4.1.1. Filter paper dried blood spot collection, processing, storage, and shipment

Dried blood spot (DBS) is an easy and inexpensive means of collection and storage of peripheral blood specimens in settings where collection and storage of plasma is not optimal. DBS can be used for molecular biology techniques and other diagnostic assays. The cost and difficulty of cold chain shipping of plasma samples are greatly reduced by the use of DBS, which can be shipped as non-dangerous goods.

Always wear gloves when handling filter papers, and hold them only by the upper corner, marked out for labelling. Do not allow the card to come into contact with any unclean surface (e.g. bench, base of hood). The procedure should be performed in accordance with the relevant health and safety practices specific to specimen handling and waste disposal. A minimum level of training is required to perform the procedure.

Reagents and materials required:

•

finger prick device (e.g. Unistik 2 device; Fisher Scientific, 22-0227);

•

alcohol swab;

•

Whatman Protein Saver Card (e.g. Whatman, 903; 10534612);

•

gas-impermeable storage bag (e.g. Fisher Scientific, NC9307519);

•

desiccant pack (e.g. Whatman, 10548234);

•

humidity indicator cards (e.g. Multisorb Des Manufacture, MS200032);

•

card drying rack (e.g. VWR, 89015-592) (optional; cards can be placed on a dry worktop if a drying rack is not available);

•

gloves, preferably powder-free; and

•

sample label.

4.1.2. Blood pellets (white blood cells)

Blood pellets can be used for the isolation of DNA (from EDTA or ACD tubes)

i.

Transfer blood from the original tube to a labelled 50 mL tube.

ii.

Fill the tube with Tris-EDTA buffer (formula) and mix vigorously. Place on ice for 5–10 minutes.

iii.

Spin at 1200g for 10 minutes.

iv.

Carefully pour off the supernatant into a waste container containing chlorine bleach. Briefly vortex the pellet and add 50 mL of Tris-EDTA buffer. Shake vigorously.

v.

If division of the sample is necessary, at this point pour 25 mL of the sample into another centrifuge tube.

vi.

Spin both tubes at 1200g for 10 minutes.

vii.

Repeat the washing if red blood cells persist.

viii.

Carefully pour off the supernatant.

ix.

Using a swirling motion, remove the pellet with a pipette and transfer it to a labelled cryovial.

x.

Store at −80 °C or in LN₂ until further use.

As an alternative, red blood cells can be lysed by using an ammonium-containing lysis buffer.

4.1.3. Plasma

Plasma collected in EDTA or ACD tubes can be used for bioassays, plasma DNA isolation, proteomic analysis, and biomarker discovery.

i.

Spin the vacutainer (about 9 mL) at 815g for 10 minutes at 4 °C to separate plasma from blood cells.

ii.

After wiping each tube with 70% alcohol, remove about 3 mL of plasma. (The tube can be retained for white blood cell extraction.)

iii.

Transfer to a labelled 15 mL tube, and centrifuge at 2500g for 10 minutes at 4 °C.

iv.

Aliquot plasma into 1 mL labelled cryovials (three or four aliquots).

v.

Place in LN₂ dewar to snap-freeze.

vi.

Store at −80 °C or in LN₂.

The purpose of double-spinning the plasma is to remove all cellular contaminants so that the plasma is suitable for plasma DNA analysis. Therefore, it is extremely important not to disturb the buffy coat after the first spin and any pellet after the second spin.

4.1.4. Platelet-poor plasma

Platelet-poor plasma can be used for the isolation of plasma DNA (from EDTA tubes)

i.

Spin blood at 3200g for 12 minutes at room temperature.

ii.

Pipette off plasma using a plastic Pasteur pipette. Transfer into a tube.

iii.

Spin plasma at 2000g for 10 minutes at 4 °C.

iv.

Aliquot into 1 mL aliquots in labelled cryovials.

v.

Store at −80 °C.

4.1.5. Serum

The blood is collected into tubes without addition of anticoagulants. Then, two phases are distinguishable: a solid phase containing fibrin and cells, and a fluid phase containing the serum.

This process should be completed after 30 minutes at room temperature, after which the process described below should start.

i.

Spin blood at 1500g for 10 minutes at room temperature.

ii.

Aliquot 1 mL portions of the supernatant into labelled cryovials.

iii.

Place into LN₂ dewar or dry ice to snap-freeze.

iv.

Transfer to −80 °C freezer or LN₂.

4.1.6. White blood cells

White blood cells collected in EDTA or ACD tubes can be used for DNA extraction and the creation of cell lines.

i.

Transfer the remaining blood from the plasma spin to a labelled 50 mL tube containing 10 mL of RPMI 1640.

ii.

After swabbing the lid of this tube with alcohol, aliquot 3 mL of Ficoll into each of two clearly labelled 15 mL tubes.

iii.

Carefully layer 9 mL of diluted blood onto each tube of Ficoll. Treat gently, and do not mix, but spin as soon as possible.

iv.

Spin at 450g for 30 minutes. Note: When centrifuging, do not use the brake.

v.

Remove most of the top layer (RPMI 1640) using a 1 mL Eppendorf tip, and discard about 3–4 mL into a waste container containing chlorine bleach.

vi.

Collect white blood cells with the same Eppendorf tip using a swirling motion to “vacuum up” white blood cells. Do not take too much Ficoll (third layer), because it is toxic to the cells. Place the white blood cells into a labelled 15 mL tube containing 10 mL of RPMI 1640.

vii.

Spin at 450g for 10 minutes.

viii.

Pour off the supernatant into a waste container containing chlorine bleach. Add 3 mL of cold freezing mix (10% DMSO, 20% fetal calf serum [FCS], RPMI 1640) and resuspend.

ix.

Dispense the white blood cells into three 1 mL labelled cryovials that have been sitting on ice.

x.

Place on ice. Place vials in a controlled-rate freezer so as to cryopreserve cells under conditions that maintain cell viability. This should be done as soon as possible, because DMSO is toxic at room temperature.

xi.

Transfer on a weekly basis to LN₂ tanks.

Instead of a separation based on Ficoll, a Percoll separation can be used.

4.1.7. Buffy coat cells

The buffy coat is a thin, greyish-white layer of white blood cells (leukocytes and lymphocytes) and platelets covering the top of the packed red blood cells after centrifugation at 450g (from EDTA- or ACD-containing blood tubes).

i.

After having spun the blood, take the buffy coat off with about 100 μL of plasma using a disposable sterile Pasteur pipette; be careful not to lift red blood cells.

ii.

Lyse the remaining red blood cells by addition of red blood cell lysis buffer at room temperature.

iii.

Spin the tube at 450g for 10 minutes at room temperature.

iv.

Resuspend the pellet.

v.

Aliquot as appropriate into labelled cryovials.

vi.

Place in LN₂ to snap-freeze.

vii.

Store in LN₂.

4.1.8. Whole blood

Whole blood is to be prepared from EDTA tubes. The anticoagulated blood can be snap-frozen as it is. If the blood cells are needed intact, DMSO is needed to keep them alive while freezing.

i.

Dispense 50 μL of DMSO into two 1 mL sterile cryovials.

ii.

Invert the EDTA tube twice, and then add 450 μL of blood to each cryovial.

iii.

Invert the cryovial to mix the whole blood with the DMSO. Note: DMSO is cytotoxic at room temperature; therefore, as soon as it is mixed with blood, it should be placed in a controlled-rate freezer.

iv.

Transfer to −80 °C after at least 4 hours.

4.2.1. Snap-freezing

Snap-freezing is the process by which samples are lowered to temperatures below −70 °C very rapidly using dry ice or LN₂. This method can provide excellent sample integrity and a wide array of options for tissue analysis, including extraction of proteins, RNA, and DNA for use in research diagnosis. Before tissues are stabilized by freezing, the protein, RNA, and DNA profiles can change, and these changes depend on the duration of warm and cold ischaemia and the ambient temperature before freezing. All the different pre-analytical conditions and durations should be documented.

4.2.2. Storage of tissue

Storage of tissue can be done according to different protocols according to the equipment available in the facility. Options for storage:

i.

Transfer the snap-frozen sample from the isopentane to a pre-chilled storage container for transfer to either a locked −80 °C freezer or a LN₂ storage facility in the liquid or vapour phase. For storage for longer than 5 years, LN₂ in the liquid or vapour phase is recommended.

ii.

Place cryostraws in a designated visotube within a goblet (removable LN₂ storage elements) and place in the locked LN₂ repository.

a.

Store duplicate samples in a different storage facility if this is available.

b.

Check the backup system for the storage repository – either a backup freezer running constantly or adequate supplies of LN₂.

c.

Record the storage details in the inventory system, and check earlier data that were entered. At a minimum, the information recorded will include the inventory number (local sequential code), the location, the pathology number, the type of tissue (the site, and also whether the sample is tumour, unaffected/normal, and/or pre-malignant), the lag time between excision and freezing, and the date.

4.2.3. Storage of FFPE blocks and slides

i.

FFPE blocks and sections mounted on slides can be stored at room temperature. Prevent exposure of blocks to sun or extreme temperature variation or humidity.

ii.

Store blocks in moisture-resistant cardboard boxes or plastic storage boxes.

iii.

Transfer details to the computerized database system.

iv.

Update the database when samples are moved or depleted.

4.2.4. Formalin fixation

Formalin fixation is standard practice in most routine histopathology laboratories. The following guidelines address specific issues related to preservation of formalin-fixed specimens in biobanks. Table 6 provides information on the composition of neutral buffered formalin.

Table 6

Composition of neutral buffered formalin and 70% ethanol.

i.

Tissue specimens should not be bigger than 3 cm × 2 cm × 0.5 cm.

ii.

Specimens should be fixed in fresh 10% neutral buffered formalin for a minimum of 4 hours and a maximum of 48 hours, after which they should be embedded in paraffin in accordance with conventional techniques.

iii.

All reagents should be DNase- and RNase-free (e.g. prepared using diethylpyrocarbonate [DEPC] water).

iv.

Fixation media, such as Bouin’s solution, that contain picric acid should be avoided, because this compound interferes with subsequent PCR analysis of extracted nucleic acids.

v.

Alcohol fixation may be used as an alternative to formalin fixation. For this, tissue is placed into 70% alcohol (diluted with DEPC water) for a minimum of 4 hours.

Because of the chemical hazards of formalin, it can be desirable to use alternatives to formalin as a routine fixative. However, the effect of long-term storage with these alternative fixatives on the desired macromolecules is not always known and should be established empirically.

4.2.5. RNAlater

This substance protects RNA in fresh specimens. It eliminates the need to immediately process or freeze samples.

4.2.6. Shipment of tissues and slides

FFPE tissues and slides are shipped at ambient temperature in accordance with the established shipment guidelines and protocols of the sending and recipient institutions. Please refer to Section 3.9 for more details.

4.2.7. Quality control for tissue samples

i.

Ensure that the reagents have not expired and are of the correct composition and volumes.

ii.

Keep high-quality records on all variables related to specimens, FFPE tissues, and slides, including the time of tissue collection, the processing time, and the period of storage before shipment and/or use.

4.2.8. Data to be recorded

i.

Date and time of tissue collection.

ii.

Number of unprocessed samples, FFPE blocks, and slides prepared.

iii.

Date and time of shipping.

iv.

Any variations or deviations from the protocol, problems, or issues related to the collection and storage.

4.3.1. Urine

i.

Plastic or glass containers for collection of urine should be clean and dry, should have a 50–3000 mL capacity, a wide mouth, and a leakproof cap, and should be clearly labelled.

ii.

When in transit, urine collections should be maintained on ice or refrigerated.

iii.

Urine should be aliquoted according to the volume needed for analysis or storage.

iv.

Depending on the analyte to be measured, a preservative may be added during collection or before aliquoting.

v.

Store urine at −80 °C or below in LN₂.

4.3.2. Buccal cells

i.

A collection kit (containing mouthwash, a 50 mL plastic tube, a plastic biohazard bottle, and courier packaging) is mailed or given to the participant, along with an instruction sheet. The participant is to brush their teeth as usual, rinse their mouth out well with water twice, and then wait 2 hours. The participant should not eat or drink anything other than water during this time.

ii.

After 2 hours, 10 mL of commercial mouthwash should be poured into the tube, and then 10 mL of tap water should be added. This diluted mouthwash should be placed into the mouth (without swallowing) and swished around vigorously for 30 seconds.

iii.

The mouthwash should then be spat back into the plastic tube, and the tube should be sealed tightly.

iv.

The sample should be sent back to the biobank immediately for processing, or stored at 4 °C until it is sent, but it should be sent within 24 hours.

v.

When the sample arrives at the laboratory, transfer the mouthwash to 15 mL conical test tubes.

vi.

Add 35 mL of Tris-EDTA to the mouthwash sample and spin at 450g for 5 minutes.

vii.

Decant the supernatant and discard.

viii.

Wash the cells twice, each time with 45 mL of Tris-EDTA.

ix.

Resuspend the cell pellet in 50 μL of Tris-EDTA and transfer to 2 mL labelled cryovials.

x.

Store the sample at −80 °C or in LN₂.

Note: Buccal cells can also be collected with other means, such as brushes.

4.5.1. Taking the sample of cells

Insert the long tip of the spatula into the cervical os, and rotate the spatula through a full circle (360°). If the cervical broom brush is used, place the tip of the brush into the cervical os, and rotate the brush gently through three 360° circles.

4.5.2. Taking the Pap smear

i.

Smear both sides of the spatula (or the contents of the brush) onto the glass slide with one or two careful swipes. If any abnormalities are seen outside the area sampled, take a separate specimen and smear it onto another slide.

ii.

Immediately fix each slide. Either use spray fixative, at a right angle to, and at a distance of 20 cm from, the slide, or immerse the slide in a container of 95% ethanol for at least 5 minutes. If the slide is not fixed immediately, the cells will dry and become misshapen; it will then not be possible to read the slide accurately in the laboratory.

iii.

Gently close and remove the speculum.

iv.

Place all used instruments in decontamination solution.

4.5.3. After taking the smear

i.

Label the frosted edge of each slide carefully.

ii.

On the patient record, note and illustrate any features you have noted: visibility of the transformation zone, inflammation, ulcers or other lesions, or abnormal discharge. Note whether other samples were taken, for example Pap smears of other areas, and if the woman has been referred elsewhere, note to whom and when.

4.6.1. Hair

Head hair may be collected as follows.

i.

Along an imaginary line drawn across the middle of the back of the head from the centre of one ear to the centre of the other, gather a lock of hair at least the thickness of a pencil, and tie it together near the root end (near the scalp) using a small string or a rubber band.

ii.

Cut the hair as close to the scalp as possible without cutting the scalp.

iii.

Maintain the horizontal position of the hairs in the bundle by wrapping the cut section in aluminium foil or plastic wrap.

iv.

Indicate the root end and the tip end by marking the foil or plastic wrap with a permanent marker or with a paper label. Do not use tape on the hair itself.

v.

Place the specimen in a clean, dry, labelled paper envelope for shipment to the laboratory. Note whether bleaches, hair dye, or medications (e.g. selenium or minoxidil) were used.

Please note that hair from other sources (pubic, axillary, beard, moustache, chest, etc.) may also be analysed if head hair is not available.

4.6.2. Nails

A clean pair of nail clippers should be used. To clean nail clippers thoroughly, they should be rubbed with alcohol swabs. Nails should be clean of all polish, dirt, and debris. Nail clippings from each finger or toe should be collected and packaged separately in plastic bottles. Each bottle should be labelled with the mass of the nail collected and its source (e.g. right index finger) (NMS Labs and ExperTox).