Patent Application
20220348876
Any and all applications for which a foreign or domestic priority claim is identified in the Application Data Sheet as filed with the present application are hereby incorporated by reference under 37 CFR 1.57.
A method of generating definitive hematopoietic cells from source cells, the definitive hematopoietic cells including at least one of: differentiating iPS cells, cells directly reprogrammed to pre-cursors of hematopoietic cells, cells directly reprogrammed to definitive hematopoietic cells, and adult or neonatal hematopoietic cells from bone marrow, cord blood, placenta, or mobilized peripheral blood, the method including using a metabolic regulator to activate a tricarboxylic acid cycle of the source cells.
In the developing embryo, primitive hematopoiesis gives rise to erythrocytes, megakaryocytes and macrophages in the blood islands of the yolk sac (YS) (Palis, J. et al. Development 126, 5073-5084 (1999)). Next, a definitive wave of hematopoiesis produces more mature erythro-myeloid (Palis, J. et al. Development 126, 5073-5084 (1999)) and lymphoid (Yoder, M. C. et al. Immunity 7, 335-344 (1997); and Böiers, C. et al. Cell Stem Cell 13, 535-548 (2013)) progenitors. Around Carnegie stage (CS) 12-13, hematopoietic stem cells (HSCs) emerge in the aorta-gonad-mesonephros (AGM) region through a second definitive hematopoietic wave (Medvinsky, A. & Dzierzak, E. Cell 86, 897-906 (1996); and Ivanovs, A. et al. J Exp Med 208, 2417-2427 (2011)). Primitive erythrocytes, erythro-myeloid progenitors (EMPs) and HSCs derive from a hemogenic endothelial (HE) cell (Lancrin, C. et al. Nature 457, 892-895 (2009); Frame, J. M. et al. STEM CELLS 34, 431-444 (2016) and Stefanska, M. et al. Sci Rep 7, 1-10 (2017)) by a process known as endothelial to hematopoietic transition (EHT) (Boisset, J.-C. et al. Nature 464, 116-120 (2010); and Kissa, K. & Herbomel, P. Nature 464, 112-115 (2010)). Studies on hematopoietic emergence during embryonic development have not only described EHT in spatial and temporal contexts in several animal models (Boisset, J.-C. et al. Nature 464, 116-120 (2010); and Kissa, K. & Herbomel, P. Nature 464, 112-115 (2010)) but also led to a deep understanding of the growth and transcription factors regulating this process (Chen, M. J. et al. Nature 457, 887-891 (2009); Zhou, F. et al. Nature 533, 487-492 (2016); and Swiers, G. et al. Nat Commun 4, 2924 (2013)). However, the role of metabolites and metabolic pathways in the emergence of hematopoietic cells has not been evaluated during development.
Growing evidence points to the fact that metabolic pathways can control cell fate (Oburoglu, L. et al. Cell Stem Cell 15, 169-184 (2014); Moussaieff, A. et al. Cell Metabolism 21, 392-402 (2015); and Folmes, C. D. L. et al. Cell Metabolism 14, 264-271 (2011)). Specifically, the fate of bone marrow HSCs is regulated by several metabolic pathways. The hypoxic niche of the bone marrow pushes HSCs to activate a minimal energy-providing pathway, anaerobic glycolysis, and ensures their quiescent state (Takubo, K. et al. Cell Stem Cell 12, 49-61 (2013)). HSC self-renewal and maintenance rely on fatty acid oxidation (Ito, K. et al. Nat Med 18, 1350-1358 (2012)) and differentiating HSCs switch to oxidative phosphorylation (OXPHOS) to meet their energetic requirements (Yu, W.-M. et al. Cell Stem Cell 12, 62-74 (2013); and Simsek, T. et al. Cell Stem Cell 7, 380-390 (2010)).
The EHT process has been modeled extensively in vitro using pluripotent stem cells (PSCs) and the HE intermediate which arises in this context can give rise to both primitive and definitive hematopoietic cells (Garcia-Alegria, E. et al. Stem Cell Reports 11, 1061-1074 (2018)). Many studies have focused on obtaining HE with solely definitive potential in vitro (Kennedy, M. et al. Cell Reports 2, 1722-1735 (2012); Sugimura, R. et al. Nature 545, 432-438 (2017); Ng, E. S. et al. Nature Biotechnology 34, 1168-1179 (2016); and Sturgeon, C. M. et al. Nat Biotech 32, 554-561 (2014)), in an effort to produce functional and transplantable HSCs for therapeutic use.
As EHT implicates tight-junction dissolution, gain of stem cell-like properties and leads to extensive transcriptional and phenotypic changes in the transitioning cell (Zhou, F. et al. Nature 533, 487-492 (2016); Swiers, G. et al. Nat Commun 4, 2924 (2013); and Guibentif, C. et al. Cell Reports 19, 10-19 (2017)), metabolism may contribute to regulating these processes. Previously, in animal models, the emergence of HSCs was shown to be regulated by adenosine signaling and the PKA-CREB pathway (Jing, L. et al. J Exp Med 212, 649-663 (2015); and Kim, P. G. et al. J Exp Med 212, 633-648 (2015)), which are tightly controlled by ATP levels and availability; suggesting a change in energy demand during EHT. Moreover, glucose metabolism has been shown to induce HSC emergence in zebrafish (Harris, J. M. et al. Blood 121, 2483-2493 (2013)).
As will be understood by one of skill in the art, currently there are limitations in the availability of suitably matched hematopoietic cells for transplantation or transfusion procedures required in the routine treatment of over 100 hematological diseases, malignancies, and other life-threatening indications. The current sources of hematopoietic cells and hematopoietic stem cells are limiting because they typically rely on donations by healthy individuals as part of blood drives (e.g., Red Cross) and stem cell donor registries for bone marrow, cord blood, and mobilized peripheral blood. These shortages of suitable donor blood products limit the ability to perform necessary therapies, therefore up to 30% of patients in need of hematopoietic stem cell transplantation for treatment of malignancies do not have a suitably matched donor, and a complicated infrastructure of transportation of transfusable blood cells and good will donor blood drives address shortages in supply as need varies over time and geographical area. Thus, there is a significant need for a more robust and reliable system for acquiring both hematopoietic stem cells and transfusable blood cell products.
There are also risks associated with using donor derived products, such as transmission of infections to the recipient patients, and tissue rejection complications such as graft versus host disease, both of which are potentially life-threatening. Therefore, development of an alternative source of these hematopoietic cells, with practically unlimited self-renewal ability, that could be perfectly matched to the recipient, without risk of transmission of infections, is needed.
We have determined that metabolic modulations prompt HE cells to preferentially adopt a definitive hematopoietic fate. We show a gradual and global increase in metabolism during human EHT, fueled by glucose, glutamine and pyruvate. By dissecting the use of these nutrients, we have elucidated their roles in hematopoietic lineage specification.
Some aspects relate to a method of generating definitive hematopoietic cells from source cells, the source cells including at least one of:
differentiating iPS cells,
cells directly reprogrammed to pre-cursors of hematopoietic cells,
cells directly reprogrammed to definitive hematopoietic cells, and
adult or neonatal hematopoietic cells from bone marrow, cord blood, placenta, or mobilized peripheral blood; and
the method including using a metabolic regulator to activate a tricarboxylic acid cycle of the source cells.
In some examples, the metabolic regulator inhibits Pyruvate dehydrogenase kinases (PDK).
In some examples, the metabolic regulator activates Pyruvate Dehydrogenase complexes (PDH).
In some examples, the metabolic regulator increases uptake of Pyruvate into mitochondria.
In some examples, the metabolic regulator accelerates conversion of Pyruvate to acetyl coenzyme A (Ac-CoA).
In some examples, the metabolic regulator is dichloroacetate (DCA).
In some examples, the concentration of the dichloroacetate in a culture media for the source cells is at least 30 μM.
In some examples, the DCA induces lymphoid/myeloid-biased definitive hematopoiesis.
In some examples, the metabolic regulator is an LSD1 inhibitor.
In some examples, the LSD1 inhibitor includes at least one of GSK2879552 or RO7051790.
In some examples, the LSD1 inhibitor generates definitive hematopoietic cells of the erythroid lineage.
In some examples, the metabolic regulator increases production of α-ketoglutarate.
In some examples, the metabolic regulator is glutamine.
In some examples, the metabolic regulator results in the generation of CD43+ cells from a hemogenic endothelial (HE) source cell.
In some examples, the method further includes using nucleoside triphosphates.
In some examples, the metabolic regulator is a more potent or more stable equivalent of α-ketoglutarate.
In some examples, the metabolic regulator is Dimethyl α-ketoglutarate (DMK).
In some examples, the concentration of Dimethyl α-ketoglutarate in a culture media for the differentiating iPS cells is at least 17.5 μM.
In some examples, the metabolic regulator is used in combination with Nucleosides.
In some examples, the concentration of Nucleosides is at least 0.7 mg/L.
In some examples, the Nucleosides include at least one of Cytidine, Guanosine, Uridine, Adenosine, Thymidine.
In some examples, the definitive hematopoietic cells include definitive hematopoietic stem cells.
In some examples, the definitive hematopoietic stem cells have lymphoid and/or myeloid repopulating potential.
In some examples, the definitive hematopoietic cells include definitive lymphoid and/or myeloid cells.
In some examples, the definitive lymphoid cells include at least one of T-cells, modified T-cells targeting tumor cells, B-cells, NK cells and NKT cells.
In some examples, the definitive hematopoietic cells include mast cells.
In some examples, the definitive hematopoietic cells include erythroid cells suitable for production of adult hemoglobin.
In some examples, cells directly reprogrammed to pre-cursors of hematopoietic cells include at least one of mesodermal precursor cells, hemogenic endothelium cells, and cells undergoing endothelial to hematopoietic transition.
In some examples, adult or neonatal hematopoietic cells include hematopoietic stem cells or hematopoietic progenitor cells.
Some aspects relate to a method of generating primitive hematopoietic cells from source cells, the source cells including at least one of:
differentiating iPS cells,
cells directly reprogrammed to pre-cursors of hematopoietic cells,
cells directly reprogrammed to definitive hematopoietic cells, and
adult or neonatal hematopoietic cells from bone marrow, cord blood, placenta, or mobilized peripheral blood; and
the method including using a metabolic regulator to inhibit a tricarboxylic acid cycle of the source cells.
In some examples, the metabolic regulator inhibits uptake of Pyruvate into mitochondria.
In some examples, the metabolic regulator inhibits conversion of Pyruvate to Ac-CoA.
In some examples, the metabolic regulator inhibits MPC.
In some examples, the metabolic regulator is UK5099.
In some examples, the concentration of UK5099 in a culture media for the source cells is at least 100 nM.
In some examples, the metabolic regulator inhibits PDH.
In some examples, the metabolic regulator is 1-Aminoethylphosphinic acid (1-AA).
In some examples, the concentration of 1 Aminoethylphosphinic acid in a culture media for the source cells is at least 4 μM.
Some aspects relate to a metabolic regulator for activation of a tricarboxylic acid cycle of source cells for the production of definitive hematopoietic cells.
Some aspects relate to a metabolic regulator for activation of a tricarboxylic acid cycle of source cells for the production of primitive hematopoietic cells.
One of skill in the art will understand that the figures below represent examples of data and diagrams showing the information as described below.
During embryonic development, hematopoiesis initially occurs through primitive and definitive waves primarily in the yolk sac (YS) and the aorta-gonad-mesonephros (AGM) regions, giving rise to distinct blood lineages (Palis, J. et al. Development 126, 5073-5084 (1999); and Medvinsky, A. & Dzierzak, E. Cell 86, 897-906 (1996)). The first hematopoietic stem cells (HSCs) emerge from hemogenic endothelial (HE) cells in the AGM, through endothelial to hematopoietic transition (EHT) (Boisset, J.-C. et al. Nature 464, 116-120 (2010); and Kissa, K. & Herbomel, P. Nature 464, 112-115 (2010)). In the adult, HSC quiescence, maintenance and differentiation are closely linked to changes in metabolism (Takubo, K. et al. Cell Stem Cell 12, 49-61 (2013); and Yu, W.-M. et al. Cell Stem Cell 12, 62-74 (2013)). In certain examples disclosed herein, de novo emergence of blood may be regulated by multiple metabolic pathways that directly induce or modulate hematopoietic specification and lineage commitment during human EHT. EHT may be accompanied by a metabolic switch, with concomitant increases in glycolysis and oxidative phosphorylation (OXPHOS). Moreover, the OXPHOS fuel glutamine may be essential for hematopoietic emergence and, through its different pathway intermediates, is able to direct distinct lineage outcomes. In both in vitro and in vivo settings, steering pyruvate use towards glycolysis or OXPHOS may differentially skews commitment of HE cells to either a primitive erythroid fate or a definitive fate with lymphoid/myeloid potential, respectively. In certain examples, the commitment to primitive or definitive fates in this context may be controlled by distinct mechanisms. During EHT, metabolism may be a major determinant of hematopoietic specification, lineage commitment and primitive versus definitive fate decisions. The disclosure provided herein may provide a basis for using modulation of metabolic pathways to generate definitive HSCs in vitro, in examples thereby providing an invaluable source of treatment for hematological disorders and malignancies.
Induced pluripotent stem (iPS) cells, because of their functional equivalence to embryonic stem cells may have unlimited self-renewal potential, and because they can be generated from somatic cells of the patient him/herself (e.g., skin cells, or amniotic fluid MSCs etc.) and thus recognized as self, is one such ideal source and perhaps the most feasible. In some examples, the ability to generate hematopoietic stem cells from patient derived iPS cells enables the generation of an unlimited supply of human leukocyte antigen (HLA) matched cells, capable of reconstituting the hematopoietic system of patients with hematological disorders or patients undergoing chemotherapy for hematopoietic and some non-hematopoietic solid tumor malignancies. In some examples, depending on the source of somatic cells for deriving the iPS cells, iPS derived hematopoietic stem cells may be superior to traditionally harvested hematopoietic stem cells in terms of: 1) reduced acquired mutations (e.g., if iPS cells were derived from neonatal cell sources), 2) unlimited expansion ability, 3) reduced rejection issues, 4) no contaminating cells from the original tumor present, and 5) the ability to correct congenital mutations in iPS cell lines from patients using existing gene editing technologies such as Crispr/Cas.
Moreover, recent advances in the ability to generate T-cells specifically designed to target and destroy malignant cells following their differentiation from iPS cells means that transplantations may be performed with simultaneous administration of stem cells and anti-tumor T-cells (Trounson et al. Nature Reviews 2016, Vizcardo et al. Cell Stem Cell 2013). As such, in some examples, the ability to generate iPS derived hematopoietic stem cells provides an immediate demand for donor cells for many patients, and potentially offers an exponential increase in use as the surrounding technologies advance. Thus, iPS derived hematopoietic cells offer a reliable and robust new treatment modality for patients with the aforementioned life-threatening diseases.
Further, the ability to generate therapeutically valuable mature or differentiated hematopoietic cells from iPS for transfusion into patients is another facet that perhaps is even greater in terms of serving a public need. In some examples, functional red cells can be generated en mass for all blood groups to be able to address the shortages of transfusion products for patient who have suffered blood loss as a result of injury, who require transfusions during surgery, or suffer from various forms of anemia. In addition, other blood cells differentiated from the iPS may also be useful in the treatment of cancer, such NK or T-cells programmed with antitumor activities.
The tricarboxylic acid (TCA) cycle, also known as the Krebs or citric acid cycle, is the main source of energy for cells and an important part of aerobic respiration. The cycle harnesses the available chemical energy of acetyl coenzyme A (acetyl CoA) into the reducing power of nicotinamide adenine dinucleotide (NADH). The TCA cycle is part of the larger glucose metabolism whereby glucose is oxidized to form pyruvate, which is then oxidized and enters the TCA cycle as acetyl-CoA.
Differentiating iPS cells function like embryonic stem (ES) cells. Unlike ES cells, iPS cells are more readily obtainable for therapy and research, and their isolation does not carry the same ethical concerns. Human iPS cells may be an ideal source for patient-specific therapy since they can be derived from the patients themselves. In addition, iPS cells can serve as useful research tools by providing models of human disease to use for screening new drugs or for studying mechanisms of pathogenesis and toxicology, and models of normal development.
Hematopoietic stem cells (HSCs) are undifferentiated cells whose progeny reconstitute blood cells lineages such as monocytes/macrophages or T and B lymphocytes through a process called hematopoiesis. HSCs possess an indefinite self-renewal potential explaining the interest in these cells for transplantation for the sustained reconstitution of blood cells. B cells are a type of lymphocytes responsible for the humoral immunity (immunity mediated by antibodies).
Definitive hematopoietic stem cells (HSCs) are responsible for the continuous production of all mature blood cells during the entire adult life span of an individual. They are clinically important cells in transplantation protocols used in therapies for blood-related diseases. Experimentally, HSCs can confer long-term reconstitution of the entire hematopoietic system of an irradiated adult recipient.
In certain examples, specific metabolic pathway regulators of glycolysis and the TCA cycle (principle means of energy production in the cell) may directly activate transcriptional changes in the precursors of hematopoietic cells (cells undergoing endothelial to hematopoietic transition) that allows for directed hematopoietic lineage biasing and generation of definitive hematopoietic cells. The ability to generate definitive hematopoietic cells from reprogrammed cells is critical for therapeutics because only definitive cells give rise to the lymphoid blood lineages, (NK, B-cells and T-cells), hematopoietic stem cells, and erythroid (red) cells that express adult hemoglobins. These are the cell types that are currently provided by donors that are already widely used or being developed for use hematopoietic cell-based therapies, including the millions of red cells transfusions that patients receive worldwide each year.
In addition to the modulation of the metabolic pathways described above in iPS derived production of definitive hematopoietic cells, metabolic modulation may be an important means for generating definitive blood from cells sources other than iPS cells. For example, de novo generation of definitive hematopoietic cells may be achieved by direct reprogramming of somatic cells into precursor cells of blood including mesodermal cells and cells undergoing endothelial to hematopoietic transition, also in addition to directly reprogrammed blood cells. Metabolic modulation may provide a basis to guide definitive blood production in all these cases. Moreover, the ability for self-renewal of already committed definitive blood cells (i.e., from bone marrow, cord blood, mobilized peripheral blood, which are currently used in hematopoietic stem cell transplantation therapies worldwide today) benefits from metabolic pathway manipulation for the self-renewal and expansion of the therapeutic hematopoietic stem cell or other definitive hematopoietic cells.
Pyruvate dehydrogenase kinase family members (PDK1, PDK2, PDK3, PDK4) are serine kinases that catalyze phosphorylation of the E1α subunit of the pyruvate dehydrogenase complex (PDC). Pyruvate dehydrogenase kinase is activated by ATP, NADH and acetyl-CoA. It is inhibited by ADP, NAD+, CoA-SH and pyruvate. Biochemicals that inhibit PDK may be used to direct hematopoietic lineage biasing and to generate definitive hematopoietic cells. For example, inhibitors of Pyruvate dehydrogenase kinases (PDK) include Leelamine HCl, a weak CB1 receptor agonist and PDK inhibitor; Quercetin Dihydrate, a natural flavonoid antiproliferative kinase inhibitor; Sodium dichloroacetate, an inhibitor of mitochondrial pyruvate dehydrogenase kinase; SB 203580 (hydrochloride), a MAPK inhibitor; Dichloroacetic acid, a mitochondrial PDK (pyruvate dehydrogenase kinase) inhibitor; PDK1/Akt/Flt Dual Pathway Inhibitor, which is a cell-permeable compound that selectively induces apoptosis; BX 795, an inhibitor of PDK1, TBK1, and IKKε SB 203580; a pyridinyl imidazole and specific inhibitor that suppresses p38 mediated activation of MK2; KT 5720, a potent, specific and cell-permeable inhibitor of PKA; BX-912, a potent and selective PDK-1 inhibitor that induces apoptosis; GSK 2334470, a potent and selective PDK1 inhibitor that subsequently induces apoptotic cell death; and OSU 03012, a PDK1 inhibitor and inducer of caspase and p53-independent apoptosis.
Pyruvate dehydrogenase (PDH) is the first component enzyme of pyruvate dehydrogenase complex (PDC). The pyruvate dehydrogenase complex contributes to transforming pyruvate into acetyl-CoA by a process called pyruvate decarboxylation (Swanson Conversion). Acetyl-CoA may then be used in the citric acid cycle to carry out cellular respiration. Thus, pyruvate dehydrogenase links the glycolysis metabolic pathway to the citric acid cycle and releasing energy via NADH. Pyruvate dehydrogenase may be allosterically activated by fructose-1,6-bisphosphate and is inhibited by NADH and acetyl-CoA. Phosphorylation of PDH is mediated by pyruvate dehydrogenase kinase. Metabolic regulators may be used that activate Pyruvate Dehydrogenase complexes (PDH).
The PDH inhibitor 1-aminoethylphosphinic acid (1-AA) may be used in a culture media for source cells, wherein the concentration of the 1-AA is preferably at least 4 μM, but may be in the range of about 0.5 μm to 50 μm, for example, about: 0.5 μm, 0.6 μm, 0.7 μm, 0.8 μm, 0.9 μm, 1.0 μm, 5 μm, 10 μm, 20 μm, 30 μm, 40 μm and 50 μm.
In some examples, metabolic regulators may be used to increase uptake of pyruvate into mitochondria. Transport of pyruvate across the outer mitochondrial membrane (OMM) is accomplished via large non-selective channels, such as voltage-dependent anion channels/porin, which enable passive diffusion (Benz R. Biochim Biophys Acta. 1994; 1197: 167-196). Voltage-Dependent Anion Channel (VDAC) is the most abundant protein in the OMM and serves as the main pathway for metabolite/ion transport between the cytosol and the intermembrane space (IMS) of mitochondria. Deficiencies in these channels have been suggested to block pyruvate metabolism (Huizing M. et al. Pediatr Res. 1996; 39: 760-765). Inhibitors of voltage-dependent anion channels/porin may be used to inhibit uptake of pyruvate. VDAC phosphorylation by protein kinases, GSK3β, PKA, and protein kinase C epsilon (PKCε), blocks or inhibits association of VDAC with other proteins, such as Bax and tBid, and also regulates VDAC opening. PKA-dependent VDAC phosphorylation and GSK3β-mediated VDAC2 phosphorylation increase VDAC conductance.
The movement of metabolites, such as pyruvate, through the inner mitochondrial membrane (IMM) may be more restrictive than across the OMM, however. Many metabolites have specific mitochondrial inner membrane transporters that have been identified and studied (Palmieri F. et al. Biochim Biophys Acta. 1996; 1275: 127-132).
Metabolic regulators may be used that accelerate conversion of pyruvate to acetyl coenzyme A (Ac-CoA). Dichloroacetate (DCA) promotes pyruvate entry into the Krebs cycle by inhibiting pyruvate dehydrogenase (PDH) kinase and thereby maintaining PDH in the active dephosphorylated state. In instances where the metabolic regulator is dichloroacetate (DCA), the concentration of dichloroacetate in a culture media for the source cells may be at least about 30 μM and can vary from 10 μM to 100 μM, including concentrations of about: 10 μM, 20 μM, 30 μM, 40 μM, 50 μM, 60 μM, 70 μM, 80 μM, 90 μM and 100 μM.
A metabolic regulator used in the disclosed methods herein may inhibit conversion of pyruvate to Ac-CoA. For example, UK-5099 is a potent inhibitor of the mitochondrial pyruvate carrier (MPC). UK-5099 inhibits pyruvate-dependent O2 consumption with an IC50 of 50 nM. The concentration of UK5099 in a culture media for the source cells may be at least 100 nM, but may be in the range of from 10 nM to 1 μm, including about: 10 nM, 20, nM, 30 nM, 40, nM, 50 nM, 60, nM, 70 nM, 80 nM, 90 nM, 100 nM, 0.1 μm, 0.2 μm, 0.3 μm, 0.4 μm, 0.5 μm, 0.6 μm, 0.7 μm, 0.8 μm, 0.9 μm and 1 μm.
Lysine-Specific Demethylase 1 (LSD1) may be used for EHT and particularly the erythroid lineage. Numerous LSD1 inhibitors have been reported such as TCP, ORY-1001, GSK-2879552, IMG-7289, INCB059872, CC-90011, ORY-2001 and RO7051790. One or more of these inhibitors may be used in combination, such as two or more, three or more, four or more, five or more, or combinations of six or more.
Metabolic regulators may be used that increase production of α-ketoglutarate. For example, L-glutamine is a nutritionally semi-essential amino acid for proper growth in most cells and tissues, and plays an important role in the determination and guarding of the normal metabolic processes of the cells. With the help of transport systems, extracellular L-glutamine may cross the plasma membrane and be converted into alpha-ketoglutarate (AKG) through two pathways, namely, the glutaminase (GLS) I and II pathways. Different steps of glutamine metabolism (the glutamine-AKG axis) may be regulated by several factors (Xiao, D. et al. 2016 Amino Acids 48: 2067-2080), rendering the glutamine-AKG axis a potential target to regulate generation of definitive hematopoietic cells from source cells by activation of a tricarboxylic acid cycle of the source cells. α-Ketoglutarate is membrane-impermeable, meaning that it is usually added to cells in the form of esters such as dimethyl α-ketoglutarate (DMKG), trifluoromethylbenzyl α-ketoglutarate (TFMKG) and octyl α-ketoglutarate (O-KG). Once these compounds cross the plasma membrane, they may be hydrolyzed by esterases to generate α-ketoglutarate, which remains trapped within cells. All three compounds increase intracellular levels of α-ketoglutarate. Thus, these compounds are metabolic regulators of α-ketoglutarate. In some examples, the concentration of dimethyl α-ketoglutarate in a culture media for the differentiating iPS cells may be at least about 17.5 μM, but may be from about 10 μM to 100 μM, including concentrations of 10 μM, 20 μM, 30 μM, 40 μM, 50 μM, 60 μM, 70 μM, 80 μM, 90 μM and 100 μM.
Various metabolic regulators disclosed herein may be used in combination with nucleosides, wherein the concentration of nucleosides may be at least 0.7 mg/L, but may be from about 0.1 mg/L to about 10 mg/L, including concentrations of about: 0.1 mg/L, 0.2 mg/L, 0.3 mg/L, 0.4 mg/L, 0.5 mg/L, 0.6 mg/L, 0.7 mg/L, 0.8 mg/L, 0.9 mg/L, 1 mg/L, 1.5 mg/L, 2 mg/L, 2.5 mg/L, 1 mg/L, 1.5 mg/L, 2 mg/L, 2.5 mg/L, 1 mg/L, 1.5 mg/L, 2 mg/L, 2.5 mg/L, 3 mg/L, 3.5 mg/L, 4 mg/L, 4.5 mg/L, 5 mg/L, 5.5 mg/L, 6 mg/L and 6.5 mg/L, 7 mg/L, 7.5 mg/L, 8 mg/L, 8.5 mg/L, 9 mg/L, 9.5 mg/L, 10 mg/L and 10.5 mg/L. The nucleosides comprise at least one of Cytidine, Guanosine, Uridine, Adenosine and Thymidine, but may include any potential combination such as two, three, four, or all five nucelosides.
In an example to obtain both primitive and definitive hematopoietic waves in culture, two previously described small molecules were combined during human iPSC differentiation (
Next, the hematopoietic potential of both HE and EHT populations was verified. Both cell types gave rise to hematopoietic cells (CD43+) (
In examples, in order to describe the metabolic processes occurring in EHT populations, glycolysis was assessed in HE, EHT and HSC-like cells. A gradual increase in glycolytic capacity and glycolysis with differentiation was shown (
In an example exploring whether glycolytic activity is required during EHT, HE cells were treated with a glucose analog, 2-Deoxy-D-glucose (2-DG), which blocks glycolysis (
Along with increased glycolysis and proliferation, HSC-like cells also had increased glucose uptake compared to HE and EHT cells (
Even in glucose-free medium, HE and EHT cells had high basal respiration levels (
To determine whether glutamine is important for EHT, glutaminase (GLS) enzyme was blocked, which catalyzes the deamidation of glutamine to glutamate (
Glutamine also participates in several metabolic pathways including nucleotide and non-essential amino acid (NEAA) syntheses (DeBerardinis, R. J. & Cheng, T. Oncogene 29, 313-324 (2009)). Therefore, to get a better grasp of its role during EHT, HE cells in its absence. Glutamine deprivation abolished CD43+ cell output (>80% decrease) from HE cells at day 3 of subculture (
As pyruvate, another fuel for the TCA cycle, could replace glutamine, we treated HE cells with a pyruvate dehydrogenase kinase (PDK) inhibitor, dichloroacetate (DCA), to increase pyruvate dehydrogenase (PDH) activity during glutamine deprivation. Without glutamine, in examples, DCA treatment alone could not restore the CD43+ cell levels seen in the control (
In certain examples, the percentage of more mature CD43+ cells that have lost CD34+ expression was significantly decreased in the glutamine-free DMK-treated condition as compared to the control (
Previously, erythroid differentiation has been shown to require glutamine-fueled nucleotide synthesis (Oburoglu, L. et al. Cell Stem Cell 15, 169-184 (2014)). Thus, HE cells were stained with a proliferation dye (Cell Trace Violet, CTV) and the proliferation status of newly-formed GPA+ or CD45+ cells was assessed 3 days later. While GPA+ cells clustered to the divided cells (low CTV MFI values), interestingly, CD45+ cells deriving from HE cells had few to no divisions (high CTV MFI values;
Interestingly, a significant increase in the percentages of CD43+CD45+ cells in conditions was restored by DMK or DMK/nucleosides (
Modulation of Pyruvate May Reshape Hematopoietic Output from HE
In some examples, HE cells take up glucose at similar levels as EHT cells (
Although the levels of total CD43+ cells were unchanged between UK5099-treated and untreated conditions at day 6 (
In certain examples, the opposite effect may be induced by increasing pyruvate flux into mitochondria. Using DCA, PDKs were blocked which repress the PDH complex: this allows pyruvate to be converted to acetyl-coA and potentially fuel the TCA cycle (
In examples, following 3 or 6 days of MPC inhibition with UK5099 in HE cells, erythroid colony (CFU-E) formation was significantly increased compared to the untreated condition, while granulocyte and macrophage colonies were decreased (CFU-G, GM and M) (
In certain examples, to understand whether DCA induces the formation of definitive hematopoietic cells, lymphoid differentiation was induced in day 3 HE cells in OP9-DL1 stroma co-cultures. While UK5099 treatment impaired NK cell formation, DCA treatment significantly increased NK cell differentiation as compared to untreated HE cells (
To verify these findings in an in vivo setting, pregnant mice were injected with UK5099 or DCA at embryonic day (E) 9.5, to influence hemogenic endothelium which gives rise to definitive hematopoiesis (both the second and third waves) occurring at E9-9.5 and E10.5, but not primitive hematopoiesis which takes place at E7-7.25 (Palis, J. et al. Development 126, 5073-5084 (1999); and Medvinsky, A. & Dzierzak, E. Cell 86, 897-906 (1996)). In some examples, the blood lineage output in embryos was assessed by characterizing the cellular composition of fetal liver (FL) at E14.5 when the FL is the prime site of hematopoiesis. In some examples, the frequency of phenotypic long-term HSCs (LT-HSCs) were not affected by UK5099 or DCA (
Furthermore, in examples, LT-HSCs from DCA-treated embryos, sorted according to the gating strategy indicated in
In certain examples in order to assess definitive hematopoietic potential of iPS-derived cells, 3-day DCA-treated HE cells co-cultured with OP9-DL1 stroma were intravenously injected into irradiated NSG mice. Engraftment levels comparable to previous studies were obtained (Rahman, N. et al. Nat Commun 8, 1-12 (2017)), with around 1% human CD45+ cells in the peripheral blood (PB) at week 8 (
In certain examples, to dissect the molecular effects of pyruvate manipulation, the transcriptmic profiles of HE cells were assessed at an early time point of treatment (day 2), at the single cell level, in control and UK5099- or DCA-treated cells. First, all conditions were grouped together and separated the cells into 7 clusters (
In examples focusing on isolated clusters 6 and 7 (
In some examples, while the percentage of cells in cluster 6 was constant between conditions, there were 38% more UK5099-treated HE cells and 35% less DCA-treated HE cells in cluster 7 compared to the control (
In certain examples, in clusters 6 and 7, the average expression levels of erythroid lineage genes RYK, KLF3, TAL1, GATA2, ZFPM1, KLF1, NFE2, ANK1 and HBQ1 were higher in UK5099-treated HE cells as compared to the untreated HE cells and these factors were nearly absent in DCA-treated HE cells (left-hand dot plot,
In some examples to further assess the effect of pyruvate manipulation on a single cell level, single HE cells were sorted onto OP9-DL1 stroma and GPA+ clones were scored at day 14. From a total of 552 single cells per condition, 12 GPA+ clones were detected in the UK5099-treated condition and 7 GPA+ clones in the DCA-treated condition as compared to 9 GPA+ clones in the control (
Previous studies have shown that Lysine-Specific Demethylase 1 (LSD1) may be important for EHT and particularly the erythroid lineage (Takeuchi, M. et al. PNAS 112, 13922-13927 (2015); and Thambyrajah, R. et al. Nat Cell Biol 18, 21-32 (2016)). During EHT, LSD1 acts in concert with HDAC1/2 (Thambyrajah, R. et al. Stem Cell Reports 10, 1369-1383 (2018)) and GFI1/GFI1B (Thambyrajah, R. et al. Nat Cell Biol 18, 21-32 (2016)) to induce epigenetic changes. In some examples, it was shown that HDACs may be important for EHT using an HDAC1/2 inhibitor (Trichostatin A, TSA) which impaired the emergence of CD43+ hematopoietic cells (
In some examples, dichloroacetate may be directly used as a precursor of acetylation marks: acetate is converted to acetyl-coA by ACSS2 and transferred onto histones via histone acetyltransferases (HATs) (
As explained above, during EHT, transitioning cells may go through radical changes in energy use and metabolism, with simultaneous increases in glycolysis and TCA cycle/OXPHOS. The disclosure and results presented herein demonstrate for the first time that glutamine is important for the EHT process, playing distinct roles in the specification of primitive erythroid and definitive hematopoietic cells. On the other hand, in examples, there may be a role for glucose in both glycolysis and the TCA cycle. Blocking its use with 2-DG may impair hematopoietic differentiation of HE cells. In quiescent HSCs, glycolysis was shown to be regulated by hypoxia through the stabilization of hypoxia-inducible factor-1α (HIF-1α) (Takubo, K. et al. Cell Stem Cell 7, 391-402 (2010)). The transition from HE to HSCs was also shown to be regulated by HIF-1α (Harris, J. M. et al. Blood 121, 2483-2493 (2013); and Imanirad, P. et al. Stem Cell Research 12, 24-35 (2014)). Thus, in examples, a HIF-la-dependent induction of glycolysis may be important for EHT.
As shown herein and explained above, glycolysis is sufficient to provide energy for primitive hematopoiesis. Indeed, at early embryonic stages, oxygen is not systemically available and glycolysis is the pathway of choice to produce energy (Gardner, D. K. et al. Semin Reprod Med 18, 205-218 (2000)). In developing embryos, primitive erythroid cells were shown to perform high rates of glycolysis to fuel their rapid proliferation (Baron, M. H. et al. Blood 119, 4828-4837 (2012)). Similarly, in the setting described herein, GPA+ cells deriving from HE proliferate faster than CD45+ cells and rely on glutamine for providing nucleotides for this process. Likewise, a crucial role for glutamine in supplying nucleotides for erythroid differentiation has been previously described in the context of HSCs obtained from cord blood (Oburoglu, L. et al. Cell Stem Cell 15, 169-184 (2014)). Blocking MPC may redirect HE commitment towards primitive erythropoiesis at a very early stage of EHT, as shown by an increased frequency of committed cells at the single-cell level as well as higher levels of erythroid factors and embryonic/fetal-specific globins in this condition.
In some examples, the results herein and shown above may unravel a role for the TCA cycle and OXPHOS in specifying definitive hematopoietic identity. Fueling the TCA cycle with DMK during glutamine deprivation or DCA treatment may lead to an increased differentiation of HE cells toward a definitive CD45+ lineage. While PDK inhibition with DCA does not affect primitive erythroid cell formation, it may induce lymphoid/myeloid-biased definitive hematopoiesis, as shown herein both in vitro and in vivo. DCA-treatment of HE cells may lead to an increased lymphoid reconstitution in NSG mice. The results presented herein are in agreement with previous findings in Pdk2/Pdk4 double knockout mice, which were shown to be anemic but retained normal frequencies of T, B and myeloid populations (Takubo, K. et al. Cell Stem Cell 12, 49-61 (2013)). In examples, the results herein show that DCA may promote CD45+ cell formation by fueling cholesterol biosynthesis. This result is corroborated by an elegant study in zebrafish demonstrating that Srebp2-dependent regulation of cholesterol biosynthesis is essential for HSC emergence (Gu, Q. et al. Science 363, 1085-1088 (2019)). As shown here, a direct metabolic change in HE cells, namely increased acetyl-coA content, can promote cholesterol metabolism and control definitive hematopoietic output.
Others have reported previously that distinct EHT cell subsets or pre-HSCs present different lineage propensities (Zhou, F. et al. Nature 533, 487-492 (2016); and Guibentif, C. et al. Cell Reports 19, 10-19 (2017)). In certain examples such as shown herein, metabolism can rewire the fate of HE cells, suggesting that lineage propensities may be decided at the HE level. In line with the results herein, a recent study combining scRNAseq with lentiviral lineage tracing revealed that cell fate biases appear at a much earlier stage during hematopoietic development than previously described with conventional methods (Weinreb, C. et al. Science. 2020 Feb. 14; 367(6479)). Furthermore, murine HSCs were shown to present lymphoid or myeloid hematopoietic lineage biases due to epigenetic priming which is established prior to their formation (Yu, V. W. C. et al. Cell 167, 1310-1322.e17 (2016)). In effect, linking epigenetic changes to metabolism is a newly emerging field which reconciliates metabolic alterations with transcriptional regulation of cellular processes. In accordance, as shown in examples herein, erythroid fate induction by MPC inhibition may be dependent on an epigenetic factor, LSD1.
The examples and results herein may indicate that the lineage propensities of primitive and definitive hematopoietic waves are shaped by nutrient availability in the YS and AGM niches. Due to scarcity of oxygen in early embryonic stages, the primitive hematopoietic wave may depend on glycolysis to form erythroid cells expressing embryonic globins with high affinity for oxygen (
As explained herein, in examples, using metabolic determinants to direct definitive HSC development in vitro from PSCs may provide a way to produce transplantable cells, able to reconstitute the hematopoietic system of patients with hematological malignancies and disorders.
hiPSC Culture, Hematopoietic Differentiation and Cell Isolation Methods
One of skill in the art will understand that the methods and materials described below and elsewhere in the specification are merely examples, and such examples may be performed using different combinations of methods and materials. Further, elements of the methods and materials described herein may be optional. The RB9-CB1 human iPSC line was co-cultured with mouse embryonic fibroblasts (MEFs, Millipore), passaged every six days and processed to form embryoid bodies (EBs) as described previously (Guibentif, C. et al. Cell Reports 19, 10-19 (2017)). The differentiation protocol used in this study was previously described (Ditadi, A. & Sturgeon, C. M. Methods 101, 65-72 (2016)), however, small modifications were made to induce both primitive and definitive hematopoiesis, as indicated below and in
Sorted HE (40,000), EHT (30,000) and HSC-like (5-20,000) cells were plated onto Matrigel (16 μg/cm2, Corning)-coated 96-well flat bottom plates in HE medium (30) with 1% penicillin-streptomycin and kept in a humidified incubator at 37° C., 5% CO2, 4% O2 overnight. The following day (day 0), wells were washed twice with PBS and fresh HE medium was added, together with 2-DG (1 mM), UK5099 (10 μM), DCA (3 mM), BPTES (25 μM), TSA (60 nM), TCP (300 nM), ACSS2i (5 μM), C646 (10 μM), CP-640186 (5 μM), Atorvastatin (0.5 μM), or in glutamine-free medium with DMK (1.75 mM), nucleosides (1×) or NEAAs (1×), where indicated. Media was changed and drugs were added every 2 days and cells were kept in a humidified incubator at 37° C., 5% CO2, 20% O2 for 6-7 days. Pictures were taken using an Olympus IX70 microscope equipped with a CellSens DP72 camera and CellSens Standard 1.6 software (Olympus).
For comparisons between HE, EHT and HSC-like cells, day 10 FACS-sorted cells (≥40,000) were directly plated onto Seahorse XF96 Cell Culture Microplate wells coated with CellTak (0.56 μg/well) in 2-4 replicates and extracellular flux was assessed immediately on a Seahorse XF96 analyzer. For comparisons between HE and EHT cells, day 8 FACS-sorted cells (≥40,000) were plated onto Matrigel (16 μg/cm2, Corning)-coated Seahorse XF96 Cell Culture Microplate wells in 3-4 replicates and extracellular flux was assessed 2 days after plating, on a Seahorse XF96 analyzer. To assess glycolytic flux, ECAR was measured in XF medium with 2 mM glutamine under basal conditions (after 1-hour glucose starvation as per manufacturer's instructions) as well as after the addition of 25 mM glucose, 4 μM oligomycin and 50 mM 2-DG and data was normalized to cell number. The levels of glycolytic capacity (ECARoligomycin-ECAR2-DG) and glycolysis (ECARglucose-ECAR2-DG) were calculated. To assess oxidative phosphorylation, OCR was measured in XF medium with 10 mM glucose, 2 mM glutamine and 1 mM sodium pyruvate under basal conditions as well as after the addition of 4 μM oligomycin, 2 μM Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP) and 1 μM rotenone/40 μM Antimycin A and data was normalized to cell number. The levels of basal respiration (OCRbasal-OCRRotenone/AntimycinA), ATP production (OCRbasal-OCROligomycin) and maximal respiration (OCRFCCP-OCRRotenone/AntimycinA) were calculated.
On days 3 and 6 of subculture, cells were collected after a 2-minute incubation at 37° C. with StemPro Accutase Cell Dissociation Reagent and stained with CD34-FITC, CD14-PE, CD33-PC7, CD11b-APC, CD45-AF700, CD43-APCH7, GPA-eF450, CD90-BV605 and the viability marker 7AAD and fluorescence was measured on a BD LSRII. To measure mitochondrial activity, cells were incubated with Tetramethylrhodamine ethyl ester (TMRE, 20 nM) for 30 minutes at 37° C. Negative controls were incubated with 100 μM FCCP for 30 minutes at 37° C., prior to TMRE staining. Fluorescence was measured on a BD FACSARIA III and MFI levels—MFI FMO were calculated. To measure glucose uptake, cells were incubated with 2-(N-(7-Nitrobenz-2-oxa-1,3-diazol-4-yl)Amino)-2-Deoxyglucose (2-NBDG) for 30 minutes at 37° C. and fluorescence was measured on a BD FACSARIA III. To measure proliferation, cells were processed with the CellTrace Violet (CTV) kit according to manufacturer's instructions (10-minute incubation) and fluorescence was measured on a BD LSRFortessa. To measure EdU incorporation, HE cells were assessed on day 1 or 2 of subculture after 24 h EdU pulses, using Click-iT EdU Flow Cytometry Cell Proliferation Assay (Thermo Fisher Scientific, C10424), according to manufacturer's instructions. Flow cytometry outputs were analyzed on FlowJo Software, with initial gatings on SSC-A/FSC-A, FSC-H/FSC-A, SSC-H/SSC-A and 7-AAD to exclude doublets and dead cells in all experiments.
Subcultured HE cells were treated with StemPro Accutase Cell Dissociation Reagent for 2 minutes at 37° C. and dissociated cells were resuspended in 3 ml Methocult H4230 (STEMCELL Technologies, France) (prepared according to manufacturer's instructions, with 20 mL Iscove's Modified Dulbecco's Medium containing 2.5 μg hSCF, 5 μg GM-CSF, 2.5 μg IL-3 and 500 U EPO). Each mixture was divided onto 2 wells of a non-tissue culture treated 6-well plate. Following a 12-day incubation in a humidified incubator at 37° C., 5% CO2, 20% O2, colonies were morphologically distinguished and scored. For globin analysis, colonies in Methocult wells were harvested with PBS, washed thoroughly and frozen in RLT buffer with β-mercaptoethanol. Following RNA extraction and RT (Qiagen), gene expression was assessed with taqman probes by q-PCR. The taqman probes used in this study are HBA1/2 (Hs00361191_g1), HBE1 (Hs00362216_m1), HBG2/1 (Hs00361131_g1) and KLF1 (Hs00610592_m1).
Subculture day 3 HE cells cultured in the presence of UK5099 (10 μM), DCA (3 mM), or in glutamine-free medium with DMK (1.75 mM), nucleosides (1×) or NEAAs (1×), as indicated, were collected after a 2-minute incubation at 37° C. with StemPro Accutase Cell Dissociation Reagent and seeded onto 80% confluent OP9-DL1 stroma. Cells were cultured in OP9 medium with SCF (10 ng/ml), FLT3-L (10 ng/ml), IL-2 (5 ng/ml), IL-7 (5 ng/ml, first 15 days only) and IL-15 (10 ng/ml) with passaging onto new OP9-DL1 stroma every week, as described previously (Renoux, V. M. et al. Immunity 43, 394-407 (2015)). At day 35 of co-culture, cells were analyzed on a BD LSRFortessa.
Sorted HE, EHT and HSC-like cells as well magnetically selected (Miltenyi Biotec) cord blood CD34+ cells were plated onto Matrigel (16 μg/cm2, Corning)-coated 96-well flat bottom plates in HE medium (Ditadi, A. & Sturgeon, C. M. Methods 101, 65-72 (2016)) with 1% penicillin-streptomycin and kept in a humidified incubator at 37° C., 5% CO2, 4% O2 overnight. The following day (day 0), wells were washed twice with PBS and fresh HE medium was added, together with UK5099 (10 μM) or DCA (3 mM), where indicated. On day 1 and 2 (as indicated), cells were washed twice with PBS 0.04% UltraPure BSA and collected after a 2-minute incubation at 37° C. with StemPro Accutase Cell Dissociation Reagent. Cells were spun down, resuspended in PBS 0.04% UltraPure BSA, counted (yield between 8,000-18,000 cells) and library preparation was conducted according to the Chromium Single Cell 3′ Reagent kit v3 instructions (10× Genomics). Sequencing was performed on a NOVASeq 6000 from Illumina with the run parameters (28-8-0-91) recommended by 10× Genomics with a final loading concentration of 300 pM of the pooled libraries. Human Umbilical Cord Blood samples were collected from Skane University Hospital (Lund and Malmo) and Helsingborg Hospital with informed consents according to guidelines approved by the regional ethical committee.
The data was processed and analyzed using Seurat v3.1.0, where cells were allowed to have up to 20% mitochondrial reads prior to log-normalization and finding the top 500 variable genes using the “vst” method. Cell cycle scores were calculated and the data was scaled regressing on mitochondrial content and the difference of the S and G2M score. Principal components were calculated prior to calculating a UMAP. Pseudotime trajectories describing two developmental routes were identified in our EHT dataset using Slingshot (Street, K. et al. BMC Genomics 19, 477 (2018)) along which the cells were ordered. The cells were then binned along each trajectory where the cell-type composition of each bin was calculated as percentages. Cord blood CD34+ cells were mapped to our data and labeled using scCoGAPS (Stein-O'Brien, G. L. et al. Cell Syst 8, 395-411.e8 (2019)). CS13 data from Zeng et al. (Zeng, Y. et al. Cell Res 1-14 (2019)) was read and processed to make a UMAP from which the cells they name as “AEC” and “Hem” were identified. These 99 cells were mapped to our data and labeled using SCMAP (Kiselev, V. Y. et al. Nat Methods 15, 359-362 (2018)). Our EHT data was mapped to data from Zeng et al. (Zeng, Y. et al. Cell Res 1-14 (2019)) and vice versa using scCoGAPS where 10 patterns were identified in each data set and then projected to each other using projectR. Each cell was assigned to the group that achieved the highest weight. An overview showing the relationship between cell-types and patterns was done by forming a contingency table on which correspondence analysis was performed using the ca package for R. Differentially expressed genes were found using the FindAllMarkers function. Cell numbers for day 1 samples are as follows: HE=1451, EHT=1523, HSC-like=732. Cell numbers for day 2 samples are as follows: HE ctrl=1195, HE+UK5099=718, HE+DCA=2309. All assessed endothelial and hematopoietic genes were previously used in several publications to validate the EHT process (Zhou, F. et al. Nature 533, 487-492 (2016); Swiers, G. et al. Nat Commun 4, 2924 (2013); Ng, E. S. et al. Nature Biotechnology 34, 1168-1179 (2016) and Guibentif, C. et al. Cell Reports 19, 10-19 (2017)). For gene expression analyses, gene sets for glycolysis, oxidative phosphorylation, glutamine transport and cholesterol efflux were downloaded from The Molecular Signatures Database (MSigDB).
Downregulation Via shRNAs
Short-hairpin sequences recognizing the genes of interest were cloned into GFP-expressing pRRL-SFFV vectors, embedded in a microRNA context for minimal toxicity, as described previously (Fellmann, C. et al. Cell Reports 5, 1704-1713 (2013)). Each lentivirus batch was produced in two T175 flasks of HEK 293T cells by co-transfecting 22 μg of pMD2.G, 15 μg of pRSV-Rev, 30 μg of pMDLg/pRRE and 75 μg of the shRNA vector using 2.5 M CaCl2. Medium was changed 16 hours after transfection and viruses were harvested 48 hours after transfection, pelleted at 20,000×g for 2 hours at 4° C., resuspended in 100 μl DMEM, aliquoted and kept at −80° C. The downregulation efficiency of each shRNA was measured by assessing the corresponding gene expression by qPCR in sorted GFP+ cells, 3 days after lentiviral transduction of cord blood CD34+ HSPCs. HE cells were transduced by direct addition of lentivirus particles into the culture medium on the day after the sort.
Pregnant female C57Bl/6xB6.SJL mice were injected intraperitoneally at E9.5 with UK5099 (4 mg/kg) or DCA (200 mg/kg) or PBS (control). Embryos were harvested at E14.5 and individually weighed and processed. Fetal livers were dissected and homogenized in 800 μL ice cold PBS supplemented with 2% fetal bovine serum (FBS) and FL cells were washed in PBS with 2% FBS. For the differentiated lineage panel, cells were stained with B220 and CD19 (B-cell markers)-PE, CD3e-APC, Ter119-PeCy7 and CD71-FITC and analyzed on a BD FACSARIA III. For the HSC panel, samples were first treated with ammonium chloride solution (STEMCELL Technologies, France) to lyse red blood cells, washed twice in ice cold PBS with 2% FBS, stained with CD3e, B220, Ter119, Gr1 (Lineage)-PeCy5, c-Kit-Efluor780, Sca1-BV421, CD48-FITC, CD150-BV605 and 7-AAD (for dead cell exclusion) and analyzed on a BD FACSARIA III. Flow cytometry outputs were analyzed on FlowJo Software, with initial gatings on SSC-A/FSC-A and FSC-H/FSC-A to exclude doublets. For plating of CFU assays, 100 LT-HSCs were sorted (gating strategy shown in
Sorted human HE cells (350,000) were mixed with OP9-DL1 stroma (60,000) and subcultured for 3 days with or without DCA (3 mM) on Matrigel (16 μg/cm2, Corning)-coated 12-well plates in HE medium30. Between 100,000-150,000 cells from control or DCA samples were transplanted into sub-lethally irradiated (300 cGy) 8-week-old female NOD/Cg-Prkdcscid Il2rgtm1Wjl/SzJ mice (NSG, The Jackson Laboratory) together with 20,000 whole bone marrow support cells from C57Bl/6.SJL mice (CD45.1+/CD45.2+, in house breeding). Cells were transplanted in single cell solution in 250 μL PBS with 2% FBS through intravenous tail vein injection. Drinking water of transplanted NSG mice was supplemented with ciprofloxacin (125 mg/L, HEXAL) for 3 weeks after transplantation to prevent infection. Mice were housed in a controlled environment with 12-hour light-dark cycles with chow and water provided ad libitum. Experiments and animal care were performed in accordance with the Lund University Animal Ethical Committee.
Peripheral blood (PB) was collected from the tail vein into EDTA-coated microvette tubes (Sarstedt, Cat #20.1341.100). Peripheral blood was lysed for mature erythrocytes in ammonium chloride solution (STEMCELL technologies) for 10 minutes at room temperature, washed and stained for cell surface antibodies for 45 minutes at 4° C., washed and filtered prior to flow cytometry analysis on the FACS AriaIII (BD). Flow cytometry outputs were analyzed on FlowJo Software, with initial gatings on SSC-A/FSC-A and FSC-H/FSC-A for doublet exclusion, on DAPI for dead cell exclusion and on huCD45/muCD45.1 for murine cell exclusion.
Bone marrow was analyzed at the 12-week transplantation endpoint. Mice were euthanized by spinal dislocation followed by the dissection of both right and left femurs, tibias and iliac bones. Bone marrow was harvested through crushing with a pestle and mortar and cells were collected in 20 mL ice-cold PBS with 2% FBS, filtered and washed (350×g, 5 min). Bone marrow cells were lysed for red blood cells (ammonium chloride solution, STEMCELL technologies) for 10 minutes at room temperature, washed and stained for cell surface antibodies for 45 minutes at 4° C., washed and filtered prior to FACS analysis on the FACS AriaIII (BD). Flow cytometry outputs were analyzed on FlowJo Software, with initial gatings on SSC-A/FSC-A and FSC-H/FSC-A for doublet exclusion, on DAPI or 7AAD for dead cell exclusion and on huCD45/muCD45.1 for murine cell exclusion.
Whole thymus was harvested at the 12-week transplantation endpoint. The thymocytes were mechanically dissociated from connective tissue in the thymus by pipetting up and down in PBS with 2% FBS, followed by filtration through a 50 μm sterile filter. Erythrocyte contamination was removed by lysing the sample in ammonium chloride solution (STEMCELL technologies) for 10 minutes at room temperature. Samples were washed and spun down after and the pellet of thymocytes was resuspended in FACS buffer and stained for cell surface antibodies for 45 minutes at 4° C., washed and filtered prior to FACS analysis on the FACS AriaIII (BD). Flow cytometry outputs were analyzed on FlowJo Software, with initial gatings on SSC-A/FSC-A and FSC-H/FSC-A for doublet exclusion, on DAPI for dead cell exclusion and on huCD45/muCD45.1 for murine cell exclusion.
For TMRE staining, on day 3 of subculture, half of the culture medium was removed and cells were stained with 20 nM TMRE (Thermo Fisher Scientific, T669) by direct addition into the culture medium of a 2× concentrated solution. After a 20-minute incubation at 37° C., wells were carefully washed with PBS and fresh HE medium was added. During acquisition, cells were kept in a humidified incubator at 37° C., 5% CO2, 20% O2. For immunocytochemistry, subculture day 2 HE cells (plated on coverslips) were washed twice in PBS, fixed with 4% PFA for 15 minutes at RT and washed three times with PBS. For filipin staining, fixed cells were incubated with 100 μg/ml filipin III (Sigma-Aldrich, F4767) for 1 hour, washed three times with PBS and rinsed with distilled water before mounting with PVA/DABCO. For H3K9 and H4 acetylation staining, fixed cells were permeabilized and blocked 1 hour at RT with PBS+0.25% Triton X-100+5% normal donkey serum (blocking solution) followed by incubation overnight at 4° C. with primary antibodies diluted in blocking solution. Cells were then washed 2×5 min with PBS+0.25% Triton X-100 (TPBS) and 5 min with blocking solution before incubation with secondary antibodies 2 hours at RT diluted in blocking solution. Cells were later washed 5 min with TPBS containing 1 μg/ml Hoechst and twice with PBS before being rinsed with distilled water and mounted with PVA:DABCO. Images were obtained with the 10× (TMRE) or 20× (Filipin and acetylation) objective of a Zeiss LSM 780 confocal microscope using the Zen software and a 1.5× zoom (TMRE) or 0.6× zoom (Filipin and acetylation). Acquisition settings were the same for all images of each experiment, taking the same number of stacks. Intensity quantification was performed using the Fiji software as follows. For TMRE, using the brightfield channel, ROIs were selected for 5 spindle-shaped and 5 round cells (randomly chosen) and average intensity for each ROI was calculated in a summatory Z-stack of the TMRE channel. For filipin and acetylation, the summatory Z-stack for the filipin channel was obtained and average intensity calculated. A total of 2-3 independent experiments with 2-3 replicate wells were quantified. For each replicate well, 4-6 images were acquired.
Significance of differences between conditions were calculated using paired/unpaired t-tests, 1/2-way analysis of variance (ANOVA) tests or Kruskal-Wallis tests with multiple comparisons in GraphPad Prism 6 software, as indicated. p values are indicated in figures with the following abbreviations: ns, not significant, *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001.
While the present description sets forth specific details of various embodiments, it will be appreciated that the description is illustrative only and should not be construed in any way as limiting. Furthermore, various applications of such embodiments and modifications thereto, which may occur to those who are skilled in the art, are also encompassed by the general concepts described herein. Each and every feature described herein, and each and every combination of two or more of such features, is included within the scope of the present invention provided that the features included in such a combination are not mutually inconsistent. All figures, tables, and appendices, as well as patents, applications, and publications, referred to above, are hereby incorporated by reference.
Some embodiments have been described in connection with the accompanying drawing. However, it should be understood that the figures are not drawn to scale. Distances, angles, etc. are merely illustrative and do not necessarily bear an exact relationship to actual dimensions and layout of the devices illustrated. Components can be added, removed, and/or rearranged. Further, the disclosure herein of any particular feature, aspect, method, property, characteristic, quality, attribute, element, or the like in connection with various embodiments can be used in all other embodiments set forth herein. Additionally, it will be recognized that any methods described herein may be practiced using any device suitable for performing the recited steps.
For purposes of this disclosure, certain aspects, advantages, and novel features are described herein. It is to be understood that not necessarily all such advantages may be achieved in accordance with any particular embodiment. Thus, for example, those skilled in the art will recognize that the disclosure may be embodied or carried out in a manner that achieves one advantage or a group of advantages as taught herein without necessarily achieving other advantages as may be taught or suggested herein.
Although these inventions have been disclosed in the context of certain preferred embodiments and examples, it will be understood by those skilled in the art that the present inventions extend beyond the specifically disclosed embodiments to other alternative embodiments and/or uses of the inventions and obvious modifications and equivalents thereof. In addition, while several variations of the inventions have been shown and described in detail, other modifications, which are within the scope of these inventions, will be readily apparent to those of skill in the art based upon this disclosure. It is also contemplated that various combination or sub-combinations of the specific features and aspects of the embodiments may be made and still fall within the scope of the inventions. It should be understood that various features and aspects of the disclosed embodiments can be combined with or substituted for one another in order to form varying modes of the disclosed inventions. Further, the actions of the disclosed processes and methods may be modified in any manner, including by reordering actions and/or inserting additional actions and/or deleting actions. Thus, it is intended that the scope of at least some of the present inventions herein disclosed should not be limited by the particular disclosed embodiments described above. The limitations in the claims are to be interpreted broadly based on the language employed in the claims and not limited to the examples described in the present specification or during the prosecution of the application, which examples are to be construed as non-exclusive.
| Number | Date | Country | Kind |
|---|---|---|---|
| 1930385-8 | Nov 2019 | SE | national |
| 2030046-3 | Feb 2020 | SE | national |
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/SE2020/051139 | 11/27/2020 | WO |
