Abstract
Background:
Hypoxic conditions preserve the multipotency and self-renewing capacity of murine bone marrow and human cord blood stem cells. Blood samples stored in sealed blood gas tubes become hypoxic as leukocytes metabolize and consume oxygen. Taken together, these observations suggest that peripheral blood stem cell samples stored under airtight conditions become hypoxic and that the stem cells contained may undergo qualitative or quantitative changes.
Objectives:
To determine the effect of storage for 8 hours in a sealed system on peripheral blood stem cell samples.
Study design:
Prospective collection of G-CSF mobilized peripheral blood stem cell samples from nine patients with myeloma or amyloidosis prior to apheresis followed by measurement of CO2, O2, hydrogen ion (pH), lactate, and glucose concentrations in the blood and immunophenotyping of stem cell and multipotent progenitor cell populations before and after 8 hours of storage in a sealed blood collection tube.
Results:
Blood concentrations of O2 and glucose, and pH measurements were significantly decreased while concentrations of CO2 and lactate were significantly increased after storage. A significantly higher concentration of CD34+ (552 ± 84 cells/106 total nucleated cells (TNC) versus 985 ± 143 cells/106 TNC, P = .03), CD34+CD38- (98 ± 32 versus 158 ± 52 cells/106 TNC, P = .03), CD34+CD38+ (444 ± 92 versus 789 ± 153 cells/106 TNC, P = .03), and CD34+CD38-CD45RA-CD90+ (55 ± 17 cells/106 TNC versus 89 ± 25 cells/106 TNC, P = .02) cells were detected after 8 hours of storage. The change in concentration of CD34+CD38+ and CD34+ cells was inversely associated with change in glucose concentration (P = .003 and P < .001) and positively associated with change in lactate concentration (P = .01 and P <.001) after 8 hours of airtight storage.
Conclusions:
Storage in a sealed, airtight environment of peripheral blood stem cell samples is associated with microenvironmental changes consistent with hypoxia and increased concentrations of immunophenotypically defined stem cells. These results may have clinical implications with regard to the collection and processing of stem cell products and warrant confirmation with functional and mechanistic studies.
Keywords: Hematopoiesis, hypoxia, graft collection, graft processing
Introduction
Adequate numbers of peripheral blood stem cells (PBSC) are required for the successful rescue of hematopoiesis after high dose chemotherapy for multiple myeloma and lymphoma (reviewed in [1, 2]). Approximately 2×106 CD34+ cells/kg body weight are commonly used to ensure adequate reconstitution of hematopoiesis with the lower limit being approximately 1×106 CD34+ cells/kg body weight [3]. Many factors affect the number of hematopoietic stem cells (HSCs) collected, including the prior use of chemotherapy and radiation [4].
Peripheral blood stem cell products are collected by apheresis after growth factor mobilization of HSCs from the bone marrow to the peripheral blood. Due to the use of granulocyte colony stimulating factor (G-CSF), a high number of leukocytes often accompanies the HSCs. In a closed system, such as a blood gas tube, leukocytes continue to consume oxygen at a rate that correlates with the total leukocyte count after blood has been collected [5, 6]. Leukocytes may also metabolize glucose resulting in a hypoglycemic environment [7]. Thus, high leukocyte counts can alter the microenvironment of blood stored in airtight tubes through their continued metabolism.
Exposure of murine bone marrow and human cord blood HSCs to normoxic conditions (21% O2) for as little as 30 minutes can trigger HSC differentiation from quiescent pluripotent long term hematopoietic stem cells (LT-HSC) into activated short term repopulating stem cells. [8] Hypoxia has been shown to preserve the multipotency of murine bone marrow and human cord blood HSCs. Harvesting and maintaining murine bone marrow and human cord blood in constant hypoxia (3% O2) increased the number of phenotypically defined LT-HSCs by 5- and 3-fold, respectively [8].
Taking into consideration the effects of high leukocyte quantity on a closed environment and the clinical significance of stem cell numbers, we evaluated the impact of storing GCSF mobilized peripheral blood in a sealed environment on oxygen, carbon dioxide, lactate, and glucose concentrations, pH measurements, and stem cell immunophenotypes. Based upon the effects of hypoxia on mouse bone marrow and human cord blood HSCs, we hypothesized that the concentration of human PBSCs would increase in sealed storage. We tested this hypothesis by immunophenotyping HSC and multipotent progenitor (MPP) cell subsets in human PBSC samples before and after eight hours of storage at room temperature in airtight blood collection tubes. We demonstrated that hypoxic conditions affect the immunophenotypes of cells in human PBSC grafts. This observation could offer new opportunities for intervention to improve the quality of human PBSC grafts.
Materials and Methods
Clinical Materials
PBSC samples were obtained from adults with multiple myeloma or amyloidosis while undergoing PBSC collection as part of a treatment plan to receive high dose chemotherapy followed by autologous stem cell rescue. Patients were enrolled onto a prospective, non-interventional, bio-sample collection protocol which was approved by the Roswell Park Comprehensive Cancer Center Institutional Review Board. Written informed consent was obtained from all patients. Patient characteristics are shown in Table 1. The median (min-max) age of all patients was 64 (53–78) years. The median (min-max) number of CD34+ cells in the PBSC grafts collected from patients was 6.84 (4.52 to 14.64) x 106/μL. The median (min-max) day of neutrophil engraftment was 11 (11–13). The median (min-max) day of platelet engraftment was 16 (11–26).
Table 1.
Patient Characteristics and Clinical Outcomes
| Subject | Age (year) | Sex | Body weight (kg) | Diagnosis | Prior therapy | Sample WBC (*103cells/μL) | Total CD34+ cells collected on the sample day (*106cells/kg) | Blood volume processed by apheresis (mL) | Days of stem cell collection | CD34+ dose received by patient (*106cells/kg) | Day of neutrophil engraftment | Day of platelet engraftment |
|---|---|---|---|---|---|---|---|---|---|---|---|---|
| 1 | 61 | M | 71.1 | Multiple myeloma | RVD | 86.93 | 5.79 | 26067 | 2 | 5.56 | 11 | 11 |
| 2 | 53 | F | 71 | Multiple myeloma | RVD | 80.46 | 6.00 | 27194 | 2 | 4.52 | 11 | 21 |
| 3 | 74 | M | 73.7 | Multiple myeloma | RVD | 46.07 | 1.02 | 75175 | 5 | 14.64 | 11 | 13 |
| 4 | 64 | M | 64.10 | Renal amyloidosis | CVD | 49.12 | 1.72 | 53039 | 4 | 8.25 | 13 | 26 |
| 5 | 63 | F | 65.8 | Multiple myeloma | RVD | 66.48 | 5.80 | 26169 | 2 | 6.84 | 11 | 16 |
| 6 | 56 | M | 80.5 | Multiple myeloma | Dara- RVD |
60.62 | 2.52 | 60820 | 4 | 7.23 | 12 | 20 |
| 7 | 71 | M | 83.1 | Multiple myeloma | CVD | 60.91 | 3.84 | 30278 | 2 | 5.4 | 11 | 17 |
| 8 | 78 | F | 39.8 | Multiple myeloma | RVD | 30.53 | 5.02 | 16000 | 2 | 8.58 | 11 | 15 |
| 9 | 67 | M | 87.6 | Multiple myeloma | RVD | 37.39 | 3.67 | 45462 | 3 | 5 | 11 | 11 |
Abbreviations: CVD – cyclophosphamide, bortezomib, dexamethasone; Dara – daratumumab; RVD – lenalidomide, bortezomib, dexamethasone
Patients undergoing stem cell mobilization received G-CSF (10 μg/kg) each morning starting on day+1 and continuing until the last day of stem cell collection by apheresis. Plerixafor (0.24 mg/kg) started on day+4 in the evening and continued daily until the end of apheresis for a maximum of 4 doses. Daily apheresis for PBSCs started the morning of day+5 and continued for a maximum of 5 days until the target dose of 8×106 CD34+ cells/kg recipient body weight was collected. CD34+ cells were enumerated at the end of each collection day. At least two days of apheresis were performed regardless of CD34+ cell counts.
All study samples were collected on day+6 of stem cell mobilization (the second day of PBSC apheresis) except one (subject 4) that was collected on day+5 (the first day of PBSC apheresis). The blood sample processing schema is shown in Figure 1. From each participant, three blood samples were collected serially and directly from the efferent lumen of the apheresis catheter flowing from the patient to the apheresis machine (Optia [Terumo] or Amicus [Fresenius Kabi]): 3 mL for blood gas measurement, 5 mL for immediate fixation into a Cyto-Chex Blood Collection Tube (BCT, Streck, La Vista, Nebraska), and 5 mL for storage into a lavender top airtight EDTA tube. Cyto-Chex fixation stabilizes white blood cell immunogenic markers and maintains cellular morphology and surface antigen expression, including cluster of differentiation markers prior to analysis by flow cytometry. Whole blood samples collected in Cyto-Chex BCTs are stable for up to 7 days at room temperature [9]. CD3, CD4, CD8, and CD45 are stable after storage in Cyto-Chex tubes for up to 7 days [9] and the stability of CD45RA expression is maintained for 3 and 10 days after fixation [10, 11]. CD34, CD38, and CD90 epitope expression before and after Cyto-Chex fixation was tested and found to vary less than 5% between each timepoint (Supplemental Material). Samples in EDTA lavender and Cyto-Chex tubes were stored at room temperature for eight hours. At the end of eight hours, a blood gas was obtained from the lavender tube and the remaining sample was fixed in a Cyto-Chex tube. Both samples were then immunophenotyped and analyzed by multicolor flow cytometry.
Figure 1. Schema for PBSC blood samples processing.
PBSCs were harvested directly into a blood gas, Cyto-Chex, or airtight EDTA tube from the efferent lumen coming from the patient and flowing to the apheresis machine. The airtight sample was incubated at room temperature for 8 hours. After 8 hours the EDTA sample was transferred to a blood gas tube or fixed in a Cyto-Chex BCT. Both samples were immunophenotyped as described in the methods within 2 hours of each other. Key: EDTA - ethylenediaminetetraacetic acid, PBSC - peripheral blood stem cells.
Sample Processing
To lyse red blood cells (RBCs), 10 mLs of 1X ammonium chloride buffer (A10492–01, Gibco) were added for each 1 mL of harvested peripheral blood. Samples were incubated for 10–15 minutes at room temperature and then centrifuged at room temperature. The supernatant was decanted and cells were washed once with phosphate buffer saline (PBS) containing 0.5% bovine serum albumin, 0.1% sodium azide, and 0.0004% tetrasodium EDTA. Samples were finally centrifuged again at room temperature, the supernatant decanted, and the pellet was re-suspended in PBS before staining.
Antibody staining
The cell concentration in each sample was enumerated using an automated cell counter (Beckman Coulter AcT diff hematology analyzer). Twenty million mononuclear cells (MNCs) were stained for 20-minutes with Fixable Live Dead Aqua 405 nm for dead cell exclusion and a monoclonal antibody (mAb) cocktail containing saturating amounts of the following mAbs: CD45RA PE, CD90 BV421, CD45 APC-H7, CD38 PerCP-Cy5.5, CD127 PE-Cy7, CD34 APC (Table 2). After incubation with mAbs, the cells were washed once with PBS. Samples were then centrifuged at room temperature and the pellet was re-suspended in PBS.
Table 2.
Antibody Reagents
| Epitope | Fluorochrome | Manufacturer | Catalog | Clone |
|---|---|---|---|---|
| CD45RA | PE | BD Biosciences | 555489 | HI100 |
| CD90 | BV421 | BioLegend | 328122 | 5E10 |
| CD45 | APC-H7 | BD Biosciences | 641408 | 2D1 |
| CD38 | PerCP-Cy5.5 | BD Biosciences | 656050 | HIT2 |
| CD127 | PE-Cy7 | BD Biosciences | 560822 | HIL-7R-M21 |
| CD34 | APC | Miltenyi | 130-113-176 | AC136 |
Flow Cytometry Analysis
Flow cytometric acquisition was performed on an LSRFortessa (BD Biosciences) equipped with three laser excitation sources (405 nm 50 mw; 488 nm 50 mw; 640 nm 40 mw) that was quality-controlled on a daily basis using CS&T beads and FACS DiVA software (BD Biosciences). The filter configurations for the PMTs measuring fluorescence emission of the applied fluorochromes were 575/26 nm (PE), 450/50 nm (BV421), 780/60 nm (APC H7), 695/40 nm (PerCP-Cy5.5), 780/60 nm (PE Cy7) and 670/14 nm (APC). Autofluorescence and single-color controls were acquired to perform spectral overlap compensation using the automated compensation matrix feature in FACS DiVA software. Fluorescence minus one controls were used to set the threshold to identify positively-stained cells. Flow cytometric data was plotted using bi-exponential plots that include axes <0 to assure all data was visible and properly compensated. Data analysis was performed with Winlist 3D, version 9.0.1 (Verity Software House). HSC were phenotypically defined as CD34+CD38-CD45RA-CD90+ and MPP were defined as CD34+CD38-CD45RA-CD90- (Figure 2) [12]. The frequency of phenotypically defined stem cell populations in each sample were determined using a sequential gating strategy (Figure 3).
Figure 2. Schema for immunophenotyping of stem and progenitor cells.
The hematopoietic stem cell is self-renewing and differentiates into the multipotent progenitor cell. Further differentiation of the multipotent progenitor cell gives rise to the megakaryocyte erythroid, common myeloid, and common lymphoid progenitor cell. In this immunophenotyping schema, the hematopoietic stem cell can be differentiated from the multipotent progenitor cell with the CD90 antigen. The CD34+ cell fraction contains the less differentiated CD34+CD38- fraction (hematopoietic stem cells and multipotent progenitor cells) and the more differentiated CD34+CD38+ fraction (megakaryocyte erythroid, common myeloid, and common lymphoid progenitor cells). Bolded and italicized immunophenotypes were assayed in this study. Key: Lin - lineage marker negative.
Figure 3. The frequency of phenotypically defined stem cell populations in each sample were determined using a sequential gating strategy.
A bivariate plot of FSC-A versus SSC-A (Plot 1) was employed to resolve debris from leukocytes; which were circumscribed by rectangular region (R1). Leukocyte events within (R1) were then gated to a bivariate plot of Live/Dead Aqua-A versus SSC-A (Plot 2), where live events (negative for the viability reagent) were circumscribed by polygonal region (R2). Events meeting the Boolean definition of (R1&R2) were gated to a bivariate plot of CD45 APCH7-A versus SSC-A (Plot 3); where cellular events were identified based on their positivity for CD45. These CD45+ events were circumscribed by a rectangular region (R3). To subsequently evaluate the light scatter characteristics of gated lymphocytes, an elliptical region (R4) was employed on Plot 3 to circumscribe lymphocytes; which exhibited bright CD45 labeling, with low side scatter. Viable cellular events (R1&R2&R3) were subsequently gated to a plot of CD34 APC-A versus SSC-A (Plot 4), where CD34+ stem cells were circumscribed by rectangular region (R5). Live, CD34+ cells (R1&R2&R3&R5) were gated to a bivariate plot of CD45 APCH7-A versus SSC-A (Plot 5), where rectangular region (R6) was employed to exclude aggregate and platelet events. This refined, live, CD34+ cellular population (R1&R2&R3&R5&R6) was gated to a bivariate plot of FSC-A versus SSC-A (Plot 6) to further elucidate the stem cell population based on its light scatter characteristics.
Statistical Considerations
Patient characteristics were described with median and range (min-max). The paired T-test was used to compare PBSC blood sample characteristics before and after eight hours of storage. The Spearman rank correlation test was used to determine the correlation between pH, CO2, O2, glucose, and lactate concentrations and the number of CD34+CD38-CD45RA-CD90+ (HSC), CD34+CD38-CD45RA-CD90- (MPP), CD34+CD38+, CD34+CD38-, and CD34+ cells. P values <.05 were considered statistically significant. Data is shown as mean ± standard error of the means or median (min-max). Data visualization and statistical analysis was performed in R v. 4.1.3 [13].
Separately, a bivariate plot of FSC-A versus SSC-A (Plot 7) was gated (R1&R2&R3&R4) to display the light scatter characteristics of lymphocytes; which were circumscribed by rectangular region (R8). This was accomplished to serve as a reference for defining the light scatter characteristics of live stem cells (R1&R2&R3&R5&R6); and in this manner, rectangular region (R7) was employed to define this population on Plot 6. The Cartesian boundaries of (R7) were established by ‘mirroring’ region (R7) to region (R8).
Live stem cells (R1&R2&R3&R5&R6&R7) were subsequently evaluated for their expression of CD38, employing a bivariate plot of CD34 APC-A versus CD38 PerCPCy5.5-A (Plot 8); where rectangular region (R9) defined CD38+ stem cells, and rectangular region (R10) defined CD38- stem cells. CD38- stem cells (R1&R2&R3&R5&R6&R7&R10) were then gated to a bivariate plot of CD45RA PE-A versus CD90 BV421-A (Plot 9) to characterize their expression of CD90; where CD90+ events were circumscribed by rectangular region (R11) and CD90- events were circumscribed by rectangular region (R12).
Results
PBSC blood samples stored in airtight tubes for eight hours become hypoxic. The median (min-max) WBC in the blood samples collected the day of apheresis was 60.62 (30.53 – 86.93) x 103/μL. (Table 1) Blood gas measurements before and after storage of PBSC blood samples for eight hours inside airtight tubes demonstrated significant microenvironmental changes (Table 3). The partial pressure of oxygen (pO2) was significantly lower (Figure 4A). Other blood gas parameters suggested a shift from aerobic to anerobic metabolism characterized by glycolysis and consistent with a hypoxic environment. pH was significantly lower (Figure 4B) as was glucose (Figure 4C). The partial pressure of carbon dioxide (pCO2) was significantly higher (Figure 4D) as was the concentration of lactate (Figure 4E).
Table 3.
Sealed Storage Affects the Peripheral Blood Stem Cell Sample Microenvironment (N=8)
| Parameter | 0 hours Mean ± SE | 8 hours Mean ± SE | P value |
|---|---|---|---|
| Partial pressure of oxygen | 42 ± 3.9 mm Hg | 22 ± 3.2 mm Hg | .003 |
| pH | 7.26 ± 0.06 | 7.07 ± 0.05 | .003 |
| Glucose | 112 ± 16 mmol/L | 49 ± 17 mmol/L | <.001 |
| Partial pressure of carbon dioxide | 49.9 ± 2.1 mm Hg | 57 ± 2.8 mm Hg | .04 |
| Lactate | 17 ± 1.4 mmol/L | 67 ± 9.0 mmol/L | <.001 |
Key: SE – standard error of the means
Figure 4. Storage of PBSC samples for 8 hours results in a hypoxic environment.
The hypoxic environment may have resulted in a switch from aerobic to anaerobic metabolism as evidenced by the decrease in glucose concentration increase in lactate concentration. The donor of the stem cells from which samples were derived is indicated by the number to the side of the circle. The numbers correlate with the subject numbers in Table 1. Donor 8 is not shown due to missing data. A. pO2. Impact of PBSC storage on pO2 at 0 and after 8 hours (42 ± 3.9 mm Hg versus 22 ± 3.2 mm Hg, P = 0.003). B. pH. Impact of PBSC storage on pH at 0 and after 8 hours (7.26 ± 0.06 versus 7.07 ± 0.05, P=0.003). C. Glucose. Impact of PBSC storage on glucose at 0 and after 8 hours (112 ± 16 mmol/L versus 49 ± 17 mmol/L, P < 0.001). D. pCO2. Impact of PBSC storage on pCO2 at 0 and after 8 hours (49.9 ± 2.1 mm Hg versus 57 ± 2.8 mm Hg, P = 0.04). E. Lactate. Impact of PBSC storage on lactate at 0 and after 8 hours (17 ± 1.4 mmol/L versus 67 ± 9.0 mmol/L, P < 0.001). Key: PBSC- peripheral blood stem cells
Storage in a sealed environment is associated with increased quantities of immunophenotypically defined stem cell populations.
When compared to a freshly collected sample, PBSC blood samples stored in airtight blood collection tubes at room temperature for eight hours contained a significantly higher concentration of CD34+ cells, CD34+CD38- cells, CD34+CD38+ cells, and CD34+CD38-CD45RA-CD90+ defined hematopoietic stem cells (Table 4, Figures 5A-D). The concentration of CD34+CD38-CD45RA-CD90- defined multipotential progenitor cells before and after eight hours of storage was not statistically different (Table 4, Figure 5E). The ratio of CD34+CD38+ to CD34+CD38- cells before and after eight hours of storage was not significantly different. The ratio of CD34+CD38-CD45RA-CD90+ to CD34+CD38CD45RA-CD90- cells before and after eight hours of storage was not significantly different. The viability of MNCs from the PBSC blood sample (N=9) was unaffected by storage (median 99.07% (min-max 95.51–99.65%)) at zero hours versus 99.26% (min-max 95.31 – 99.93%) at eight hours).
Table 4.
Sealed Storage Affects the Quantity of Immunophenotypically Defined Peripheral Blood Stem Cell Populations (N=9)
| Immunophenotypic population | 0 hours (cells / 106 TNC) Mean ± SE | 8 hours (cells / 106 TNC) Mean ± SE | P value |
|---|---|---|---|
| CD34+CD38-CD45RA-CD90+ Hematopoietic stem cells | 55 ± 17 | 89 ± 25 | .02 |
| CD34+CD38-CD45RA-CD90- Multipotent progenitor cells | 16 ± 6 | 28 ± 11 | NS |
| CD34+CD38- cells | 98 ± 32 | 158 ± 52 | .04 |
| CD34+CD38+ cells | 444 ± 92 | 789 ± 153 | .02 |
| CD34+ cells | 552 ± 84 | 985 ± 143 | .01 |
Key: SE – standard error of the means
Figure 5. Impact of eight hours storage in a sealed environment on peripheral blood stem cell immunophenotypes.
The donor of the stem cells from which samples were derived is indicated by the number to the side of the circle. The numbers correlate with the subject numbers in Table 1. A. CD34+ cells. CD34+ cell concentration is increased after storage (552 ± 84 versus 985 ± 143 cells / 106 total nucleated cells (TNC), P=0.01). B. CD34+CD38- cells. CD34+CD38- cell concentration is increased after storage (98 ± 32 versus 158 ± 52 cells /106 TNC, P=0.04). C. CD34+CD38+ cells. CD34+CD38+ cell concentration is increased after storage (444 ± 92 versus 789 ± 153 cells / 106 TNC, P=0.02). D. CD34+CD38-CD45RA-CD90+ cells. CD34+CD38-CD45RA-CD90+ defined hematopoietic stem cell concentration is increased after storage (55 ± 17 versus 89 ± 25 cells / 106 TNC, P = 0.02). E. CD34+CD38-CD45RA-CD90+ cells. CD34+CD38-CD45RA-CD90+ defined multipotential progenitor cell concentration is increased but not significantly changed after storage (16 ± 6 versus 28 ± 11 cells / 106 TNC, P = not significant).
After airtight storage of PBSC samples for 8 hours, the change in glucose concentration was significantly correlated with the change in CD34+CD38-CD45RA-CD90+ (MPP) cells (p=.02), CD34+CD38+ cells (p=.02), and CD34+ cells (p<.001), and the change in lactate concentration was correlated with the change in CD34+CD38+ cells (p=.01) and CD34 cells (p<.001) (Figure 6). No other significant associations between other microenvironmental parameters (pH, pCO2, pO2) and cell subtypes (CD34+CD38-CD45RACD90+ (HSC), and CD34+CD38-) were found.
Figure 6. Association between microenvironmental parameters and cell concentrations.
The change in microenvironmental parameters is plotted against the change in cell concentrations after 8 hours of airtight incubation in blood collection tubes. Statistical testing of correlation performed with Spearman rank test. The donor of the stem cells from which samples were derived is indicated by the number to the side of the circle. The numbers correlate with the subject numbers in Table 1. A. Glucose vs CD34+CD38+ cell concentration. Change in CD34+CD38+ concentration negatively correlates with change in glucose concentration (rho=−0.89, p=0.003). B. Glucose vs CD34+ cell concentration. Change in CD34+ cell concentration negatively correlates with change in glucose concentration is associated with change in CD34+CD38+ cell concentration (rho=−0.99, P<.001). C. Lactate vs CD34+CD38+ cell concentration. Change in CD34+CD38+ cell concentration positively correlates with change in lactate concentration (rho=0.86, P=0.01). D. Lactate vs CD34+ cell concentration. Change in CD34+ cell concentration positively correlates with change in lactate concentration (rho=0.98, p<.001). Key: [CD34+] – concentration of CD34+ cells, [CD34+CD38+] – concentration of CD34+CD38 cells, [CD34+CD38-CD45RA-CD90-] – concentration of CD34+CD38-CD45RA-CD90- cells, [Glucose] – concentration of glucose, [Lactate] – concentration of lactate.
Discussion
In this study, we demonstrate that sealed storage of mobilized peripheral blood stem cell samples results in a hypoxic, hypoglycemic, and acidic microenvironment that is associated with increased concentrations of all immunophenotypically defined CD34+ cells including CD34+CD38-CD45RA-CD90+ hematopoietic stem cells. The altered microenvironment may be a metabolic byproduct of the high number of leukocytes in a sealed airtight environment. The changes in the concentration of immunophenotypically defined stem cells may have been due to hypoxia as suggested by the decreased oxygen level and work by others demonstrating the effect of oxygen concentration on stem cells. Other parameters such as hypoglycemia and increased acidity may have also contributed to the changes in cell quantities.
Prospectively collected human blood specimens containing PBSCs were analyzed in this study. Samples were drawn from the efferent apheresis catheter lumen leaving the patient and flowing to the apheresis machine, not from the afferent lumen leaving the apheresis machine and flowing to the patient. Thus, the cell sample collected is a reasonable approximation of the state of cells in the patient without the influence of the apheresis machine. To preserve the immunophenotype prior to analysis by flow cytometry, PBSC apheresis samples were harvested directly into Cyto-Chex tubes capable of maintaining cellular morphology and surface antigen expression, including cluster of differentiation markers [9–11].
The overall viability of samples before and after 8 hours of storage was not significantly different. However, selective cell death could still explain the changed proportions of specific immunophenotypes. Fixation with Cyto-Chex may have also affected the viability although granulocyte viability has been reported to be >90% after 2 days of fixation. [14, 15]
The demonstration that storage of peripheral blood stem cell samples under hypoxic conditions is associated with increased concentrations of all immunophenotypically defined CD34+ cells has clinically significant implications. First, apheresis products are currently exposed to normoxic environments during collection and storage. In contrast, diagnostic samples are collected into and transported in airtight tubes. Our work suggests there may be a discrepancy between the stem cell populations quantified in diagnostic samples and the populations infused into patients.
Second, adequate numbers of healthy human HSC are necessary for a successful bone marrow transplant (reviewed in [1, 2]). Heavily pretreated multiple myeloma (MM) and lymphoma patients undergoing peripheral blood stem cell mobilization are at risk for collecting an inadequate cell dose for transplant [16, 17]. In patients with MM, prior lenalidomide therapy is associated with failure of stem cell mobilization with filgrastim in 25% of patients [18]. Enhancing the number of CD34+, CD34+CD38-, CD34+CD38+ and CD34+CD38-CD45RA-CD90+ stem cells by storage under hypoxic conditions may lower the risk of graft failure in heavily pretreated MM and lymphoma patients by providing a better-quality stem cell product.
A recent murine study demonstrated that collection of bone marrow cells under hypoxic conditions led to a partial reversal of the dysfunction observed with aging (decreased long term repopulation potential and a shift to increased myeloid versus lymphoid differentiation) such that bone marrow cells from aged (20–28 months old) mice collected under hypoxic conditions performed as well as bone marrow cells from young (8–12 weeks old) mice collected under normoxic conditions [19]. Hematopoietic cell dysfunction with aging also occurs in humans, as suggested by experimental studies [20, 21] and a retrospective clinical study which demonstrated that peripheral blood stem cells from older allogeneic stem cell donors are qualitatively different from those from younger donors as indicated by an increase in overall survival by 3% for every decade of difference in prospective donor age [22]. In this context, our data showing that hypoxia can affect PBSC graft immunophenotypes may have additional impact if later investigations can demonstrate that hypoxic conditions can reverse the dysfunction observed in aging human hematopoietic stem cells, leading to interventional strategies to improve the quality of PBSC grafts and overcoming the decrease in overall survival associated with stem cells obtained from older donors.
Third, this study utilized stem cells collected from the peripheral blood. Compared to the bone marrow microenvironment, the peripheral blood is less hypoxic [23], suggesting that the effects of increased O2 concentration on stem cells might have already occurred by the time they were collected in our study. The increase in CD34+CD38-CD45RA-CD90+ defined stem cells after storage in hypoxia suggests that the effect of increased O2 concentration may be reversible. Changes in glucose concentration and acidity of the microenvironment may have also resulted in an increase in CD34+CD38-CD45RA-CD90+ defined stem cells.
Finally, the maintenance of hypoxia has been very difficult to achieve in the past requiring the construction of large airtight work chambers. Various methods have been developed to generate and maintain low oxygen environments during cell culture experiments. Hypoxia modular chambers, hypoxia incubators, and hypoxic sub-chamber systems are widely used to perform hypoxic experiments. However, these items of equipment may be expensive and difficult to use. In this study, we present a simple low-cost and portable method for exposing samples to hypoxic conditions that could be utilized in exploratory studies prior to experimentation using more comprehensive methods of inducing hypoxia.
Many caveats apply regarding the translational significance of our findings. First, we quantified the number of hematopoietic stem cells using immunophenotypic rather than functional criteria. Thus, the clinical implications of our work will not be fully understood until additional functional characterization of human peripheral blood stem cells exposed to hypoxic conditions is performed. Second, the present study used purple top (EDTA) blood collection tubes to store peripheral blood containing mobilized stem cells as an abstraction of the storage and processing conditions peripheral blood stem products experience during the collection process. Peripheral blood undergoing apheresis is mechanically separated into its constituents and anticoagulants such as anti-coagulant citrate dextrose, solution A (ACD-A) and heparin are often added which may also affect the metabolism of stem cells. Apheresis does not occur under airtight conditions. Studies replicating the conditions present during apheresis will be necessary to determine the true impact of hypoxia on stem cell collection. Third, collection in Cyto-Chex tubes was used to preserve the immunophenotype of the peripheral blood stem cell samples. Pre-hypoxia samples were preserved for 8 hours while post-hypoxia samples were preserved for less than 2 hours. Finally, this study is limited by the small number of subjects. Despite these caveats, the present data demonstrate how microenvironmental conditions in blood collection tubes may change as a function of storage time and how it may affect the numbers and phenotypes of human hematopoietic stem cells.
Conclusions
Storage of stem cells under increasingly hypoxic conditions may be associated with changes in stem cell immunophenotype. Additional stem cell clonality and functional studies to further characterize this immunophenotypic phenomenon are warranted.
Supplementary Material
Highlights.
Storage of peripheral blood stem cell samples in airtight blood collection tubes results in a hypoxic microenvironment.
The concentration of CD34+, CD34+CD38-, CD34+CD38+, and CD34+CD38-CD45RA-CD90+ immunophenotypically defined stem cells increased after 8 hours of storage in airtight blood collection tubes.
Acknowledgements
This study was supported by the Frawley Research Fellowship from the University at Buffalo (AE), Roswell Park Comprehensive Cancer Center (GLC), and National Cancer Institute grants P30CA016056 involving the use of Roswell Park Comprehensive Cancer Center’s Flow and Image Cytometry Shared Resource and R50CA211108 (H.M.)
We gratefully acknowledge the patients described in the study and the clinical team providing care to the patients; without their support, this study could not have been completed.
Footnotes
Conflict of Interest Disclosures
None of the authors have conflicts of interest.
Financial disclosures statement: The authors do not have any relevant financial disclosures.
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