The immuassay handbook parte45


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The immuassay handbook parte45

  2. 2. 427© 2013 David G. Wild. Published by Elsevier Ltd. All rights reserved. Sample Collection, Including Participant Preparation and Sample Handling Colin Wilde1 Dorothée Out2 Sara Johnson2 Douglas A. Granger2 ( 1Previous edition. 2Revisions for this edition, including comprehensive new section on Oral Fluid. Although great emphasis is placed on the accuracy and precision of immunoassays, some of the largest potential sources of error concern sample collection, handling meth- ods, and the way the subject is prepared before the sample is taken. Unfortunately, this critical area is often neglected in laboratory investigations and overlooked in quality con- trol procedures and troubleshooting investigations. This chapter describes collection methods for various types of samples, together with some of the types of prob- lems to which they are susceptible. Examples are given to enable the reader to be aware of the types of difficulties that may arise, the impact on the interpretation of results, and an understanding of how to avoid them. A compre- hensive listing of published references can be found in Effects of Preanalytical Variables on Clinical Laboratory Tests by D. S. Young (2007). Participant State and Preparation Certain aspects of the participant that may affect the inter- pretation of a test result are not under the control of the clinician or laboratory staff. These include age, sex, ethnic origin, health status or diagnoses, physiological states such as pregnancy and stage of menstrual cycle, and they must be well documented at the sample collection stage so that the test results may be interpreted accordingly. There are other variables, however, which often require active inter- vention and control if test results are to be meaningful. STRESS The stress response involves acute and chronic changes in activity of the central nervous system, the hypothalamic– pituitary–adrenal axis and the parasympathetic and sympa- thetic branches of the autonomic nervous system (ANS), and the immune system. Secretory products of these systems, such as glucocorticoids, neurotransmitters, and cytokines, bind to receptors and affect the function of a variety of cells and physiological systems. Stress, whether mental or physical, can alter bodily functions, including secretions into the body fluids that are to be tested. For standard laboratory purposes, the participant should not therefore be anxious or tense when samples are collected but should be made to feel relaxed and at ease in a comfort- able environment. Fear and stress are potent stimuli of growth hormone (GH), prolactin, cortisol, catecholamines, cytokines, dehydroepiandrosterone (DHEA), aldosterone, and plasma renin activity. Many other intermediary metab- olites, and also carrier proteins such as transferrin, may be affected by longer term stress, and after a major stressful event, such as myocardial infarction, the results of assays of such analytes should be interpreted accordingly. Postsurgical stress can be particularly severe, and immu- noassay tests should be avoided if possible until the patient has stabilized. For example, transferrin can fall after about 3h, and ferritin starts to rise shortly afterward. Thyroid hormone levels are also often depressed after surgery. EXERCISE Physical exercise stimulates the production and secretion of a number of hormones including GH, prolactin, lactate, cortisol, testosterone, and plasma renin. The extent of the increase depends on the amount of exercise taken and on the physical fitness of the individual. When resting-state levels of affected hormones are required, it is important to ensure that the subject has not recently exercised, including running up the stairs to the collection center immediately before sample collection. Physical challenge, however, may be used in stimulation tests to assess a patient’s reserve capacity for hormone production, during which controlled exercise is undertaken immediately before sample collection as, for instance, in the assessment of GH deficiency. FOOD AND DRINK Dietary state may also be relevant to the investigation required. Many of the commonly measured constituents of plasma vary in concentration according to the time elapsed since the last meal, but these changes have been quantified for only a few analytes normally measured by immunoas- say. Serum insulin, gastrin, and calcitonin are examples of hormones with levels that significantly alter following food intake, and fasting samples are required unless a stimula- tion test is being done. The levels of circulating therapeu- tic drugs are also influenced by the timing of meals, because of slowing of the drug absorption from the gut. Caffeine from coffee, tea, and soft drinks has a strong effect on some analytes and can increase plasma cortisol levels by up to 50% after 3h. Malnutrition results in a lowering of levels of insulin- like growth factor-1 (IGF1), albumin, caeruloplasmin, C H A P T E R 6.1
  3. 3. 428 The Immunoassay Handbook transferrin, and prolactin, indeed IGF1 and albumin are often used for monitoring nutritional status. Lipids There are well-documented changes in lipids after a fatty meal, when the blood serum or plasma may have a milky appearance due to the presence of triglycerides and chylo- microns. Such hyperlipidemic serum may interfere with antibody binding in immunoassay procedures and, although not always practicable, it is desirable to collect specimens from subjects after a period of fasting, usually overnight, for any type of investigation. Alternatively, ultracentrifugation or enzymatic cleavage is sometimes used to remove or break down lipids. Alcohol Alcohol ingestion induces changes to body fluid composi- tion that vary depending on whether the subject is an abuser or casual drinker and on the timing of collection after alcohol intake. Examples are ferritin and the liver enzymes alkaline phosphatase, aspartate aminotransferase, and γ-glutamyl transferase, although the latter are seldom measured in immunoassays. Smoking Smoking can influence the levels of a number of analytes commonly measured by immunoassay. Both cortisol and GH rise as an acute response to smoking, and it is advis- able not to take blood within 30min of smoking. Long- term effects are seen as increases in IgE, androstenedione, insulin, C-peptide, placental alkaline phosphatase, and carcinoembryonic antigen, which can give false positives for the latter two tumor markers. Significantly lower levels of IgG and prolactin can be expected in smokers. The metabolism of some drugs, e.g., theophylline and tricyclic antidepressants, is stimulated by smoking, result- ing in reduced half-life and increased body clearance. In pregnant smokers, the levels of human chorionic gonadotropin (hCG) and estradiol are significantly lower than those in nonsmokers. POSTURE When an individual assumes an upright position after a period of recumbency, there is movement of ultrafiltrate from the intravascular to the extravascular compartment of the extracellular fluid. This produces a 10–20% hemo- concentration with a concomitant concentration of large molecules and those substances that are bound to them. Plasma concentrations of proteins, peptides, enzymes, and protein-bound substances, such as cortisol, thyrox- ine, and drugs, are affected. When patients rise from a sitting position, similar changes occur but to a lesser degree. For ambulatory patients, blood should be collected after the subject has been seated for 15min, but results from seated subjects are not directly comparable to those from hospital in-patients whose blood is often collected after prolonged recumbency. The renin–aldosterone–angiotensin system is strongly influenced by posture, and it is normally recommended that blood samples are taken after overnight recumbency, without sitting or standing before collection. Even a short period of sitting will produce significant increases in aldo- sterone. Measurement before and after 4h of ambulation may be used to differentiate between adrenal hyperplasia and adrenal adenoma. MEDICAL PROCEDURES Some medical procedures have short-term effects on levels of circulating analytes, and it is important to allow suffi- cient time to elapse after such procedures before sample collection. Surgery or intramuscular injection results in increased creatine kinase activity, and rectal examination or pros- tatic manipulation may cause an increase in circulating prostate-specific antigen (PSA). Care must be taken not to collect blood proximate to a site of intravenous injection as the concentrations of many components are likely to be misleadingly low. Samples should never be taken from the arm in which a drip has been inserted because of the dilution effect on blood constituents. Transfusion results in analyte concentrations that are a composite between recipient and donor. Also folate and ferritin concentrations may be increased. It is therefore preferable to avoid taking samples for testing for a few days after the transfusion if possible. DRUGS Drugs may cause difficulties in the interpretation of immu- noassay results through their effect on in vivo physiological and biochemical mechanisms or by in vitro effects in the analytical process. There are now many thousands of ref- erences in the literature testifying to these problems and over 80% of these relate to the in vivo type. Drug Interactions Oral contraceptives have a profound effect through their estrogenic activity, leading to increased levels of many binding proteins, including those for thyroxine, cortisol, and the sex hormones (sex hormone-binding globulin, SHBG). Other drugs, such as barbiturate and phenytoin, cause hepatic enzyme induction with increased levels of liver enzymes. Amiodarone is one of the more well-known drugs that interfere with thyroid hormone levels, raising the level of thyrotropin (TSH) and thyroxine. A novel type of interaction is the one caused by high concentrations of β-lactam antibiotics (penicillins and cephalosporins), which inactivate aminoglycosides both in vivo and in vitro. Samples for aminoglycoside assay that contain β-lactams should therefore either be assayed immediately or stored frozen. The list of drug interactions is so large that it may only be accommodated on constantly updated computer data banks. Although nobody can be expected to know more than a small fraction of the documented drug–test interac- tions, it is important to record all drug therapy at the time
  4. 4. 429CHAPTER 6.1 Sample Collection, Including Participant Preparation and Sample Handling of the request or sample collection so that anomalous results may be checked by reference to drug–test interac- tion data banks. It is, of course, impractical to cease drug therapy before sample collection for all tests, but for some, it is necessary if the results are to be interpreted without ambiguity. For instance, because the levels of aldosterone and renin vary inversely with the activity of the sympathetic nervous sys- tems, diuretic, and hypertensive drugs must not be taken for 3 weeks before analysis if meaningful results are to be obtained. PREGNANCY The effects of pregnancy on a number of analytes are well documented elsewhere in this book. In addition to the well- known rises in hCG, estriol, human placental lactogen, etc. (see PREGNANCY), the serum concentrations of corticotropin- releasing hormone (CRH), cortisol, and binding proteins for glucocorticoids and thyroid hormones also rise causing the total concentrations of these hormones to increase. AGE Changes occur in the levels of many components with aging; some of these are gradual, others occur quite rapidly at certain times of life such as the neonatal period, puberty, and the menopause. Different reference ranges must be used for different age groups. Thus, TSH and free thyroxine levels peak at birth but settle to normal levels within 6 days. The concentration of 17-hydroxyprogesterone is very high at term, due to enzy- mic inhibition by placental steroids, but levels decline to normal within 48h. Vitamin D levels are low at birth but rise in the first 2 days, this corresponds to an increase in parathyroid hormone (PTH) that often occurs in response to early hypocalcemia. At adrenarche (DHEA, androstenedione) and puberty (testosterone, estrogen, progesterone), there are well- known changes in sex hormone levels, but this is also the time of peak levels of IGF1 and of GH and prolactin in females. The menopause in women results in increases in follicle- stimulating hormone (FSH) and luteinizing hormone (LH) followed by a postmenopausal lowering of estradiol. Bone markers generally change with aging with steeper changes at the menopause in women; this is particularly true of osteocalcin levels. Thus, there are increases in PTH and urinary pyridinoline and decreases in 25-hydroxy vitamin D and calcitonin. In midlife, there are decreases in testosterone in men and an increase in PSA that is due to physiological benign prostatic hyperplasia. RACE & ETHNICITY There are ethnic differences for some analytes, the most notable being the lower levels of 25-hydroxy vitamin D and osteocalcin levels in some ethnic/racial minority popula- tions and higher levels of PSA in African and West Indian men. Asian-Pacific populations do not show the age-related increases in PSA described in the above section on AGE. Timing BIOLOGICAL RHYTHMS Changes that follow well-defined rhythms (diurnal, circa- dian, ultradian) occur in a number of biological systems. Common rhythms relating to biochemical substances are the menstrual cycle, with a periodicity of about 28 days in female humans (i.e., progesterone, estradiol) and diurnal or circadian rhythms (e.g., melatonin, cortisol), with a periodicity of about 24h. It is important therefore to understand the rhythmic patterns of any biological sub- stances measured by immunoassay and to time the collec- tion of samples carefully, making best use of this knowledge in the interpretation of the test results. It is also important to take samples at the same time in the rhythm if intra- or interindividual results are to be compared. Menstrual Cycle Female reproductive hormones, e.g., LH, FSH, estradiol, and progesterone, follow a monthly cyclical pattern, and it is important to know the timing in relation to the men- strual cycle and to have a series of two or more samples at known times in the cycle before any interpretation of the result is possible. Levels of sSHBG are significantly higher during the luteal phase as are those of interleukin- 1α, a finding which may be due to the rise in body tem- perature during this phase. Circadian Rhythms Of the circadian rhythms, it is probably the hormonal ones that have been most studied. Those of adrenocorticotropic hormone (ACTH) and corticosteroids are perhaps the most striking with the peak levels in body fluids occurring between 06:00 and 08:00 with a nadir at midnight in humans. In the rat, a nocturnal animal, this rhythm is almost exactly 12h out of phase with that of man. Prolactin, corti- costeroid, and aldosterone levels follow a similar pattern of building up during sleep, but a GH peak occurs earlier, dur- ing the first few hours of sleep, and unlike other hormones, the GH peak is abolished if the subject remains awake. A rhythm that is related to subject activity rather than photo- periodic effect occurs with plasma testosterone. This again normally peaks at 08:00 but with a much steadier fall during the day to a trough at 20:00. Interleukin-1α peaks early in the sleep cycle, being related to the onset of slow wave sleep, but perhaps the most striking circadian rhythm occurs with melatonin, which is almost entirely produced during the night with very low blood levels during the day. Rhythms may vary at different stages of life, and this is, of course, most pronounced with the female menstrual cycle, but a nocturnal rise in LH is characteristic of puberty in males. A loss of rhythm is often diagnostic of a disease pro- cess (e.g., cortisol in Cushing’s disease), but sometimes, the rhythm may become more marked, as with 17-hydroxypro- gesterone in congenital adrenal hyperplasia in neonates. Pulsatile Secretion Within the well-defined rhythms, there is sometimes a pronounced episodic or pulsatile secretion. This occurs
  5. 5. 430 The Immunoassay Handbook with ACTH and the corticosteroids where strong pulses occur about 12 times every 24h. Androstenedione, which shows a prominent circadian rhythm, also has marked epi- sodic secretions with 25% swings in circulating levels. LH is another hormone with pulsed secretions, but the peaks are much less pronounced than with corticosteroids. DYNAMIC TESTS Stimulation and suppression tests are often used to investi- gate the capacity of organs to respond to positive or nega- tive stimuli. These usually involve the collection of timed samples following the stimulus. For reliable interpreta- tion, it is important that the prescribed timings are strictly adhered to and that each sample is clearly labeled with the time of collection. The dose of the stimulating substance should always be recorded. PATHOLOGICAL CHANGES Timing is sometimes important in the confirmation of a pathological event. A good example is that of the creatine kinase MB isoenzyme where, for reliable diagnostic infor- mation to be obtained, a blood sample should be collected within a predetermined time interval (usually between 6 and 30h) following the suspected myocardial infarction. NON-THYROIDAL ILLNESS Severe illness or injury can induce changes in thyroid hor- mone levels. See THYROID. THERAPEUTIC DRUG MONITORING The timing of sample collection is especially important when measuring the circulating levels of therapeutic drugs. The appropriate timings during the dose interval vary for different drugs depending on their absorption and distri- bution characteristics. At the start of the therapy or follow- ing a change in dosage regimen, there is an initial absorption phase with the average drug concentration continuing to rise after repeated doses until the steady state condition is reached. This may not be until at least five half-lives and several dose intervals have elapsed. As a general rule, samples should be taken when the con- centration of drug is at the lowest point (or trough level), and this is usually immediately before the next dose. This trough level normally relates to the steady state concentra- tion. For digoxin, for instance, it is preferable to wait at least 8h after administration because this drug has slow accumulation and excretion rates. Procainamide is a drug that is absorbed slowly, causing the highest blood levels at the end of the dose interval. Ideally in such cases, a number of samples should be taken at intervals or, at the very least, one at the end and one 2h into the dose interval. With some drugs where toxic levels are close to thera- peutic levels, e.g., the aminoglycosides, it is recommended that both peak and trough concentrations within the dose interval are measured. This practice helps to prevent toxic- ity and ensure therapeutic efficiency. For similar reasons, theophylline is normally measured at the time of peak levels. It should also be remembered that food intake may delay the attainment of peak concentrations by slowing absorption. Blood Collection by Venepuncture PRECAUTIONS RELATING TO THE PATIENT Errors in interpretation of immunoassay results may occur if the sample is not collected correctly and under the best possible conditions. Taking a blood sample can be very stressful for the patient and stress causes major changes in some blood constituents (see SUBJECT STATE AND PREPARA- TION). Considerable attention must therefore be paid to the environment, to the facilities available, and to reassur- ing the patient during the procedure. Patient Identification Correct patient identification is obviously essential, but it is not unusual for errors to be made. The venepuncturist must always ensure that the blood specimen is being drawn from the patient designated on the request form and that it is placed in a correctly and unambiguously labeled container. The use of a single barcoded primary tube for sample collection and testing is well established now that automated immunoassay analyzers are fitted with barcode readers. The test request can be logged onto the main laboratory computer, which is connected to the analyzer via a two-way link. This avoids the risk of errors when transferring samples or test-request information. Position The patient must be positioned comfortably, preferably in a special phlebotomy chair with adjustable armrests or on a bed or examination couch. Patients should not be startled, such as by sudden wakening, because this may affect the levels of some analytes, as would sudden changes in posture, especially for investigations involving the renin–angiotensin–aldosterone axis (see SUBJECT STATE AND PREPARATION – POSTURE). Puncture Site There are a number of veins in the arm that may be used, but the larger median cubital and cephalic veins are used most frequently. The puncture site should be carefully chosen avoiding any area of hematoma and any area of extensive scarring. A specimen taken from the side on which a mastectomy has recently been done may not be truly representative because of lymphostasis; specimens should never be taken from an arm being used for intra- venous therapy because hemodilution is likely. Lowering the arm over the side of the armrest or bed will cause the veins to distend. Stroking in an upward direction usually makes the veins more visible. The veins become more prominent and easier to enter when the patient forms a fist, but vigorous hand exercise should not be allowed as
  6. 6. 431CHAPTER 6.1 Sample Collection, Including Participant Preparation and Sample Handling this may change some blood constituent levels. The puncture site should be cleaned with an isopropanol swab and wiped dry to minimize contamination of the specimen. Tourniquet A tourniquet is often applied to enhance venous filling, increase vein distention, and aid vein location. Its use is contraindicated when analytes affected by hemoconcen- tration are being measured, because the occlusion of the upper arm results in ultrafiltration of blood in the forearm to produce spurious increases in the concentration of large molecular mass substances and anything bound to them. Such analytes include proteins and peptides, bound com- ponents, cellular elements, and enzymes. If a tourniquet has to be used in such instances for preliminary vein selec- tion, it should be released and reapplied after an interval of at least 2min. The tourniquet must never be left on for longer than 1min immediately before venepuncture, and it should be removed as soon as the blood begins to flow, otherwise, hemoconcentration will occur, and local stasis is likely. PHLEBOTOMY TECHNIQUES For detailed descriptions of phlebotomy techniques, see CLSI document H3-A6 (see FURTHER READING). BLOOD COLLECTION Blood Collection Systems Blood collection may be achieved by the conventional needle and syringe method followed by transfer of the blood from the syringe to a suitable container or alterna- tively by using an evacuated tube system. Evacuated or vacuum tubes are manufactured to withdraw a predeter- mined volume of blood in a closed system. The needle used is double ended, one end pierces the stopper or dia- phragm that seals the tube, the other end is inserted into the vein. With these systems, there is no transfer from syringe to tube, and the blood comes into contact with anticoagulant or preservative immediately when it is drawn. The likelihood of hemolysis and microclot forma- tion is reduced, and there is less deterioration of metaboli- cally labile substances. If the investigation requires multiple sampling, as in stimulation or suppression tests, it is often helpful to insert a cannula which can stay in position for the duration of the investigation and allow blood to be withdrawn at intervals with minimum stress to the subject. However, great care must be exercised to avoid sample contamination with heparin, which may be used to flush the cannula. Common Types of Blood Container The main types of blood container available are listed in Table 1 below. The codes and colors are those used according to the advice of US and European Clinical Lab- oratory Standards Committees, although some countries use different codes and colors. Withdrawal of Blood The needle must be inserted carefully into the chosen vein bevel side up. The desired amount of blood is drawn either by withdrawing the syringe plunger or by allowing the blood to flow into the evacuation tube until the vacuum is exhausted. If additional blood is required, a further syringe or evacuated tube should be attached to the needle, which should remain in the vein. If the syringe method is used, the blood should be transferred to an appropriate con- tainer after separation from the needle and never through the needle as this causes hemolysis. Filling Tubes Containing Additives Whichever method is used, tubes containing anticoagulant must be filled to the mark, otherwise, the concentration of anticoagulant will be too high, and this may affect the assay system, particularly the antibody-binding characteristics. If several specimens are to be drawn at the same time, the plain nonadditive tube should always be filled first, with additive- containing tubes being filled later, the recommended order of fill being plain tube, citrate, heparin, ethylene diamine tet- raacetic acid (EDTA), and finally oxalate fluoride. Care must still be taken to avoid cross-contamination between different additive tubes as this may result in factitious test results. PREPARATION OF SERUM Blood serum is often preferred for immunoassay. For this the blood must be collected into a plain tube, and the serum separated from the blood cells by centrifuga- tion following clot formation and retraction, which may normally take up to 1 h. This time can be reduced using axial centrifugation where the blood collection tube is rotated about its own longitudinal axis, separating the blood into three concentric cylinders, cells against the tube walls, serum inside the cells, and air in the center. A separator may be introduced while the tube is spinning to permanently isolate the serum from the cells. The DuPont Axial Separation Technology system utilized this technique. TABLE 1 Blood Sample Containers: Anticoagulant Codes and Colors Anticoagulant Code Color Potassium EDTA KE Lavender Sodium EDTA NE Lavender Potassium oxalate KX Trisodium citrate 9NC* Blue 4NC* Black Fluoride oxalate FX Gray Ammonium and potassium oxalate AKX Lithium heparin LH Green Sodium heparin NH Green Acid citrate dextrose (ACD) ACD Yellow None Z Red *Figures denote ratio between blood and anticoagulant.
  7. 7. 432 The Immunoassay Handbook Clotting Activators Clotting activators are used to speed up the process to 10–15min, and these are present in many commercial blood-collecting devices. Minute silica or glass particles are commonly used. These may be introduced in the form of small beads or attached to the tube wall with a water-soluble silicone coating or to a “carrier” such as a paper disc or polypropylene cup. Thromboplastin has been substituted for glass particles when very rapid clotting is required. These materials accelerate the clotting process and help produce a clean, well-defined clot. They also diminish latent fibrin formation in the separated serum. If fibrin is allowed to form, it may interfere with the accuracy of pipetting devices or with the efficiency of solid-phase binding in the assay system. Serum produced using clot- ting activators is less likely to be hemolyzed although the clotting process itself may cause the release of some eryth- rocyte contents, including erythrocyte enzymes, which are usually present in higher concentrations in serum than in plasma. There is unlikely to be interference with immuno- assay from this source unless the enzymes themselves are being assayed using immunological techniques. Serum samples are not always suitable. For instance, samples for antithrombin III assay should be taken into the anticoagulant EDTA, otherwise, the analyte will be lost in the clotting process. The serum levels of thrombospondin are 100 times higher than plasma levels due to the release of this putative breast tumor marker from platelets during the clotting process. PREPARATION OF PLASMA Anticoagulants When plasma is required blood must be collected into an anticoagulant. Anticoagulants work by interfering with the clotting process and the principal ones in general use are heparin and EDTA salts. Immediate gentle mixing of the anticoagulant with the blood after phlebotomy is essential if clotting is to be prevented but too vigorous a treatment will cause hemolysis. Separation of the plasma from the cells by centrifugation may be effected immediately. If the samples are centrifuged for a long period or at too high a temperature, hemolysis will occur, and cell contents leak into the plasma. A speed equivalent to 1000–1500g for 5–10min, preferably at 20–25°C in a temperature-con- trolled centrifuge, is recommended. Some analytes require cooled centrifugation at 2–8 °C, e.g., ACTH. Axial cen- trifugation (see PREPARATION OF SERUM) may also be used for preparation of plasma. Interference by Anticoagulants If plasma is to be used for immunoassay, care must be taken to select an appropriate anticoagulant. Chelation of essential metal ions by EDTA may inhibit the enzyme activity at the signal generation stage in an enzyme immu- noassay, especially if alkaline phosphatase is used. Antico- agulants may also interfere with some antibody–antigen reactions. The use of heparin decreases the rate of reaction of some antibodies, particularly at the precipitation stage in second-antibody systems. Solid-phase systems and careful selection of antibodies have virtually eliminated, this problem but some assays still exhibit interference by anticoagulants, including heparin, and care must be taken to identify these and use appropriate specimens. Heparin should not be used for the investigation of cryoproteins because the anticoagulant precipitates cryofibrinogen. WHOLE BLOOD If whole blood is required for a laboratory assay, an antico- agulant is used to prevent coagulation. EDTA is often pre- ferred for this purpose because of its efficiency in preserving the integrity of the sample for blood cell investigations. In the laboratory, immunoassays are rarely carried out on whole blood but if they are, then the effect of anticoagula- tion on cellular components, as well as matrix effects on the immunological reaction, must be considered. In con- trast, recently introduced point-of-care devices utilize fresh whole capillary blood applied directly to the device following skin puncture. Interference by Tube and Stopper Components Soluble substances may be leached out of tubes and their closures, and in certain cases, the substances may pro- foundly affect the results of assay procedures. INTERFERENCE There is much literature concerning such problems, par- ticularly with the rubber stoppers used in some commercial blood collection devices. Plastics such as tris(2-butoxy- ethyl) cause displacement of some drugs and other analytes from protein-binding sites with consequent redistribution between erythrocytes and plasma. They may also strip antibody from coated solid-phase systems such as coated tubes, interfere with the binding of antigen and antibody and inhibit enzyme activity at the signal generation stage of an assay. There has been considerable concern regarding interference with drug–protein interactions, particularly with the lipophilic drugs bound to acidic α1-glycoprotein. PRECAUTIONS The manufacturers of blood collection tubes recognize these problems, and attempts have been made to reformu- late the closure material using specially selected low- extractable rubbers to minimize interference. It is, however, good practice always to fill the tubes to their designated volume so that any material leached out is not concentrated in a low volume. Ensure also that the length of time the blood is in contact with the stopper is minimal. Inversion of the tube to mix the contents should be done gently not more than five times and roller mixing of samples should be avoided. The tubes and samples should always be stored upright to minimize contact with the stopper. Storage in the tubes should be for no longer than 24h and should be at low temperature (2–8°C) because leach- ing is greater at higher temperatures. The tubes should
  8. 8. 433CHAPTER 6.1 Sample Collection, Including Participant Preparation and Sample Handling always be stored upright. Such precautions will reduce but not eliminate the problem, particularly if a poor batch of stoppers is being used. As it is almost impossible to predict interference with particular methodologies and because variation between batches is not uncommon, it is a wise precaution to test the batch of tubes before using them for a specific purpose, recognizing that a new analyte may invalidate the assump- tion that the system is appropriate for all assay purposes. It is also helpful to ask manufacturers for any data that they may hold and to consult references on the specific type of tube and stopper. It is also necessary to consider possible adsorption of the analyte on to the tube. This is particularly pertinent with ACTH which is readily adsorbed on to glass and for which plastic tubes must be used. The Use of Serum Separators Serum separators are frequently used to assist the rapid and efficient separation of cell-free serum from clotted whole blood, although they may also be used for the separation of plasma from the cells in anticoagulated blood. CHARACTERISTICS OF SEPARATORS Separators are usually silicone gels or polyester formula- tions, although glass beads and plastic and fiber devices have been used. They all essentially have a specific gravity that is intermediate between that of serum and that of the cells or the clot. During centrifugation, while the cell-clot coagulum settles to the bottom of the tube, the gel viscos- ity decreases, and the gel migrates to the surface of the clot. Following centrifugation, the original viscosity is restored, and the gel forms a nonpermeable barrier at the serum– clot interface. The separators are generally best used at 20–25°C because chilling may impair the flow characteris- tics and too high a temperature may cause breakdown of the gels as well as vitiation of the serum components. Provided the gel barrier is visually checked for integrity, it is generally safe to store the serum on the gel for up to 2–3 days without significant contamination from cellular components. The tubes should never be recentrifuged as this causes mixing of separated serum with serum that has remained in contact with the cells. INTERFERENCE Pieces of gel or droplets of oil may be seen on or within the separated serum with some batches of gel-containing tubes. There is evidence that this can interfere with liquid-sensing sampling devices and that it may coat tubes and cuvettes, possibly leading to physical interference with binding in solid-phase systems. Some laboratories routinely filter gel- separated serum before assay to prevent such contamina- tion. Although undoubtedly some batches of gel have posed particular problems, it is important to follow the manufac- turer’s instructions and not use the gels at excessive tem- peratures or inappropriate centrifuge speeds or to subject the tubes to rough handling or unusual orientations. Despite the usefulness of separators in helping to pro- vide a good clear serum specimen, they may interfere with the measurement of some analytes. Progesterone shows a time-related absorption or adsorption phenomenon with decreases of up to 50% in measurable hormone when stored over a gel for 6 days. Other hormones investigated at the same time showed no significant change. Lignocaine, pentobarbital, phenytoin, and carbamaze- pine are among the drugs that have shown similar reduc- tions in levels on some gels even after only a few hours’ storage. If there is any doubt about the storage character- istics of a particular analyte in gel-containing tubes, the serum should be decanted into a plain tube as soon as pos- sible after separation. OTHER ADDITIVES Some of the low-molecular mass polypeptide hormones such as ACTH, glucagon, gastrin, and other gastrointesti- nal hormones are rapidly destroyed by enzymes present in blood and may require protection by the addition of suit- able antiproteolytic agents such as Trasylol (aprotinin) to the tubes into which the blood is collected. Even when antiproteolytic agents are used, it is still necessary to cen- trifuge the sample at low temperature within 10min of col- lection and to store the serum or plasma immediately at −20°C, only thawing it out immediately before assay. If any transport is necessary to a distant laboratory, this must also be at −20°C. FUTHAN-EDTA should be used as a sample stabilizer for some complement components such as the labile C2 and C5–9, which additionally should be stored at −70°C. It must also be added to samples for com- plement breakdown products, e.g., C3d, otherwise, spuri- ously high levels will be recorded due to in vitro destruction of C3 on storage. HEMOLYSIS Noticeable hemolysis will interfere with antibody–antigen reactions and with some signal generation stages. Heavily contaminated samples should always be discarded and not used for immunoassay. Even small degrees of hemolysis may be unacceptable, principally due to the release of pro- teolytic enzymes, which destroy small peptides. Such pep- tides include insulin, glucagon, calcitonin, PTH, ACTH, and gastrin. Samples with any sign of hemolysis are not acceptable for such assays. Anomalous results may be obtained in other systems because of leakage of cellular components. Such an assay is serum folate where red cell values are approximately 30 times greater than those in serum. Collection of Blood by Skin Puncture Small but adequate amounts of capillary blood may be obtained by skin puncture. Such collection procedures are especially important in babies but are also often useful for collection from adults where multiple sampling is required, when venesection may constitute a hazard to the patient or
  9. 9. 434 The Immunoassay Handbook where no suitable superficial veins are available, for exam- ple, in gross obesity or in cases of severe burns. It must be remembered, however, that capillary blood is not the same as venous blood, indeed, it more closely resembles arterial blood, and the differences in composition must be consid- ered when interpreting test results. There is also a greater risk of hemolysis when capillary blood is taken. SKIN PUNCTURE SITES Usual sites for collecting blood by skin puncture are G the most lateral or most medial plantar heel surfaces (recommended in infants); G the medial plantar surface of the big toe; G the distal digit of a finger, preferably the third or fourth; G the earlobe. It is essential to obtain a good blood flow; otherwise, con- stituents may be diluted with tissue fluid. A similar dilution will occur if the skin puncture site is edematous or if excess pressure is exerted at the site in an attempt to increase the flow rate. Adequate flow rate is best obtained by covering the site with a warm, moist towel at a temperature no higher than 42°C for at least 3min. The site should be cleaned with an isopropyl alcohol swab and then thoroughly dried with a sterile gauze pad before being punctured. Residual alcohol causes rapid hemolysis. The skin is normally punctured with a special sterile lancet or puncture device that pene- trates to a depth of 2.4mm. Drops of blood should well out spontaneously without excessive rubbing or pressing. The first drop of blood must be discarded, as this is likely to contain excess tissue fluid. COLLECTION INTO CAPILLARY TUBES The blood may be conveniently collected into a capillary tube by allowing the tip of the tube to touch the drop forming over the puncture site. Blood will flow into the tube by capillary action. Capillary tubes with no additives or containing heparin or EDTA salts may be used depending on whether serum, plasma, or whole blood is required. The capillary tube or tubes are then plugged with sealing clay and, if anticoagulant is used, must be inverted gently at least 10 times to prevent coagulation and hemolysis. Mixing may also be done by placing a small magnetic mixing bar in the barrel of the tube and manipulating it from the outside. To prevent identifica- tion error, the tubes should be individually labeled or all tubes for each patient or timed sample placed in a single labeled test tube. Urine Collection CONTAINERS Collecting containers should be clean and made of inert disposable plastic for one use only. They should have a lid that can be tightened securely to prevent leakage of the contents and should be sterile if the sample is collected for microbiological studies. PRESERVATIVE If the sample is to be stabilized because of delayed analy- sis or an unstable constituent, chemical preservatives or stabilizers should be added to the container or the urine. Sodium merthiolate, 150 g per 24 h in the container, or boric acid, 27 g per 24 h in the container, are preferred preservatives for immunoassay. Their main function is to prevent bacterial destruction of the analyte. When- ever material is added, it should be noted on the con- tainer as it may be hazardous or interfere with the assay system. TYPES OF COLLECTION There are several types of urine specimen: G A random specimen may be collected at any unspecified time during a 24-h period and is generally used where only a qualitative result is required. G Midstream specimen, normally required for microbio- logical investigations where precautions have to be taken to provide a clean specimen by careful washing of the genitalia beforehand, voiding the first portion of urine into the toilet, and catching the midportion in an appropriate sterile container. G The early morning, overnight specimen is normally col- lected immediately the patient rises from an overnight sleep. It is often used when more concentrated urine is necessary because of a low concentration of analyte. G Timed specimens are collected between specified times and include those associated with dynamic tests where the urine is collected into a series of containers at specified times; 2 or 3h specimens that are collected at specific times during the day or within a certain period following a meal or ingestion of pharmaceuti- cals; and the more usual 12 or 24h urine collection that provides an average excretion value over a long period. INSTRUCTIONS The patient should always be given written instructions, and these should also be explained orally. This is particu- larly important when a timed specimen is required. The following are instructions for a 24h sample: G Always use the collection bottle provided. G At the beginning of the 24h collection, the bladder should be voided, and this specimen must be discarded. The exact time should be noted on the bottle label. G All the urine passed during the next 24h must be col- lected in the bottle, which should be kept in a cool place, preferably a refrigerator. G On the following day at exactly the time noted on the label on the previous day, the bladder should again be voided and the urine added to the bottle. This is the end of the 24h collection. G If any urine is accidentally discarded, a new 24h collec- tion should be started in an empty bottle. G The urine must not be contaminated by bowel move- ments. If this happens, a new 24h collection should be started.
  10. 10. 435CHAPTER 6.1 Sample Collection, Including Participant Preparation and Sample Handling SAMPLE HANDLING When the sample reaches the laboratory, it must be thor- oughly mixed before a specimen is taken for analysis. Urine is often cloudy, particularly if stored for any length of time and should be centrifuged before use. It may be necessary to take the pH of the sample and adjust it if appropriate. β2-Microglobulin, for instance, is destroyed at above pH 6.0, and catecholamines and related substances must be collected at a pH less than 2.0. It is often useful, particularly if there is doubt about the completeness of the collection, to measure the creatinine. Although creatinine excretion is related to body mass, it is relatively constant in individuals from day to day, and any gross changes throw doubt on the integrity of the sample. Oral Fluid The assessment of biomarkers in saliva has several advan- tages compared to other biospecimens such as urine and blood. Saliva collection is generally stress and pain free, and samples can therefore be collected repeatedly, at regu- lar intervals and also in reasonable quantities. It does not require skilled professionals and special laboratory equip- ment, and samples can be taken while the patient continues normal activity, possibly at home or at work. This makes it an attractive alternative to blood for use in investigations involving immunoassays, especially when these focus on neonates, infants, children, and vulnerable populations. The biospecimen historically called “saliva” is actually a composite of oral fluids secreted from many different glands. The major source glands are located in the upper posterior area of the oral cavity (parotid gland area), lower area of the mouth between the cheek and jaw (submandibu- lar gland area), and under the tongue (sublingual gland area). There are also many minor secretory glands in the lip, cheek, tongue, and palate. A small fraction of oral fluid (crevicular fluid) also comes from serum leakage in the cleft area between each tooth and its surrounding gums or via leakage from serum due to mucosal injury or inflammation. Each secretory gland produces a fluid that differs in vol- ume, composition, and constituents, and thus, each source gland’s contribution to the pool of oral fluid varies. For instance, mucins make saliva viscous, elastic, and sticky to protect tooth enamel against wear and to encapsulate microorganisms. These glycoproteins are not present in oral fluid secreted by the parotid gland. Oral fluid is water-like in composition and has a pH (acidity) range between 6 and 9. Foods and substances placed in the mouth are capable of changing salivary acid- ity very quickly because this water-like fluid has minimal buffering capacity. Immunoassays (see below) are a method of choice for assaying many salivary analytes. The anti- body–antigen binding step during an immunoassay is compromised when the specimen is highly acidic (pH<3) or basic (pH>9). This unique characteristic of saliva inter- acts with procedures used to collect it and can compromise measurement accuracy. Many of the salivary analytes are serum constituents (e.g., steroid hormones) transported into saliva either by filtration through the tight spaces between acinus or duct cells in the salivary glands or diffusion through acinus or duct cell membranes. Some of the analytes found in oral fluids are synthesized, stored, and released from the gran- ules within the secretory cells of the saliva glands (i.e., enzymes, mucins, cystatins, histatins). Still others are com- ponents of humoral (antibodies, complement) immunity or compounds (cytokines) secreted by cells (neutrophils, macrophages, lymphocytes) of the mucosal immune sys- tem. Furthermore, saliva contains sufficient cellular mate- rial to obtain a high quantity and quality of DNA. An understanding of whether an analyte is transported into oral fluid by filtration or passive diffusion, secreted from salivary glands, or released or derived from cells locally in the oral mucosa is essential to interpreting the meaning of individual differences in that measure. The secretion of oral fluids is influenced by the day– night cycle; chewing movement of the mandibles; taste and smell; iatrogenic effects of medications that cause xerostoma (dry mouth); as well as medical interventions (e.g., radiation) and conditions (e.g., Sjögren’s syndrome) that affect saliva gland function. Saliva glands are directly innervated by parasympathetic and sympathetic nerves, and activation of the ANS component of the psychobiol- ogy of the stress response affects saliva flow rates. The lev- els of salivary analytes that are produced in the mouth, like alpha-amylase (sAA) and secretory IgA, and the levels of those that migrate into saliva from blood by filtration through the junctions between acinar or duct cells in the salivary gland (e.g., DHEA-sulfate [DHEA-S] and other conjugated steroids) are influenced by the rate of saliva secretion. For these saliva analytes, a correction must be made by multiplying the measured concentration or activ- ity of the analyte (e.g., U/mL, pg/mL, µg/dL) by the flow rate (mL/min) to express the measure as output as a function of time (e.g., U/min, pg/min, µg/min; for example, 0.50µg/ dL×0.5mL/min=1µg/min). Even under normative-healthy conditions, more than 250 species of bacteria are present in oral fluids (Paster et al., 2001). During upper respiratory infections, for example, oral fluids are highly likely to contain agents of disease. Oral fluid specimens should therefore be handled with universal precautions. SAMPLE COLLECTION In the past, saliva collection devices have involved cotton- based absorbent materials. Placed in the mouth for 2–3min, oral fluids rapidly saturate the cotton; the specimen is expressed into collection vials by centrifugation or com- pression. Most of the time, this approach is convenient, simple, and time efficient. However, when the absorbent capacity is large and sample volume small, the specimen absorbed can be diffusely distributed in the cotton fibers, making sample recovery problematic. Poor saliva volume recovery is linked to higher rates of missing data and can also be associated with artificially low cortisol estimates. The process of absorbing oral fluid with cotton and other materials also interferes with the immunoassay of several salivary analytes.
  11. 11. 436 The Immunoassay Handbook Early studies employed serum assays modified for use with saliva by, among other things, requiring large saliva test vol- umes (200–400µL). To collect sufficient test volumes, saliva flow was often stimulated using techniques that involved chewing (gums, dental wax) or tasting (sugar crystals, pow- dered drink mixes, citric acid drops) substances. When not used minimally and/or consistently, some of these methods can alter immunoassay performance. Indirectly, stimulants also influence the levels of salivary analytes that are depen- dent on saliva flow rate (SIgA; DHEA-S; neuropeptide Y, NPY; vasoactive intestinal peptide, VIP). We advise avoid- ance of these techniques unless pilot studies show that their application does not adversely affect measurement validity of the salivary analytes of interest. Depending on where in the mouth an absorbent device is placed, a different fluid type may be collected. There- fore, the placement of oral swabs in different areas of the mouth can introduce variation in the measured levels or activity of some salivary analytes. If this specialized collec- tion issue is not controlled, it may contribute to measure- ment error across sampling occasions, both within and between subjects. Standardizing instructions to research staff and participants about the placement of saliva collec- tion devices is essential, as is monitoring compliance. Threats to measurement validity related to swab place- ment can also be avoided by collecting whole saliva by passive drool. Briefly, participants are asked to imagine that they are chewing their favorite food, slowly move their jaws in a chewing motion, and allow the oral fluid to pool in their mouth without swallowing. Next, they gently force the spec- imen through a short device (e.g., SalivaBio LLC, Baltimore, MD) into a vial. The advantages of this procedure include (1) a large sample volume may be collected within a short collection timeframe (3–5min); (2) target collection volume may be confirmed by visual inspection in the field; (3) the fluid collected is a pooled specimen mixture of the output from all salivary glands; (4) the procedure does not introduce interference related to stimulating or absorbing saliva; (5) samples can be aliquoted and archived for future assays. The use of saliva, rather than urine or blood, enables sam- ple collection without significantly interrupting both everyday and laboratory activities. Most of the saliva col- lection techniques that have been studied have unique benefits as well as shortcomings that prevent universal application. When possible, saliva collection methods should always be piloted in the field to ensure that they do not contribute to measurement error, using the exact assay protocols to be employed. Blood Leakage into Oral Fluid To meaningfully index systemic biological activity, quantita- tive estimates of an analyte (e.g., hormone) in saliva need to be highly correlated with the levels measured in serum. The magnitude of this serum–saliva association depends, in part, on consistency in the processes used to transport circulating molecules into oral fluids. When the integrity of diffusion or filtration is compromised, the level of the serological marker in saliva will be affected. Most serum constituents are present in serum in much higher levels (10- to 100-fold) than in saliva. Blood and blood products can leak into oral fluids via burns, abrasions, or cuts to the cheek, tongue, or gums. Blood in oral fluid is more prevalent among individuals who suffer from poor oral health (i.e., open sores, periodontal disease, gingivitis), in children with loose deciduous teeth, those with certain infectious diseases (e.g., human immuno- deficiency virus, HIV), and those who engage in behavior known to influence oral health negatively (e.g., tobacco use). Spiking whole blood into saliva reveals that samples visibly contaminated with blood will present varying degrees of yellow-brownish hue. Kivlighan et al. (2004) pre- sented a simple 5-point Blood Contamination in Saliva Scale (BCSS) with the following response options: (1) saliva appears clear, no visible color; (2) saliva has a hint of color, a little brown or yellow tint is barely visible; (3) saliva has a clearly visible yellow or brown tint; (4) yellow or brown coloring is more than just a tint, color is obvious but not very deep; (5) saliva is very colored, deep, rich, dark yellow or brown is very apparent. Under healthy conditions, BCSS ratings (n=42) averaged 1.33; after microinjury caused by vigorous tooth brushing, ratings averaged 2.42. We recommend: (1) participants should be screened for events in their recent history that could cause blood leakage into saliva by asking questions related to oral health (i.e., “Do your gums bleed when you floss or brush your teeth?”), shedding teeth, or open sores or injury to the oral cavity; (2) sampling saliva within 45min of microinjury to the oral cavity (e.g., brushing teeth) should be avoided; (3) samples should be systematically inspected at the collection point and if visibly contaminated with blood excluded from analyses. Particulate Matter and Interfering Substances The integrity of oral fluid samples can also be influenced by items placed in the mouth. Food residue in the oral cavity after drinking or eating may include particulate matter, may change salivary pH or composition (viscosity), and/or con- tain substances (e.g., bovine hormones, active ingredients in medications, enzymes) that cross-react in immune or kinetic reaction assays. We recommend a simple solution: research participants should not consume food or drink within the 20 min prior to sample donation. If anything has been eaten within this time window, participants should rinse their mouth with water prior to providing a specimen. Impor- tantly, however, they must wait at least 10min after drink- ing before a specimen is collected to avoid diluting it with water and artificially lowering concentration/volume (µg/ dL, ng/mL, pg/mL) or activity/volume (U/mL) estimates of salivary analytes. Access to food and drink should be care- fully planned and scheduled when multiple samples are col- lected over a specific time period.
  12. 12. 437CHAPTER 6.1 Sample Collection, Including Participant Preparation and Sample Handling Sample Handling, Transport, and Storage Typically, once specimens are collected, they should be kept cold or frozen. Refrigeration prevents degradation of some salivary analytes and restricts the activity of proto- lytic enzymes and growth of bacteria. Significant declines occur in the levels of some salivary analytes when samples are stored at room temperature or 4°C. Thus, the way in which samples are handled, stored, and also transported after collection has the potential to influence sample integ- rity and measurement validity. Our recommendation is conservative. After collection, saliva samples should be kept frozen (at least −20°C). If freezing is not possible, then at a minimum samples should be kept cold (on ice or refrigerated) until they can be fro- zen later that day. Repeated freeze–thaw cycles should be avoided with saliva samples. In our experience, DHEA, estradiol, and progesterone are very sensitive to freeze– thaw, whereas DNA, cortisol, testosterone, and sAA are robust (up to at least three cycles). This position is consis- tent with aliquoting and archiving frozen samples in antic- ipation that biotechnology advances will enable different markers to be assayed in the future. It should also be noted that some salivary analytes (e.g., neuropeptides) may require specimens to be directly collected into storage vials that are chilled (e.g., oxytocin) or treated with inhibitors (such as EDTA or aprotinin) to minimize rapid degrada- tion. For large-scale national surveys, investigators work- ing in remote areas or patients collecting samples at home, freezing and shipping these frozen samples can be logisti- cally complex and cost-prohibitive. MEDICATIONS Medications that are applied intranasally, inhaled, or applied as oral topicals (e.g., teething gels) are a primary source of concern. These substances have the potential to change saliva composition due to residue left by their use in the oral cavity. They also deliver specific and/or non- specific molecules directly into oral fluids that have the potential to cross-react or interfere with antibody– antigen binding in immunoassays. Many medications reduce salivary flow, including diuretics, hypotensives, antipsychotics, antihistamines, barbiturates, hallucino- gens, cannabis, and alcohol. Reduced salivary flow changes the physical composition (viscosity) of saliva, alters salivary pH, influences the movement of molecules from serum into oral fluid, and changes the measured levels of salivary analytes produced and secreted by the salivary glands. Each individual medication has the potential to influ- ence salivary analytes by more than one mechanism or pathway. Estimating the nature of the effects is further complicated by the fact that individuals may be taking more than one medication simultaneously. Investigators may be interested in monitoring use of OTC or prescrip- tion medications that: (1) influence the subjective experience of “stress,” emo- tion, novelty, threat or pain; (2) have agonistic or antagonistic effects within the physiological system of interest; (3) interfere with the biosynthesis of the analyte; (4) affect physiological systems networked with the subsystem that secretes the analyte of interest; (5) alter levels of binding globulins and the fraction of biologically active analyte; (6) have active ingredients that cross-react or cause nonspecific interference in immunoassays for the salivary analyte in question. Finally, it is important to account for the medical condi- tion for which the medication is prescribed; this may also explain individual differences in the analyte levels or activ- ity, confounding attempts to control for medication effects on salivary analytes. In summary, we recommend that the name, dosage, and schedule of all prescription and OTC medications taken within the last 48h are recorded in the field. This information should be employed (covaried, controlled) to statistically rule out the possibility that med- ication use is driving the primary salivary analyte–outcome relationships of interest. ANALYTES IN SALIVA OF INTEREST We suspect that most readers may not know that the National Institute for Craniofacial and Dental Research (NIDCR) initiated a multisite program project charged with characterizing the salivary proteome. The list includes more than 1000 analytes (Hu et al., 2007). Some analytes are present in saliva because oral fluid represents an ultra- filtrate of serum constituents. This group of analytes has high value because their levels in saliva are highly corre- lated with and reflect levels in general circulation. These measures enable investigators to make inferences about systemic physiological states. Adrenal and gonadal hor- mones are exemplars of this category of salivary markers (see Table 2). The majority of analytes in oral fluid are produced locally in the oral cavity and are secreted from salivary glands. These salivary analytes may reflect features of and variations in oral biology rather than systemic physiology. Many salivary immune and inflammatory markers such as neopterin, beta-2-microglobulin, cytokines (see Table 2) fall into this category. Individual differences may reflect systemic immune function or status, but more likely than not, a major contributor is local inflammatory processes related to oral health and disease. Markers that represent local oral or compartmentalized physiological processes may be less interesting to sciences outside the specific fields of (developmental) oral biology and health. A subset of analytes is produced locally by salivary glands, but the levels vary predictably with systemic physi- ological activation. The activation of the ANS affects the release of catecholamines from nerve endings, and these compounds’ action on adrenergic receptors influences the activity of the salivary glands. Salivary alpha-amylase (sAA) is considered a surrogate marker of ANS activation, with the majority of findings linking it to sympathetic activation via beta-adrenergic pathways. Salivary mea- sures of NPY and VIP may also serve as surrogate markers of ANS. Antibodies to specific antigens are also measurable in oral fluids. Antibodies to HIV and hepatitis C virus are
  13. 13. 438 The Immunoassay Handbook the exemplars in this category of salivary analytes, and Table 2 offers several additional examples. The presence of an antibody in oral fluids reflects immunological history of pathogen/microbe exposure, and depending on the specific antibody measured may represent local and/or systemic immune activity or current/prior exposure. A variety of pharmaceuticals, abused substances, and environmental contaminants can be quantitatively moni- tored in oral fluids. For example, cotinine, a metabolite of nicotine is routinely measured in oral fluid to estimate pri- mary and secondary exposure to nicotine. Within the past 2 years, technical advances confirm that high quantity and quality DNA can be extracted from whole saliva. That is, when a saliva sample is centrifuged to remove mucins and particulate matter and prepare the clear top phase (i.e., supernatant) to be pipetted into test wells for immunoreaction or kinetic reaction assay (i.e., assays for hormones, enzymes, or antibodies), genetic material may be found either in the “pellet” at the bottom of the centrifuged vial or adhered to the saliva collection device. Genetic polymorphisms can be determined from the same specimens already in use, or planned for use, to assess individual differences in salivary analytes and biomarkers. Cerebrospinal Fluid FORMATION OF CEREBROSPINAL FLUID Cerebrospinal fluid (CSF) is mainly produced by passive filtration from the plasma across the blood–CSF barrier formed by the blood vessels (choroid plexi) of the ventri- cles in contact with CSF. Proteins are filtered with some selectivities according to size so that the low-molecular mass proteins pass more readily than those of high-molec- ular mass, but all plasma proteins commonly measured have been found in CSF, demonstrating the lack of a molecular size exclusion limit by the blood–CSF barrier. About 500mL CSF is produced per day and the CSF occupies a volume of about 135mL in the normal adult. Approximately two thirds of this is produced by the cho- roid plexi lying in the lateral ventricles. The remainder is produced at extrachoroid sites. Outflow of CSF is by reab- sorption into the superior sagittal and other adjacent sinuses following an upward flow over the cerebral cortex. Circulation downward into the spinal cord is much slower as the spinal canal constitutes a “dead-end.” Lumbar CSF has a protein concentration up to three times greater than the fluid found within the ventricles, which also has a higher proportion of the low molecular mass proteins than does lumbar fluid. This is due to the progressive equilibration of CSF with plasma through the capillary walls during passage down the spinal canal. Immobilized patients may exhibit increased protein levels in lumbar CSF as the reduced flow rate also allows increased equilibration with plasma. COLLECTION OF CSF CSF is normally collected by lumbar puncture, which must only be done by an experienced medical practitioner. It is contraindicated in patients with increased intracranial pressure due to space-occupying lesions and by those with untreated blood clotting defects. It is normally collected with the patient in a lateral recumbent position under local procaine anesthesia. A sitting position may be preferable in some patients, especially neonates. The puncture should be done under sterile conditions with the needle inserted into the lumbar subarachnoid space, usually at the midline of the L3–4 interspace. When the needle is positioned cor- rectly, the CSF should slowly flow out. Blood-stained fluid indicates contamination with plasma constituents and invalidates the results of immunoassays of protein fractions. This may be due to a traumatic tap such as transfixation of a vessel or because of a recent subarachnoid hemorrhage. A subarachnoid hemorrhage is characterized by xanthochromic CSF due to the accumulation of bilirubin. Amniotic Fluid COLLECTION OF AMNIOTIC FLUID Amniotic fluid bathes the fetus during its development in the amniotic sac within the womb. From early pregnancy, fetal skin cells are shed into the fluid and may be retrieved. At later stages, the fluid consists mainly of fetal urine and lung fluid. TABLE 2 Salivary Analytes of Interest to Health Sciences Endocrine Aldosterone Estradiol, estriol, esterone Androstenedione Progesterone, 17-OH progesterone Cortisol Testosterone DHEA and DHEA-S Melatonin Immune/Inflammation Secretory immunoglobulin A (SIgA) Beta-2-microglobulin (B2M) Neopterin Cytokines, chemokines Soluble tumor necrosis factor receptors C-reactive protein Autonomic Nervous System Alpha-amylase (sAA) NPY VIP Chromogranin A Nucleic Acids Human genomic mRNA Mitochondrial Microbial Bacterial Viral Antibodies Specific for Antigens Measles Hepatitis A, B, C, E HIV Mumps Herpes simplex CMV Rubella Epstein–Barr Pharmaceuticals, Drugs of Abuse and Stimulants Cotinine Alcohol Meth-, amphetamine Lithium Methadone Cocaine Opioids Marijuana (THC) Caffeine Phenytoin Bisphenol-A (BPA) Barbituates
  14. 14. 439CHAPTER 6.1 Sample Collection, Including Participant Preparation and Sample Handling AMNIOCENTESIS Amniocentesis, the procedure by which amniotic fluid is withdrawn from the amniotic sac using a needle through the mother’s abdominal wall and uterus, must be done by an experienced medical practitioner. It is normally done under local anesthetic with ultrasound scanning control and not normally before 16 weeks’ gestation, as the volume of amniotic fluid is small in early pregnancy but increases rapidly from about 16 weeks onward. SPECIAL PRECAUTIONS There is a risk of spontaneous abortion after amniocentesis of about 1 in 200. This should be explained to the patient. Amniotic fluid should be placed in a plain tube without additives and should be centrifuged before immunoassay. Care should be taken to avoid contamination with blood, particularly fetal blood, if this is likely to contain the ana- lyte of interest. This is a particularly pertinent consider- ationinalphafetoprotein(AFP)assays,whereblood-stained amniotic fluid samples may give factitious results. Con- versely, if serum AFP is to be assayed in conjunction with amniocentesis, the blood sample must be taken before amniocentesis, as fetomaternal hemorrhage induced at amniocentesis may produce spuriously high maternal serum levels. It is possible, and not unknown, for urine to be obtained by percutaneous puncture in the mistaken belief that it is amniotic fluid. This should be borne in mind when interpreting apparently anomalous results, e.g., very low AFP levels. It is useful to note the gestational age, preferably assessed by ultrasonography, as this is helpful in interpretation of the results of most assays for amniotic fluid constituents, which generally change as pregnancy progresses. Sweat Sweat samples are used for monitoring intake of drugs of abuse, particularly cocaine. Insensible perspiration may be collected over a period of days using a sweat patch. The PharmChek™ patch (Sudormed, Santa Ana, CA) is a non- occlusive dressing consisting of a cellulose blotter collec- tion pad, covered by a thin polyurethane and acrylate adhesive membrane. The water component of the sweat passes through the polyurethane and solids, salts, and drugs excreted in the sweat are trapped on the collection pad, which will collect a minimum of 300µL/day in a 22°C environment. Exercise or higher temperatures will increase the amount collected. The drugs are eluted at the end of the collection period by placing the cellulose collection pad in a 5mL capped tube together with 2.5mL of a pH 5.0 acetate buffer with methanol. Drugs deposited on the skin prior to wearing the patch may cause false positives. Semen Immunological tests on semen, such as the assessment of anti-sperm antibody, depend on a reasonable sperm count and on the integrity and viability of the sperm in the sample. It is therefore advisable for the subject to abstain from intercourse for 3 days before production of the speci- men, which may be obtained by hand masturbation or coitus interruptus (withdrawal). A normal sheath or condom should not be used as these may contain spermicidal agents. However, there are special reusable condoms spe- cifically designed for the collection of semen. The speci- men should be delivered into a clean bottle provided by the laboratory. It is essential that the sample is tested within 2h of ejac- ulation, and it is helpful to allow 30min to elapse because the initial gel formation requires this time for dissolution, although the semen sample may be liquified by warming in a sealed plastic container for 20min at 37°C. The sample should be maintained at body or room tem- perature before assay because cooling damages the sperm and reduces motility. Frozen sperm must not be used as freezing alters the surface of the sperm and may interfere with antibody binding. Hair Sectional hair analysis can be utilized for toxicological and illicit drug investigations as it can provide an assessment of the intake of certain analytes over a period of many months. It is increasingly being used to monitor drug abuse by sen- sitive immunoassay. There are a number of problems that have to be addressed. The first is surface contamination of the hair from the surrounding environment of the subject. The second is the release of the drug that was deposited in the hair structure during the keratinization process. A third is to ensure that the drug has not been lost during cosmetic hair treatment as it is known that there can be an 80% loss during bleaching and that repeated shampooing can decrease the levels by up to 60%. The carefully selected hair sections undergo alkaline digestion before neutraliza- tion and buffering for analysis. Milk Milk, whether from humans or animals, may be collected either by expressing the milk manually or by use of a breast pump into plain, sterile, glass tubes. The milk should be thoroughly mixed before aliquots are taken for storage or for immediate analysis. If analysis is to be delayed, milk should be stored at −20°C to suppress bacterial action, and, after thawing, thorough mixing is again necessary before analysis. As an alternative to freezing, potassium dichromate may be used to preserve milk, which may then be stored at 4°C while awaiting analysis. This is a conve- nient way of storing bovine milk prior to assay for progesterone. If the analytes are not lipid soluble, e.g., the immuno- globulins to food proteins, removal of the lipid layer after centrifugation minimizes lipid interference with the immunological reaction. If the analyte is lipid soluble, e.g., morphine, then extraction of the substance from the milk is usually necessary to eliminate possible matrix effects due to the high lipid and protein concentrations when the milk is assayed undiluted.
  15. 15. 440 The Immunoassay Handbook Storage and Transportation Most analytes are more stable when the sample is main- tained in a cool or frozen condition. For some, especially the small peptide hormones, storage at −20°C and trans- portation in the frozen state is usually necessary for reli- able results. Such hormones include insulin and insulin C- peptide, gastrin, glucagon, ACTH, and vitamin D. Some complement components need to be maintained at a tem- perature of −70°C if spurious results are not to be obtained. The need for low temperature preservation of the sam- ple depends not only on the analyte but on the method being used for assay, and in particular the antibodies and the epitopes to which they are directed. Thus, the sample for a particular assay may require different handling depending on the method to be used in the investigation. While it is strongly recommended that samples should be separated from red cells as soon as possible following venesection, and stored at low temperature, it has been reported (Diver et al., 1994), that there was little change in the assayed levels of commonly measured hormones after storage at ambient temperature in whole blood for up to 1 week and that these changes were unlikely to be of clini- cal significance. The specimen may be transported in the frozen state by packing in solid carbon dioxide (dry ice) in an insulated con- tainer or vacuum flask or, if such a low temperature is not necessary, in an insulated box containing a bottle of 20% sodium chloride which has been frozen solid over a number of days in a deep freeze capable of reaching −21.6°C (the freezing point of the sodium chloride solution). The frozen sample, enclosed in a sealed polythene bag and placed next to a bottle of frozen sodium chloride solution and cushioned with foam plastic in an insulated box, will stay at a tempera- ture of below −8°C for a period of 40h, approximately twice as long as with the same volume of solid carbon dioxide. For safety reasons, the transport of samples outside the laboratory is subject to legislation in many countries, and local rulings should always be applied. Typically, they should ensure that every sample is enclosed in a primary container, which is securely sealed. This must be wrapped in sufficient absorbent material to absorb all possible leak- age in the event of damage. The container and absorbent material must be sealed in a leakproof bag. This package should further be placed in a polypropylene, metal, or strong cardboard container, which is securely closed and properly labeled for transit. Conclusions In this chapter, we have attempted to describe the proce- dures for the collection of samples and to draw attention to the many preanalytical factors that may influence the results of immunoassays. We have not tried to specifically define those factors that are clinically relevant as these will vary according to circumstances and the purpose for which the assay has been requested. But for reliable results, practitioners must know of these factors and set standards of control that minimize their effect on the test results and on the inter- pretation of the test outcome. Author Notes In the spirit of full disclosure, DAG is the Chief Scientific and Strategy Advisor of Salimetrics, LLC (State College, PA) and SalivaBio LLC (Baltimore, MD), and these relationships are managed by the policies of the Conflict of Interest Com- mittee at the Johns Hopkins University School of Medicine. DO was supported by a Rubicon award (446-10-026) from the Netherlands Organization for Scientific Research. Further Reading CLSI (formerly NCCLS): Collection, Handling, Transport, and Storage for Body Fluids Quick Guide. CLSI document C49-A/H56-A (Clinical and Laboratory Standards Institute, Pennsylvania, 2007a). CLSI. Collection, Handling, Transport, and Storage for Hemostasis Quick Guide. CLSI document H21–A5 (Clinical and Laboratory Standards Institute, Pennsylvania, 2010a). CLSI. Collection, Transport, and Processing of Blood Specimens for Testing Plasma-Based Coagulation Assays and Molecular Hemostasis Assays; Approved Guideline—Fifth Edition. CLSI document H21–A5 (Clinical and Laboratory Standards Institute, Pennsylvania, 2008a). CLSI. Handling, Transport, and Storage of Specimens Quick Guide. CLSI document H18–A4 (Clinical and Laboratory Standards Institute, Pennsylvania, 2010b). CLSI. Procedures for the Collection of Diagnostic Blood Specimens by Venipuncture; Approved Standard—Sixth Edition. CLSI document H3–A6 (Clinical and Laboratory Standards Institute, Pennsylvania, 2007b). CLSI. Procedures and Devices for the Collection of Diagnostic Capillary Blood Specimens; Approved Standard—Sixth Edition. CLSI document H04–A6 (Clinical and Laboratory Standards Institute, Pennsylvania, 2008b). CLSI. Procedures for the Collection of Arterial Blood Specimens; Approved Standard— Fourth Edition. CLSI document H11–A4 (Clinical and Laboratory Standards Institute, Pennsylvania, 2004). CLSI. Procedures for the Handling and Processing of Blood Specimens for Common Laboratory Tests; Approved Guideline—Fourth Edition. CLSI document H18–A4 (Clinical and Laboratory Standards Institute, Pennsylvania, 2010c). CLSI. Quality Venipuncture Quick Guide. CLSI document H03–A6 (Clinical and Laboratory Standards Institute, Pennsylvania, 2010d). CLSI. Technique for Skin Puncture in Adults and Older Children Quick Guide. CLSI document H04–A6 (Clinical and Laboratory Standards Institute, Pennsylvania, 2010e). CLSI. Tubes and Additives for Venous and Capillary Blood Specimen Collection; Approved Standard—Sixth Edition. CLSI document H01–A6 (Clinical and Laboratory Standards Institute, Pennsylvania, 2010f). CLSI. Urinalysis; Approved Guideline – Third Edition. CLSI document GP16–A3 (Clinical and Laboratory Standards Institute, Pennsylvania, 2009). Cone, E.J. and Huestis, M.A. Interpretation of oral fluid tests for drugs of abuse. Ann. N. Y. Acad. Sci. 1098, 51–103 (2007). Diver, M.J., Hughes, J.G., Hutton, J.L., West, C.R. and Hipkin, L.J. The long term stability in whole blood of 14 commonly requested hormone analytes. Ann. Clin. Biochem. 31, 561–565 (1994). Granger, D.A., Hibel, L.C., Fortunato, C.K. and Kapelewski, C.H. Medication effects on salivary cortisol: Tactics and strategy to minimize impact in behav- ioral and developmental science. Psychoneuroendocrinology 34, 1437–1448 (2009). Hu, S., Loo, J.A. and Wong, D.T. Human saliva proteome analysis. Ann. N. Y. Acad. Sci. 1098, 323–329 (2007). Kivlighan, K.T., Granger, D.A., Schwartz, E.B., Nelson, V., Curran, M. and Shirtcliff, E.A. Quantifying blood leakage into the oral mucosa and its effects on the measurement of cortisol, dehydroepiandrosterone, and testosterone in saliva. Horm. Behav. 46, 39–46 (2004). Malamud, D. and Tabak, L. Saliva as a diagnostic fluid. Ann. N. Y. Acad. Sci. 694, 216–233 (1993). National Institutes of Health. U.S. Surgeon General’s Report on Oral Health (2000) Nemoda, Z., Horvat-Gordon, M., Fortunato, C.K., Beltzer, E.K., Scholl, J.L. and Granger, D.A. Assessing genetic polymorphisms using DNA extracted from cells present in saliva samples. BMC Med. Res. Methodol. 11, 170 (2011). Nieuw Amerongen, A.V., Ligtenberg, A.J.M. and Veerman, E.C.I. Implications for diagnostics in the biochemistry and physiology of saliva. Ann. N. Y. Acad. Sci. 1098, 1–6 (2007). Sreebny, L.M. and Schwartz, S.S. A reference guide to drugs and dry mouth – 2nd edn. Gerodontology 14, 33–47 (1997). Vining, R.F. and McGinley, R.A. Hormones in saliva. CRC Crit. Rev. Clin. Lab. Sci. 23, 95–114 (1986). Wu, A.H.B. Tietz Clinical Guide to Laboratory Tests (Saunders, Philadelphia, 2006). Young, D.S. Effects of Preanalytical Variables on Clinical Laboratory Tests, 3rd edn. (American Association for Clinical Chemistry Press, Washington DC, 2007).