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Volume: 20 Issue: 10 October 2022


Effect of CD34+ Total/Myeloid CD34+ Cell Progenitors and B-Lymphoid Progenitors Within the Bone Marrow Grafts on the Hematopoietic Recovery After Hematopoietic Stem Cell Transplantation

Objectives: The influence of CD34+ cells on transplant outcomes following hematopoietic stem cell transplantation remains controversial. A minimum of 2.0 to 2.5 million CD34+ cells/kg of patient weight is requested for a rapid and durable engraftment. The aim of this study was to detect the ratio of CD34+ B-lymphoid progenitors (hematogones) in bone marrow grafts and investigate their effects on hematopoietic recovery after transplant.
Materials and Methods: Our study included 41 patients who received a bone marrow graft from their HLA-matched donor from 2016 through 2019. The CD34+ cell numbers within the graft were detected using Stem-Kit (Beckman Coulter). The ratio of CD34+ hematogones was determined either by their light scatter characteristics or by the detection of CD34, CD19, or CD10 coexpressing cells in a separate tube.
Results: The median number of CD34+ cells was 5.9 × 106/kg (0.8-14.3 × 106/kg). The CD34+ cells consisted of 71% (range, 35.7%-100%) and 29% (range, 5.7%-64.3%) myeloid and B-lymphoid progenitors, respectively. Percentage of CD34+ (P < .001) and total (P < .001) hematogones in correlation with donor age. Time of neutrophil engraftment was significantly longer (P = .039) when total infused CD34+ cell content wa <3 × 106/kg.
Conclusions: A remarkable population of hematogones within the CD34+ cell pool was detected in bone marrow grafts. Detection of the ratio of hematogones and most primitive stem cells (CD34+CD90+CD38-) may overall provide more information to build a better correlation between CD34+ cell content and the recovery of bone marrow.


Hematopoietic stem cell transplantation (HSCT) is a treatment modality used to cure many hematologic and nonhematologic diseases. The only source of hematopoietic stem cells (HSCs) was known to be the bone marrow (BM). Although peripheral blood stem cells have replaced BM HSCs after the 1990s, BM is still being used, especially in pediatric age group donors.1 Regardless of their origin, HSCs are capable of starting BM regeneration after high-dose therapy used for hematologic diseases.1,2 Bone marrow is traditionally procured without the requirement of an initial growth factor application, creating less risk for graft-versus-host disease but has the disadvantage of a slower engraftment.2

Hematopoietic stem and progenitor cells (HSPCs) can be detected by their CD34 antigen expression. The faster and reliable method to detect the engraftment potential of the graft is the enumeration of CD34-expressing cells by flow cytometry.3-5 A minimum of 2.0 to 2.5 million viable CD34+ cells/patient weight (kg) is accepted to be essential for a rapid and durable engraftment.6 CD34 expression has been linked to adhesion properties and proliferation and differen-tiation capacities of HSPCs.7

The BM CD34+ cells are extremely heterogeneous, where most are progenitors that are committed to differentiating to distinct cell lineages.8 Early B-cell progenitors, namely, stage 1 hematogones, are characterized by the expression of CD38, CD19, and CD10 molecules on their surface and represent a significant part of BM CD34+ cells.9 Stage 1 hematogones are very small lymphocytes and create a lower forward scatter signal differing them from other HSPCs (Figure 1). Based on this light scatter pattern, CD34+ hematogones can be detected during quantitation of CD34+ cells with the use of the International Society of Hematotherapy and Graft Engineering (ISHAGE) protocol without the requirement of an additional reagent.10 The most immature B-cell precursors (stage 1 hematogones) express CD34 in combination with CD38, CD19, and CD10. Stage 2 hematogones differ from stage 1 hematogones by the expression levels of CD10 and CD19 and a progressive upregulation of CD20 accompanied by a downregulation of CD34. Stage 3 hematogones are characterized by CD20 expression and a lower CD10 and CD38 expression. The percentage of each stage may vary. Most hematogones (65%) in BM samples are stage 2 hematogones. Stage 1 hematogones represent a minority cell population in the BM of most healthy individuals.9 The ratio of stage 1 hematogones is higher in children and infants younger than 4 years old and may increase after high-dose chemotherapy.9,11-13 Hematogones are localized in BM and blood samples, and they can also be detected in extramedullary sites in fetuses and in children.10,14-16 Because healthy individuals rarely undergo BM biopsies, establishing the normal range of hematogones in healthy BM is still very challenging.17 Lúcio and colleagues analyzed hematogones in the BM of volunteers and reported a percentage of 7.5% within nucleated BM cells of healthy children.17 Although this ratio was reported as 1.99% in 1 study, no reference ranges for adults have yet been identified.18

The aim of our study was to investigate the total and CD34+ stage 1 hematogone content of BM grafts and to evaluate their effect on neutrophil and platelet engraftment kinetics after HSCT.

Materials and Methods

Our study included engraftment data of 36 patients who received a BM graft from their HLA-matched donor at the Ankara University Medical School Pediatric and Adult Stem Cell Transplantation Units between 2016 and 2019 and 5 patients who received an HLA-matched HSCT transplantation at the Memorial Hospital (Ankara, Turkey). Three additional BM grafts were collected but not infused for medical reasons and cryopreserved upon collection. Ten of the patients were transplanted in the adult HSCT unit, whereas the remaining 31 patients received transplants in the pediatric HSCT unit. Most of the patients were diagnosed with thalassemia major, acute leukemia, or primary immunodeficiencies (severe combined immuno-deficiency and immunodysregulation syndromes). The demographic characteristics of the patients and their donors are summarized on Table 1. Before procurement, granulocyte colony-stimulating factor (GCSF) was given to 3 donors in the pediatric patient group (2 haploidentical matched donors and 1 matched related donor). For the adult patients, all of the products were unmanipulated. Erythrocyte separation was required in 2 patients, and plasma separation was required in 1 patient.

Flow cytometric detection of total CD34+ hematopoietic stem and progenitor cells
After BM aspirates were retrieved, CD34+ cell content was immediately measured using Stem-Kit (Beckman Coulter). The samples were diluted to obtain a maximum of 30 000 white blood cells (WBCs) per microliter of sample before the assay procedure was started. After we performed a daily control measurement of the instrument, a minimum of 75 000 CD45+ events were acquired in the “Navios” flow cytometer instrument (3L10C, Beckman Coulter). Collected data were analyzed based on the ISHAGE “single platform” protocol (Navios software, Beckman Coulter)19 (Figure 2). We calculated CD34+ cell numbers with the following formula: CD34+ cell absolute count (in cells/μL) = (CD34+ event count in the run × bead count per μL) × dilution factor/bead event count for the run, where the bead count for each lot was written on the vial label present within Stem-Kit.

As an internal control, the absolute counts of CD45+ events were also calculated and compared with the WBC counts obtained from the whole blood count.

Flow cytometric detection of CD34+ hematogones
The ratio of the hematogones was determined by 2 approaches: (1) selection of cells having a lower forward scatter within the CD34+ cell population (Figure 1) and (2) selection of the CD19+ cells within the WBC population and detection of CD19+, CD10+, CD45dim+, and CD34+ stage 1 hematogones within the population (Figure 3).

Statistical analyses
We used Microsoft Excel 2016 and GraphPad Prism 6 (GraphPad Software) for data analyses. Based on Shapiro-Wilks normality test for equal variances, normal distribution was not found in any of the tested or log-transformed data in GraphPad Prism 6. Therefore, we used nonparametric 2-tailed tests for correlation analyses and comparisons between 2 (Mann-Whitney U test) or 3 (Kruskal-Wallis test) groups. We used the Spearman r test for correlation analyses. We classified patient data into 2 groups: age ≤10 years and age >10 years (using Microsoft Excel 2016). These data were then log-transformed, and we used Mann-Whitney U test in GraphPad Prism 6 for comparisons between the 2 age groups. For comparison of neutrophil engraftment (NE) and platelet engraftment (PE) time in total CD34+ cells, we classified NE and PE time into 3 groups: (1) total CD34+ cells <3 × 106/kg (group 1), (2) total CD34+ cells ≥3 and <6 × 106/kg (group 2), and (3) total CD34+ cells ≥6 × 106/kg (group 3). For comparison of NE and PE time in myeloid CD34+ cells, we classified NE and PE times into 3 groups: (1) myeloid CD34+ cells <2 × 106/kg (group 1a), (2) myeloid CD34+ cells ≥2 and <4 × 106/kg (group 2a), and (3) ≥4 × 106/kg (group 3a). In both sets of groups, we used Microsoft Excel 2016 to classify the groups. We log-transformed the NE and PE time results for the groups and compared the groups using Kruskal-Wallis test in GraphPad Prism 6. P < .05 was considered statistically significant.

Ethical approval
Our study was performed in accordance with the Declaration of Helsinki and was approved by the internal review board of the Ankara University Faculty of Medicine (number I8-553-20). Clinical information, laboratory results, and engraftment data were retrieved retrospectively from patient medical records. All data remained anonymous with no identifying personal information.


Cellular graft composition
The BM grafts contained 5.9 × 106/kg total CD34+ cells (range, 0.8-14.3 × 106/kg) and 4.35 × 106/kg myeloid CD34+ cells (range, 0.71-11.14 × 106/kg) (Table 2). Among 41 patients, 10 received <2 × 106/kg myeloid CD34+ cells. The median CD34+ cell content of all grafts consisted of 70.6% myeloid CD34+ cells (range, 35.7%-100%) and 29.4% stage 1 hematogones (range, 5.7%-64.3%). The percentage of total hematogones within WBCs (P < .001) and the percentage of stage 1 hematogones within CD34+ cells (P < .004) were shown to be 2.3-fold and 1.5-fold higher in donors aged ≤10 years, respectively (Table 2). The median number of infused total and myeloid CD34+ cells were significantly different within the ≤10-year and >10-year donor age groups. Grafts from donors ≤10 years old yielded significantly higher total (P < .001) and myeloid (P = .006) fractions (Table 2). In our study group, grafts obtained from donors who were ≤10 years old contained a significantly higher total (P < .001) and myeloid (P = .006) CD34+ cell content. Amounts of infused total CD34+ (P < .001) and myeloid CD34+ (P = .001) cells were 1.6-fold and 1.5-fold higher, respectively, in the younger donor age group.The percentages of total CD34+ cells and total hematogones within WBCs and CD34+ stage 1 hematogones within the CD34+ cell fraction were also higher in the younger age group, with a median ± SD of 1.98 ± 1 versus 1.3 ± 0.6 (P = .008), 4.05 ± 2.2 versus 1.8 ± 1.6 (P < .001), and 36.3 ± 8.3 versus 23.6 ± 13.4 (P = .004), respectively. The graft composition was similar for male and female patients (data not shown).

Neutrophil engraftment time
For 39 of 41 patients, NE was achieved after 17 days (range, 11-29 days). Four patients with severe combined immunodeficiency did not receive a conditioning regimen. We also did not detect NE in 1 patient with aplastic anemia who received 1.87 × 106/kg myeloid CD34+ cells and 1 patient with myelodysplastic syndrome who received 0.71 × 106/kg myeloid CD34+ cells (data not shown).

The adult patients with acute lymphoblastic leukemia and aplastic anemia received GCSF until engraftment. Three patients in the pediatric group received GCSF before engraftment. When we grouped patients according to CD34 content of the graft (Table 3), we observed that the median of detected NE time for total CD34+ cell numbers were 20.5, 16, and 16 days for groups 1, 2, and 3, respectively. We found NE time to be significantly longer in group 1, in which patients received <3 × 106/kg of total CD34+ cells (P = .039). The median of detected NE time for the myeloid CD34+ cells was 19, 16 and 16 days for groups 1a, 2a, and 3a, respectively. Differences between these groups were not significant (P = .375).

The median of detected NE time was 16 days (range, 11-29 days) in patients who underwent transplant from a matched related donor and 19 days (range, 11-28 days) in patients who underwent transplant from a matched unrelated donor (data not shown).

Platelet engraftment time
For 35 of 41 patients, a stable PE (blood platelet count >20 × 109/L for at least 7 days, no platelet transfusion) was achieved after 22 days (range, 11-55 days). Four patients with severe combined immunodeficiency did not receive a conditioning regimen, and their platelet count did not decrease. We did not detect PE in 2 patients: 1 patient with aplastic anemia who received 1.87 × 106/kg myeloid CD34+ cells and 1 patient with myelodysplastic syndrome who received 0.71 × 106/kg myeloid CD34+ cells. The detected PE time was 22, 23.5, and 19 days for groups 1, 2, and 3 and 20, 26, and 19 days for groups 1a, 2a, and 3a, respectively (Table 3). Detected PE times for the total and myeloid CD34+ cell numbers were not significantly different between groups. The age of the donor and the time to NE or PE were not correlated (data not shown).


Hematopoietic stem cell transplant is a treatment modality used for treatment of many hematologic and nonhematologic diseases. Initially, BM was the only source for obtaining stem cells. Thereafter, peripheral blood stem cells have almost replaced the BM; however, BM grafts are still in use, especially in pediatric age groups. Hematopoietic stem and progenitor cells responsible for engraftment can be detected by measuring CD34 antigen-expressing cells in the graft. Enumeration of CD34+ cells by flow cytometry (using the ISHAGE protocol) is the reference method used to detect the engraftment potential of HSPCs.3-5 The infusion of at least 2.0 to 2.5 million viable CD34+ cells per kilogram is essential for a rapid and persistent engraftment.6

The main difference between CD34+ HSCs and committed progenitors is their expression of CD90, CD133 along with CD34, and the absence of CD45RA, CD38, HLA-DR, and hematopoietic cell lineage markers on their surface.20,21 Because this study was a retrospective analysis, only the ratio of total CD34+ cells and CD34+ stage 1 hematogones could be evaluated.

Healthy BM CD34+ stage 1 hematogones may represent a considerable CD34 subfraction. This subfraction accounted for a median of 29.4% in our study, which is in accordance with the study of Dmytrus and colleagues, who reported 32% CD34+ stage 1 hematogones within BM grafts collected from donors who were aged between 2 and 48 years.21 Similar to the work of Pichler and colleagues,20 we grouped patients according to CD34 content of the graft; our results were also in accordance with the study of Pichler and colleagues, where 27% CD34+ stage 1 hematogones were detected in the pediatric age group.

In our study group, grafts obtained from donors ≤10 years old contained significantly higher total (P < .001) and myeloid (P = .006) CD34+ cell fractions. This observation is similar to Ince and colleagues, who used 16 years as the threshold to define younger donors.22 In our study, 7 of 27 donors were >10 and ≤16 years old (but were classified in the older donor group); however, when we used 16 years as the threshold, we did not observe a significant difference between the 2 age groups.

We found infused total CD34+ and myeloid CD34+ cells to be 1.6-fold higher and 1.5-fold higher in the younger donor age group, respectively. Furthermore, the percentage of both total CD34+ cells within WBCs and CD34+ stage 1 hematogones was also 1.5-fold higher in the younger donors. In contrast, Pichler and colleagues reported the lowest myeloid CD34+ cell fraction in grafts from donors younger than 15 years old.20 Our findings imply that BM collected from a donor younger than 10 years old might yield approximately 1.5 times more myeloid CD34+ cells compared with a donor older than 10 years of age. Accordingly, it may be possible to decrease the necessity of multiple collections, reduce the operating room/anesthesia time for donors <10 years old, and prevent unnecessary transfusions of pediatric donors.

Although several studies reported a correlation between infused BM CD34+ cell content and the NE time, other results did not find such a correlation.20 The BM CD34+ compartment consists of variable CD34+ cell subgroups, including the stage 1 hematogones, and we investigated the hypothesis that infused non-hematogone CD34+ stem cells reliably predict NE time. Accordingly, the only significant difference that we detected was when we compared the group with infused total CD34+ cell numbers, and the group infused with <3 × 106/kg had a longer NE time (20.5 vs 16 days). A similar difference was not detected for PE time. These results were in accordance with the results of Alanazi and colleagues, who reported a significantly higher neutrophil recovery within day 28, if the CD34+ cell dose was ≥3 × 106/kg per kg of recipient weight.23 In this manner, our results were not in accordance with the results from Pichler and colleagues.20

The size of the groups, priming with GCSF, and post-HSCT complications may affect the engraftment kinetics. Increasing the number of cases by including BM and mobilized stem cell grafts and evaluating all of these parameters along with different CD34+ subgroups, including the HSCs and progenitors in future studies, may facilitate our understanding of engraftment kinetics. The restriction of this study was that it is heterogeneous and some data requested from the clinics during the analysis that could have affected the engraftment kinetics were missing. We cannot add the exact time for cytomegalovirus reactivation. However, the patients with myelodys-plastic syndrome older than age 60 years receive reduced intensity conditioning, and all patients with adult acute lymphoblastic leukemia and acute myeloid leukemia receive myeloablative con-ditioning (cyclophosphamide plus total body irradiation or busulfan plus cyclophosphamide, respectively). In the pediatric group, 10 patients received a myeloablative regimen, 4 patients received a reduced intensity regimen, and the remaining patients did not receive a conditioning regimen. None of the pretransplant patients had active cytomegalovirus infection.

The CD34+ cell population, excluding the stage 1 hematogones, still constitutes a heterogeneous cell group. The use of additional monoclonal antibodies to define this fraction may help our understanding of the engraftment kinetics. Until these antibodies are defined, we can easily measure/report total CD34+ and non-hematogone CD34+ cell fractions and limit the collected BM graft volume to achieve 3 × 106/kg per kg of recipient weight. It is noteworthy that the identification of hematogones within the CD34+ cell population can be done solely based on their very low light scatter properties using the CD45 side scatter and forward side scatter graphs (Figure 1).

Our findings suggested the introduction of an additional calculation into the ISHAGE protocol by grouping the CD34+ cells according to their light scatter properties, especially if a BM graft contains <3 × 106 CD34+ cells per kilogram patient weight. Although our patient numbers were low for allogeneic transplantations, we propose that hematogone analysis could be considered as a routine part of HSC analysis in donor stem cell collections for better prediction of transplant outcomes.


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Volume : 20
Issue : 10
Pages : 937 - 944
DOI : 10.6002/ect.2022.0210

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From the 1Department of Hematology, Ankara University Faculty of Medicine, the 2Department of Pediatric Immunology and Allergy, Ankara University Faculty of Medicine, the 3Biotechnology Institute, Ankara University, the 4Department of Pediatric Hematology-Oncology, Memorial Ankara Hospital, and the 5Pediatric BMT Unit, Ankara University Faculty of Medicine, Ankara, Turkey
Acknowledgements: The authors have not received any funding or grants in support of the presented research or for the preparation of this work and have no declarations of potential conflicts of interest.
Corresponding author: Klara Dalva, Ankara Üniversitesi Tıp Fakültesi Ibni Sina Hastanesi Hematoloji Bilim Dalı Laboratuvarı Poliklinik Blokları Asma Kat, 06100 Ankara, Turkey
Phone: +90 312 3082739