cells (fluorescence-tagged) are introduced into the cytometer (injected into the sheath fluid within the flow chamber) and made to flow in a single file past a laser (light amplification by stimulated emission of radiation) beam. The stream transporting the cells should be positioned in the center of the laser beam. The portion of the fluid stream where the cells are located is called as the sample core. Only a single cell or particle should pass through the laser beam at one time. Flow cytometers use the principle of hydrodynamic focusing (process of centering the sample core within the sheath fluid) for presenting cells to the laser.
(2) Optics: This system consists of lasers for illumination of cells in the sample, and filters to direct the generated light signals to the appropriate detectors.
The light source used in most flow cytometers is laser.
The laser most commonly used in flow cytometry is Argon-ion laser. The light signals are generated when the laser beam strikes the cell, which are then collected by appropriately positioned lenses. A system of optical mirrors and filters then directs the specified wavelengths of light to the designated detectors. Two types of light signals are generated when the laser beam strikes cells tagged with fluorescent molecules: fluorescence and light scatter. The cells tagged with fluorescence emit a momentary pulse of fluorescence; in addition, two types of light scatter are measured: forward scatter (proportional to cell diameter) and side scatter (proportional to granularity of cell).
(3) Electronics: The optical signals (photons) are converted to corresponding electronic signals (electrons) by the photodetectors (photodiodes and photomultiplier tubes). The electronic signal produced is proportional to the amount of light striking a cell. The electric current travels to the amplifier and is converted to a voltage pulse. The voltage pulse is assigned a digital value representing a channel by the Analog-to Digital Converter (ADC). The channel number is then transferred to the computer which displays it to the appropriate position on the data plot.
Further Reading:


Published in Hemotology
Friday, 28 July 2017 17:59
  1. Leukemias and lympomas: Immunophenotyping (evaluation of cell surface markers), diagnosis, detection of minimal residual disease, and to identify prognostically important subgroups.
  2. Paroxysmal nocturnal hemoglobinuria: Deficiency of CD 55 and CD 59.
  3. Hematopoietic stem cell transplantation: Enumeration of CD34+ stem cells.
  4. Feto-maternal hemorrhage: Detection and quantitation of foetal hemoglobin in maternal blood sample.
  5. Anemias: Reticulocyte count.
  6. Human immunodeficiency virus infection: For enumeration of CD4+ lymphocytes.
  7. Histocompatibility cross matching.


Published in Hemotology
Friday, 28 July 2017 10:05
Platelet aggregation tests are carried out in specialized hematology laboratories if platelet dysfunction is suspected. These tests are usually indicated in patients presenting with mucocutaneous type of bleeding and in whom screening tests reveal normal platelet count, prolonged bleeding time, normal prothrombin time, and normal activated partial thromboplastin time. Platelet aggregation studies are carried out on platelet-rich plasma using aggregometer. When a platelet aggregating agent is added to platelet-rich plasma, platelets form aggregates and optical density falls (or light transmission increases); this is recorded by a chart recorder on a strip chart. Commonly used platelet aggregating agents are ADP (adenosine 5-diphosphate), epinephrine (adrenaline), collagen, arachidonic acid, and ristocetin. ADP (low dose) and epinephrine induce primary and secondary waves of aggregation (biphasic curve). Primary wave is due to the direct action of aggregating agent on platelets. Secondary wave is due to thromboxane A2 synthesis and secretion from platelets. Collagen, arachidonic acid and ristocetin induce a single wave of aggregation (monophasic curve) Normal aggregation curve is shown in Figure 804.1. Aggregation patterns in various platelet function defects are shown in Figures 804.2 to 804.4, and in Table 804.1.
Figure 804.1 Normal platelet aggregation curves
Figure 804.1 Normal platelet aggregation curves
Figure 804.2 Platelet aggregation curves in von Willebrand disease and Bernard Soulier syndrome absent aggregation with ristocetin normal aggregation with ADP epinephrine and arachidonic acid
Figure 804.2 Platelet aggregation curves in von Willebrand disease and Bernard-Soulier syndrome (absent aggregation with ristocetin, normal aggregation with ADP, epinephrine, and arachidonic acid)
Figure 804.3 Platelet aggregation curves in storage pool defect absent second wave of aggregation with ADP and epinephrine absent or greatly diminished aggregation with collagen and normal ristocetin aggregation
Figure 804.3 Platelet aggregation curves in storage pool defect (absent second wave of aggregation with ADP and epinephrine, absent or greatly diminished aggregation with collagen, and normal ristocetin aggregation)
Figure 804.4 Platelet aggregation curves in Glanzmanns thrombasthenia absent aggregation with all agonists except ristocetin
Figure 804.4 Platelet aggregation curves in Glanzmann’s thrombasthenia (absent aggregation with all agonists except ristocetin)


Published in Hemotology
Thursday, 27 July 2017 11:22
A blood smear is examined for:
A peripheral blood smear has three parts: Head, body, and tail. Also read: PREPARATION OF BLOOD SMEAR BY WEDGE METHOD.
A blood smear should be examined in an orderly manner. Initially, blood smear should be observed under low power objective (10×) to assess whether the film is properly spread and stained, to assess cell distribution, and to select an area for examination of blood cells. Best morphologic details are seen in the area where red cells are just touching one another. Low power view is also helpful for the identification of Rouleaux formation, autoagglutination of red cells, and microfilaria. High power objective (45×) is suitable for examination of red cell morphology and for differential leukocyte count. A rough estimate of total leukocyte count can be obtained which also serves to crosscheck the total leukocyte count done by manual counting or automated method. Oil-immersion objective (100×) is used for more detailed examination of any abnormal cells.
Further Reading:


Published in Hemotology
Wednesday, 26 July 2017 17:45
Box 802.1 Role of blood smear in thrombocytopeniaPlatelets are small, 1-3 μm in diameter, purple structures with tiny irregular projections on surface. In blood films prepared from non-anticoagulated blood (i.e. direct fingerstick), they occur in clumps. If platelet count is done on automated blood cell counters using EDTA-anticoagulated blood sample, about 1% of persons show falsely low count due to the presence in them of EDTA dependent antiplatelet antibody. Examination of a parallel blood film is useful in avoiding the false diagnosis of thrombocytopenia in such cases. Occasionally, platelets show rosetting around neutrophils (platelet satellitism) (see Figure 802.1). This is seen in patients with platelet antibodies and in apparently normal persons. Blood smear examination can be helpful in determining underlying cause of thrombocytopenia such as leukemia, lymphoma, or microangiopathic hemolytic anemia (Box 802.1).
Also Read:

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