Erythrocyte Sedimentation Rate
The rate at which erythrocytes sediment in anticoagulated whole blood is an indirect measure of fibrinogen and globulin. Thus, it is a poor man’s interleukin-1 assay. This rate is elevated in a wide variety of conditions of infection, inflammation, tissue necrosis, and neoplasia. Because of its nondiagnosticity for single diseases, it has lost favor; it is now possible to distinguish, for example, angina pectoris from infarction or rheumatoid arthritis from osteoarthritis on other grounds. Still, the test has utility for screening large numbers of patients and for following therapy in inflammatory conditions such as rheumatoid arthritis. I also use it on a yearly basis when following hypochondriacal patients.
Place blood, anticoagulated with ethylenediamine tetraacetic acid (EDTA) or oxalate, in a Wintrobe tube, which is placed in a rack or taped to the wall behind the patient’s bed. Be sure the tube is perfectly vertical.
One hour later, note, in millimeters, how much settling has taken place. (Measure from the top of the plasma to the top of the red cell column.)
Correct the sedimentation rate according to the hematocrit (Fig. 28-1
). If the hematocrit is not already known, it can be determined on the same sample in the Wintrobe tube.
False-positive elevations in the sedimentation rate occur in old age and pregnancy.1
False negatives (normal sedimentation rates) occur in typhoid fever, brucellosis, and 2% of cavitary tuberculosis cases.
Some viral infections do not increase the sedimentation rate, whereas some mild viral infections may do so (Ham et al., 1957
; Wintrobe, 1967
FIGURE 28-1 Reference chart for correcting the depth of sedimentation in 1 hour for the hematocrit, using the Wintrobe-Landsberg method. (Modified from Hynes M, Whitby EH. Correction of the sedimentation rate for anemia. Lancet. 1938;2:249-251, with permission.) The sedimentation rate is to be corrected from the observed hematocrit to a sedimentation that corresponds to a hematocrit of 45%. The normal range for the corrected sedimentation rate is 0 to 10 mm. Plot the point corresponding to the observed hematocrit and the observed sedimentation rate. This will fall in one of the zones indicating the approximate degree of increase in the rate. (Follow the nearest curve to the point where it intersects the line corresponding to a hematocrit of 45%. Read the corrected sedimentation rate. For example, if the observed sedimentation is 50 mm and the observed hematocrit is 25%, then the sedimentation in 1 hour corrected to a hematocrit of 45% is approximately 11 mm. If the observed sedimentation is 8 mm and the observed hematocrit is 56%, then the sedimentation in 1 hour corrected to a hematocrit of 45% is approximately 30 mm.) (From Wintrobe MM. Sedimentation rate vs. hematocrit. Lancet. 1938;4:30, with permission.)
Lee-White Clotting Time
Although replaced in most laboratories by the various partial thromboplastin times, this test is still useful for diagnosing decreased amounts of procoagulants and for monitoring heparin therapy, if the partial thromboplastin time is not available. It can also be used in the “50/50” mixing test (vide infra), even if a partial thromboplastin time is available, simply as a matter of convenience.
Draw blood from a vein and place 2 mL in a Pyrex tube 8 × 15 × 100 mm. (Wider tubes yield longer clotting times, and smaller volumes yield shorter clotting times.) Note “zero time,” defined as the moment that blood first appears in the syringe.
. The blood should be allowed to run down the side of the tube and specifically should not be “jet-sprayed” into the tube or handled in any way that will produce bubbles (Waldron and Duncan, 1954
After 5 minutes have elapsed, begin tilting the tube to a 45-degree angle at 1-minute intervals.
The clotting time is the interval at which the tube can be inverted without displacing the clot. The normal time is 5 to 8 minutes (Todd and Sanford, 1948
Variations. This test has been modified to use two, three, and even four tubes. One begins the stopwatch immediately after filling the tubes and starts tilting the first tube. When that one has clotted, proceed to the next one in turn. The intertilting interval may be shortened to 30 seconds but increased handling shortens the clotting time.
In some versions of the test, the tubes are placed in a 37°C water bath. In others, the tubes are prechilled by rinsing in iced saline. Under warmer conditions, the clotting time is slightly shorter.
. When two tubes are used, the clotting time is the average unless the tubes vary by more than 5 minutes, in which case only the clotting time of the second tube is reported. The normal value is 4 to 12 minutes (Frommeyer and Epstein, 1957
For three or four tubes, the clotting time is taken to be the time at which the last tube can be inverted. The normal is 5 to 15 minutes for three tubes and less than 17 minutes for four tubes (Wintrobe, 1967
The most important issue in interpreting results is that the test should be performed under the same conditions each time.
. The importance of drawing blood from the vein so as to avoid contact with tissue thromboplastin was already known to Howell in 1905. The test had been performed using a coagulometer by Pratt in 1903, and Morawitz and Bierch performed it using a glass tube in 1907. Addis had added the step of inverting the test tube for determining the end point at least by 1910. Other workers had performed essentially the same test, as noted by Lee and White, who wrote, “the method described is not completely new…” (Lee and White, 1913
“50/50” Mixing Test for Circulating Anticoagulants
A patient with a prolonged clotting time may have a circulating anticoagulant instead of a simple deficiency of a clotting factor. To make this distinction, the clotting time (or other test of impaired coagulation, such as the automated partial thromboplastin time) is repeated, using a mixture of equal parts of the patient’s blood and that of a normal person. The abnormal clotting test will become normal if the patient has insufficient clotting factors but not if a circulating anticoagulant is present. Preincubation at 37°C will increase the positive yield (Clyne and White, 1988
The rationale for this test is as follows: If one views the clotting factors as operating in an enzymatic cascade, an improvement from near zero to 50% of normal will provide sufficient material to run the reaction. However, if there is an anticoagulant sufficient to impair normal levels of enzymes, the clotting test will still be abnormal if the anticoagulant is only diluted out by one half.
Blood Clot Tests
Observe a perfectly vertical tube of blood that has been taped to the wall next to the patient’s bed.
If no clot ever forms, the fibrinogen content of the blood is thought to be very low, less than 60 mg per dL.
If the clot dissolves within 20 or 30 minutes, the patient has hypofibrinogenemia or accelerated fibrinolysis.
Clot retraction begins at about 1 hour and reaches a maximum in less than a day. With normal clot retraction, the clot becomes progressively smaller, leaving a space of completely clear serum behind it. Impaired clot retraction occurs more slowly than normal. Also, the red cells are not trapped effectively, so they leak out of the clot, fall through the serum, and coat the floor of the test tube.
Clot retraction depends on normal platelet number and function. If the platelets are decreased in number, there will be a decrease in clot retraction. Impaired clot retraction in the presence of a normal platelet count makes the diagnosis of “thrombasthenia” (weak platelets), or what we would now refer to as the family of “numerically adequate but functionally inferior” platelet syndromes. In fact, this is the only bedside test that distinguishes “thrombasthenia” from vascular defects (such as those found in scurvy or amyloidosis).
False Negatives. These are falsely normal-appearing clot retractions despite impaired platelet function. With hypofibrinogenemia, fibrinolysis, or severe anemia, the abnormally small clot may mimic a normal clot that has retracted.
. Polycythemia may cause the clot to appear “too big” despite normal retraction. (For more on platelets, see “Rumpel-Leeds Test,” Chapter 7
, and the section on “Platelets” in this chapter.)
Leave the tube of blood up for more than a day to check for normal fibrinolysis.
Test for Heparin-Induced “White Clot” Syndrome
Occasionally, heparin induces immune-mediated platelet aggregation, causing thrombocytopenia and paradoxic thromboemboli. The emboli are ghostly white and consist of platelet aggregates and fibrin.
Mix platelet-rich plasma from a control subject with plateletpoor plasma from a patient. If the patient has white clot syndrome, his plasma will sensitize the platelets of the control subject. With the addition of 1 unit of heparin per cubic centimeter of plasma, plus adenosine diphosphate solution, a platelet aggregate will form. This may be obvious to inspection or can be quantitated with a platelet aggregometer (Stanton et al., 1988
Formed Elements of the Blood
First, clinical microscopy was purged from the curriculum because there was no National Board Examination in it. Then, manual differential counts were automated, to the further detriment of patients, as certain kinds of findings were lost. This section is concerned with what was lost (and can be regained) rather than with cost effectiveness. It is intended to show neophytes and those who instruct them that there is both utility and pleasure in finding things for one’s self in the laboratory, in exact analogy to interviewing and examining one’s own patient.
How to Make a Blood Smear
Place a thick glass slide on a solid surface, frosted side up. Write the patient’s name and the date in pencil on the frosted area.
Place a drop of blood very near one end of the slide. Hold the other end with the fingers of your nondominant hand.
With your dominant hand, pick up a second glass slide to be used as a spreading device. Hold it at a 45-degree angle to the first slide, and place its edge near the drop of blood, between the drop and your nondominant hand. (The drop will now be in the 45-degree angle formed by the two slides; see Fig. 28-2
Bring the second slide back toward the drop of blood. When it touches the drop, surface tension will cause the drop suddenly to spread out all along the length of the touching slides (see Fig. 28-2
Quickly move the second slide toward your nondominant hand. This will pull the blood behind it and spread the blood over the glass without pressing on the blood corpuscles (Fig. 28-2
A perfect blood film will have a feathery edge. Usually, it takes two or three practice runs under supervision to produce such an edge, but once you learn the skill you will never lose it.
Allow the slide to dry (fix). Stain it according to the local ground rules, or the following procedure.
Staining a Peripheral Blood Smear
Place the smear on a support such as a cork nailed to a board or two rods hung over a sink.
When it is completely air dried (fixed), pour Wright stain on it so as to cover the entire slide. Wait 3 to 5 minutes.
Add the prepared buffer. Use enough to make the surface of the stain appear iridescent but not so much that the overlying fluid becomes transparent enough to see the smear beneath it. Mix in the traditional manner by gently blowing on the slide, not with a dowel rod or wooden applicator, lest the smear be disturbed.
After a few minutes, rinse with a wash bottle or under the tap and shake dry. Remaining topside bubbles may be blown off.
Blot with absorbent paper from the side. (Try not to touch the smear itself.)
The order of examination is as follows: (a) red cells, (b) white cells, and (c) platelets.
FIGURE 28-2 Preparing a blood smear (see text). When the second slide touches the drop on the first slide, surface tension will cause the drop suddenly to spread out all along the length of the touching slides.
The red cell number has already been implicitly measured by estimating the hematocrit from the nail beds and conjunctiva. Keep in mind such estimates of anemia and polycythemia when attempting to estimate the white cell count and platelet count from the relative numbers of cells on the smear (vide infra).
The diameter of the normal red cell is approximately the same as the nucleus of a small lymphocyte.
Information about red cell size can also be obtained from the mean corpuscular volume (MCV), which the electronic cell counter determines by displacement of an electrolyte solution, provided that there is no elliptocytosis, rouleaux formation (vide infra
), or cold agglutinins (Rappaport et al., 1988
). (In fact, the electronic cell counter’s measurement of MCV is more accurate than the hematocrit; the latter is not measured but rather calculated from the MCV and the red cell count, giving an error of 3% to 5% compared with the spun hematocrit.)
Considerable variation in the shape of the red cells (poikilocytosis) is a significant finding, not available from an inspection of the electronic hemogram. Abnormal cells include drepanocytes (also called banana cells and sickle cells), spherocytes, microspherocytes, stomatocytes, elliptocytes, burr cells, spur cells, helmet cells, bite cells, teardrop cells, basophilic cells, and schistocytes. Cabot rings, Howell-Jolly bodies, malarial parasites, nucleated red cells, and so forth are examples of red cell inclusions that cannot be detected without inspecting the smear.
Rouleaux formation is the appearance of the red cells as rolls or “stacks of coins.” A normal phenomenon in the thick portion of the smear, rouleaux formation is “called” only in the thin part of the smear and therefore requires much experience. It results from changes in the surface charge of the red cells due to coating with increased amounts of globulin. Dr Eugene Robin once diagnosed a case of multiple myeloma from noting the rouleaux formation and so can you.
Cold agglutinins can also cause rouleaux formation on slides prepared in the customary manner. This can be diagnosed with confidence if such rouleaux are not seen on a second slide made from blood kept warm through the smearing and fixing stages, as with an incandescent lamp.
Anisocytosis is an abnormal variation in the size distribution of the red cells.
Unless your electronic counter has an accurate measure of the size distribution of the red cells (red cell distribution width [RDW]), one would not know whether the patient had a dimorphic population of red cells without examining the smear. If the electronic counter does provide both these measures (MCV and RDW), there is no need to make size estimates from the smear, except to learn how to do it against the day when such a device might not be available. (The same principle applies, although unstated, to the other tests in this section.)
One source of excellent slides of blood cells is the online pathology lectures, online pathology lectures, www.medicalschoolpathology.com/,
by Dr John Minarcik (see “Red Blood Cells and Bleeding Disorders” and “Diseases of White Blood Cells, Lymph Nodes, Spleen, and Thymus”). For extensive material on histopathology, with videos, questions, and narrations, see www.medicalschoolpathology.com/HistoPathLabx.htm.
With practice, one can estimate the white blood cell (WBC) count from the peripheral smear, if one has estimated the hematocrit.
An Arneth lobe count, looking for hypersegmented polymorphonuclear leukocytes, is performed by counting only those lobes that are seen to be connected by a thin thread of chromatin, not those that are simply overlapped. In counting 100 to 200 cells, one only need see 5% five-lobed polymorphonuclear leukocytes to know that an abnormality is present. If you find a single six-lobed polymorphonuclear leukocyte, you can stop counting instantly.
The causes of hypersegmented polymorphonuclear leukocytes are well known and are found in standard texts, with two important exceptions.
The most common cause of hypersegmented polymorphonuclear leukocytes in the present hospital population is the uremic syndrome. This is immediately reversed by the administration of supplemental folate, even though pretreatment serum folate levels may have been borderline or even normal (Siddiqui et al., 1970
Patients with severe iron deficiency may have hypersegmented neutrophils that disappear with iron therapy. It is likely that iron deficiency inhibits formiminotransferase, thus producing a functional folate deficiency despite normal levels of folate in both the serum and red blood cells (Beard and Weintraub, 1969
A Barr body is a tiny projection from the nucleus, which is thought to represent the “extra” X chromosome in female cells. Males may appear to have Barr bodies on up to 5% of their neutrophils. More than 10% of leukocytes from women (genotype XX) will have Barr bodies. Although the Barr body has been likened to a drumstick (with the thick part distal to the nucleus proper), many XY nuclei will have drumsticks. A true Barr body looks more like a lollipop or a balloon on a stick.
The payoff for examining blood smears is to find the 1 of 600 phenotypically male patients whose hypogonadism is suddenly explicable based on the XXY chromosomes of Klinefelter syndrome.
are pale blue inclusions seen in Wright-stained polymorphonuclear leukocytes. By histochemical analysis, they are composed of ribonucleoprotein and/or ribonucleic acid. Once thought to be diagnostic of scarlet fever, they appear in the blood of 7% of hospitalized patients, specifically in the conditions listed in Table 28.1
. They are not seen in healthy student nurses (Abernathy, 1964
) or patients with urticaria or serum sickness (Granger and Pole, 1913
Döhle bodies with red granules within them are called Amato bodies
. These have the same significance as Döhle bodies, although they too were once thought to be diagnostic of scarlet fever (Toomey and Gammel, 1927
Structures identical to Döhle bodies in light microscopic appearance have also been seen in various congenital syndromes, including the May-Hegglin anomaly, Chédiak-Higashi syndrome, the pseudo-Pelger-Huet syndrome, the Fechtner syndrome, and so forth (Peterson et al., 1985
are pink (by Wright stain) intracytoplasmic rods or clumps that come from the azure granules that in turn come from lysosomes.
They are most frequent in immature cells. They were discovered by Auer, who reported seeing them in lymphocytic leukemia (Auer, 1906
), and this continues to be reported (Juneja et al., 1987
). Both reported seeing Auer rods in tuberculosis in 1913 (Freeman, 1960
), and this too has been replicated (Leavell and Twomey, 1964
). They are seen in 21% of acute myelogenous leukemia patients, or 66% to 75% if peroxidase staining is used (Jain et al., 1987
TABLE 28.1 Döhle bodies: Associations
Scarlet fever (100% to first 2 days)
(Granger and Pole, 1913)
Tuberculosis (11% to “practically all”)
(Bachman and Lucke, 1918)
Lobar pneumonia (67%-100%)
(Bachman and Lucke, 1918)
Viral upper respiratory infection
(Bachman and Lucke, 1918)
(Granger and Pole, 1913)
German measles (rare)
(Bachman and Lucke, 1918)
Chicken pox (rare)
(Bachman and Lucke, 1918)
Measles (45% in the first 5 days)
(Granger and Pole, 1913)
Diphtheria (68% in the first 5 days)
(Granger and Pole, 1913)
(Granger and Pole, 1913)
After the infusion of diphtheria toxin
After blood transfusion
After cyclophosphamide treatment
From Abernathy MR. Incidence of Döhle bodies in physiologic and pathologic conditions. Lab Digest. 1964;28:3-5 and Abernathy MR. Döhle bodies associated with uncomplicated pregnancy. Blood. 1966;27: 380-385, with permission.
The eosinophil count obtained by multiplying the white cell count by the percentage of eosinophils is only accurate when the percentage of eosinophils is at least 5% to 6%, even if the white cell count is very high. Accuracy can be improved by counting 200 white cells instead of the usual 100.
The term eosinophilia refers to a count of more than 310 eosinophils per mm3. The differential diagnosis of eosinophilia (which includes parasites, allergic conditions, some infectious conditions such as tuberculosis, malignant conditions, and a number of other causes, all diagnosable on other grounds) is out of fashion in most medical centers. Thus, there is little awareness of a frequent new cause of eosinophilia in the modern medical center—therapeutic radiation-induced eosinophilia (Ghossein et al., 1975
Any degree of eosinophilia rules out the diagnosis of Cushing syndrome, and the absence of eosinophils rules out the diagnosis of chronic adrenocortical insufficiency.
An estimation of platelet number should routinely be made from the smear, and the examiner’s skill should be augmented by feedback from the electronic platelet count. Begin by estimating 25,000 platelets per mm3 for every platelet seen in a high-power field.
Electronic cell counters may produce a spurious thrombocytopenia in three circumstances (Kjeldsberg and Hershgold, 1974
): (a) platelet satellitism, which occurs in blood collected in EDTA as platelets adhere to polymorphonuclear leukocytes and are excluded by size from the count; (b) in the presence of platelet agglutinins, which can cause the platelets to clump together—the clump is also optically excluded by the counter because of its size; and (c) giant platelets, which are excluded for the same reason.
Spurious thrombocytosis may be produced by the electronic cell counter in three circumstances (Rappaport et al., 1988
), all of which can be recognized on the smear: (a) fragmented white cells (smudge cells), as in leukemia or sepsis; (b) fragmented red cells, as in thrombotic thrombocytopenic purpura, disseminated intravascular coagulation, microangiopathic hemolytic anemia, or cardiopul-monary bypass; and (c) marked microcytosis of the red cells as in microspherocytosis. (Also see the “Rumpel-Leede Test” section in Chapter 7
and “Blood Clot Tests” above.)
Earlobe Histiocytes (Low Power)
In the first
drop of blood (Smith, 1964
) taken from the lanced earlobe of a patient with subacute bacterial endocarditis, it is often, but not always, possible to find large histiocytes, which are at least one and a half times as large as monocytes (Daland et al., 1956
; Van Nuys, 1907
). (This is an example of empiric but not scientific fact as we do not know why this should be true, only that it is.)
Earlobe histiocytes are not pathognomonic for bacterial endocarditis. If one uses strict criteria and requires at least 10% of the differential count to be these unusual cells, the differential diagnosis also includes malaria and trypanosomiasis.
Differential Diagnosis of Lower Counts of Histiocytes
Large histiocytes (also called macrophages) may constitute 2% of the differential count in septicemia, rheumatic fever with active carditis, tuberculosis, Hodgkin disease, chronic sinusitis, subsiding hepatitis, mediastinal tumor, agranulocytosis, localized bacterial infection (Smith, 1964
), and systemic lupus erythematosus.
If one accepts 1% histiocytosis as evidence of a positive sign, one will start to include perinephric abscess, infectious mononucleosis, mastoiditis, ulcerative colitis, subsiding acute appendicitis, trichinosis, acute and chronic myelocytic leukemia, typhoid fever, paroxysmal nocturnal hemoglobinuria, cholera, severe hemolysis, transfusion reactions, chronic meningococcemia, sickle cell anemia (Greenberg, 1964
), and anemia of the newborn (erythroblastosis fetalis) (Hill and Bayrd, 1960
; Smith, 1964
Leukocytes Containing Bacteria
In patients with septicemia, a low-power scan of a thin blood film prepared from the first drop of earlobe blood and stained with Gram stain (or Jenner-Giemsa, Leishman, or May-Grünwald-Giemsa) may reveal intracellular organisms. The ear that the patient has not been lying on gives better results. The feathered edge of the smear is the best place to find the infected leukocytes.
Gram Stain of the Buffy Coat
Gram stain of the buffy coat is an incredibly simple technique (see later in this chapter for the Gram stain) that permits the identification of bacteria in the blood about 24 to 48 hours before the laboratory reports a positive culture, and rarely, in cases with negative culture results (Humphrey, 1944
). See Table 28.2
for the diagnostic value of this technique (which has also been applied to the peripheral blood film and blood aspirated from skin lesions) in various studies. No studies have yet been reported giving a comparison of buffy coat staining with the earlobe histocyte test performed in the same patients.
TABLE 28.2 Results of studies examining blood film or buffy coat for bacteria
Predictive value of a negative test
Brooks et al. (1973)
Buffy coat examined on 135 specimens sent for blood culture
Powers and Mandell (1974)
Buffy coat examined in 16 patients suspected of having endocarditis (6 of whom did not) and 6 normal controls
Earlobe blood of septicemic patients
Examination of peripheral blood film in rapidly fatal cases of meningococcemia
Hoefs and Runyon (1985)
Ascitic buffy coat in spontaneous bacterial peritonitis
Bush and Bailey (1944)
Examination of peripheral blood film in rapidly fatal cases of meningococcemiaa
McLean and Caffey (1931)
Examination of blood film obtained from hemorrhagic purpuric lesion in meningococcemia
a The patients with organisms seen on peripheral smear all had negative blood cultures. The cells containing the Gram-negative diplococci superficially resembled basophils.
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