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1 Laboratory Errors in Pre-Analytical, Analytical, and Post-Analytical Stages

1 Laboratory Errors in Pre-Analytical, Analytical, and Post-Analytical Stages

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Ionized Calcium

Analysis................. 43



Table 3.1 Common Laboratory Errors



Key Points ............. 44



Type of Error



References ............ 45



Pre-Analytical Errors

Tube filling error

Patient identification error

Inappropriate container

Empty tube

Order not entered in laboratory information system

Specimen collected wrongly from an infusion line

Specimen stored improperly

Contamination of culture tube

Analytical Errors

Inaccurate result due to interference

Random error caused by the instrument

Post-Analytical Errors

Result communication error

Excessive turnaround time due to instrument downtime



Failure at any of these steps can result in an erroneous or misleading laboratory result, sometimes with adverse outcomes. The analytical part of the analysis involves measurement of the concentration of the analyte corresponding

to its “true” level (as compared to a “gold standard” measurement) within a

clinically acceptable margin of error (the total acceptable analytical error,

TAAE). Errors can occur at any stage of analysis (pre-analytical, analytical, and

post-analytical). It has been estimated that pre-analytical errors account for

more than two-thirds of all laboratory errors, while errors in the analytical

and post-analytical phases account for only one-third of all laboratory errors.

Carraro and Plebani reported that, among 51,746 clinical laboratory analyses

performed in a three-month period in the author’s laboratory (7,615 laboratory orders, 17,514 blood collection tubes), clinicians contacted the laboratory regarding 393 questionable results out of which 160 results were

confirmed to be due to laboratory errors. Of the 160 confirmed laboratory

errors, 61.9% were determined to be pre-analytical errors, 15% were analytical

errors, while 23.1% were post-analytical errors [1]. Types of laboratory errors

(pre-analytical, analytical, and post-analytical) are summarized in Table 3.1.

In order to avoid pre-analytical errors, several approaches can be taken,

including:





The use of hand-held devices connected to the LIS that can objectively

identify the patient by scanning a patient attached barcode, typically a

wrist band.



3.2 Order of Draw of Blood Collection Tubes









Retrieval of current laboratory orders from the LIS.

Barcoded labels are printed at the patient’s side, thus minimizing the

possibility of misplacing the labels on the wrong patient samples.



When classifying sources of error, it is important to distinguish between cognitive errors (mistakes), which are due to poor knowledge or judgment, and

non-cognitive errors (commonly known as slips and lapses), which are due to

interruptions in a process during even routine analysis involving automated

analyzers. Cognitive errors can be prevented by increased training, competency evaluation, and process aids (such as checklists); non-cognitive errors

can be reduced by improving the work environment (e.g. re-engineering to

minimize distractions and fatigue). The vast majority of errors are noncognitive slips and lapses performed by the personnel directly involved in

the process. These can be easily avoided.

The worst pre-analytical error is incorrect patient identification where a physician may act on test results from the wrong patient. Another common error

is blood collection from an intravenous line that may falsely increase test

results for glucose, electrolytes, or a therapeutic drug due to contamination

with infusion fluid.



CASE REPORT

A 59-year-old woman was admitted to the hospital due to

transient ischemic heart attack. During the first day of hospitalization she experienced generalized tonic-clonic seizure

and a 1000 mg intravenous phenytoin-loading dose was

administered followed by an oral dose of 100 mg of phenytoin

every three hours for a total of three doses. For the next five

days, the patient received 100 mg phenytoin intravenously or

orally every 8 hours. On the evening of Day 5 she received

two additional 300 mg doses of phenytoin intravenously.

Beginning with Day 7 the dose was 100 mg intravenously



every 6 hours. On Day 5, phenytoin concentration was

17.0 µg/mL and on Day 7 phenytoin concentration was

13.4 µg/mL. Surprisingly on Day 8, phenytoin concentration

was at life-threatening level of 80.7 µg/mL, although the

patient did not show any symptom of phenytoin toxicity.

Another sample drawn 7 hours later showed a phenytoin

level of 12.4 µg/mL. It was suspected that a falsely elevated

serum phenytoin level was due to drawing of the specimen

from the same line through which the intravenous phenytoin

was administered [2].



3.2 ORDER OF DRAW OF BLOOD COLLECTION

TUBES

The correct order of draw for blood specimens is as follows:











Microbiological blood culture tubes (yellow top).

Royal blue tube (no additive); trace metal analysis if desired.

Citrate tube (light blue).

Serum tube (red top) or tube with gel separator/clot activator (gold top

or tiger top).



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Heparin tube (green top).

EDTA tube (ethylenediamine tetraacetic acid; purple/lavender top).

Oxalate-fluoride tube (gray top).



Tubes with additives must be thoroughly mixed by gentle inversion as per

manufacturer-recommended protocols. Erroneous test results may be

obtained when the blood is not thoroughly mixed with the additive. When

trace metal testing on serum is ordered, it is advisable to use trace element

tubes. Royal-Blue Monojects Trace Element Blood Collection Tubes are

available for this purpose. These tubes are free from trace and heavy metals.



3.3 ERRORS WITH PATIENT PREPARATION

There are certain important issues regarding patient preparation for obtaining

meaningful clinical laboratory test results. For example, glucose testing and

lipid panel must be done after the patient fasts overnight. Although cholesterol concentration is not affected significantly by meals, after meals chylomicrons are present in serum that can significantly increase the triglyceride

level.

Physiologically, blood distribution differs significantly in relation to body

posture. Gravity pulls the blood into various parts of the body when recumbent, and the blood moves back into the circulation, away from tissues,

when standing or ambulatory. Blood volume of an adult in an upright

position is 600À700 mL less than when the person is lying on a bed, and

this shift directly affects certain analytes due to dilution effects. Therefore,

concentrations of proteins, enzymes, and protein-bound analytes (thyroidstimulating hormone (TSH), cholesterol, T4, and medications like warfarin)

are affected by posture; most affected are factors directly influencing hemostasis, including renin, aldosterone, and catecholamines. It is vital for laboratory requisitions to specify the need for supine samples when these

analytes are requested. Several analytes show diurnal variations, most

importantly cortisol and TSH (Table 3.2). Therefore, the time of specimen

collection may affect test results.



3.4 ERRORS WITH PATIENT IDENTIFICATION

AND RELATED ERRORS

Accurate patient and specimen identification is required for providing ordering clinicians with correct results. Regulatory agencies like The Joint

Commission (TJC) have made it a top priority in order to ensure patient



3.4 Errors with Patient Identification and Related Errors



Table 3.2 Common Analytes that show Diurnal Variation

Analyte



Comment



Cortisol

Renin

Iron

TSH

Insulin

Phosphate

ALT



Much higher concentration in the morning than afternoon

Maximum activity early morning, minimum in the afternoon

Higher levels in the morning than afternoon

Maximum level 2 AMÀ4 AM while minimum level 6 PMÀ10 PM

Higher in the morning than later part of the day

Lowest in the morning, highest in early afternoon

Higher level in the afternoon than morning



Abbreviations: TSH, Thyroid stimulating hormone; ALT, Alanine aminotransferase.



safety. Patient and specimen misidentification occurs mostly during the preanalytical phase:













Accurate identification of a patient requires verification of at least two

unique identifiers from the patient and ensuring that those match the

patient’s prior records.

If a patient is unable to provide identifiers (i.e. neonate or a critically ill

patient) a family member or nurse should verify the identity of the

patient.

Information on laboratory requisitions or electronic orders must also

match patient information in their chart or electronic medical record.

Specimens should not be collected unless all identification discrepancies

have been resolved.



The specimens should be collected and labeled in front of the patient and

then sent to the laboratory with the test request. Non-barcoded specimens

should be accessioned, labeled with a barcode (or re-labeled, if necessary),

processed (either manually or on an automated line), and sent for analysis.

Identification of the specimen should be carefully maintained during centrifugation, aliquoting, and analysis. Most laboratories use barcoded labeling

systems to preserve sample identification. Patient misidentification can have

a serious adverse outcome on a patient, especially if the wrong blood is

transfused to a patient due to misidentification of the blood specimen sent

to the laboratory for cross-matching. In this case a patient could die from

receiving the wrong blood group.

Although errors in patient identification occur mostly in the pre-analytical

phase, errors can also occur during the analytical and even post-analytical

phases. Results from automated analyzers are electronically transferred to the

LIS through an interface, but if direct transfer of the result from a particular

instrument is not available, errors can occur during manual transfer of the



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results. Dunn and Morga reported that, out of 182 specimen misidentifications they studied, 132 misidentifications occurred in the pre-analytical stage.

These misidentifications were due to wrist bands labeled for wrong patient,

laboratory tests ordered for the wrong patient, selection of the wrong medical

record from a menu of similar names and social security numbers, specimen

mislabeling during collection associated with batching of specimens and

printed labels, misinformation from manual entry of laboratory forms, failure of two-source patient identification for clinical laboratory specimens, and

failure of two-person verification of patient identity for blood bank specimens. In addition, 37 misidentification errors during the analytical phase

were associated with mislabeled specimen containers, tissue cassettes, or

microscopic slides. Only 13 events of misidentification occurred in the postanalytical stage; this was due to reporting of results into the wrong medical

record and incompatible blood transfusions due to failure of two-person verification of blood products [3].



CASE REPORT

A 68-year-old male presented to the hospital with sharp

abdominal pain. The patient underwent an appendectomy

and received one unit of type A blood. The patient developed

disseminated intravascular coagulation and died 24 hours

after receiving the transfusion. Postmortem analysis of the



patient’s blood revealed that he was actually type O. The

patient had been sharing a room with another patient whose

blood was type A. The specimen sent to the blood bank had

been inappropriately labeled [4].



Delta checks are a simple way to detect mislabels. A delta check is a process

of comparing a patient’s result to his or her previous result for any one

analyte over a specified period of time. The difference or “delta,” if outside

pre-established rules, may indicate a specimen mislabel or other preanalytical error.



3.5 ERROR OF COLLECTING BLOOD IN WRONG

TUBES: EFFECT OF ANTICOAGULANTS

Blood specimens must be collected in the right tube in order to get accurate

test results. It is important to have the correct anticoagulant in the tube (different anticoagulant tubes have different colored tops). Anticoagulants are

used to prevent coagulation of blood or blood proteins to obtain plasma or

whole blood specimens. The most routinely used anticoagulants are ethylenediamine tetraacetic acid (EDTA), heparin (sodium, ammonium, or lithium

salts), and citrates (trisodium and acid citrate dextrose). In the optimal



3.5 Error of Collecting Blood in Wrong Tubes: Effect of Anticoagulants



anticoagulant, blood ratio is essential to preserve analytes and prevent clot or

fibrin formation via various mechanisms. Proper anticoagulants for various

tests are as follows:



















Potassium ethylenediamine tetraacetic acid (EDTA; purple top tube) is

the anticoagulant of choice for complete blood count (CBC).

EDTA is also used for blood bank pre-transfusion testing, flow cytometry,

hemoglobin A1C, and most common immunosuppressive drugs such as

cyclosporine, tacrolimus, sirolimus, and everolimus; another

immunosuppressant, mycophenolic acid, is measured in serum or

plasma instead of whole blood.

Heparin (green top tube) is the only anticoagulant recommended

for the determination of pH blood gases, electrolytes, and ionized

calcium. Lithium heparin is commonly used instead of sodium

heparin for general chemistry tests. Heparin is not recommended

for protein electrophoresis and cryoglobulin testing because of the

presence of fibrinogen, which co-migrates with beta-2 monoclonal

proteins.

For coagulation testing, citrate (light blue top) is the appropriate anticoagulant.

Potassium oxalate is used in combination with sodium fluoride and

sodium iodoacetate to inhibit enzymes involved in the glycolytic

pathway. Therefore, the oxalate/fluoride (gray top) tube should be used

for collecting specimens for measuring glucose levels.



Although lithium heparin tubes are widely used for blood collection for

analysis of many analytes in the chemistry section of a clinical laboratory, a

common mistake is to collect specimens for lithium analysis in a lithium

heparin tube. This can cause clinically significant falsely elevated lithium

values that may confuse the ordering physician.



CASE REPORT

A healthy 15-month-old female was brought in by her mother

after ingesting an unknown amount of nortriptyline and lithium carbonate at an undetermined time. The mother reported

that the patient had vomited after ingestion. Vital signs were

normal. The patient was lethargic but easily aroused, and the

physical examination was unremarkable. Initial ECG was also

normal for age. The initial lithium level in the serum was

1.4 mEq/L, and a nortriptyline concentration of 36 ng/mL

indicated that none of the drug level was in a toxic region.

The patient was treated with activated charcoal, but 13 hours



after admission her serum lithium concentration was elevated

to 3.1 mEq/L. The patient was given l mg/kg oral sodium

polystyrene sulfonate, the rate of IV fluids was doubled, and

the patient was started on an IV dopamine infusion.

However, at 15 h her serum lithium level was 1.6 mEq/L.

Review of her records revealed that the specimen was

collected in a lithium heparin tube. A 19-hour serum

lithium concentration was 0.6 mEq/L, and the patient was

discharged within 24 h after admission without further incident [5].



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3.6 ISSUES WITH URINE SPECIMEN COLLECTION

Urinalysis remains one of the key diagnostic tests in the modern clinical laboratory, and, as such, proper timing and collection techniques are important.

Urine is essentially an ultrafiltrate of blood. Examination of urine may take

several forms: microscopic, chemical (including immunochemical), and electrophoresis. Three different timings of collection are commonly encountered.

The most common is the random or “spot” urine collection. However, if it

would not unduly delay diagnosis, the first voided urine in the morning is

generally the best sample. This is because the first voided urine is generally

the most concentrated and contains the highest concentration of sediment.

The third timing of collection is the 12- or 24-hour collection. This is the preferred technique for quantitative measurements, such as for creatinine, electrolytes, steroids, and total protein. The usefulness of these collections is

limited, however, by poor patient compliance.

For most urine testing, a clean catch specimen is optimal, with a goal of collecting a “midstream” sample for testing. In situations where the patient cannot provide a clean catch specimen, catheterization is another option, but

must be performed only by trained personnel. Urine collection from infants

and young children prior to toilet training can be facilitated through the use

of disposable plastic bags with adhesive surrounding the opening.

For point of care urinalysis (e.g. urine dipstick and pregnancy testing) any

clean and dry container is acceptable. Disposable sterile plastic cups and

even clean waxed paper cups are often employed. If the sample is to be sent

for culture, the specimen should be collected in a sterile container. For routine urinalysis and culture, the containers should not contain preservative.

For specific analyses, some preservatives are acceptable. The exception to this

is for timed collections where hydrochloric acid, boric acid, or glacial acetic

acid is used as a preservative.

Storage of urine specimens at room temperature is generally acceptable for

up to two hours. After this time the degradation of cellular and some chemical elements becomes a concern. Likewise bacterial overgrowth of both pathologic as well as contaminating bacteria may occur with prolonged storage at

room temperature. Therefore, if more than two hours will elapse between

collection and testing of the urine specimen, it must be refrigerated.

Refrigerated storage for up to 12 hours is acceptable for urine samples destined for bacterial culture. Again, proper patient identification and specimen

labeling is important to avoid errors in reported results.



3.7 ISSUES WITH SPECIMEN PROCESSING AND

TRANSPORTATION

After collection, specimens require transportation to the clinical laboratory. If

specimens are collected in the outpatient clinic of the hospital and analyzed



3.8 Special Issues: Blood Gas and Ionized Calcium Analysis



in the hospital laboratory, transportation time may not be a factor. However,

if specimens are transported to the clinical laboratory or a reference laboratory, care must taken in shipping specimens. Ice packs or cold packs are especially useful for preserving specimens at lower temperatures because analytes

are more stable at lower temperature. Turbulence during transportation, such

as transporting specimens in a van to the main laboratory, can even affect

concentrations of certain analytes.

Many clinical laboratory tests are performed on either serum or plasma.

Due to the instability of certain analytes in unprocessed serum or plasma,

separation of serum or plasma from blood components must be performed

as soon as possible, and definitely within two hours of collection.

Appropriate preparation of specimens prior to centrifugation is required to

ensure accurate laboratory results. Serum specimens must be allowed ample

time to clot prior to centrifugation. Tubes with clot activators require sufficient mixing and at least 30 minutes of clotting time, Plasma specimens

must be mixed gently according to manufacturer’s instructions to ensure

efficient release of additive/anticoagulant.



3.8 SPECIAL ISSUES: BLOOD GAS AND IONIZED

CALCIUM ANALYSIS

Specimens collected for blood gas determinations require special care, as the

analytes are very sensitive to time, temperature, and handling. In standing

whole blood samples, pH falls at a rate of 0.04À0.08/hour at 37 C,

0.02À0.03/hour at 22 C, and ,0.01/hour at 4 C. This drop in pH is concordant with decreased glucose and increased lactate. In addition, pCO2

increases around 5.0 mmHg/hour at 37 C, 1.0 mmHg/hour at 22 C, and

0.5 mmHg/hour at 4 C. At 37 C, pO2 decreases by 5À10 mmHg/hour, but

only 2 mmHg/hour at 22oC. Ideally, all blood gas specimens should be measured immediately and never stored. A plastic syringe transported at room

temperature is recommended if analysis will occur within 30 minutes of collection, but a glass syringe should be used if more than 30 minutes are

needed prior to analysis and specimens are stored in ice. Bubbles must

be completely expelled from the specimen prior to transport, as the pO2 will

be significantly increased and pCO2 decreased within 2 minutes [6].

Blood gas analyzers re-heat samples to 37 C for analysis to recapitulate physiological temperature. However, for patients with abnormal body temperature, either hyperthermia due to fever, or induced hypothermia in patients

undergoing cardiopulmonary bypass, a temperature correction should be

made to determine accurate pH, pO2, and pCO2 results.

Ionized calcium is often measured with ion-sensitive electrodes in blood gas

analyzers. Ionized calcium is inversely related to pH: decreasing pH decreases

albumin binding to calcium, thereby increasing free, ionized calcium.



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Therefore, specimens sent to the lab for ionized calcium determinations

should be handled with the same caution as other blood gas samples since

pre-analytical errors in pH will impact ionized calcium results [7].



KEY POINTS



































Errors in the clinical laboratory can occur in pre-analytical, analytical, or postanalytical steps. Most errors (almost two-thirds of all errors) occur in pre-analytical

steps.

During specimen collection, a patient must be identified by matching at least two

criteria. Blood should be collected in the correct tube following the correct order

of draw.

Correct order of drawing blood: (1) microbiological blood culture tubes (yellow

top), (2) royal blue tube (no additive) if trace metal analysis is desired, (3) citrate

tube (light blue), (4) serum tube (red top) or tube with gel separator/clot activator

(gold top or tiger top), (5) heparin tube (green top), (6) EDTA tube (purple/lavender

top), and (7) oxalate-fluoride tube (gray top).

Proper centrifugation (in the case of analyzing serum or plasma specimens) and

proper transportation of the specimen to the laboratory are required, as well as

maintaining proper storage of the specimen prior to analysis in order to avoid

artifactual changes in the analyte.

EDTA (purple top tube) is the anticoagulant of choice for the complete blood

count (CBC). The EDTA tube is also used for blood bank pre-transfusion testing,

flow cytometry, hemoglobin A1C, and most common immunosuppressive drugs

such as cyclosporine, tacrolimus, sirolimus, and everolimus; another

immunosuppressant, mycophenolic acid, is measured in serum or plasma instead

of whole blood.

Heparin (green top tube) is the only anticoagulant recommended for the

determination of pH blood gases, electrolytes, and ionized calcium. Lithium

heparin is commonly used instead of sodium heparin for general chemistry tests.

Heparin is not recommended for protein electrophoresis and cryoglobulin testing

because of the presence of fibrinogen, which co-migrates with beta-2 monoclonal

proteins.

For coagulation testing, citrate (light blue top) is the appropriate anticoagulant.

Potassium oxalate is used in combination with sodium fluoride and sodium

iodoacetate to inhibit enzymes involved in the glycolytic pathway. Therefore the

oxalate/fluoride (gray top) tube should be used for collecting specimens for

measuring glucose level.

Ideally, all blood gas specimens should be measured immediately and never

stored. A plastic syringe, transported at room temperature, is recommended if

analysis will occur within 30 minutes of collection. Otherwise, a specimen must be

stored in ice. Glass syringes are recommended for delayed analysis because glass



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