New Biotechnology  Volume 00, Number 00  May 2015

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Q1

Research Paper

Metabolic profiling of hematopoietic stem and progenitor cells during proliferation and differentiation into red blood cells Hasbullah Daud1, Susan Browne1, Rasoul Al-Majmaie1,4, William Murphy2,3 and Mohamed Al-Rubeai1 1

School of Chemical and Bioprocess Engineering, University College Dublin, Belfield, Dublin 4, Ireland School of Medicine and Medical Science, University College Dublin, Belfield, Dublin 4, Ireland 3 Irish Blood Transfusion Services, Dublin, Ireland 4 Institute of Laser for Postgraduate Studies, University of Baghdad, Iraq 2

An understanding of the metabolic profile of cell proliferation and differentiation should support the optimization of culture conditions for hematopoietic stem and progenitor cell (HSPC) proliferation, differentiation, and maturation into red blood cells. We have evaluated the key metabolic parameters during each phase of HSPC culture for red blood cell production in serum-supplemented (SS) and serumfree (SF) conditions. A simultaneous decrease in growth rate, total protein content, cell size, and the percentage of cells in the S/G2 phase of cell cycle, as well as an increase in the percentage of cells with a CD71 /GpA+ surface marker profile, indicates HSPC differentiation into red blood cells. Compared with proliferating HSPCs, differentiating HSPCs showed significantly lower glucose and glutamine consumption rates, lactate and ammonia production rates, and amino acid consumption and production rates in both SS and SF conditions. Furthermore, extracellular acidification was associated with late proliferation phase, suggesting a reduced cellular metabolic rate during the transition from proliferation to differentiation. Under both SS and SF conditions, cells demonstrated a high metabolic rate with a mixed metabolism of both glycolysis and oxidative phosphorylation (OXPHOS) in early and late proliferation, an increased dependence on OXPHOS activity during differentiation, and a shift to glycolytic metabolism only during maturation phase. These changes indicate that cell metabolism may have an important impact on the ability of HSPCs to proliferate and differentiate into red blood cells. Introduction There has been considerable progress in the development of Q2 systems to generate red blood cells from hematopoietic stem and progenitor cells (HSPCs) as an alternative source for blood transfusion. Several groups have attempted to generate significant numbers of erythroid cells from the HSPCs in mobilized peripheral blood, bone marrow (BM), and umbilical cord blood (UCB), as well as from human embryonic stem cells and induced pluripotent Corresponding author: Al-Rubeai, M. ([email protected]) http://dx.doi.org/10.1016/j.nbt.2015.05.002 1871-6784/ß 2015 Published by Elsevier B.V.

stem cells [1–5]. The most successful of these to date showed enucleation levels of 100 percent with up to 1.95  106-fold expansion by using a three-phase culture system that included coculture with a stromal cell line [6]. Later, a protocol was developed to produce red blood cells (RBCs) from human cord blood in the absence of a feeder layer [7]. A different approach, undertaken by Fujimi and co-workers, used a four-phase system which incorporated stromal cells in the first phase and macrophages in the third phase [8]; this yielded more than 1013 RBCs from one unit of cord blood with a 99.4 percent enucleation efficiency. Nevertheless, an www.elsevier.com/locate/nbt

Please cite this article in press as: Daud, H. et al., Metabolic profiling of hematopoietic stem and progenitor cells during proliferation and differentiation into red blood cells, New Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.05.002

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Materials and methods Source and isolation of CD34+ cells Peripheral blood buffy coats (a donation by-product) from healthy donors were obtained from the Irish Blood Transfusion Service (Dublin, Ireland). MNCs were enriched using density gradient centrifugation with Histopaque-1077 (Sigma–Aldrich, Dublin, Ireland) followed by an additional separation step with 20 percent (w/v) sucrose to obtain a higher purity of MNCs and reduce plasma, erythrocyte, and platelet contamination. CD34+ cells were purified by magnetic bead separation using a human CD34+ MicroBead Kit with the AutoMACS separator (Miltenyi Biotech, Bergish Gladbach, Germany) as per manufacturer’s instruction. Cells were eluted with 1000 mL pre-warmed complete medium supplemented with growth factors (see below). Cell samples were taken for determination of cell concentrations using the trypan blue exclusion method, and CD34+ yield was confirmed through flow cytometric analysis using anti-CD34-PE and anti-CD45-FITC antibodies (Caltag-Medsystems, Buckingham, UK). For this work, CD34+ cells were isolated from three buffy coats and combined, with a final purity of 87  6.74 percent.

Cell culture As per Giarratana et al. [6], IMDM medium (Biochrom, Berlin, Germany) was supplemented with 1 percent bovine serum 2

1e+6 Total expansion (fold)

efficient, cost-effective, and simple system for the industrial expansion and differentiation of HSPCs into RBCs remains a challenge and far from realization. The identification and purification of hematopoietic stem cells is based on the expression of the cell surface marker CD34 (a 105to 120-kDa transmembrane cell surface glycoprotein) on hematopoietic stem cells and progenitor cells of all hematopoietic lineages [9,10]. The CD34 antigen is involved in adhesive interactions between HSCs and the stromal environment and in regulation and compartmentalization of stem cells [11,12]. CD34+ cells are found at frequencies of 0.1–0.5 percent of mononuclear cells (MNCs) in UCB, 0.5–3 percent in BM, and 0.05–0.2 percent in peripheral blood; however, this number can be increased by granulocyte–macrophage colony-stimulating factor mobilization, which results in the release of immature cells from the BM into the blood stream [13,14]. Substantial research has been conducted in relation to the therapeutic potential of CD34+ cells in areas such as cancer [15], diabetes mellitus [16], allogenic transplantation [17] and ischemic CVD [18]. Elucidating the cellular and metabolic processes involved in the production of RBCs is an important step in addressing these challenges. Understanding the metabolic changes of HSPCs as they expand and differentiate to mature RBCs will allow the manipulation of intra and extracellular environments through improved media formulations. To the best of our knowledge, the metabolism of HSPCs during their expansion and differentiation into RBCs has not received adequate attention. In this study, we compared key metabolite profiles in HSPCs between serum-supplemented (SS) and serum-free (SF) conditions at the expansion, differentiation, and maturation phases of erythroid development. Our findings will enable the optimization and identification of components for SF media formulation. Development of SF media is critical for future clinical application.

New Biotechnology  Volume 00, Number 00  May 2015

1e+5

1e+4

SCF, IL3, EPO, hydrocorsone

1e+3

0

3

9 Early proliferaon

Late proliferaon

EPO

12

Time 23 (days)

16

Differenaon

Maturaon

FIGURE 1

Typical in vitro culture profile of hematopoietic stem and progenitor cells (HSPCs) during expansion and differentiation into red blood cells in medium supplemented with 10 percent FBS. Experimental batch cultures in SS and SF medium were set up at four time-points from the expansion pool. Arrows indicate time- points for experimental set up.

albumin, 120 mg/mL iron-saturated human transferrin, 900 ng/ mL ferrous sulphate, 90 ng/mL ferric nitrate, and 10 mg/mL insulin (all from Sigma–Aldrich). Proliferation media was supplemented with 100 ng/mL stem cell factor (SCF, Millipore, Billerica, USA), 5 ng/mL interleukin-3 (IL-3, R&D Systems, Minneapolis, USA), 3 U/mL erythropoietin (EPO, Janssen-Cilag, Dublin, Ireland, and 10 6 M hydrocortisone (Sigma–Aldrich). SCF, IL-3, and hydrocortisone were omitted after day 11 for differentiation media, while EPO was omitted after day 16 for maturation media. Cells for metabolic profiling experiments were taken from a pool of HSPCs grown under normal expansion conditions and cultured in batch mode for 72 hours in either SS (10 percent FBS; Lonza, Slough, UK) or SF media. Cultures were performed in triplicate in 24-well plates with a seeding density of 3  105 cells/mL, and maintained at 37 8C, 5 percent CO2 in a humidified atmosphere. Four phases of culture were defined as early proliferation (days 6–9), late proliferation (days 10–13), differentiation (days 16–19), and maturation (days 20–23) based on cell growth and surface marker profiles (Fig. 1).

Determination of metabolite concentrations Glucose was quantified using a blood glucose meter (Ascensia Contour; Bayer Healthcare, Berks, UK); lactate, using the BM Lactate Analyser (Accutrend Lactate; Rocher Diagnostics, Penzberg, Germany); glutamine, using the L-Glutamine/Ammonia (Rapid) Assay Kit (Megazyme, Wicklow, Ireland); and ammonia, using an ammonia assay kit (Sigma–Aldrich). Primary amino acids from culture supernatants were subjected to pre-column derivatization with o-phtalaldehyde (OPA) and quantified by reverse-phase HPLC (ZORBAX Eclipse-AAA; 3.5 mm, 150 mm  4.6 mm column) on an Agilent 1100 HPLC analyser (Agilent Technologies, CA, USA). Collected samples were de-proteinized with one part 20 percent sulfosalicylic acid to four parts sample at 48C for 20 min. Samples were centrifuged at 18,000 rcf for 20 min, and supernatants were transferred to fresh tubes. Buffer A (40 mM phosphate buffer, pH 7.8) and buffer B (methanol:acetonitrile:water = 45:45:10) were used for amino acid separation with a gradient elution program as follows: 0 percent buffer

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B for 1.9 min followed by a steady increase in buffer B to 53 percent over 16.2-min, a wash step with 100 percent buffer B, and equilibration at 0 percent buffer B for a total analysis time of 26 min. A 1 mL sample was injected at a flow rate of 1 mL/min, and derivatized amino acid was detected at l = 338 nm, bandwidth: 10 nm; reference: 390 nm, bandwidth: 20 nm for OPA. The chromatogram peaks were integrated and quantified against internal and external standards. Triplicate runs were carried out for each sample, and the error was determined to be less than 10 percent.

Determination of intracellular ATP and ADP concentration Intracellular ATP was measured using the ATP Bioluminescence Assay Kit HS II (Roche Applied Science, Mannheim, Germany) as per manufacturer’s instruction. The light emitted was measured and integrated for 10 s using a luminometer (Tecan GENios Microplate Reader; Ma¨nnedorf, Switzerland). Intracellular ADP concentrations were measured using an ADP Colorimetric/Fluorometric Assay Kit (Abcam, Cambridge, MA, USA) as per manufacturer’s instructions. ADP concentration was measured at 570 nm.

Total protein content Cells (1  106) were collected from each culture phase for analysis of total protein content. Protein concentration was measured using the QuantiPro BCA Assay Kit (Sigma–Aldrich) as per manufacturer’s instructions.

Flow cytometric analyses Cell cycle analysis was performed by propidium iodide (PI; Sigma– Aldrich) staining of ethanol-fixed cells. Cells were analyzed on the Cell Lab Quanta SC (Beckmann Coulter, Fullerton, CA, USA) with excitation at 488 nm. Cell cycle distributions were analyzed using FlowJo software (FlowJo LLC, Oregon, USA). For cell surface antigen analysis 8  104 cells were stained with the following mouse anti-human antibodies: CD34-PE, CD45FITC, CD71-PE, and glycophorin A-FITC (Caltag-Medsystem, Buckingham, UK). Cells were stained for 30 min in the dark at room temperature, washed, and resuspended in PBS before analysis. CD34-PE/CD45-FITC was used to measure early-phase surface markers and CD71-PE/GpA-FITC for late-phase surface markers. Cell size was measured simultaneously during cell surface antigen analysis using the Cell Lab Quanta SC. This flow cytometer measures electronic volume instead of forward scatter and was calibrated for cell size with standard-sized latex beads of 5, 10, and 20 mm (Spherotech Inc, Illinois, USA).

Statistical analysis All results are reported as standard deviation of the mean unless indicated otherwise. Statistically significant differences were determined by Student’s T-test at p < 0.05.

Results and discussion Typically, HSPCs cultures show an initial lag phase of 2–3 days post-isolation, followed by a high proliferation rate from day 3 until approximately day 11–13 (Fig. 1). In this study, cells were taken at four time-points and grown in SS or SF media for 3 days. The selected time-points represent the early proliferation (day 6), late proliferation (day 9), differentiation (day 13), and maturation

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(day 20) phases of culture. Distinct growth profiles were seen for each phase. The highest fold-expansion in cell density was observed in early proliferation phase for both SS and SF conditions, and viability was above 90 percent (Fig. 2a,b). Compared with early proliferation, expansion was lower in late proliferation for both conditions, yet viability remained high; whereas both expansion and viability decreased in differentiation and maturation phases, particularly under SF conditions. Late proliferation was the only phase that showed a significant difference (p = 0.03) in fold-expansion between SS and SF cultures. This possibly caused by serum which inhibiting differentiation in SS cultures as shown by the cell surface marker profiles. However, interestingly, when HSPCs were cultured in expansion mode (continuous culture for 20 days) there was no difference at this point, although cell viability was considerably lower in differentiation and maturation phases under SF conditions (p = 0.0007; Fig. 2c,d). Thus, serum is not essential for growth and viability during early proliferation but is important during late proliferation, differentiation, and maturation. The specific growth rate in early proliferation was effectively the same for SS and SF cultures at 0.03 hour 1 (Table 1). Growth rate decreased significantly in late proliferation phase and was negligible during differentiation and maturation phases. These results are in agreement with previous studies [1]. Cell cycle profiles are illustrated in Fig. 3a, and show a high proportion of cells (50–60 percent) in the G2/S phase during early and late proliferation, compared with approximately 90 percent G1 phase during maturation. In late proliferation significantly more SF cells were in G1; this is likely the reason for the lower cell numbers and higher doubling times observed. Removal of serum can induce cell cycle exit [19] through the removal of mitogenic factors or through transcriptional repression of cyclins and cyclin-dependent kinases [20]. Hence, serum is vital during late proliferation phase, and there should be a focus on this stage when developing SF media. As cells were already in G0/G1 in later phases, clear differences between SS and SF cultures could not be identified. Cell surface marker profiles denote maturation of erythroid progenitor cells (Fig. 3b). Although the percentage of CD71 / GpA+ cells was lower in SS than SF cultures, this difference was not significant (p = 0.07). However, this lower expression could be due to an inhibitory factor in serum leading to a delay in the differentiation process. Myoblast differentiation and osteogenic differentiation of adipose stem cells are also reportedly inhibited by serum [21,22]. Since total serum withdrawal adversely affects growth, viability, and cell cycle, an optimized concentration of serum or a serum substitute should be sought. Freshly isolated CD34+ cells had an average diameter of 7.5– 8.5 mm (results not shown), and increased in size to a peak during late proliferation (10.4 mm SS and 9.6 mm SF), followed by a gradual decrease until maturation (6.8 mm SS and 6.7 mm SF; Table 1). Total protein content was also highest during late proliferation (250 pg/cell SS and 235 pg/cell SF), with a sharp decrease at differentiation, and lowest at maturation (143 pg/cell SS and 138 pg/cell SF; Table 1). This pattern is characteristic of erythroid differentiation both in vivo and ex vivo [1,23]. SS cultures had a higher total protein content than SF cultures, particularly during differentiation phase (p = 0.02). This may be due to the effects of serum deprivation or because SF cells were slightly more mature.

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New Biotechnology  Volume 00, Number 00  May 2015

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(c) 1000

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2.5E+06

Late proliferaon SS 2.0E+06

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Maturaon SS Maturaon SF

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Viable cell number and viability of hematopoietic stem and progenitor cells in serum free and serum supplemented batch cultures (a,b) and continuous culture (c,d). (Red) Serum supplemented and (dark grey) serum-free cultures.

TABLE 1

HSPC culture parameters in SS and SF medium Early proliferation

Growth rate, (hour 1) Doubling time (dt) Cumulative cell time (109 cell hour/L) Cell size (mm) Total protein content (pg/cell)

Late proliferation

Differentiation

SS

SF

SS

SF

SS

SF

SS

SF

0.031 22.53 117 9.5 238

0.030 22.83 106 9.9 220

0.022 31.4 82 10.4 250

0.019 41.45 64 9.1 235

0.003 n/a 20 7.6 186

0.004 n/a 19 7.0 157

0.005 n/a 19 6.8 142

0.006 n/a 18 6.7 138

As expected, there was a noticeable decrease in extracellular pH in early proliferation for both SS and SF cultures, with lower levels for SS compared with SF cultures in late proliferation (7.1 SS and 7.35 SF after 72 hours), and almost no change in pH during differentiation and maturation phases for either (Fig. 3ci–iv). This agrees well with the cell growth and metabolism profiles. Rapid cell growth and metabolite accumulation during early and late proliferation accelerates media acidification compared with nondividing cells at later phases [24,25]. This could be useful for indirect monitoring of HSPC differentiation into RBCs given the reproducible and characteristic profile observed. Since cells were only cultured for 72 hours, the extended effects of metabolite accumulation are unclear; however, extracellular acidification may affect metabolism, such as glucose and glutamine uptake [26], and higher pH levels have been shown to accelerate erythroid differentiation [27]. Thus, the higher pH in SF cultures in late 4

Maturation

proliferation phase may have contributed to the more mature surface marker profile. Previous studies have demonstrated a distinct metabolic profile during during HSPC differentiation into RBCs [28–30]. Generally, proliferative cells with a very high growth rate rely on glycolysis for energy, while differentiating cells switch to more efficient energy production through OXPHOS [28,31]. Interestingly, during RBC maturation, cells switch from glycolysis to OXPHOS and back to glycolysis. This is because mature RBCs lack the mitochondria required to generate ATP via OXPHOS. However, this is not a feature of differentiation in all cell types – mesenchymal stem cells also show glycolytic suppression during osteogenic differentiation, with a corresponding increase in OXPHOS [32]. In terms of metabolite consumption/production, in both SS and SF cultures total glucose consumption was high during early and

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FIGURE 3

Cellular attributes of hematopoietic stem and progenitor cells in serum-supplemented and serum-free cultures during the ex vivo development of red blood cells. (a)(i) Cell cycle: (upward diagonal) G0/G1 phase, (diamond-cross) S phase, (downward diagonal) G2/M phase. (b)(ii)–(iv) Typical cell cycle profiles for early and late proliferation, differentiation, and maturation phase; (b)(i) cell surface marker profiles (CD71 and GpA) using flow cytometry with anti-CD71-RPE and anti-GpAFITC antibodies, (upward diagonal) CD71-/GpA-, (dotted) CD71+/GpA , (downward diagonal) CD71+/GpA+, (solid) CD71 /GpA+, (b)(ii)–(iv) cell surface marker profiles for early and late proliferation, differentiation and maturation phase; (c) extracellular pH: (i) early and (ii) late proliferation, (iii) differentiation, and (iv) maturation phase, (diamond-red) serum-supplemented and (diamond-dark grey) serum-free cultures.

late proliferation and substantially lower during differentiation and maturation phases (Fig. 4a). There was no significant difference in cell-specific glucose uptake rate between SS and SF cultures in any phase (Table 2). This distinct pattern of high glucose consumption during proliferation followed by a sharp decrease in maturing erythroid cells is also seen in differentiating mouse FVA erythroid precursor cells [23]. Lactate production mirrored the glucose uptake profile, with no differences between SS and SF cultures at any phase (Fig. 4b), and an approximately 10-fold decrease in cell-specific lactate production between late proliferation and differentiation and a negligible value at maturation (Table 2). Table 2 also shows a high ratio of lactate production to glucose consumption (YLac/Gluc) during the early and late proliferation phases, followed by a sharp decrease

during differentiation before increasing to the highest YLac/Gluc value observed during maturation phase. This reinforces the assumption of a reliance on glycolysis during proliferation and maturation, with a switch to OXPHOS during differentiation. Cord blood mononuclear cell cultures which contain high populations of erythroid-burst forming units during proliferation also show a high YLac/Gluc [33]. Lactate accumulation during early proliferation, at 35 mM and 31 mM for SS and SF, respectively, may have exerted an inhibitory effect on cell growth, even though cell viability was not impaired. Inhibition of proliferation, metabolism, and differentiation has been shown in hematopoietic cultures at lactate concentrations above 20 mM, which is accompanied by a decrease in pH [24]. Interestingly, proliferation of other cell lines remains unaffected

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10 11 12 13 14 15 16 17 18 19 20 21 22 23

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(c)

5 4 3 2 1 0

Ammonia (mM)

(d) 3.5 3 2.5 2 1.5 1 0.5 0 Time (day)

FIGURE 4

Initial and final consumption and production of metabolites in hematopoietic stem and progenitor cells taken from different phases and cultured in serumsupplemented and serum-free medium for 72 hours. (a) Glucose consumption; (b) lactate production; (c) glutamine consumption, and (d) ammonia production. (Red) Serum-supplemented and (dark grey) serum-free culture.

by lactate levels as high as 40–50 mM provided pH is maintained at an optimal value [26,34–39]. Glutamine consumption followed a similar pattern to that of glucose (Fig. 4c), with no difference between SS and SF cultures. Glutamine was not depleted at any phase, and remained particularly high during differentiation and maturation. The glutamine cell-specific uptake rate was similar in both SS and SF cultures (Table 2). A high glutamine uptake rate during proliferation provides energy to support cell growth and maintain homeostasis,

with cells using a combination of glutamine and glucose as carbon source [40]. The more energy-efficient differentiating cells consume glutamine through OXPHOS and show lower levels of glycolysis. These results agree with the evidence that dividing progenitor cells depend less on OXPHOS for ATP production than non-dividing differentiated cells [41]. The lowest glutamine uptake levels were during maturation phase where cells relied on glycolysis for energy generation. Ammonia is considered inhibitory at concentrations above 2 mM [42]. The highest ammonia levels here were in early proliferation (2.67 mM SS and 2.58 mM SF; Fig. 4d), and although levels reached the lower end of the scale for inhibition, there was no effect on cell viability. Mirroring glutamine consumption, ammonia production dropped significantly with differentiation and maturation. The cellspecific production rate was highest during late proliferation (0.07 pmol cell 1 day 1 SS and 0.08 pmol cell 1 day 1 SF) and lowest during maturation (0.01 pmol cell 1 day 1 SS and 0.004 pmol cell 1 day 1 SF; Table 2). The yield of ammonia from glutamine was also highest during the early proliferation phase due to high metabolic activity (Table 2). This agrees with a previous study that found that ammonia production was associated with the cellular energy state [43]. Figure 5 shows cell-specific uptake and production rates for ten essential and seven nonessential amino acids. In both SS and SF cultures, all essential amino acids measured were consumed, with the nonessential amino acids glutamate, glycine, alanine, and tyrosine produced, and aspartatic acid, serine, and arginine consumed (Fig. 5). Leucine, isoleucine, and valine were probably converted to derivatives of the Kreb cycle, which is indirectly associated with energy production and directly associated with protein biosynthesis [44]. Arginine and histidine are utilized in energy production, while methionine and threonine are vital for cell and tissue development [45]. Secretion of alanine can occur in culture media with an excess of carbon sources [46], while glutamate secretion is caused by arginine, histidine, proline, and glutamine metabolism before it is fed into the TCA cycle [47]. Both SS and SF cultures showed higher specific consumption and production during the early and late proliferation phases, as with glucose/ glutamine and lactate/ammonia. In differentiation phase, amino acid consumption and production decreased considerably although the majority of essential amino acids (with the exception of leucine production in SF cultures) were still consumed in both SS and SF cultures. During the maturation phase, a similar pattern was seen, although more nonessential amino acids were consumed in both SS and SF cultures. All essential amino acid were utilized during the early and late proliferation phases, similar to mesenchymal stem cells [48]. These results are important for medium

TABLE 2

Specific metabolic rates and yields for HSPCs in SS and SF medium Early proliferation

qGluc (pmol/cell/hour) pLac (pmol/cell/hour) qGln (pmol/cell/hour) pAmm (pmol/cell/hour) YLac/Gluc (mol/mol) YAmm/Gln (mol/mol) 6

Late proliferation

Differentiation

Maturation

SS

SF

SS

SF

SS

SF

SS

SF

0.47 0.90 0.08 0.07 1.91 0.84

0.45 0.87 0.08 0.07 1.93 0.89

0.39 0.71 0.10 0.07 1.82 0.70

0.35 0.79 0.10 0.08 2.26 0.76

0.07 0.07 0.02 0.01 0.91 0.72

0.06 0.07 0.02 0.01 1.24 0.66

0.02 0.05 0.01 0.01 2.80 0.65

0.02 0.05 0.01 0.004 2.70 0.64

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FIGURE 5

Cell-specific consumption and production rates of amino acids at different phases of hematopoietic stem and progenitor cell expansion into red blood cells. (a) Early proliferation phase; (b) late proliferation phase; (c) differentiation phase, and (d) maturation phase. (Red) Serum-supplemented medium and (dark grey) serum-free medium. Glu, glutamic acid; Ser, serine; Asp, aspartic acid; Gly, glycine; Arg, arginine; Ala, alanine; Tyr, tyrosine; Val, valine; Val*, norvaline; Trp, tryptophan; Phe, phenylalanine; Ile, isoleucine; Lys, lysine; Met, methionine; Leu, leucine; His, histidine and Thr, threonine.

formulation and optimization because amino acids, such as glutamate and glycine, could be excluded from the media, since sufficient quantities are produced by cells throughout the culture. In addition, due to the low consumption rate, especially during differentiation and maturation, amino acid depletion does not occur. This implies a high degree of mass overflow in amino acid metabolism, which could result in increased levels of toxic species [48]. Hence, HSPC culture may benefit from media formulation with reduced amino acid concentrations.

Conclusion This study examined cell metabolism during ex vivo erythroid differentiation of HSPCs in SS and SF cultures; an important step in the optimization of serum-free media formulations. Both glycolytic and OXPHOS metabolic pathways were active during early and late proliferation phase in both SS and SF culture, followed by a greater reliance on OXPHOS in differentiation

phase, before a total shift to glycolysis as cells matured into RBCs. Cells in SF cultures both consumed and produced fewer metabolites than those in SS cultures. The importance of serum supplementation during differentiation and maturation phase was clearly shown. However, during the earlier phases of culture, SF was equivalent to SS medium. These findings will bring us closer to the achievement of SF HSPC culture conditions furthering preclinical research and potential therapeutic applications in this area. However, this study focused on extracellular metabolites only; further studies of intracellular metabolism using higher resolution approaches such as mass spectrometry, will prove beneficial.

Acknowledgements This work was supported by the Irish Blood Transfusion Service, Q3 Dublin, Ireland and the National Science Fellowship, Ministry of Science, Technology and Innovation, Malaysia.

References [1] Boehm D, Murphy WG, Al-Rubeai M. The potential of human peripheral blood derived CD34+ cells for ex vivo red blood cell production. J Biotechnol 2009;144(2):127–34. [2] Vannucchi AM, Bianchi L, Cellai C, Paoletti F, Rana RA, Lorenzini R, et al. Development of myelofibrosis in mice genetically impaired for GATA-1 expression (GATA-1(low) mice). Blood 2002;100(4):1123–32.

[3] Neildez-Nguyen TM, Wajcman H, Marden MC, Bensidhoum M, Moncollin V, Giarratana MC, et al. Human erythroid cells produced ex vivo at large scale differentiate into red blood cells in vivo. Nat Biotechnol 2002;20(5):467–72. [4] Lu SJ, Feng Q, Park JS, Vida L, Lee BS, Strausbauch M, et al. Biologic properties and enucleation of red blood cells from human embryonic stem cells. Blood 2008;112(12):4475–84.

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NBT 790 1–8 RESEARCH PAPER

Research Paper

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New Biotechnology  Volume 00, Number 00  May 2015

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www.elsevier.com/locate/nbt Please cite this article in press as: Daud, H. et al., Metabolic profiling of hematopoietic stem and progenitor cells during proliferation and differentiation into red blood cells, New Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.05.002