BIOSAFETY

GENERAL CONSIDERATIONS

1. The main objective is to bank as much of the specimen as possible without compromising the ability to render the definitive diagnosis. The patient always comes first.
   
   
   1. The diagnostic paraffin block can provide invaluable banked sections that can be used for in situ hybridization (ISH), immunohistochemistry (IHC), and polymerase chain reaction (PCR) studies.
      
   2. There is no need to stuff a cassette with tissue. It is wasteful and often counterproductive. A 2 mm thick piece of tissue (thickness of a quarter) is more than adequate.
      
   3. The formalin fixative should be 10% (of the 40% commercial stock) and buffered. Commercially available buffered formalin is often best for reproducibility.
      
      
2. Complete patient confidentiality must be ensured at all times.
   
   
3. Approved informed patient consent forms must be on file in your institution's Office of Human Research.
   
   
4. Standard practices presently in use by participating pathology departments should be reviewed with the needs of the patient and the ACSB in mind. Considerations include:
   
   
   1. Whether portions of a specimen need be routinely fixed in multiple fixatives
   2. Is flow cytometry routinely necessary?
   3. How much tissue is really needed for flow cytometry?
   4. Does a block really need to be over 2 mm thick?
   5. Is it necessary to freeze tissue for gene rearrangement studies on all cases?
   6. Some diagnosticians believe that frozen tissue is preferable to paraffin-embedded tissue for immunohistochemistry
   
   
5. No specimen is to be permanently deposited in the ACSR until a final diagnosis has been rendered. If necessary, the frozen tissue held for potential banking can be retrieved for further processing (e.g. IHC, gene arrangement, and paraffin embedding).
   
   
6. Close cooperation between diagnostic pathologists, bankers, and researchers is mandatory.
   
   
7. Diagnostic, banking, and research methods change. Therefore, constant review and updating of ACSB policies are mandatory.
   
   
8. Diagrams prepared at the time of specimen processing are very helpful, e.g. verifying, labeling and sampling.
   
   
9. Optimal and standardized preservation of banked specimens is mandatory.
   
   
10. Banked specimens must be in the physical possession of each bank site before inclusion in the national registry.
    
    
11. Accurate inventories of specimens must be maintained.
    
    
12. Both clinical correlation and follow-up are important to the actual specimen for the ACSR.
    
    
13. Appropriate and sufficient controls must be collected (in the order of 10%).
    These controls may include:
    
    
    
    1. "Uninvolved" portions (margins) from resection specimen
    2. "Normal" tissue from HIV patients
    3. Specimens from non-HIV patients with processes similar to those banked from HIV patients
    
    
14. A validation slide that mirrors the banked specimen must be simultaneously banked and readily accessible.
    
    
15. Virtually no specimen is so small that a portion of it cannot be frozen.
    
    
16. A lesion that has been previously diagnosed, e.g., by FNA, could have more of subsequently resected specimen banked than one that is a total unknown.

SPECIMEN LABELING - still under determination by the ACSR Committee

SPECIMEN HANDLING - "THE ZEBRA METHOD"

1. Specimens should be transported on a moist substrate but never submerged in any fluid. A saline-moistened gauze pad is adequate.
   
   
2. The specimen should be handled using an aseptic technique.
   
   
3. Considerations in larger specimen processing:
   
   
   1. Larger specimens should be sampled sequentially, i.e., a 2 mm thick piece of tissue for paraffin embedding followed by a 5 mm thick piece for freezing, and so on. This produces multiple banked paraffin blocks and frozen specimens.
   2. A thin shaving of one or more of the tissues that will be frozen should be rapidly placed in glutaraldehyde for Transmission Electron Microscopy (TEM).
   3. The frozen tissue is linked to a validation slide derived from the adjacent paraffin-embedded block.
   4. The unfixed tissue is divided into multiple 5 mm³ blocks for freezing, alternating with and without OCT-coating. Specimens thinly coated with OCT are ideal for frozen section immunophenotying and molecular studies. The snap-frozen, uncoated specimens are ideal for PCR analysis (mRNA and DNA) and molecular studies.
   5. A portion of tissue designated for freezing can also be set aside, sampled for flow cytometry or used for cell culture, if necessary.
   
   
4. Considerations in small specimen processing:
   
   
   1. A 1 mm central slice or one-half of the specimen (e.g. a 3 mm-punch biopsy) will be used for paraffin-embedding and the rest for TEM and freezing: prepare OCT-coated specimen first followed by an uncoated specimen, if there is enough.
   2. It is critical, especially with small specimens, that a fresh sharp razor blade or scalpel be used for dividing tissue (to prevent crushed-artifact).
   
   
5. Specimen processing for diagnosis and banking is best performed in the frozen section of the cutting room at the same time as processing for Surgical Pathology. Do not freeze tissue and then later divide it for the ACSR. We must get to the tissue before it is placed in formalin for transport or storage.
   
   
6. There are benefits to using "see-through" freezing tubes with volume markers and a site for labeling, e.g., Sarstedt Screw-Cap Micro Tube. This tube (approximately 2 ml in volume with a 7 mm opening) is ideal for storing 5 mm³ frozen pieces of tissue and visualizing what is available in the tube.

TRIAGING OF SPECIMENS

1. Glutaraldehyde Fixation of Specimens for Plastic Section Light Microscopy and Transmission Electron Microscopy (TEM)
   
   
   1. No specimen is so small that a TEM specimen cannot be taken. If you can see it, it is large.
   2. A thin shaving of viable, representative tissue should be placed in 5 - 10 ml of neutral-buffered glutaraldehyde in a scintillation vial (transferred directly from scalpel/razor blade).
   3. If the specimen is variegated, more than one vial should be used and labeled, noting the relationship to the paraffin section/frozen tissue.
   
   
2. OCT-Coated Specimen
   
   
   1. Place the first piece of available tissue, not to exceed approximately 5 mm³ (0.5 gram)* in a freezing tube which has an orifice of approximately 0.7 cm. If the specimen is larger than 5 mm³, it will be difficult to remove the tissue from the tube.
   2. Roll the tissue in enough OCT to produce a thin coating. Too much OCT may interfere with tissue preservation. A small piece of aluminum foil can be used as a working surface. The foil-wrapped OCT-coated specimen can then be placed in the tube.
   3. Cap tube just enough to secure a seal and label appropriately with an indelible pen, e.g., Cryopen.
   4. Place the freezing tube in liquid nitrogen to snap freeze.
      • NOTE: It is essential that the specimen not be larger than 0.5 grams (5 mm³) in order to ensure complete and rapid freezing of the entire specimen. If the piece of tissue is too large and has been stuffed into the tube, uniform freezing will not occur.
   5. If there is only a single 5 mm³ specimen available for freezing, it should be cut in half. One-half should be used for OCT and one-half for uncoated snap freezing (see #3 below).
   6. Anything smaller than 4 mm³ should only be used for OCT coating.
      
      
3. Snap-Frozen Uncoated Specimen
   
   
   1. The second 5 mm³ piece of tissue should be placed in a freezing tube WITHOUT cryopreservative i.e., no OCT, labeled and snap frozen in liquid nitrogen.
   2. The same specimen size requirements must be met in order to ensure complete and rapid freezing of the entire specimen.

PREPARATION OF PLASMA AND MONONUCLEAR CELLS

• 10% DMSO (dimethylsulfoxide)
  
• 40% fetal calf serum
  
• 50% RPMI
  
• 0.2 mm L-glutamine
  

1. Preparation of Plasma Samples
   
   
   1. The 7 ml tube of whole blood in acid citrate dextrose should be rotated gently 2 or 3 times before being centrifuged. Do not transfer before centrifugation.
   2. The cells are separated by centrifugation at 500 g for 10 minutes.
   3. Three 1 ml aliquots of plasma are removed and put into 1.5 ml screw top tubes and transferred to liquid nitrogen storage.
   
   
2. Peripheral Blood Mononuclear Cell Separation and Freezing
   
   
   1. The cells and plasma remaining from the previous step are transferred into a 15 ml conical tube, capped and re-suspended by gently tapping the bottom of the tube.
   2. Sterile PBS should be added to the suspended cells until the final volume is 8 ml; invert to mix.
   3. The 8 ml whole blood-PBS mixture should be carefully overlaid onto 4 ml of room temperature LSM or Ficoll-Hypaque solution in a sterile 15 ml conical tube. A sharp interface should exist between the LSM and the whole blood mixture. (If the layer of LSM gets mixed with the blood-PBS, the tube should be gently rotated to mix the blood, PBS, and LSM, and transfer to a 50 ml sterile conical tube. Add equal volume of PBS to separate the cells at 600 g for 15 minutes. After removal of LSM-PBS supernatant, return to Step b).
   4. The 15 ml conical tube for 30 minutes at 900 g at room temperature. The mononuclear leukocytes (principally lymphocytes and monocytes) will band at plasma/LSM interface.
   5. The fluffy white layer just below the plasma layer should be aspirated off, along with with approximately half of the LSM layer under it, and transfered to an appropriately labeled 15 ml sterile conical centrifuge tube. Mix by gentle rotation.
   6. Washed twice in sterile PBS - centrifuge at 500 g for 10 minutes.
   7. Cell pellet should be mixed well with a gentle finger-tapping action.
   8. Using a 1 ml pipette, the *DMSO freezing mixture should be added dropwise to the cell pellet suspension. Gently finger-tap between drops. If the cell pellet is small, only 1 ml of freezing media is added (and only one aliquot of cells is frozen). If the cell pellet is large, up to 4 ml of freezing media can be added in a dropwise fashion. (Cell densities of 1 - 10 million PBMC/ml are best for cryopreservation. If a hemocytometer is available, the optimal concentration is 5 million PBMC/ml).
   
   *Important-Do not put the DMSO containing media on the cell button all at once.
   
   
3. Freeze the cell suspension in 1 ml aliquots in sterile NUNC vials by placing the NUNC tubes in a room temperature, alcohol saturated, control rate freezer container and store in the -80°C freezer overnight. Transfer the cell suspension into the liquid nitrogen temperature freezer for long term storage the next working day.

PREPARATION OF CEREBROSPINAL FLUID AND CELLS

• 10% DMSO (dimethylsulfoxide)
  
• 40% fetal calf serum
  
• 50% RPMI
  
• 0.2 mm L-glutamine
  

1. The total volume of CSF should be transferred sterilely to a 15 ml conical tube and centrifuged at 500 g for 10 minutes. The supernatant should be aliquoted into 1 ml portions with a sterile 1 ml serologic pipette and placed in NUNC tubes for liquid nitrogen storage.
2. In general, no appreciable cell pellet will be visible. In fact, there may only be a few thousand cells. This is still an adequate sample for PCR. From a 1 ml pipette, add the DMSO freezing mixture dropwise to the cell button suspension. Gently finger-tap between drops.
3. Freeze the cell suspension in 1 ml aliquots in sterile NUNC vials by placing the NUNC tubes in a room temperature, alcohol saturated, control rate freezer container and store in the -80°C freezer overnight. Transfer the cell suspension into the liquid nitrogen temperature freezer for long term storage the next working day.

PREPARATION OF PLEURAL, ASCITIC, AND SYNOVIAL FLUID AND CELLS

• 10%DMSO (dimethylsulfoxide)
  
• 40% fetal calf serum
  
• 50% RPMI
  
• 0.2 mm L-glutamine
  

1. The total contents of each vacutainer should be transferred to 15 ml sterile conical tubes and centrifuged at 500 g for 10 minutes. The supernatant should be aliquoted into 1ml portions with a sterile 1 ml serologic pipette and placed in NUNC tubes for liquid nitrogen storage. No more than 5 aliquots of the liquid phase will be required.
2. In general, small cell pellets will be visible. These pellets should be washed twice in sterile PBS.
3. Using a 1 ml pipette, add the DMSO freezing mixture dropwise to the cell button obtained from each of the vacutainers. Gently finger-tap between drops.
4. Freeze the cell suspension in 1 ml aliquots in sterile NUNC vials by placing the NUNC tubes in a room temperature, alcohol saturated, control rate freezer container and store in the -80°C freezer overnight. Transfer the cell suspension into the liquid nitrogen temperature freezer for long term storage the next working day.

PREPARATION OF BONE MARROW ASPIRATE

• 20% DMSO (dimethylsulfoxide)
  
• fetal calf serum
  
• 0.2 mm L-glutamine
  

1. Do not try to separate cells or do a cell count.
2. Carefully mix aspirate 1:1 with 2x concentrated DMSO freezing mixture and freeze in 0.5 ml aliquots in NUNC tubes.
3. Place the NUNC tubes in a control rate freezer container and store in the -80°C freezer overnight. Transfer the cell suspension into the liquid nitrogen temperature freezer for long term storage the next working day.

PREPARATION OF CLARIFIED URINE AND URINARY SEDIMENT

• 10%DMSO (dimethylsulfoxide)
  
• 40% fetal calf serum
  
• 50% RPMI
  
• 0.2 mm L-glutamine
  

1. The volume of urine should be transferred under sterile conditions to 50 ml conical tubes and centrifuged at 500 g for 10 minutes. The supernatants should be aliquoted into 1ml portions and placed in NUNC tubes for liquid nitrogen storage, using sterile serologic pipettes. No more than 5 tubes of urine will be necessary.
2. In general, a small "dirty" cell pellet will be visible. In fact, there may only be a few thousand cells and a large amount of cellular sediment. This is still an adequate sample for PCR or tissue culture. Wash the cell pellet twice in PBS.
3. Using a 1ml pipette, add the DMSO freezing mixture dropwise to the cell button obtained from each 50 ml of urine. Gently finger-tap between drops.
4. Freeze the cell suspension in 1 ml aliquots in sterile NUNC vials by placing the NUNC tubes in a room temperature, alcohol saturated, control rate freezer container and store in the -80°C freezer overnight. Transfer the cell suspension into the liquid nitrogen temperature freezer for long term storage the next working day.

PREPARATION OF BUCCAL RINSES

1. Have patient rinse mouth with PBS (optional)
2. Scrub sides of cheeks firmly with a cytobrush
3. Rinse mouth with 10 ml of PBS and expels contents into a labeled, 50 ml sterile conical tube
4. Spin tube at 2,000 rpm for 10 minutes at room temperature
5. After centrifugation, carefully decant supernatant into labeled waste tube. Touch lip of sample tube to small square of lab soaker to catch last drop of PBS
6. With transfer pipette, use 0.5 ml PBS to transfer cell pellet to Eppendorf tube (try to avoid non-cellular debris). Save pipette in tube and with new pipette add 0.5 ml more PBS to rinse tube and old pipette, transfer to same Eppendorf
7. Spin Eppendorf tubes in micofuge at 3,000 rpm for 10 minutes at room temperature. Decant supernatants
8. Measure pellet size:
   

   +	=	0.1 ml
   ++	=	> 0.1 ml
   +++	=	0.25 ml
   ++++	=	> 0.25 ml

9. If pellet is larger, split into 2 tubes. Add 1 ml STM/tube and freeze at -70°C

ANO-GENITAL SPECIMEN COLLECTION PROTOCOL

1. Collection of Anal Swab Material for Anal Cytology
   1. Moisten Dacron swab with tap water or normal saline
   2. Have alcohol cytology bottle open at bedside
   3. Insert anal swab as far into the anal canal as possible
   4. With gentle pressure on the walls of the anal canal, twirl the swab as you withdraw
   5. Smear material onto labeled glass slide as quickly as possible; delay of more than 10 seconds may result in air-drying.
   
   
2. Collection of Anal Swab Material for HPV Testing
   1. The recommended collection and storage system is the HPV DNA Specimen Collection Kit (Digene Diagnostics #4203-0020)
   2. Moisten Dacron swab from kit with tap water or normal saline
   3. Insert anal swab as far into the anal canal as possible
   4. With gentle pressure on the walls of the anal canal, twirl the swab as you withdraw
   5. Insert swab into collection kit tube and cap. Freeze as soon as possible at -70°C to -80°C
   
   
3. Collection of Cervical Swab Material for HPV Testing
   1. The recommended collection and storage system is the HPV DNA Specimen Collection Kit (Digene Diagnostics #4203-0020)
   2. Moisten Dacron swab from kit with tap water or normal saline
   3. Insert cervical swab into the endocervix and rub against exocervix per kit instructions. A cervical cytobrush may also be used and may increase cellularity, but may also result in more bleeding
   4. Insert swab into collection kit and cap. Freeze as soon as possible at -70°C to -80°C
   
   
4. Collection and processing of Cervicovaginal Lavage for HPV Testing
   1. 10 ml of sterile normal saline is sprayed against the cervical and the exocervix using a syringe equipped with a 2-inch 18 gage angiocath type Teflon catheter
   2. The fluid is then aspirated from the posterior vaginal fornix using the syringe and transferred to a 15 ml sterile polypropylene tube
   3. If the volume recovered is less than 6 ml, a second lavage using 5 ml of sterile normal saline is performed. Add volume recovered to 15 ml tube
   4. Gently vortex specimen and store 1 ml aliquots in Sarstedt tube (#72.694.006)

SALIVA PROTOCOL

1. Remove lipstick and any removable dental prosthesis.
2. Subjects should rinse thoroughly with de-ionized water and rest for five minutes before saliva collection begins.
3. Collect whole saliva in a 50 ml tube by allowing saliva to accumulate in a closed mouth and spitting into a tube (on ice) every minute for about 5 minutes.
4. Saliva should be suspended in 10% DMSO preservative.
5. Can be stored on dry ice, –70° freezer, or liquid nitrogen.

• Sterile Nunc tubes
• Sterile transfer pipettes
• Dimethylsulfoxide (DMSO)
• Sterile pipette tips
• Air displacement pipettes
• Rate controlled freezer container
• Vortex
• Sterile 1x PBS

1. Follow Universal Precautions. Open and process samples under a biohazard hood.
2. Verify specimen label matches paperwork.
3. Using a sterile transfer pipette, determine the approximate volume of saliva. (If the saliva is not loose enough to pipette, add just enough sterile saline to make it loose and vortex. Then measure final volume.)
4. Add DMSO to make a final dilution of 10% DMSO.
5. Vortex briefly to mix.
6. Aliquot into Nunc tubes, 0.5 ml per tube.
7. Label the tubes as follows:
8. Place labeled tubes in a rate controlled freezer container. Place the container in a –700 C freezer overnight. If a –70 C freezer is not available, freeze and hold on dry ice until shipped to repository.
9. Next day, place tubes in freezer boxes for LN2 storage.

1. Patient #
2. Date and time of collection

• Patient #
• Date and time of saliva collection.
• Specimen type, i.e. saliva
• Specimen purpose: Donation