CD34 Enumeration in the Clinical Laboratory Learning Objectives Contents

Transcription

1 CD34 Enumeration in the Clinical Laboratory Vera S. Donnenberg, Ph.D., Assistant Professor of Surgery Deborah L. Griffin, M.S., Manager QA Cellular Therapies Learning Objectives To understand and perform daily instrument setup and quality assessment To understand the theoretical basis for single-platform flow cytometric determination of absolute counts (bead calibration, lyse no-wash) To understand the ISHAGE gating strategy for detection of CD34 cells, and the exclusion of non-viable cells by dye uptake To stain, acquire and interpret a process control sample To stain, acquire and interpret an unknown sample To relate absolute hematopoietic CD34 count to endothelial and mesenchymal lineages of the human adult bone marrow Contents 2. Introduction to CD34 Enumeration 3. Vera and Debe s short cut 4. SOP: QC Daily Start-up, Shutdown and Maintenance 5. SOP: CD34 staining CD Chex Process Control Sample 6. SOP: CD34 Staining with Stem-Kit 7. Hematopoietic, Endothelial and Mesenchymal Stem Cell Analysis of Human Adult Bone Marrow 8. Reprint: Donnenberg AD, Koch EK, Griffin DL, Stanczak HM, Kiss JE, Carlos TM, BuchBarker DM, Yeager AM. Viability of Cryopreserved BM Progenitor Cells Stored for More than a Decade. Cytotherapy 4(2): , Product Insert: Flow-Check Fluorospheres 10. Product Insert: Flow-Set Fluorospheres 11. Product Insert: CaliBRITE Beads 12. Product Insert: CD-Chex CD Product Insert: Stem-Kit Page 1 of 48

2 Introduction Vera S. Donnenberg, PhD Assistant Professor of Surgery University of Pittsburgh Hillman Cancer Research Pavilion L Centre Ave Pittsburgh PA Phone Fax Deborah L. Griffin, MS Manager, QA Cellular Therapies University of Pittsburgh Medical Center Hillman Cancer Center Suite 1a 5150 Centre Ave Pittsburgh PA Phone Fax Hematopoietic Progenitor Cell Enumeration CD34 progenitor cells usually comprise 1% or less of a bone marrow or mobilized peripheral blood progenitor cell product. According to our release criteria, a pure, potent and safe product would have an adequate progenitor cell dose (5 x 10 6 CD34/kg), high viability (>99%) and be negative for bacterial or fungal contamination. A single platform, bead-calibrated assay was used to assess CD34 purity, absolute CD34 content and CD34 viability (Figure), using the landmark gating strategy of Sutherland (1). Absolute white cell count (WBC) was also measured using CD45 to identify leukocytes. CD34 and WBC viability was determined by exclusion of the fluorescent agent 7-amino-actinomycin D (2). Single platform assays include those using a volumetric approach (3), and those that are calibrated with reference to beads of a known concentration. The later approach will be demonstrated. Product, added as precisely as possible using a positive displacement pipette, is incubated with antibodies. Red cells, if present, are lysed by addition of ammonium chloride. Before acquisition, a precise volume of uniform beads of known concentration (StemCount, Beckman-Coulter) is added. Since the sample is never washed, there is no possibility of cell loss. The number of events of interest counted Page 2 of 48

3 (CD34+ cells) is compared to the number of beads counted. Since the concentration of Single platform determination of CD34 content and viability of a peripheral hematopoietic progenitor cell graft. Graft aliquots (100 μl) were stained in duplicate with anti-cd45-fitc and anti-cd34-pe, in a bead-calibrated lyse/no wash assay (Beckman-Coulter StemKit). A sample stained with the same reagents plus great excess of unlabeled anti-cd34 was run as a negative isoclonic control. Cells were acquired on a Beckman-Coulter XL cytometer. Stem Count beads are identified by their high fluorescence in FL4 vs. time (region A). These are removed from subsequent analyses using a not gate. Collecting time as a parameter facilitates identification of fluidic and other instrument problems during sample acquisition. White blood cells are identified by CD45 expression (region B). The WBC gate, compounded with the not bead gate is passed to a histogram of anti- CD34 versus side scatter. CD34+ events with low side scatter are identified (region C). The compound gate of WBC, not bead and CD34+ is passed to a histogram of CD45 versus side scatter, where cells of intermediate CD45 expression are identified (region D). The compound gate of WBC, not bead, CD34+, and intermediate CD45 expression is passed to a histogram of forward scatter versus side scatter (region E), which is used to eliminate events with low forward scatter. The number of CD34+ cells obtained in region E, taken as a percent of WBC identified in region B, represents the percent CD34+ cells as defined by the International Society of Hematotherapy and Graft Engineering (ISHAGE, now the International Society of Cellular Therapy). For determination of the viability of CD34+ cells, as defined by ISHAGE, the compound gate of WBC, not bead, CD34+, intermediate CD45+ expression and intermediate to high forward scatter is passed to a histogram of forward scatter versus intracellular 7AAD concentration (FL3). Because 7AAD is excluded by viable cells, viable CD34+ cells are detected in region F. Even in samples with low overall viability, the proportion of dead cells within the F region is invariably low. This is because the majority of dead and dying cells have low forward scatter and are eliminated by gate E. It is for this reason that we measure and report CD34 viability using a compound gate of WBC, not bead, CD34+ and intermediate CD45 expression, where viable CD34+ cells are detected in region G and reported as a percent of cells in gate D. Similarly, viable WBC are detected in region H and reported as a percent of cells in gate B. Absolute WBC and CD34 counts are obtained by dividing the events in gates B and E, respectively, by the known StemCount bead concentration. In this sample, CD34+ cells represented more 3% of CD45+ events. The viability of total WBC (CD45+) and CD34+ cells exceeded 99%. Single Platform Quantification of CD34+ HPC WBC (NOT A) CD34+ (C NOT A) CD34+ CD45 INT (CD NOT A) CD34+CD45 INT FS INT (CDE NOT A) B 2.96% of WBC SS SS C SS D SS E CD45 FITC CD34 PE CD45 FITC FS ISHAGE CD34 VI (CDEF NOT A) TOTAL CD34 VI (CDE NOT A) CAL BEADS (UNGATED) CD45 VI (B NOT A) 7AAD 7AAD 100% CD34 Viability STEM COUNT A 7AAD 99.8% WBC Viability F G H FS FS TIME FS the beads is known, the concentration of the events of interest can be inferred with accuracy and precision not attainable in dual platform assays. Page 3 of 48

4 Quality Control Perhaps the least glamorous, but most important aspect of modern flow cytometry is the emphasis on quality control (QC). Like those pesky experimental controls, essential quality control measures are necessary for the valid interpretation of flow cytometric data. The instrument-related elements of a quality control program are shown in Table 1. The College of American Pathologists (CAP), which administers a quality assurance program for clinical flow cytometry laboratories, states that quality assurance measures must cover specimen and result integrity throughout pre-analytical, analytical and post-analytical processes. CAP provides a helpful checklist covering the areas that must be addressed by a quality assurance program. As a part of their accreditation process, laboratories also participate in surveys conducted twice or three times per year, in which blood samples are aliquoted and sent by express mail to participating laboratories. The results are tabulated and compared to the results of other laboratories using similar instrumentation and methods. Although the level of quality control required in clinical laboratories may be overkill in the research setting, the careful investigator would do well to review the CAP checklist and decide which standards should be adopted. Page 4 of 48

5 Table 1. Recommended Elements of Flow Cytometry Quality Control Element Procedure Frequency Comments Fluidics/optics Check CV of all Daily Poor CV can result parameters with from partial standard beads obstruction, drifting optical alignment, failing laser Electronics Set gain for all PMTs to place standard beads in target channels Daily Assures day-to-day consistency in fluorescence measurement. Assures that a balance between PMTs is maintained (important for compensation). Color compensation Stain single color Variable Stained controls may control samples be cells or beads and may be reagentspecific or fluorochromespecific Internal Concurrent staining Daily For standard cells, Process/Reagents of normal or mean percent preserved standard positive and absolute cells count with upper and lower limits are published by the Page 5 of 48

6 vendor Linearity/sensitivity Check fluorescence Yearly. More often Beads of graded intensity of multi- if quantitative fluorescence peak beads fluorescence intensity are run and measurements are the expected and made observed fluorescence are compared Page 6 of 48

7 The items listed in Table 1 are suggested for laboratories involved in human investigation to confirm consistent operation of the cytometer and performance of the reagents. Other QC issues such as sample integrity and consistency of data analysis are beyond the scope of this discussion, but should be considered as well. The first two steps of QC, optics and PMT setting verification, are performed with standard beads such as Beckman-Coulter Flow-Check and Flow-Set, respectively. Fluidics/optics verification is performed on a linear scale and the objective is to ensure that the beads give a coefficient of variation (CV, standard deviation divided by the mean) below a predetermined value. Tight CVs indicate proper laser alignment and good hydrodynamic focusing in the flow cell. When a partial obstruction occurs, the most sensitive parameter is forward light scatter, and this parameter will fall out of tolerance. Daily assessment of PMT gain is also important and easy to do. During your assay development phase, you will have chosen PMT settings (voltage and gain) appropriate to your test. For each fluorochrome, settings will have been chosen such that cell populations known to be negative are usually placed within the first decade, and the brightest positive populations are on scale. At that time, calibration beads, such as Beckman-Coulter Flow-Set would have been run, and the target channel (channel of bead mean fluorescence intensity) would have been noted for each PMT. During daily calibration, the same calibration beads are run again, and PMT gain is adjusted, if necessary, to place the beads at their target channel. This helps ensure that quantitative determination of fluorescence is consistent from day to day. The other ingredients necessary for quantitative fluorescence measurement are linearity of detection and calibration relative to a known standard. Linearity can be measured with multi-peak beads, and antibody binding can be calibrated to molecules of equivalent soluble fluorochrome (MESF) using beads of known antibody binding capacity. The topic of color compensation, and how often and by what method it should be confirmed, is controversial. In the two-color world, where fluorescein isothiocyanate (FITC) and phycoerythrin (PE) where the only fluorochromes used, the required amount of color compensation did not vary from day to day, providing that the instrument had been calibrated as described above. Correct two-color compensation can be verified daily, weekly, or even less often, using beads dyed with the fluorochromes of interest, such as Becton-Dickinson CaliBRITE beads. The introduction of tandem dyes such as Page 7 of 48

8 PE-Texas Red (also known as ECD), PE-Cyanine5 (PC5), PE-Cyanine7 (PC7), Allophycocyanine-Cyanine5 (APC-Cy5) and Allophycocyanine-Cyanine7 (APC-Cy7), opened up the world of polychromatic flow cytometry, but also introduced a twist to compensation. The emissions spectra of single fluorochrome dyes are always the same, regardless of the antibody to which it is conjugated, given that other relevant parameters (such as ph for FITC) are constant. This is not always the case for the tandem dyes, which can vary from manufacturer to manufacturer, lot to lot, and, over time, within the same vial. Most of this variability is explained by the amount of free PE, PE that behaves as if it were not part of a tandem dye. Tandem dyes are especially light sensitive, and the amount of apparent free PE can increase upon exposure to ambient light. Thus, the simple approach used for two-color cytometry, arriving at a single compensation solution to be used for all cells stained with the same fluorochromes, is not optimal in every setting. A method, advocated by some vendors, is to stain preserved or freshly isolated cells singly, with brightly staining antibodies representing each fluorochrome to be used in the multicolor combination. CD8 or CD45 are frequently used for this purpose. However, the assumption that the compensation required for the CD8-PC7 in your single stained standard is identical to that required by the CD4-PC7 in your multicolor stained sample may not always be correct. The alterative, to stain cells singly with each antibody used in your panel, is not ideal either, since some of the markers will stain dimly, and therefore will yield imprecise compensation settings. A solution advocated by some laboratories is the use of anti-ig capture beads, which bind all murine monoclonal antibodies equally well, regardless of their specificity or fluorochrome. Such standards could be run with each assay, or even acquired after the fact to confirm that correct compensation settings were used. As mentioned above, many new cytometers have the ability to save uncompensated high-resolution listmode data (16 or 20 bits), allowing compensation to be performed or corrected after acquisition. Page 8 of 48

9 Literature Cited 1. Sutherland DR. Anderson L. Keeney M. Nayar R. Chin-Yee I. The ISHAGE guidelines for CD34+ cell determination by flow cytometry. International Society of Hematotherapy and Graft Engineering. Journal of Hematotherapy. 5(3):213-26, Donnenberg AD. Koch EK. Griffin DL. Stanczak HM. Kiss JE. Carlos TM. Buchbarker DM. Yeager AM. Viability of cryopreserved BM progenitor cells stored for more than a decade. Cytotherapy. 4(2):157-63, O'Gorman MR. Gelman R. Inter- and intrainstitutional evaluation of automated volumetric capillary cytometry for the quantitation of CD4- and CD8-positive T lymphocytes in the peripheral blood of persons infected with human immunodeficiency virus. Site Investigators and the NIAID New CD4 Technologies Focus Group. Clinical & Diagnostic Laboratory Immunology. 4(2):173-9, Page 9 of 48

10 CD34 STAINING OF CELLS 1) Label 3 tubes with patient name and: Tube 1> 45/34, Tube 2> 45/34, Tube3> Isotube. 2) Add 20μl of Stem-Kit antibody to appropriate tube, i.e., 45/34 to tubes 1 & 2 and 45/isoclonic control to tube 3. 3) Add 20μl of 7AAD to each tube. 4) Pipet 100μl of patient sample to all 3 tubes. a) Using the same Eppendorf repeat pipettor (positive displacement) for the addition of sample and fluorospheres, draw blood into 500μl repeat tip. b) Wipe tip with Kimwipe. c) * Click pipettor once for priming, ejecting sample into original sample container. Precision is necessary for specimen pipetting or erroneous results may occur. 5) Cap all tubes and incubate for 20 minutes in the dark at room temperature. 6) After 20 minute incubation, add 2ml of lysing solution to each tube and vortex. 7) Cap tubes and incubate for 10 minutes at room temperature in the dark. 8) Add 100μl of fluorospheres to all 3 tubes a) Gently swirl the bottle of Stem Count Fluorospheres. Tap the fluorospheres lightly on the vortex but do not vortex vigorously as this may cause erroneous cell counts. Using the same Eppendorf repeat pipettor (positive displacement) as for the addition of sample, draw fluorospheres into 500μl repeat tip. b) Wipe tip with Kimwipe. c) * Click Pipettor once for priming. Precision is necessary for pipetting of beads or erroneous results may occur. 9) Before acquisition on the flow cytometer, label a 4 th tube as blank and add 2ml of distilled water to it. This blank can be used for multiple samples. 10) Samples are now ready for flow cytometric analysis. Page 10of 48

11 CD 34 ACQUISITION ON THE XL IN SYSTEM II 1. To run CD34 Stem-kit tubes: under Panel choose Stem Kit-7AAD, click Okay. 2. Vortex Tube 1, which is CD45/CD Place tube on sample port. 4. Instrument will stop acquiring when 75,000 events have been collected in Gate B or 999 seconds have elapsed. Watch event number. 5. After 1 st tube is collected, Enter Specimen ID box will appear. 6. Sample port will lower. Remove tube and wait for flashing green: Insert Sample Tube. 7. Repeat with Tube 2 Do not change any gates. The gates are set on Tube 1 and will remain the same for all remaining tubes for that patient. Just make sure beads are within gate or cells have not shifted, if necessary push Prime button and restart acquisition. 8. The next tube is the blank containing only distilled water. This will prevent any carry-over to the isotube. 9. The final tube will be the isotube, CD45/Isoclonic. This tube will stain all white blood cells but you should see little to no staining of CD Following acquisition, a report screen will appear. Click PRINT at the bottom right of screen. You should receive 4 sheets; Tube 1, Tube 2, Isotube and Final Report. Page 11of 48

12 University of Pittsburgh Medical Center University of Pittsburgh Cancer Institute/Children s Hospital of Pittsburgh HSC Laboratories Adult Program Pediatric Program 5117 Centre Ave th Avenue Pittsburgh PA Pittsburgh PA Procedure Name: R8 QC DAILY START-UP, SHUT DOWN AND MAINTENANCE PROCEDURES FOR COULTER XL-MCL PRIMARY AND BACK-UP INSTRUMENTS Date Adopted: May 26, 1999 Date Revised: September 12, 2006 Revision number: 8 Author: Griffin DL, Koch EK, Shierer-Fochler S, Albert D. Donnenberg Supersedes Procedure: NA Distribution: Adult and Pediatric HSC Laboratories Laboratory Director Medical Director Supervisor Date Date Date Technologist Annual Review Date Signature University of Pittsburgh Medical Center University of Pittsburgh Cancer Institute/Children s Hospital of Pittsburgh Page 12of 48

13 HSC Laboratories Adult Program Pediatric Program 5117 Centre Ave th Avenue Pittsburgh PA Pittsburgh PA R8 QC DAILY START-UP, SHUT DOWN AND MAINTENANCE PROCEDURES FOR COULTER XL-MCL PRIMARY AND BACK-UP INSTRUMENTS Principle A Daily Start-Up procedure including quality control checks is essential to ensure that the flow cytometer is working accurately and precisely. Flow-Check fluorospheres are used to check the stability of the optical and fluidic systems. Flow Set fluorospheres are used to set the voltage settings to predetermined values. CaliBRITE beads are used to set compensation between FL1 and FL2. The Daily Shut-Down procedure ensures that the machine is free of debris and biohazardous materials. All documentation referring to Flow Cytometry reagents and instruments shall be available in close proximity to the flow cytometer. This ensures that the technical staff operating the equipment is able to effectively utilize the instrument. Reagents and Supplies Supplies Supplier Catalog number Coulter-Clenz Coulter Clorox Bleach, unscented UPMC Stores Deionized water Millipore wall tap 12x75 Tubes Falcon 2052 CaliBRITE beads Becton Dickinson PBS GIBCO Flow Check Beads Beckman-Coulter Flow Set Beads Beckman-Coulter Page 13of 48

14 Instrumentation Flow Cytometer HSC Lab Hillman 1.54 Flow Cytometer HSC Lab Hillman 1.47 Beckman Coulter Inc Beckman Coulter Inc Primary Epics XL MCL System ID# Back-Up Epics XL MCL System ID# (green code) Reagent Preparation Cautionary Note: prepare bead tubes immediately before use. Protect beads from direct light. Discard after 1 hour. Flow Check Label a 12x75 tube Flow check. Gently mix the vial so that the beads come off of the side of the vial. Dispense ~200μl (approximately 7-10 drops) of Flow Chek into the tube. Cap until ready for use. Flow Set Label a 12x75 tube Flow Set. Gently mix the vial so that the beads come off of the side of the vial. Dispense ~200μl (approximately 7-10 drops) of Flow Set into the tube. Cap until ready for use. Unlabeled CaliBRITE Label a 12x75 tube Flow check. Gently mix the vial so that the beads come off of the side of the vial. Pipette ~1ml of PBS into the tube. One bead vial will be needed: unstained. Invert unstained bead vial completely and squeeze one drop of unstained beads into the PBS. The drop should be cloudy, indicating that beads are present. Cap until ready for use. Labeled CaliBRITE Label a 12x75 tube Flow check. Gently mix the vial so that the beads come off of the side of the vial. Pipette ~1ml of PBS into the tube. Three beads vials are necessary: unlabeled, FITC and PE. When dispensing, invert bead vial completely and squeeze one drop of each bead color into the PBS. The drop should be cloudy, indicating that beads are present. Cap until ready for use. Procedure Cautionary Notes Use Universal Precautions when handling fluids and instrumentation that comes into contact with biohazardous fluids such as the waste tank. If any of the procedures fail to produce the expected results, complete a Form 42 and initiate corrective action, including contacting the Flow Cytometry Technical Page 14of 48

15 Supervisor or the Beckman Coulter Service Center to have a Service Representative schedule service, as needed. Should it be necessary to run the back-up instrument in an emergency see below for detailed methods specific to the instrument in 1.47 Hillman. These instructions are below the section Emergency Service Call. Detailed Methods Daily Start-Up 1. Check fluid levels daily before operation. Fluid containers are located in the bottom right drawer of the instrument. If instrument is already turned on you must place it in idle mode to pull out reagent drawer. Simply press the RUN button, the button should have a yellow light. The button will flash green when in the idle mode. 2. Remove middle door panel on front of XL. If panel is not removed, the tubing in the drawer will catch on the panel. 3. Pull drawer straight out, open black cap for sheath tank. 4. Fill sheath fluid tank with deionized water. Do not overfill as air space is necessary to pressurize the tank. 5. Check cleanse tank fluid level and, if necessary, fill with Coulter Clenz 6. Tighten caps finger-tight, but do not over-tighten as it will be impossible to remove the caps later. 7. To empty waste container located in front of power supply unit: remove the cap for the waste, saturate a paper towel with bleach, hold it in one hand. Remove the cap with the other hand and place the float assembly part onto the bleach soaked towel. This is to ensure that biohazardous fluids do not drip onto the floor or the instrument. Remove the waste container and place the towel and cap into the holder. The hose has a tendency to shift (uncoil) if placed on the top of the power supply. Empty container into the sink, being careful to avoid splashing. Flush sink with copious amounts of water. Add ½ inch of Clorox to container and replace cap. 8. Press ctrl-alt-del 9. An MS-DOS menu will appear Previous version of MS-DOS is the correct selection that will start the cytometer. It will take at least 20 minutes to warm-up before being able to put samples on for acquisition. 11. Enter your operator initials. THIS IS MANDATORY. Click on initials in upper left corner. You will be prompted to enter your initials. Hit Enter. 12. Pull down Applications menu, select the QC screen. Pull down Screen menu, select Maintenance. The far right of the screen has the assignments for operator initials. If your initials are already assigned, you may begin the QC maintenance record. If not, Page 15of 48

16 chose a color block that is not assigned. There are 6 items located on the Power Supply that must be checked during Start-Up: Open Power Supply door (front) Value must be entered into Maintenance Spreadsheet for the following two gauges. If they do not fall within the normal ranges established by Coulter, make adjustments according to the Coulter Manual. System Pressure Check that the system pressure gauge reads 30+2 psi System Vacuum Check that system vacuum gauge reads at least 17 in. Hg. The following parts must be checked to ensure that they are free from damage and accumulated fluids or waste. See the Coulter Manual if these parts need adjustment or replacement. Vacuum Filter Air Filter Vacuum Trap Air Trap Record compliance with the Start-up Checklist by marking the appropriate boxes on the Maintenance Screen. 13. Pull down Application menu, select Acquisition. Pull down Settings menu, select protocol Pro2>_QFLOW CHECK BEADS. Click Region button, ensure that lot numbers are current. Modify lot numbers if necessary to match product in use. 14. Select protocol Pro2>_AStem FLOW BEADS. Click Region button, ensure that lot numbers are current. Modify lot numbers if necessary to match product in use. 15. Pull down Application menu, select Acquisition. Pull down Panel menu, select WeeklyQC/2COLOR COMP panel in Panel>Pro2> WeeklyQC/2COLOR COMP or Daily Morning QC panel in Panel>Pro2>Daily Morning QC 16. Status will read: Start-up in Process until initialization is completed. 17. Instrument is ready when status reads: Insert Sample Tube. If a Panel or Protocol is NOT selected, instrument will continue to read Start-up in Process. The instrument takes a minimum of 19 minutes to warm up the laser. Remove dih 2 O tube when sample stage drops. Press Prime at least twice to flush the system before running the first tube. 18. Gently mix the Flow Check and Flow Set vials so that the beads come off of the side of the vial. The beads have settled by gravity and will be visible as an orange band. Swirl the buffer in the vial until the band has disappeared. Do NOT over-vortex the beads. Gentle swirling by hand is usually sufficient to resuspend the beads. 19. Prepare three tubes: 200 μl Flow Check beads. 200 μl Flow Set Beads 1 ml dih 2 O Page 16of 48

17 20. Prime cytometer before running first tube. Acquire Flow Check beads on LOW flow rate, check fluidic stability as data acquires. The machine will prompt when the next tube is needed. Inspect that histograms 1-5 have PASSED. 21. Acquire Flow Set beads on MEDIUM flow rate. Ensure that the peak positin values(pkposx) are within the prespecified ranges in Region ID. The machine will prompt when the next tube is needed. 22. The machine will prompt when the next tube is needed. 23. Acquire dih 2 O. The XL will most likely display a message that read Unable to Autogate. Please Acknowledge. Click to acknowledge. Ensure that the number of events within the autogate is less than All three tubes MUST pass for samples to be run. Initial and date the printouts. Weekly Start-Up with Compensation 1. Compensation must be set on the first day that the flow cytometer is used during a normal business week. 2. Follow steps 1-15 above, preparing the cytometer. 3. Remove the beads from the refrigerator and gently swirl the container to resuspend the beads. The beads will not be visible through the vial. 4. Prepare the following tubes as above: 1) 200 l Flow Check beads 2) 200 l Flow Set Beads 3) Unstained CaliBRITE beads 4) stained CaliBRITE beads 5. Prime cytometer before running first tube. Acquire Flow Check beads on LOW flow rate, check fluidic stability as data acquires. The machine will prompt when the next tube is needed. Inspect that histograms 1-5 have PASSED. 6. Acquire Flow Set beads on MEDIUM flow rate. Ensure that the peak positin values(pkposx) are within the prespecified ranges in Region ID. The machine will prompt when the next tube is needed. 7. Acquire Unlabeled CaliBRITE beads on MEDIUM flow rate. Ensure that the population is falling within the negative quadrant in Histogram 2. The machine will prompt when the next tube is needed. 8. Acquire U/FITC/PE CaliBRITE beads on MEDIUM flow rate. Compare the mean fluorescence (MnI X) in Quadrant 1 (B1) with (MnI X) in Quadrant 3 (B3) population. Compare the mean fluorescence (MnI Y) in Quadrant 4 (B4) with (MnI Y) in Quadrant 3 (B3) population. The difference should be no greater than?? 9. The machine will prompt when the next tube is needed. 10. Acquire dih 2 O 11. All five tubes MUST pass for samples to be run. Page 17of 48

18 To Run the MCL (Multi-tube Carousel Loader) 1. At the red Acquisition Screen, click Setup Screen and select Worklist from the dropdown menu. A map of the MCL will appear as a spreadsheet-like screen. 2. To add a panel, click the Panel section of line 1. A list of panel will appear. 3. Select the panel desired by clicking on the name of the panel and click OK in the bottom left corner of the screen. 4. The panel will load with each tube occupying a line position. 5. Repeat as necessary for all control or patient panels required. Any combination of panel can be selected up to 32 tubes. 6. When the Panel column has been populated, click on Specimen ID in line 1 (ignore Patient Name). Type in required Specimen ID and hit Enter. 7. This will populate all the tubes of the first panel. If there are additional panel, click on the line containing the primary tube in that panel to enter the Specimen ID. 8. When Worklist is complete, either save the Worklist by choosing Worklist>Save As and give the Worklist an easily identifiable name such as the current date or select Setup>Run to immediately run the MCL. 9. Load the MCL in the order specified in the Worklist, ensuring that tubes are adequately vortexed prior to insertion. Don t forget that additional blank tubes will be necessary for patient panels. 10. Replace carousel in the MCL, ensuring that the carousel is seated properly. 11. Close the lid. 12. Click the large green RUN box on the Acquisition screen (lower right corner of screen). 13. Three dialogue boxes will pop up a. Do you want to start the MCL (Y/N)? Answer Y for Yes, N for No (used when restarting a stopped Worklist) b. Are you starting a new carousel (Y/N)? Answer Y for Yes, N for No (used when restarting a stopped Worklist) c. Do you want to enter specimen id (Y/N)? Answer Y for Yes, N for No. 14. The cytometer status message will read Cytometer Ready. The cytometer will acquire the tubes as specified by the Worklist. 15. Individual tubes will be printed out, but to obtain a report, it must be printed out under the Reports Screen. Select Acquisition>Reports. Select File>Select. 16. Select report to be printed. Click Okay. Results PRN box must be enabled. Click Results Output or Batch to print report(s). Shut-Down 1. Perform Shut-Down after all samples have been run or at the end of the day. Leave the instrument shut down until the following day or at least 30 minutes before restarting, to allow the Clenz to disinfect all surfaces. 2. Prepare a 10% Clorox cleaning solution: 1 part Clorox to 9 parts distilled or deionized water. Page 18of 48

19 3. Prepare fresh daily or the solution could contain contaminants or the chlorine could evaporate and decrease effectiveness. 4. You will need 9 tubes: 2 tubes with 2mL 10% Clorox 7 tubes with 2mL of distilled or deionized water. 5. Under acquisition, choose panel>select>change dir>..>pro3>click okay 6. Click on Cleaning panel 7. Check that cytometer status reads: Insert sample tube 8. Panel order will be: Clorox, water, water, water 9. Place the 1 st tube on the sample port. When tube is done, Specimen ID window will pop up. Hit return to get rid of window. 10. Continue with the next water tube when status reads: Insert Sample Tube and is flashing 11. Continue until all tubes have been run. 12. Place the remaining four tubes in a carousel in the MCL, same order as before: Clorox, water, water, water. 13. Press AUTO button. Two prompt windows will appear, hit return to eliminate the windows. 14. When MCL is done, remove the tubes. 15. To clean the Vacuum Line press the RUN button to take the system out of Run mode. The RUN light blinks green when the unit is in IDLE mode. 16. Fill a cleaning adapter reservoir with ~ 5 ml of distilled water. 17. Install the adapter just as a sample tube is installed on the sample stage. The water is aspirated in the vacuum line. Repeat. 18. Prepare two adapters with dih 2 O and place in carousel. Simultaneously hit the AUTO and CLEANSE buttons. Both adapters will be aspirated and the cleanse cycle will be run. The Cleanse cycle affects both the Manual and MCL sample ports and does not need to be run in each mode. When the cleanse cycle ends the CLEANSE button indicator turns off and the RUN button indicator flashes. The cytometer status message reads Cytometer Ready. Place a fresh tube of dih2o on the sample stage for shut down. 19. Wipe down all exposed surfaces of the instrument with 10 % Clorox or Dispatch and then with 70% ethanol. 20. The Sample Probe for the Manual and MCL sample ports must be wiped with a gauze pad saturated with Clorox and then wiped clean with gauze saturated with dih2o. Push up on the manual sample stage to expose the probe. For the MCL, push up the round sample head to expose the sample probe. 21. Record Shut-Down procedures done in Maintenance: Page 19of 48

20 Bleach Vacuum Line Sample Head Cleanse 22. Under Applications click Exit 23. Exit to DOS> hit Y 24. Type XLOFF to shutdown the XL at the MS DOS prompt. Weekly/Monthly/Bimonthly Cleaning/Yearly PM Air Filters 1. Clean the air filters once a week after the shut down procedure. There are 5 air filters: Cytometer back panel-2 Power supply-inside front door-1 Power supply-back panel-2 2. Make sure instrument is turned off at the computer. 3. Pull off each filter cover (they are not screwed in). They are made of flexible plastic and will snap out when pulled. 4. Pinch and pull out each filter. Handle gently to avoid damage. 5. Rinse each filter in water and squeeze excess water out. 6. Let dry for 30 minutes. Use paper towels to check that each filter is completely dry. 7. Return each filter to its holder and put each filter cover back on. Fluid Supply Containers Remove and clean the sheath fluid container once a month. 1. To remove a reagent container, put the instrument in idle mode by pressing the RUN button. It will flash green. 2. Remove the cytometer center front panel. 3. Pull out the reagent drawer. 4. Disconnect the tubing on the top of the container by pushing in on the metal clips on the connectors. 5. Disconnect the sensor at the back of the drawer by sliding its sleeve to the right (slide to the left if removing Clenz container). The sensor for the sheath container is on the right; for the cleaning agent container, on the left. 6. Remove container and empty. 7. Fill with mL of sterile deionized water. 8. Cap and swirl container, rinsing all surfaces. Page 20of 48

21 9. Empty and replace the container into the cytometer. 10. Reconnect sensor and top tubing. 11. Fill with fresh, sterile deionized water. Remove and clean the cleaning agent container every 60 days. 1. Follow steps 1-6 above to remove and clean Clenz container. 2. Fill with mL of fresh Coulter Clenz cleaning agent. 3. Cap and swirl container, rinsing all surfaces. 4. Empty and replace the container into the cytometer. 5. Reconnect sensor and top tubing. 6. Fill with fresh Coulter Clenz. 7. Record on the XL s Computer Maintenance Checklist all maintenance performed. The instrument will code the checklist with your initials and personal color. Yearly Preventative Maintenance (PM) Service Call 1. Refer to the HSC Laboratory QC Master List for the date that the PM is due. 2. Schedule the PM with Coulter by calling Give them our System ID # and the CMI Insurance number #CMI After the Coulter Service Representative completes the PM, record the date and your initials on the HSC Laboratory Master List. 4. FAX the PM service report to Rich Davis at File the original service report in the Flow Cytometry Equipment and QC Procedure Manual. Emergency Service Call 1. Determine that a service call is necessary after reviewing XL error messages and Troubleshooting manual. 2. Schedule the service with Coulter by calling Give them our System ID # and the CMI Insurance number #CMI5375. Indicate that this is an emergency and the machine is for clinical use. 3. After the Coulter Service Representative completes the service, sign and date the service report. 4. FAX the service report to Rich Davis at File the original service report and any supporting documentation in the Flow Cytometry Equipment and QC Procedure Manual. Page 21of 48

22 Operation of Back-Up Instrument 1. Check the green machine (room 1.47 Hillman) schedule and block out time as soon as possible. This schedule is found on-line in Outlook. Open public folders and then open UPCI Flow Cytometry. Schedule time on the green machine. 2. The instrument will always require our weekly 2-color compensation start-up panel. Take all necessary beads for this procedure to room The green machine is located in the farthest left corner on room 1.47 Hillman and is system ID# When you arrive the machine should be on the windows (Windows 98) operating system. 4. Exit out of the Expo 32 Software by hitting the X in the upper right hand corner of the window. 5. Double click on the System II Software Icon. (A red and green square with the Roman Numeral II) Our familiar System II Screen will appear. 6. Click on Panel. The directory that appears is Pro3 and not our familiar Pro2. 7. Click on the select directory blue button. 8. Click on the double dots (..) to go up a directory. 9. Select Pro2 directory and then click on the OK blue button. Our Flow Cytometry Directory appears. 10. Run the weekly 2-color compensation panel according to the detailed methods listed above in the section Weekly Start-Up with Compensation. 11. Run CD Chex CD34 and /or CD Chex Plus Controls as needed. 12. Run patient samples. 13. The next morning ask Cassie or Erin in Room 1.45 Hillman to burn a CD of our data from the back-up machine. 14. Load the data from the CD on to our primary instrument for permanent storage. References Operator s Manual Epics XL-MCL, Coulter Corporation Special Procedures and Troubleshooting Manual, Coulter Corporation Appendices Page 22of 48

23 Revision Change Rationale CFR/FACT/CAP/PA DH Standards 1 NCCLS format 2 Address Change 3 Address Change 4 New Terminolog y, Revision Chart Modificatio n 5 Addition of CAP terminolog y 6 Addition of MCL procedure 7 Addition of Flow Set and Flow Check beads to reagents 8 Title page format change Move from MUH to SHY hospital. Move from SHY to HCC FACT standards, second edition; addition of the inclusive dates of use Lack of information in procedure Inadvertently omitted D5.212 D8.230, A3.000, D5.600 FLO.30250; FLO.30260; FLO Start Date Annual review 09/12/06 End Date 05/13/03 10/21/03 10/21/03 01/30/04 01/30/04 11/05/04 11/05/04 3/29/05 3/29/05 09/12/06 Page 23of 48

24 University of Pittsburgh Medical Center University of Pittsburgh Cancer Institute/Children s Hospital of Pittsburgh HSC Laboratories Adult Program Pediatric Program 5117 Centre Avenue th Avenue Pittsburgh, PA Pittsburgh, PA Procedure Name: R7 QC PROCEDURE FOR CD34 AND CD3 STAINING WITH CD CHEX (PLUS) Date Adopted: March 1999 Date Revised: July 12, 2006 Revision number: 8 Author: Griffin DL, Koch E Supersedes Procedure: NA Distribution: Adult and Pediatric HSC Laboratories Laboratory Director Medical Director Supervisor QA Manager Date Date Date Date Technologist Annual Review Date Signature Page 24of 48

25 University of Pittsburgh Medical Center University of Pittsburgh Cancer Institute/Children s Hospital of Pittsburgh HSC Laboratories Adult Program Pediatric Program 5117 Centre Avenue th Avenue Pittsburgh, PA Pittsburgh, PA Page 25of 48

26 X:\UPCIFLOW\FLOW COURSE 2007\CD34 lab 2007\for Jada\temp.doc R7 QC PROCEDURE FOR CD34 AND CD3 STAINING WITH CD CHEX (PLUS) Principle CD34 + cell dose is the best available predictor of graft quality. In autologous transplantation the dose of CD34 + cells correlates with the time to neutrophil and platelet engraftment. CD34 + cell content is used to define cell harvest goals and guides growth factor administration and the number of leukapheresis sessions necessary for a graft. Stem-Kit is a set of reagents containing CD34 and CD45 monoclonal antibodies and Stem-Kit fluorospheres. This is a single-platform assay which directly measures the absolute (cells/μl) and relative (%) count of CD34 + cells in human marrow, blood, and blood derived samples by flow cytometry. CD-Chex is a stabilized preparation of human placental blood to be used as a complete process control when evaluating CD34 positive cells.when stained with monoclonal antibodies for CD34 positive cell enumeration, CD-Chex CD34 control will provide reference values for CD34 positive cells within the ranges on the assay sheet. The College of American Pathologists requires that 2 levels of positive cellular controls be analyzed daily to verify the performance of reagents, preparation methods and staining procedures for quantitative tests. CD Chex for CD34 enumeration is produced in Level I, Level II, and Level III forms. CD Chex Plus for CD3 enumeration is produced in Normal and High Levels. This assay is to be run every day that CD34 cells are enumerated. It does not need to be run when only CD3 cells are being enumerated. On days that only CD34 is required, CD3 controls do not need to be performed. All control specimens must be tested in the same manner and by the same personnel as patient samples. Specimen or Component Requirement CD Chex contains placental cord blood cells in a preservative medium, and is shipped in vials, 2 vials to one box of Level III, and 4 vials to one box of Level I and Level II; ~3 tests per vial. Reagents and Supplies Supplies Supplier Catalog Number Sterile Water for Irrigation, USP Baxter 2F7114 CD-Chex Streck Laboratories Page 26 of 48

27 X:\UPCIFLOW\FLOW COURSE 2007\CD34 lab 2007\for Jada\temp.doc CD-Chex Plus Streck Laboratories Stem-Kit Coulter 2390 CD3 PE Immunotech IM1282 CD45 FITC Immunotech IM0782 IgG PE Immunotech IM0670 Tip Stack Pack, yellow tips Sarstedt Redi-Tip General Purpose, μl Fisherbrand A Repeat Pipettor tips, CombitipsPlus 0.5ml Eppendorf mm polyethylene caps BD, Falcon 2032 Polystyrene round bottom tube BD, Falcon 2052 Reagent Preparation Ammonium chloride lysing solution from Stem-Kit: 1. This is a 10X solution. Dilute by adding 1 part lysing solution from Stem-Kit to 9 parts distilled water. 2. Keep at room temperature. Make fresh daily. Page 27 of 48

28 X:\UPCIFLOW\FLOW COURSE 2007\CD34 lab 2007\for Jada\temp.doc Instrumentation Equipment Manufacturer Model Number Flow Cytometer Beckman Coulter Epics XL-MCL Eppendorf Positive Brinkman Displacement Repeat Pipettor Eppendorf adjustable Brinkman 4810 Micropipettors (5-100mcl) Gilson adjustable Gilson Pipetman Micropipettors (5-100mcl) Refrigerator (2-8 o ) Forma Scientific 3881 Vortex Genie Fisher Scientific Blood Rotator Adams Nutator Procedure Cautionary Notes CD-Chex is made from human placental blood and should be considered potentially infectious and therefore handled with universal precautions. It has tested negative for antibodies to HBV, HIV, and HCV. It must be disposed with infectious medical waste. Do not dilute. The gloves to be worn should be powder-free to prevent any contamination either in specimen staining or with the instrumentation during analysis. Detailed Methods CD34/CD3 STAINING OF CD-CHEX/Plus 1. Start XL-MCL in accordance with QC Daily Start-Up and Shut-down Procedures for Coulter XL-MCL. 2. Label 3 tubes with CD-Chex or CD Chex Plus and: Tube 1> 45/34, Tube 2> 45/34, Tube3> Isotube or Tube 1> 45/3, Tube 2> 45/3, Tube3> Isotube, whichever is appropriate. 3. When staining for CD34 + assay, add 20μl of Stem-Kit antibody to appropriate tube, i.e., 45/34 to tubes 1 & 2 and isoclonic control to tube 3. When staining for CD3 + assay, add 10μl each of antibody to appropriate tube, i.e., 45 and 3 to tubes 1 & 2 and 45 and IgG control to tube 3. Page 28 of 48

29 X:\UPCIFLOW\FLOW COURSE 2007\CD34 lab 2007\for Jada\temp.doc 4. Set Eppendorf repeat pipettor to 5, which will deliver 50μl/click. Using the same Eppendorf repeat pipettor (positive displacement) for both the addition of CD Chex and fluorospheres, draw CD-Chex/Plus into 500μl repeat tip. 5. Wipe tip with Kimwipe. 6. Click pipettor once (50μl), returning CD-Chex to vial, for priming. Precision is necessary during this step or erroneous results may occur. 7. Add 2 clicks (100μl) of CD-Chex/Plus to all 3 tubes. 8. Cap all tubes and incubate for 20 minutes in the dark at room temperature. 9. Remove fluorospheres from the Stem-Kit and vortex gently. Leave at room temperature for 30 minutes prior to addition. 10. Dilute Stem-Kit lysing solution using a 1:10 dilution with distilled water (see Reagent Preparation). 6ml of lysing solution is needed 11. After 20 minute incubation, add 2ml of lysing solution to each tube and vortex. 12. Cap tubes and incubate for 10 minutes at room temperature in the dark. 13. After 10 minutes, tubes can be held at this point for 2 hours at in the dark or processed immediately. 14. When ready to acquire, using the same Eppendorf repeat pipettor (positive displacement) as for the addition of sample, draw fluorospheres into 500μl repeat tip. 15. Wipe tip with Kimwipe. 16. Click Pipettor once, returning fluorospheres to vial, for priming. Precision is necessary during this step or erroneous results may occur. 17. Add 100μl of fluorospheres to all 3 tubes using the same pipette that was used to add CD-Chex to tubes. 18. After addition of fluorospheres, tubes must be acquired within 1 hour. Store at 2-8 o C or place tubes in ice tray. 19. Before acquisition on the flow cytometer, label a 4 th tube as blank and add 2ml of distilled water to it. This blank can be used for multiple samples. 20. Confirm that the Stem Count Fluorospheres concentration factor has been correctly entered into the cytometer. This number can be found on the fluorosphere vial as a whole number (ex: 994 fluorospheres/ul.) Refer to R9 QC Daily Start-Up, Shut Down and Maintenance Procedures for Coulter XL-MCL Primary and Back-Up Instruments for instructions to perform this function. 21. CD-Chex sample is now ready for flow cytometric analysis. CD 34 ACQUISITION AND ANALYSIS OF CD-CHEX ON THE XL-MCL Page 29 of 48

30 X:\UPCIFLOW\FLOW COURSE 2007\CD34 lab 2007\for Jada\temp.doc 11. To run CD34 Stem-kit tubes: under Panel choose CD CHEX CONTROL or CD CHEX PLUS CONTROL, whichever is applicable, Hit Okay. 12. On the right of the screen (gray section), the panel you selected will appear with the order in which the tubes should be run. 13. Vortex Tube 1, which is CD45/CD Check Status (lower right gray section) to make sure Insert Sample Tube is flashing green. 15. Open door to sample port and place tube on sample port. Close door (instrument will proceed even if door remains open). Instrument will lift sample port up into sampling chamber. 16. Status will read: Sample tube detected. 17. You will see a series of 8 histograms. CD-Chex cells are expected to have dim CD45+, bright CD34+ and low to moderate side scatter (SS). Gating strategy is setup according to ISHAGE Guidelines for CD34+ Cell Determination by Flow Cytometry. CD-Chex Plus cells are expected to have dim CD45+, bright CD3+ and low to moderate side scatter (SS). Gating strategy is also setup according to ISHAGE Guidelines for CD34+ Cell Determination by Flow Cytometry 18. The 1 st histogram is CD45 vs. SS and is gated on not M to remove beads from view. CD45 stains all human white blood cells but not red cells, platelets and other debris. Click on this 1 st Region to enlarge. Adjust Region A to include only those cells that stain CD45+. Click on the letter, this will allow you to adjust Region. Instrument will automatically update data so you do not need to restart. Region E is tracking Region C and can not be adjusted. This may help to determine how far to the left to adjust Region A. 19. If at any point you want to go back and view a Region, click on 8 Hist button at bottom of screen in blue and then touch the screen to be viewed, this will enlarge it. 20. Click on Next Hist button at bottom of screen in blue. 21. This 2 nd histogram is CD34 or CD3 vs. SS. It is gated on Region A not M so you are looking at only the cells that are CD45+. The population of interest now is CD34+ or CD3+. Adjust Region B to include only CD34+ or CD3+ with low to moderate SS. 22. Click on Next Hist button at bottom of screen in blue. 23. This screen is gated on A, B not M. Region C should include a cluster of dim CD45+ events with low to moderate SS. Cells that tend to have brighter staining with slightly higher SS should be avoided. 24. Click on Next Hist button at bottom of screen in blue. 25. This histogram is gated on Regions A, B and C not M. Adjust Region D if necessary to include all clustered events with low SS and intermediate to high forward scatter. If specimen has low viability you will see these dead cells clustering to the left of the Region having very low forward scatter. The cells located in this Region are considered to be the true CD34 or CD3 cells. The %positive of CD-Chex will vary Page 30 of 48

31 X:\UPCIFLOW\FLOW COURSE 2007\CD34 lab 2007\for Jada\temp.doc from lot to lot, but will average about 0.5%, which should give a visible population. The %positive of CD-Chex Plus will vary from lot to lot, but will average about 70%. 26. Click on Next Hist button at bottom of screen in blue. 27. This histogram is gated on Region not M. It shows CD34+or CD3+/dim CD45 cells located in the upper right quadrant, I Click on Next Hist button at bottom of screen in blue. 29. This histogram is gated on Region not M. It is a threshold check to determine if valid events are being lost due to the discriminator. 30. Click on Next Hist button at bottom of screen in blue. 31. This histogram is ungated. Region G should include the Stem-Count Fluorospheres singlets only. It will be labeled as CAL to allow automatic calculation of absolute numbers of CD34+ or CD3+. If beads are falling out of this region, push Prime button located on right of sample port. Region M includes all beads and cannot be adjusted during acquisition. Events within Region M will be excluded in all other gates because they would be counted as white blood cells and affect the absolute counts. 32. Click on Next Hist button at bottom of screen in blue. 33. This histogram is ungated. It shows FSC v. Stem-count beads. If beads start to fall out of Region H, Prime and Restart. 34. Instrument will stop acquiring when 75,000 events have been collected in Region B or 999 seconds have elapsed. 35. All adjustments need to be made before 75,000 events have been collected. You may need to watch event number. This is located in gray area under STOPS, HIST 2 in yellow. If it is getting close to 75,000 and adjustments still need to be made, HIT Restart at bottom of screen and acquisition will start again. 36. After 1 st tube is collected, Enter Specimen ID box will appear. Enter CD CHEX, the lot number and the expiration date and hit Return. 37. Sample port will lower. Remove tube and wait for flashing green: Insert Sample Tube. 38. Place Tube 2 on sample port. It is also CD45/CD34 or CD45/CD3. Do not change any Regions. The Regions are set on Tube 1 and will remain the same for all remaining tubes for that patient. Just make sure beads are within region or cells have not shifted, if necessary push Prime button and restart acquisition. 39. When acquisition is complete, sample port will lower. Remove tube and wait for flashing green: Insert Sample Tube. 40. The next tube is the blank containing only distilled water. This will prevent any carry-over to the isotube. 41. When acquisition is complete, sample port will lower. Remove tube and wait for flashing green: Insert Sample Tube. Page 31 of 48