Specimen Collection Guides and Resources

Definitions That May Be Helpful:

• Anticoagulant – A substance that prevents blood from clotting.
• Hemolyze – The rupturing of the red blood cells in the specimen. This can often interfere with accurate analysis of blood chemistries. Avoid hemolysis as it interferes with many tests.
• Plasma – The fluid part of anticoagulated blood after it has been separated from the cells.
• Serum  – The yellow fluid that is separated from the cells when blood is clotted and centrifuged.
• Whole blood – Blood that has an anticoagulant and is not separated by centrifuging.
  
  

Basic Phlebotomy Guidelines:

• Correct Order of Draw – It is very important to collect blood tubes in proper sequence to avoid contamination and anticoagulant carry-over to the next tube drawn. Please refer to chart in educational module, Collecting Blood Specimens.
• Inversion of Tubes – In general, all the tubes are to be mixed well by gently inverting several times after filling with blood. Tubes are full when the vacuum quits drawing blood into the tube.
• Plasma – Draw a sufficient amount of blood with the indicated anticoagulant to yield the necessary plasma volume. Gently mix the blood collection tube by inverting six to ten times immediately after collection. Never shake the tubes to mix. Shaking will fragment the blood cells and could result in false abnormal hematology findings. If required, separate plasma from cells by centrifugation within 20-30 minutes.
• Serum – Draw a sufficient amount of blood to yield the necessary serum volume. Serum volume is approximately half of the whole blood volume. Invert the tube several times to activate the clotting. Let stand for 20-30 minutes, or until a clot is well formed. Centrifuge for ten minutes. If frozen serum is required, pour off serum into plastic vial and freeze.
• Syringe Collection – If using a syringe to collect the blood, it is best to fill the tube through the stopper. If the stopper has been taken off, it may slide loose at a later time, resulting in a blood spill.
• Whole Blood – Draw a sufficient amount of blood with the indicated anticoagulant. Gently mix the blood collection tube by inverting six to ten times immediately after collection.

The following is a brief overview of the requirements involved for collecting blood specimens. Each of the different tubes has individual characteristics. In general, all the tubes are to be mixed well by gently inverting several times after filling with blood. Tubes are full when the vacuum quits drawing blood into the tube.

Serum Type Tubes

• Red-Top Tube:
  This tube is a plain Vacutainerâ containing no anticoagulant, used for collection of serum specimens. Tests using this tube may include therapeutic drugs, selected chemistry tests and some blood bank procedures. Any chemistry sample may be collected in these tubes. However, the yellow-top SST tubes are superior as the serum is automatically separated. When a test is designated to be collected in a red-top tube, an SST tube should not be substituted. The gel barrier may interfere with analysis.
• Yellow-Top Serum Separator Tube (SST):
  This tube contains a clot activator and a serum gel separator, used for various laboratory tests, including the majority of chemistry tests. Invert the tube six to ten times in order to activate the anticoagulant.

Plasma and Whole Blood Tubes

• Blue-Top Tube (Sodium Citrate):
  This tube contains sodium citrate as an anticoagulant, used for collection of blood for coagulation studies. NOTE: It is imperative that the tube be completely filled. The ratio of blood to anticoagulant is critical for valid coagulation studies. Immediately after draw, invert the tube six to ten times in order to activate the anticoagulant.
• Gray-Top Tube (Potassium Oxalate/Sodium Fluoride):
  This tube contains potassium oxalate as an anticoagulant and sodium fluoride as a preservative, used to preserve glucose in whole blood and for some special chemistry tests. The minimum volume should not be less than 1.0 mL due to the anticoagulant present in the tube.
• Green-Top Tube (Lithium Heparin):
  This tube contains lithium heparin, used for the collection of heparinized plasma or whole blood. The minimum volume should not be less that 1.0 mL due to the anticoagulant present in the tube.
• Green-Top Tube (Sodium Heparin):
  This tube contains sodium heparin, used for the collection of heparinized plasma or whole blood for special tests. The minimum volume should not be less than 1.0 mL due to the anti-coagulant present in the tube.
• Lavender-Top Tube (EDTA):
  This tube contains EDTA as an anticoagulant, used for most hematological procedures. Mix the tube well to dissolve the powdered anticoagulant. The blood level should at least reach the bottom of the tube’s label.

  NOTE: Never shake the tubes to mix. Shaking will fragment the blood cells and could result in false abnormal hematology findings.

• Light Yellow-Top Tube (ACD):
  This tube contains ACD, used for the collection of whole blood for special tests.

  NOTE: This tube is very similar in appearance to the SPS tube. Read the label for verification of correct anticoagulant.

• Light Yellow-Top Tube (SPS):
  This tube used for the collection of blood cultures contains, SPS, is always drawn first to avoid bacterial contamination.

  NOTE: This tube is very similar in appearance to the ACD tube. Read the label for verification of correct anticoagulant.

• Pink-Top Tube (EDTA):
  This tube contains EDTA as an anticoagulant, used for most blood bank procedures. Mix the tube well to dissolve the powdered anticoagulant.
• Royal Blue-Top Tube:
  There are two types of royal blue-top Monojectâ tubes, one with the anticoagulant EDTA and the other plain. These are used in the collection of whole blood or serum for trace element analysis. Refer to the individual metals in the specific test listing to determine the tube type necessary.
• Tan-Top Tube (Sodium Heparin):
  This tube is lead-free and thus used for collection of blood lead levels.
• White-Top Tube (K2 EDTA):
  This plasma preparation tube (PPT) contains a gel barrier in the plastic tube. After centrifugation the plasma may be frozen without opening the tube.

Miscellaneous tubes Microtainers
These are designed for fingersticks, which are commonly used for children or other difficult venipunctures. (Some vials may contain a powdered form of the anticoagulant. This must not be shaken out as the vial is being mixed.)

Peach/red – no anticoagulant

Yellow top; Amber tube – no anticoagulant, protects specimen from light.

Green – heparinized anticoagulant

Lavender – EDTA anticoagulant

NOTE: All hematology collections (CBC, hemoglobin, hematocrit, etc.) must be well mixed. Any clotting will cause the results to be invalid. When collecting the specimen and the blood flow is slow, cap the vial and mix quickly midway through the blood collection. This will help prevent a clot formation. Mix again at the completion of the collection. The minimum collection is at the second purple line (500 ml).

Pediatric Tube (Half-draw)
The half-draw tubes, which have clear lids, are designed for children or patients who are difficult to draw. These tubes contain a smaller amount of vacuum, thus less trauma to the vein. Because of the reduced amount of anticoagulant, as well as vacuum, it is important not to overfill these tubes. This is possible if filled by a syringe. Overfilling could cause rejection of the specimen. On some tubes, the proper fill line is marked with small blocks halfway up the edge of the label.

Special Collection Tubes
Some tests require specific tubes for proper analysis, (such as metals). Please contact Meadville Medical Center Laboratory prior to patient draw to obtain the correct tubes as identified in the individual test listings

Clostridium Difficile:
Optimum specimens are from fresh diarrheal stool. Non-diarrheal specimens will not be tested.
One stool specimen positive for C. difficile is sufficient for diagnosis. Up to three stool specimens per week may be submitted for testing, though no more frequent than one every 24 hours.

Cryptosporidium and Giardia Antigen:
Antigen tests are more sensitive than conventional ova and parasite exam for the detection of Giardia and Cryptosporidium; therefore it is only necessary to submit one specimen. If antigen tests are negative, additional specimens may be submitted. The laboratory will keep a portion of the specimen in preservative for up to 5 days if traditional ova and parasite exam is desired on antigen negative specimens. Stool samples submitted in PVA fixative are not acceptable for antigen testing. SAF and MIF preserved specimens, and unpreserved specimens are acceptable.

Ova and Parasites:
A maximum of three specimens should be submitted, one every other day. If a patient has been hospitalized for three or more days, ova and parasite may not be ordered. The likelihood of acquiring infection by parasitic agents during hospitalization is extremely low.
Cryptosporidium and Giardia antigen test will be ordered unless a pertinent travel history or medical condition is specified by the ordering physician.

Stool Culture:
Up to three stool specimens per week may be submitted for testing, though no more frequent than one every 24 hours. If a patient has been hospitalized for three or more days, stool cultures may not be ordered.
Organisms detected include Campylobacter, E. coli 0157, Salmonella and Shigella. The following organisms will be reported if found in predominance; Pseudomonas, Aeromonas, Bacillus, Plesiomonas, Staph aureus and yeast. Examination of the stool for other enteric pathogens such as Yersinia will be done by special request. Cultures for Vibrio are forwarded to a reference lab.

Clean Catch Urine Collections

1. Wash hands thoroughly.
2. Care must be taken to avoid bacterial contamination of the specimen container. After removing the lid, place it on a flat surface with the inside facing up. Do not touch the inside of the specimen lid or the container. Any contact by your skin may cause a bacterial contamination that may interfere with the test.
3. Open the towelette package. Unfold the towelette.

   Females – Using one stroke, wash by wiping the urinary area from the front to the back.

   Males – Using one stroke, wash by wiping the urinary area from top to bottom.

4. Start to urinate directly into the toilet. While urinating, pass the collection cup into the stream of urine. Do not fill the specimen cup to the top. Remove the cup before the stream of urine has stopped. Screw the cap securely on the container.
5. Place the label on the cup after securely closing the lid. Do not put label on the lid.
6. Refrigerate specimen immediately if unable to deliver specimen to lab promptly. Deliver the refrigerated culture to the lab within 12 hours. If a urinalysis is also ordered, the specimen must be delivered to the lab within 2 hours if unpreserved.
7. If preservative tube is used, the specimen is stable at room temperature up to 72 hours.

Preservative (Urinalysis & Culture)
If using the blue-capped preservative tube available from Meadville Medical Center Laboratory, refrigeration is not necessary. The urinalysis/culture preservative system contains a mercuric oxide-based tablet and has a maximum fill capacity of 10 mLs. Please fill to the maximum fill line. Shake specimen to dissolve the tablet and label the specimen tube.

NOTE: Urinalysis preservative system is stable up to 72 hours without refrigeration.
Preservative tubes are NOT acceptable for common urine chemistry tests.

Random Collections
For routine analysis and microscopic evaluation, have the patient void into a clean container. A clean-catch specimen is preferred. The specimen should be capped, labeled and refrigerated. If specimen is not transferred into a urine preservative tube, it must be transported to the lab within 2 hours.

24-Hour Urine Collections
For detailed collection information refer to the specific test ordered in the Patient Information section in the back of manual.

Some 24-hour urine tests require a preservative. If an additive is needed, please notify the lab. We will prepare the container according to the proper requirements. Please be careful that this preservative does not come in contact with your patient’s skin. If a splash does occur, wash the affected area with large amounts of water. Use a separate container to collect the urine sample and then pour the urine into the 24-hour container. This will help to control splashing of the preservative.

Care should be taken when collecting a 24-hour urine. It is essential that all the urine passed during a 24-hour period is totally collected. If the collection is not complete, our reference values are not valid. Use the following procedure for the correct specimen collection and preparation.

1. Warn the patient of the presence of potentially hazardous preservatives in the collection container. Preservative type and quantity must be written on the bottle.
2. Label the container with the patient’s name prior to collecting the specimen.
3. Instruct the patient to discard the first morning specimen and to record the time of voiding.
4. The patient should collect all subsequent voided urine for the remainder of the day and night.
5. Collect the first morning specimen on day two at the same time as noted on day one.
6. The collection is now complete and the sample may now be delivered to the laboratory for processing.

During the collection period, it is important that the urine sample be kept cold. Urine may be stored in a refrigerator, an ice chest, or in a pail of ice to prevent the growth of bacteria in the sample.

NOTE:
If two tests requiring different additives are ordered on the same patient, contact the lab. A special container will be provided to accommodate this special collection need.

24 Hour Urine Tests Not Requiring an Additive

• Aminolevulinic Acid
• Calcium
• Chloride
• Citrate
• Cortisol Free
• Creatinine Clearance, Serum and Urine
• Creatinine
• Cystine
• Electrolyte
• Microalbumin
• N-Telopeptide-Collagen Cross-Linked
• Phosphorus
• Potassium
• Protein, Total
• Sodium
• Urea Nitrogen
• Uric Acid

24 Hour Urine Tests Requiring an Additive

• 5-Hydroxyindoleacetic Acid (5-HIAA)
• 17-Ketosteroid
• Catecholamine Fractionated
• Homovanillic Acid (HVA)
• Magnesium
• Metanephrines
• Oxylate
• Vanillylmandelic Acid (VMA)

Centrifuge all red-top and SST tubes must be completely clotted prior to centrifuging. Pair tubes of equal volume to be placed opposite of each other in centrifuge holders. Perfect balance is necessary to prevent tube breakage. A “balance tube” of water may be used to equal volume of the tubes to be spun. Unless otherwise indicated, spin for 10 minutes.

Separating:
SST tubes contain a gel separator that provides a barrier. No additional steps are necessary after centrifugation. For plasma or frozen specimens, label plastic vial with needed patient information. Using plastic pipette, remove plasma or serum from tube, placing it into vial.

Packaging:

1. Place specimen in zip-lock Biohazard bag and seal securely. If specimen is STAT or FROZEN, please apply appropriate label to outside of bag.
2. Place form(s) in outer pouch of bag.
   NOTE: All specimens must be placed in a bag for delivery. If patient has urine and blood, place urine in separate biohazard bag, secure, and place in biohazard bag with blood.

Temperature Storage:

MMC Lab is required to maintain proper storage and transport temperatures for all specimens from time of collection to time of analysis to ensure specimen integrity. Please view the temperature control within the education module.

NOTE: Do not freeze glass Vacutainer® tube.

The information listed below is given only as a guideline. The information is not all-inclusive.

Hazard Identification
The following areas are to be identified with a Biohazard sticker:

• Refrigerators or freezers containing laboratory specimens
• Waste containers used for body substances (the containers should have closeable lids)
• Centrifuge
• Disposable sharps containers
• Phlebotomy trays
• Transport container

Personal Safety
Hepatitis B vaccine should be provided to all employees having occupational exposure to bloodborne pathogens.

Safety Procedures

General Cleaning:
Work areas in which blood is processed are to be cleaned with a 10% dilution of household bleach. The bleach mixture is to be fresh daily. Areas are to be cleaned at the end of each day or after they have been contaminated.

Handwashing:
Frequent handwashing is an important safety precaution. Hands are to be washed:

• After completion of laboratory-related work
• After removing gloves
• Before and after all other activities which entail hand contact with mucous membranes or breaks in the skin
• Immediately after accidental skin contact with blood, body fluids or tissues
• If the contact occurs through a break in the gloves, the gloves are to be immediately removed and the hands washed thoroughly.

Blood Spills
The following procedure is recommended for decontaminating blood spills:

1. Wear gloves, (heavyweight, such as are used for dishwashing), and a disposable gown.
2. Absorb the blood with paper towels.
3. Using a detergent solution, clean the spill site of all visible blood with paper towels.
4. Wipe down the spill site with paper towels soaked in a fresh 10% dilution of household bleach. (Bleach solutions are less effective in the presence of a high concentration of protein. Therefore, remove as much liquid blood or serum as possible before decontaminating or disinfecting.)
5. Place the contaminated material in a biohazard container.

10% Formalin Spills

For your safety information, we have included Formalin Safety Data Sheets (SDS). They are located in the “Additional Information“ section in the back of this manual. Most Pathology specimens contain formalin.

10% formalin is considered to be a toxic substance, causing respiratory tract problems. Care is needed when cleaning up a spill.

1. Wear gloves, disposable gown, face mask and face shield.
2. Using the Polyform F®, pour the granules to encircle the spill, creating a “dam”.
3. Sprinkle the Polyform F® granules to completely cover the spill.
4. Wait several minutes until the formalin has gelled. Remove the remains with the scoop and scraper.
5. Dispose of all contaminated material and specimen matter directly into a biohazard container.
6. If necessary, wipe up remains with paper towels. You may spray the area with Formalex® or a detergent to aid you in cleaning the remaining residue. (The formalin has already been neutralized at this time.)
7. If formalin is spilled on a rug, rinse the spill with large amounts of water.

Safety Equipment

Sharps Container:
Used, disposable needles and other sharps are to be placed into a rigid, plastic, puncture-resistant disposable container with a lid. The container is to be clearly marked “Biohazard”.

Vacutainer Needles:
Disposable single-use, locking, needle adapters are required and are to be disposed in the sharps containers. Prior to disposal, press orange guard against a hard surface, locking it around the needle.

Butterfly
Prior to disposal, slide clear plastic guard over exposed needle.

NOTE: As a required safety precaution, needles must never be recapped.

Gloves:
Gloves are to be worn when…

• …it can be reasonably anticipated that the employee may have hand contact with blood or other potentially infectious material.
• …procuring blood specimens and changed between patients.
• ….processing blood. This is especially important when separating serum or plasma, and transferring specimens into tubes.

Replace worn gloves if they become grossly contaminated with blood.

Facial Protection:
Facial protection is to be used if any splashing may occur. Splashing may result when removing rubber stoppers from specimen tubes. (Covering the tube with a gauze pad while removing the stopper may help minimize splashing.) The gauze pad is to be discarded into the biohazard waste container after use.